<figure><img src="image_1.png" /></figure>

HAL
open science

# Fatty acid desaturases in non-photosynthetic bacteria: classification, regulation, and roles in plasma membrane function and cellular homeostasis

Ye Tao, Yilin Sun, Yidan Chai, Luminita Duma, Yannick Rossez

<figure><img src="image_2.png" /></figure>

► To cite this version:

Ye Tao, Yilin Sun, Yidan Chai, Luminita Duma, Yannick Rossez. Fatty acid desaturases in non-photosynthetic bacteria: classification, regulation, and roles in plasma membrane function and cellular homeostasis. Biochimica et Biophysica Acta Molecular and Cell Biology of Lipids, 2025, Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids, 1870 (5), pp.159630. 10.1016/j.bbalip.2025.159630. hal-05106843

HAL Id: hal-05106843

https://lilloa.hal.science/hal-05106843v1

Submitted on 11 Jun 2025

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_3.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
Journal Pre-proof

Fatty acid desaturases in non-photosynthetic bacteria:
classification, regulation, and roles in plasma membrane function
and cellular homeostasis

Ye Tao, Yilin Sun, Yidan Chai, Luminita Duma, Yannick Rossez

PII: S1388-1981(25)00038-1

DOI: https://doi.org/10.1016/j.bbalip.2025.159630

Reference: BBAMCB 159630

To appear in: *BBA - Molecular and Cell Biology of Lipids*

Received date: 4 March 2025

Revised date: 25 April 2025

Accepted date: 11 May 2025

Please cite this article as: Y. Tao, Y. Sun, Y. Chai, et al., Fatty acid desaturases in non-photosynthetic bacteria: classification, regulation, and roles in plasma membrane function and cellular homeostasis, *BBA - Molecular and Cell Biology of Lipids* (2024), https://doi.org/10.1016/j.bbalip.2025.159630

This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

<figure><img src="image_4.png" /></figure>
# Fatty acid desaturases in non-photosynthetic bacteria: classification, regulation, and roles in plasma membrane function and cellular homeostasis

Ye Tao¹, Yilin Sun¹, Yidan Chai¹, Luminita Duma²* and Yannick Rossez³*

¹ School of Food and Pharmacy, Zhejiang Ocean University, Zhoushan, 316022, PR China;
taoye@zjou.edu.cn

² Université de Reims Champagne-Ardenne, CNRS, ICMR UMR 7312, 51097 Reims, France;
luminita.duma-catoire@cnrs.fr

³ Univ. Lille, CNRS, UMR 8576 - UGSF - Unité de Glycobiologie Structurale et Fonctionnelle,
Lille, France; yannick.rossez@univ-lille.fr

* Correspondence: yannick.rossez@univ-lille.fr, luminita.duma@univ-reims.fr

## Abstract

Fatty acid desaturases are key enzymes that catalyze the desaturation of acyl lipid chains by regioselectively introducing double bonds into saturated or pre-existing alkene fatty acids to produce unsaturated fatty acids. While extensive studies have focused on desaturases in higher organisms, research on these enzymes in non-photosynthetic bacteria remains limited, largely due to insufficient structural and functional data. This review aims to summarize the current knowledge on the structural features and catalytic mechanisms of desaturases in non-photosynthetic bacteria, shedding light on the sophisticated regulatory strategies underlying unsaturated fatty acid biosynthesis. In addition, factors influencing desaturase activity are presented and analyzed. This comprehensive review provides insights into bacterial homeostatic adaptation to environmental changes and highlights potential applications in nutraceutical development and therapeutic target identification.

## Keywords

Non-photosynthetic bacterial desaturases, unsaturated fatty acids, architecture, catalytic mechanism, biotechnology

## 1. Introduction

The bacterial cell membrane serves as a dynamic and adaptive interface, providing the first line of defense against an ever-changing external environment. From the extreme heat of deserts to the freezing temperatures of polar ice, bacteria have evolved sophisticated mechanisms to fine-tune the fluidity of their membrane lipid bilayers [1,2]. The adaptability of bacterial cell membranes is largely dependent on the fatty acids (FAs) that make up the membrane. Non-photosynthetic bacterial FAs exhibit considerable structural diversity [3]. In *Escherichia coli* (*E. coli*) and related species, FAs typically have acyl chain lengths of 10 to 19 carbons with two cyclopropane FA (cyC17:0 and cyC19:0) [4,5]. In other species, different possible modifications include branching (branched-chain FA), hydroxylation, epoxy group formation, or beta-amino FAs [6–8]. In contrast to non-photosynthetic bacteria, certain photosynthetic and psychrophilic bacteria are capable of synthesizing long-chain polyunsaturated fatty acids (PUFAs) with up to 22 carbon atoms [9,10]. Cyanobacteria, representative photosynthetic bacteria, can produce fatty acids with up to four double bonds, although their chain length typically does not exceed 20 carbon atoms [9]. In these organisms,
fatty acid desaturases (FADs) contribute to membrane fluidity and play a critical role in the functioning of plant-like oxygenic photosynthesis. However, the biosynthesis of long-chain PUFAs in psychrophiles is primarily carried out by polyketide synthases rather than FADs [10]. Because the roles and biosynthetic pathways of PUFA production in photosynthetic bacteria are relatively well understood and often linked to photosynthetic function, our review focuses instead on non-photosynthetic bacteria. These organisms rely more heavily on FADs for adaptive responses to environmental stress, yet the diversity and function of these enzymes remain underexplored. Desaturation, a stereospecific dehydrogenation process that converts single bonds (C-C) to double bonds (C=C) in FA acyl chains [11,12], is a mechanism commonly observed in all organisms. In most species of non-photosynthetic bacteria, this modification is typically limited to a single double bond per molecule. However, under certain conditions, such as in persister cells undergoing unique metabolic states or stress responses, FAs with two or three unsaturations have been detected [13,14]. Deep-sea piezophilic bacteria can produce polyunsaturated FAs (PUFAs) independently of desaturases, utilizing the polyketide synthase (PKS) pathway via acetyl-CoA [15]. These FAs are broadly categorized into saturated fatty acids (SFAs) and unsaturated fatty acids (UFAs), and their roles are fundamental to membrane structure and function. SFAs increase membrane rigidity, while UFAs contribute to flexibility by influencing the regulation of headgroup spacing through their structural conformation [16]. The balance between these FA types is critical for maintaining membrane functionality and allowing adaptation to changing environmental conditions [7]. In this review, we explore how diverse non-photosynthetic bacterial species, particularly human pathogens, exploit FADs to survive in hostile and fluctuating environments. To address the fragmented terminology in the field, we propose a harmonized nomenclature for FADs that enhances clarity and facilitates cross-species comparisons. Our framework extends beyond well-characterized desaturases to include lesser-known FAD-like enzymes involved in the biosynthesis of vinyl ether bonds, furan fatty acids, and polyynes, highlighting their emerging biological importance and the need for an integrated classification system. We examine the structural and functional diversity of these enzymes, dissect their molecular mechanisms, and emphasize their critical roles in bacterial physiology. In addition, we investigate the regulatory networks that control FAD expression and activity, providing insight into how these pathways contribute to bacterial resilience and adaptability. To our knowledge, this is the first comprehensive review to synthesize current findings on FAD-mediated strategies employed by non-photosynthetic bacteria to cope with extreme and dynamic environments.

## 2. Fatty acid desaturation in non-photosynthetic bacteria

The UFAs can be synthesized through two different pathways: oxygen-independent and the oxygen-dependent, respectively [17]. Initially, FA desaturation was described to occur anaerobically in *E. coli* by introducing double bond into a ten-carbon intermediate by the FabA dehydratase/isomerase of the bacterial type II FA biosynthetic system (FASII) [18–20]. In some species, like *Acinetobacter baumannii* (*A. baumannii*), an unusual FASII has been found, as it lacks a FabA orthologue, one of two enzymes essential for UFA production via the FASII pathway [12,21]. However, it's only true for some strains as a recent paper has identified the aforementioned genes in sequence type 2 (ST2), which is the predominant carbapenem-resistant *A. baumannii* lineage worldwide [22]. The pathway for UFA synthesis involving desaturases is exclusive to aerobic organisms and primarily utilizes integral membrane-bound desaturases. Soluble acyl-acyl carrier protein (acyl-ACP) desaturases, mainly found in plants, have been less identified in non-photosynthetic bacteria. Nonetheless, a soluble acyl-ACP,
DesA2 from *Mycobacterium tuberculosis* (*M. tuberculosis*) has been recognized via structural similarity to plant FAD [23,24]. Integral membrane-bound desaturases can be further subdivided into acyl coenzyme A (acyl-CoA) and acyl-lipid desaturases, which are widely spread among living organisms. They can bind the endoplasmic reticulum in eukaryotes [25], the plant chloroplast membrane [26] and the plasma and thylakoid membranes of cyanobacteria [27]. These widely distributed FADs are essential for the maintenance of cellular homeostasis.

**3. Controlling cellular homeostasis**

Lipid unsaturation plays a critical role in homeostatic adaptation, maintaining membrane fluidity and supporting essential cellular functions such as electron transport. Inadequate fluidity can lead to phase separation, impaired membrane potential, and defects in processes such as cell growth and division. FA composition, which strongly influences the biophysical properties of glycerophospholipids (GPLs), is critical for environmental adaptation and cellular homeostasis. In vivo studies on various bacterial species show that insufficient membrane fluidity disrupts key cellular processes, but induces reversible lipid phase separation without compromising the membrane barrier function [28,29]. The presence of double bonds in FA introduces kinks in their acyl chains, affecting their packing and interactions. These changes affect membrane properties such as fluidity, viscosity, permeability, curvature, and thickness (Figure 1) [2,30,31]. These physical properties, determined by the stereospecific conformations of SFA and UFA and their interactions with headgroups, influence critical cellular processes such as nutrient transport [32], signal transduction [33], cell division [15], motility [34], and stress responses [35,36].

*Methylobacterium extorquens* (*M. extorquens*), an environmental bacterium, synthesizes a distinct set of four GPLs that include a narrow range of saturated FAs, as well as mono- and diunsaturated FAs, resulting in the formation of up to 27 lipid species [37]. In comparison, *A. baumannii* is more complex, by including seven plasma membrane lipids with eight FAs, three of which are monounsaturated. These microorganisms can adjust the ratio of SFAs and UFAs by changing desaturase activity, thereby changing the fluidity of the plasma membrane to adapt to shifting in temperature, pH, and atmospheric pressure [38]. Using whole-genome design and synthesis based on mycoplasmas, a minimal synthetic genome with 492 genes was created, retaining essential functions despite its reduced size. This model highlights the complexity of identifying truly essential genes and demonstrates that acyl chain diversity plays a more critical role in growth than head group diversity [39].
<figure><img src="image_5.png" /><figcaption>**Figure 1.** Illustration of bacterial desaturases controlling membrane homeostasis. The cartoon representation of crystal structure of desaturase (PDB: 1ZA0, *M. tuberculosis*) is used for the illustration.</figcaption></figure>

Interestingly, membrane properties can also be modulated by altering fatty acid unsaturation through the activity of cis-trans isomerases (CTIs). While FADs introduce or adjust double bonds in FAs to regulate membrane fluidity de novo, CTIs rapidly reconfigure pre-existing unsaturated lipids by converting *cis* to *trans* isomers, enabling immediate responses to environmental fluctuations without requiring oxygen or additional biosynthesis [40,41]. This dynamic cooperation ensures membrane integrity under both aerobic and anaerobic conditions, a critical advantage in polymicrobial infections where oxygen availability varies. A recent study has highlighted that CTI-mediated isomerization increases membrane rigidity, serving as a rapid adaptation mechanism to environmental stressors [42], while FADs maintain baseline fluidity essential for various cellular functions [43].

**4. Nomenclature and classification of bacterial desaturases**

The most used nomenclature for desaturases is primarily based on the double bond (C=C) position in FAs [11]. In this regard, two methods are employed to label desaturase activities: one utilizes delta (Δ) [44], and the other one employs omega (ω) [45]. The former signifies that the counting of the double bond starts from the carboxyl end (-COOH) of a FA chain, while the latter is determined from the methyl end (-CH₃) of the FA chain (Figure 2A). For example, a desaturase capable of catalyzing the conversion of myristic acid (C14:0) to myristoleic acid (C14:1) by placing a double bond between the ninth and tenth carbon atoms can be named C14:Δ9 if counted from the carboxyl end, or C14:ω5, if the double bond position is counted from the methyl end. For clarity, we will exclusively use the Δ nomenclature in this review. The corresponding desaturase could be annotated Δ9-Des. Bacteria can exhibit two types of double bonds: *cis*, where the hydrogen atoms are on the same side of the bond, and *trans*, where they are on opposite sides. To achieve greater accuracy in determining double bond orientation, additional recording of Δ9C-Des (Δ9T-Des) or ω5C-Des (ω5T-Des) can be performed on the relevant desaturase (Figure 2B).
To date, desaturases from non-photosynthetic bacteria are characterized by their ability to introduce the first double bond into saturated FA chains. However, ω-desaturases, which insert a new double bond between an existing unsaturation site and the methyl terminus, and front-end desaturases, which establish a double bond between preexisting alkanes and the carboxyl terminus, have not been identified in non-photosynthetic bacteria.

<figure><img src="image_6.png" /><figcaption>**Figure 2.** Schematic representation of myristoleic acid and the insertion position of double bond by specific desaturase (Δ9-Des/ω5-Des) (A). Particularly, in response of orientation about H atoms, *cis* and *trans*, Δ9T-Des, ω5T-Des, Δ9C-Des, ω5C-Des would be annotated in panel (A) and (B).</figcaption></figure>

To ensure consistency, consideration should be given by the scientific community to the establishment of a standardized nomenclature for bacterial desaturases. One proposed approach is to distinguish between desaturases acting on endogenous (designated as DesA) and exogenous (designated as DesB) FAs, as elucidated in the little existing literature addressing these protein types. For example, *Pseudomonas aeruginosa* (*P. aeruginosa*) possesses two FA desaturases. One is a Δ9-desaturase associated with the membrane (endogenous) and introduces a double bond into the Δ9-position of fatty acyl chains linked to the sn-2 position of existing glycerophospholipids. The other enzyme operates similarly, but it works with acyl-CoA derived from externally supplied SFAs (exogenous) [46]. It is advisable to specify the site of desaturation in the nomenclature whenever possible. For example, a DesA enzyme that introduces a double bond at the Δ9 position should be named DesA9, while a DesB enzyme that catalyzes desaturation at the Δ9 position would be named DesB9. To further distinguish homologous enzymes across bacterial species, a species-specific prefix should be included. For example, in *P. aeruginosa*, a membrane-associated Δ9-desaturase that acts on endogenous FAs would be named PaDesA9, whereas a Δ9-desaturase that modifies exogenous SFAs could be named PaDesB9.

**5. FADs, which modify fatty acids to produce lipid species that do not directly contribute to plasma membrane flexibility**

A recently identified FAD, CarF, was described in *Myxococcus xanthus* (*M. xanthus*), where it catalyzes the conversion of an ether bond into a vinyl ether bond. These lipids are referred to as plasmenylphospholipids (plasmalogens) and plasmanylphospholipids, respectively [47,48].
Plasmalogens are glycerophospholipids characterized by a hydrocarbon chain linked via a vinyl ether bond at the glycerol sn-1 position. They are present in animals and some anaerobic bacteria but are absent in plants, fungi, and most aerobic bacteria, except for myxobacteria and a few other groups, such as *Leptospiraceae* and certain *Alphaproteobacteria*. In anaerobic bacteria like *Clostridium perfringens* (*C. perfringens*), plasmalogen biosynthesis occurs via an oxygen-independent mechanism involving a multidomain enzyme complex similar to benzoyl-CoA reductase/hydroxyacyl-CoA dehydratase [49].

Furan fatty acids (FuFAs) are distinct lipid components found in plants, fish, and microbes. These fatty acids, which can have zero, one, or two methyl groups, act as second messengers in pathways protecting cells from membrane-damaging agents. UfaD, a FA-modifying enzyme isolated from *Cereibacter* spp. (formerly known as *Rhodobacter* spp.), contains a cofactor (FAD/Nicotinamide adenine dinucleotide (NAD))-binding domain. It shares significant amino acid similarity with reductases and dehydrogenases, including phytoene dehydrogenases and desaturases involved in carotenoid biosynthesis. UfaD modifies dimethylated vaccenic acid (trans-C18:1) to produce a dimethylated diunsaturated C18 FA, which is subsequently processed by UfaO, a potential new class of fatty acyl monooxygenases, to generate FuFA [50].

Additionally, a unique type of FAD, initially identified in plants, basidiomycetes, and insects, has been discovered in plant-associated bacteria, including both pathogenic species (e.g., *Trinickia caryophylli* (*T. caryophylli*)) and plant-protective species (e.g., *Pseudomonas protegens* (*P. protegens*)and *Paraburkholderia graminis* (*P. graminis*)). This FAD operates through either a single desaturase or a combination of desaturase and acetylenase activities, introducing C-C double bonds and subsequently C-C triple bonds to form bacterial polyynes. These specialized FAs, known for their bioactivities such as antibiotic or antifungal properties, fall outside the scope of this review [51–53].

**6. How to identify desaturases in non-photosynthetic bacteria?**

Several features can help identify FADs among uncharacterized protein sequences, including histidine-rich motifs, transmembrane domains, and their electron source (see Table 1). However, these features are not exclusive to FADs, as they are also found in other enzyme families such as hydroxylases and monooxygenases. Genetic organization can provide additional clues, as predicted oxidoreductases are often located in adjacent chromosomal regions, assisting in the identification of FADs.

**6.1 Histidine-metal system**

The few available crystal structures of membrane-bound FAD remain limited and are recent, primarily due to the challenges posed by their membrane association, which complicates protein extraction, solubilization, stabilization, and crystal formation [54]. Despite these difficulties, advances in technology have enabled the crystallization and structural elucidation of membrane-associated desaturases. These crystal structures have provided valuable insights into the structure, binding mechanisms, and biocatalytic activity of the enzyme. For example, the X-ray structure of Δ9 mouse stearoyl-CoA desaturase was solved at 2.6 Å resolution [55], and the 3.25 Å crystal structure of human stearoyl-coenzyme A desaturase-1 (SCD1) in complex with its substrate was determined [56]. In these structures, the predicted three conserved histidine-rich motifs - 'HX3 (or X4)H', 'HX2 (or X3)HH', and 'H/QX2 (or X3)HH' (Table 1) - were clearly identified as critical for enzymatic activity by coordinating the dimetal catalytic center. Specifically for non-photosynthetic bacteria, a diiron cluster, positioned deep
inside the center of acyl-lipid desaturase Δ5-Des from *Bacillus cereus* (*B. cereus*) has been discovered by topological details. It adopts a symmetrical style to coordinate with histidine box regions: His72-His76, His108-His112 and His270-His274 (Figure 3) [57].

The *M. tuberculosis* acyl-ACP DesA2, a soluble FAD, was successfully characterized in 2005 at 2.0 Å resolution due to its solubility. Interestingly, its metal-binding motif, (D/E)X±40(D/E)X2HX±60(D/E)X±40(D/E)X2H, differs from those found in membrane-bound desaturases, but is also catalytically essential through coordination of a diiron center [23]. Recently, homologues of DesA2 have been identified in mycolata, including species of *Nocardia* and *Rhodococcus*, where DesA2 is associated with the biosynthesis of long-chain mycolic acids [24].

## 6.2 Transmembrane domains

Membrane-bound FADs feature multiple transmembrane (TM) domains that anchor the protein within the lipid bilayer of the cell membrane [58]. The number and arrangement of these TM regions may differ among desaturase enzymes. Until now, the crystal structures of bacterial desaturases have not been fully resolved [59,60]. However, some structural predictions of bacterial desaturases have been established. Δ5-Des from *Bacillus subtilis* (*B. subtilis*), which introduces a cis-double bound at the Δ5 position (Δ5C-Des) of a wide range of SFAs located in bilayer phospholipids, was the first identified desaturase in non-photosynthetic bacteria [61]. It was reported with one peripheral membrane-associated domain and six TM segments with both N- and C-terminal parts oriented towards the cytoplasm [61–63]. The abundance of hydrophobic stretches in the amino acid sequences of Δ5-Des is consistent with the existence of many TM regions. The deduced hydropathy profile of this desaturase is similar to that of eukaryotic membrane-bound desaturases [63]. The conformation of membrane desaturases, featuring a crack, allows substrates to approach horizontally to the binding cavity. This conformation has been recognized to host distinct active sites compared to soluble desaturases [64]. A similar observation has been noted in the Δ5-Des<sub>Bcereus</sub> [57]. The proposed topological model of Δ5-Des<sub>Bcereus</sub> suggests four transmembrane segments (TM1-4) and two peripheral helices (P1-2). Among them, one constitutes the cytosolic catalytic domain, primarily composed of helical secondary structures, as depicted in Figure 3. The N- and C-termini of Δ5-Des<sub>Bcereus</sub> are located on the cytosolic side, and each segment of TM1, TM2, TM3 and TM4 comprises 19 (Val25-Tyr43), 20 (Tyr50-Ile69), 21 (Tyr185-Trp203) and 20 (Leu207-Val226) amino acids, respectively. This structure aligns with the hypothesized crystal homology model of the human SCD1, featuring a mushroom-like architecture [56,64]. One notable difference in the Δ5-Des<sub>Bcereus</sub> structure is the position of the helix between residues 146 and 160, which is parallel to the bilayer membrane but embedded into it. This structural difference between P2 and P1 may have functional implications for its activity.

The study also unveiled the crucial role of Tyr40, demonstrating its indispensable contribution to the accurate orientation of the TM1 within the plasma membrane [57]. It was speculated that the amphiphilic amino acid Tyr40, which anchors the TM helix to cell membrane, may orient towards the water phase, allowing the acyl chain to access the substrate tunnel directly from the lipid bilayer to cytosolic domain of the desaturase [65]. According to these mechanisms, Tyr40 could guarantee the stereospecificity and regioselectivity for the desaturation reaction. Moreover, all these analogous residues (yellow) may contribute to the localization of the desaturase to the membrane (Figure 3).
<figure><img src="image_7.png" /><figcaption>**Figure 3.** Schematic representation of the topological model of membrane-bound desaturase DesA from *B. cereus* [57].</figcaption></figure>

**6.3 Electron transport system**

It has been shown that domains located near desaturase genes are involved in electron transport. For example, the *desB* gene in *P. aeruginosa* and *desA* in *M. tuberculosis* have predicted oxidoreductases in adjacent chromosomal regions that play roles in the electron transport chain required for FA desaturation [46,66]. In *A. baumannii* AB5075, only the putative desaturase gene *ABUW_3842* has an obvious adjacent putative oxidoreductase (*ABUW_3843*) [12]. Four desaturases, encoded by *MSMEG_1886*, *MSMEG_1743*, *MSMEG_6835*, and *MSMEG_1211*, were identified in *Mycobacterium smegmatis* (*M. smegmatis*). Genes encoding putative oxidoreductases are located upstream of *MSMEG_1886* (*desA3*), *MSMEG_1743*, and *MSMEG_6835*, but not upstream of *MSMEG_1211* [67]. The existence of an electron transport mechanism, enabling the transfer of electrons from the nicotinamide adenine dinucleotide phosphate (NAD(P)H) cofactor, is a critical requirement for FA desaturation processes [43]. The general catalytic mechanism of FA desaturases involves a dehydrogenation reaction [68]. Further studies have shown that the high-valent metal center, often containing diiron clusters, can form reactive radical intermediates to dehydrogenate the methylene substrate of FAs [68]. In bacteria, three structurally distinct but functionally comparable electron donor systems have been identified: ferredoxin (Fer) and two flavodoxin (Fld) units, YkuN and YkuP [69,70]. YkuN and YkuP are isofunctional with Fer and serve as redox partners, providing electrons for Δ5-desaturase activity in *B. subtilis* [71]. Disruption of either the *fer* gene or the *ykuNOP* operon significantly decreases the production of palmitoleic acid (C16:1). The electron transfer mechanism in bacteria is generally similar to that of plant desaturases [66]. Two distinct but functionally equivalent electron transfer systems have been identified for bacterial FA desaturation: the NAD(P)H/ferredoxin-NAD(P)+ oxidoreductase (FNR)/Fer system and the NAD(P)H/FNR/Fld system, as illustrated in Figure 4. In these systems, NAD(P)H provides the two electrons required for double bond formation [69,72]. Electrons are transferred from NAD(P)H to the carrier protein FNRox, which acts as a 'step-down transformer'. Each of the reduced carrier proteins, Fer<sub>red</sub> or Fld<sub>red</sub>, can transfer one electron at a time to the metal catalytic center of the desaturase. This sequential electron transfer reduces O<sub>2</sub> and ultimately allows the formation of double bonds in acyl lipid chains.
<figure><img src="image_8.png" /><figcaption>**Figure 4.** Putative schematic representation of the electron transport mechanism of bacterial FA desaturation reaction [69]. According to the type of desaturases there are two possible electron donor systems involved in the catalytic process: NAD(P)H/Fer-NAD(P)⁺-FNR/Fer and NAD(P)H/FNR/Fld.</figcaption></figure>
<table><caption>**Table 1.** An overview of potential catalytic mechanisms factors in significant desaturase studies from various non-photosynthetic bacterium species. Ferredoxin (Fer) and two flavodoxin (Fld). When no specific details are provided, it is because they have not been clearly demonstrated.</caption><thead><tr><td>Organism</td><td>Desaturase</td><td>Catalytic motif</td><td>Metal center</td><td>Redox partners</td><td>Ref</td></tr></thead><tbody><tr><td>Acinetobacter baumannii</td><td>Δ9-Des</td><td>HX<sub>4</sub>H, HX<sub>2</sub>HH, HX<sub>3</sub>HH (DesA) HX<sub>3</sub>H, HX<sub>2</sub>HH, HX<sub>2</sub>HH (DesB) HX<sub>3</sub>H, HX<sub>3</sub>HH, HX<sub>3</sub>HH (DesC)</td><td>-</td><td>-</td><td>[12][21]</td></tr><tr><td>Bacillus spp.</td><td>Δ5-Des</td><td>HX<sub>3</sub>H, HX<sub>2</sub>HH, HX<sub>3</sub>HH</td><td>Diiron</td><td>Fer, Fld</td><td>[61][57][73][69]</td></tr><tr><td>Mycobacterium spp., Nocardia spp., Rhodococcus spp.</td><td>MSMEG_1211, MSMEG_1743 (homologue of P. aeruginosa DesB), MSMEG_6835, DesA1, DesA2, DesA3</td><td>HX<sub>3</sub>H, HX<sub>3</sub>HH, QX<sub>2</sub>HH</td><td>Diiron, Mn<sup>2+</sup></td><td>-</td><td>[23][24][74]</td></tr><tr><td>Pseudomonas aeruginosa</td><td>Δ9-Des</td><td>HX<sub>4</sub>H, HX<sub>2</sub>HH, HX<sub>2</sub>HH (DesA) HX<sub>3</sub>H, HX<sub>3</sub>HH, HX<sub>3</sub>HH (DesB)</td><td>-</td><td>-</td><td>[46]</td></tr><tr><td>Pseudomonas sp.</td><td>Δ9-Des</td><td>HX<sub>4</sub>H, HX<sub>2</sub>HH, HX<sub>2</sub>HH HX<sub>3</sub>H, HX<sub>2</sub>HH, HX<sub>2</sub>HH</td><td>Diiron</td><td>-</td><td>[75][76][77]</td></tr><tr><td>Pseudoalteromonas sp.</td><td>Δ9-Des</td><td>HX<sub>4</sub>H, HX<sub>2</sub>HH, HX<sub>2</sub>HH</td><td>-</td><td>-</td><td>[78][79]</td></tr><tr><td>Psychrobacter sp.</td><td>Δ9-Des</td><td>HX<sub>4</sub>H, HX<sub>2</sub>HH, HX<sub>2</sub>HH</td><td>-</td><td>-</td><td>[79]</td></tr><tr><td>Rhodococcus sp.</td><td>Δ6-Des</td><td>HX<sub>3</sub>H, HX<sub>3</sub>HH, QX<sub>2</sub>HH</td><td>Diiron</td><td>-</td><td>[80]</td></tr><tr><td>Myxococcus xanthus</td><td>CarF (vinyl ether bond synthesis)</td><td>HX<sub>3</sub>H, HX<sub>2</sub>HH</td><td>Diiron</td><td>-</td><td>[47]</td></tr><tr><td>Cereibacter spp. (Formerly known as Rhodobacter spp.)</td><td>Δ10-Des (associated with methylated furan FAs, named UfaD)</td><td>HX<sub>3</sub>H, HX<sub>2</sub>H</td><td>-</td><td>-</td><td>[81]</td></tr><tr><td>Trinickia caryophylli</td><td>CayC, CayB, CayE (polyynes synthesis)</td><td>HX<sub>4</sub>H, HX<sub>2</sub>HH</td><td>-</td><td>-</td><td>[51]</td></tr></tbody></table>
Q/HX₂HH

<figure><img src="image_9.png" /></figure>
# 7. Regulatory mechanisms of FAD and their impact on bacterial pathogenicity

Fine regulation of UFAs biosynthesis in response to the adjustments of intracellular and extracellular environment is one of the survival rules for microorganisms [82]. The biochemical relevance of bacterial FAs desaturation is mainly related to two factors: temperature fluctuation and feedback regulation mechanism. Fluctuation in temperature could induce membrane GPLs to undergo a reversible change between liquid ordered (Lo) and liquid disordered (Ld) phases [83,84]. The Ld state is caused in part by UFAs, which can counteract the rigid state brought by the close packing of SFAs. Expression of desaturases in bacteria increases the flexibility of membrane lipids toward FAs to maintain intracellular homeostasis in response to changes in ambient temperature (Figure 5A) [85].

## 7.1 Regulatory mechanisms of FA desaturation

A well-studied example is the *B. subtilis* Δ5-DesB, which is controlled by the DesK/DesR two-component system (TCS). This system comprises a membrane-bound histidine kinase (DesK) lacking an extracellular domain and a cytoplasmic response regulator (DesR) (Figure 5B) [61,86,87]. DesK senses changes in membrane fluidity, triggered by either temperature fluctuations or the saturation level of membrane phospholipids. When membrane fluidity decreases, DesK adopts a kinase-dominant state, autophosphorylating a conserved histidine residue. The phosphate group is then transferred to DesR, activating its DNA-binding domain. This activation induces the expression of the *des* gene, enabling Δ5-DesB to introduce double bonds into fatty acyl chains. This desaturation reduces membrane thickness, increases fluidity, and restores optimal membrane function. Conversely, when the membrane becomes too fluid, DesK transitions to a phosphatase-dominant state, dephosphorylates DesR. This prevents DesR from binding to the *des* promoter, repressing *des* gene expression. As a result, UFA production decreases the membrane thickness and rigidity. Similar regulatory mechanisms have been observed in *B. cereus*, where DesA and DesB catalyze double bond formation at Δ5 and Δ10 positions, respectively [88]. In mutants lacking the CasK/R system, the downregulation of *desA* and *desB* genes results in reduced UFA production, underscoring the importance of CasK/R in regulating DesA activity. Recent studies in *B. subtilis* have shown that the TCS YvfT/YvfU regulates the expression of the *yvfRS* operon, encoding a putative ABC transporter, through a synergistic interaction with the TCS DesK/DesR. This cross-regulation allows the integration of environmental signals and ensures that transporter expression is coordinated with membrane fluidity conditions. The *yvfRS* operon was confirmed as the regulatory target of YvfT/YvfU, and its induction requires the combined action of YvfU and DesR, as neither response regulator is sufficient alone. Notably, the promoter of *yvfRS* (*PyvfR*) contains binding motifs that closely resemble those of *Pdes*, the DesR-regulated promoter [89]. In *P. aeruginosa*, DesT functions as an on-off switch that responds with high affinity to different ligand structures. DesT shows strong affinity for its specific palindromic DNA binding site when bound to unsaturated acyl-CoA, but fails to bind this site when bound to saturated acyl-CoA. This ability to respond to different ligand shapes allows DesT to act as a rheostat, maintaining membrane lipid homeostasis based on the ratio of UFAs to SFAs in phospholipids. The presence of similar compositional sensors, such as *E. coli* FabR and *Streptococcus pneumoniae* (*S. pneumoniae*) FabT, suggests that these regulatory mechanisms may be widespread in nature [90]. In mycobacteria, the desaturation of mycolic acids, which are essential components of their distinctive cell envelope, is tightly regulated by an acyl-CoA-dependent mechanism involving the protein MadR [91]. MadR serves as a critical regulator of mycolic acid desaturation, effectively linking lipid biosynthesis to metabolic signals through its
interaction with acyl-CoA. This regulatory system plays an important role in maintaining precise control over the structure and function of mycolic acids, which is essential for the integrity and adaptability of the mycobacterial cell envelope. By ensuring the appropriate level of desaturation, MadR contributes to the resilience of mycobacteria under diverse environmental conditions, highlighting the importance of lipid metabolism for bacterial survival and pathogenicity. Collectively, these findings reveal that such sensors and regulatory pathways are crucial for bacterial physiology.

## 7.2 Impact of the oxygen

Oxygen availability is a critical determinant of the activity of FAD because they perform dehydrogenation reactions that require oxygen. For example, in the amoeba *Acanthamoeba castellanii* (*A. castellanii*), increasing the oxygen concentration from very low levels to near air saturation resulted in a 2.3-fold increase in n-6 desaturase activity, accompanied by increased lipid unsaturation [92]. Conversely, low oxygen conditions (hypoxia), such as those found in the tumor microenvironment, can negatively affect desaturase activity [93]. In the yeast *Schizosaccharomyces japonicus* (*S. japonicus*), oxygen deprivation triggers the production of asymmetric acyl tails that replace UFAs, enabling cells to preserve membrane properties and maintain fitness. This adaptation is accompanied by modifications in GPL headgroups and sphingolipid precursors, such as ceramides. Interestingly, deletion of the DesΔ9 FAD led to a comparable shift toward fully saturated, asymmetric lipids. At 37°C, neither the absence of oxygen nor the deletion of desaturases compromised cell fitness, suggesting that saturated, asymmetric lipids can effectively substitute for unsaturated counterparts under these conditions. However, this compensatory mechanism is insufficient at lower temperatures, as cells grown at 24°C exhibited growth defects. Thus, while asymmetric lipids represent an adaptive strategy to maintain membrane fluidity in the absence of molecular oxygen, their effectiveness is limited, particularly under suboptimal thermal conditions [94]. In *P. aeruginosa*, the transcription of the desA gene is significantly enhanced under anaerobic conditions, suggesting a potential adaptation mechanism for scavenging the limited oxygen in its environment [17,46]. Conversely, when *Lactobacillus helveticus* (*L. helveticus*: a facultative anaerobe) is exposed to oxidative stress, the activity of FAD increases, promoting oxygen consumption and leading to an increase in UFAs [95].
<figure><img src="image_10.png" /><figcaption>**Figure 5.** Model of signal transduction pathway for regulation of desaturases functionality in bacteria. A) Desaturase expression and activity sensed by temperature regulation. B) Typical feedback mechanism of regulation of UFAs biosynthesis in *B. subtilis*.</figcaption></figure>

7.3 The role of FA desaturation in bacterial virulence

*Staphylococcus aureus* (S. aureus) strains that are resistant to thrombin-induced platelet microbicidal proteins have higher levels of UFAs than their sensitive parental strains. This suggests that resistance mechanisms may be related to alterations in FA desaturation, which may affect the ability of bacteria to withstand the antimicrobial effects of these proteins [96]. Similarly, resistant strains of *Listeria monocytogenes* (*L. monocytogenes*) to class IIa bacteriocins have increased levels of unsaturated phosphatidylglycerol (PG), which leads to increased membrane fluidity that can hinder the effective insertion of bacteriocins. In contrast, susceptible strains have a higher concentration of saturated PGs, which promote greater susceptibility to bacteriocin action [97]. In *P. aeruginosa*, the *desB* mutants displayed notable deficiencies in the production of key virulence factors, such as proteolytic enzymes, pyocyanin, and rhamnolipid. Additionally, these mutants exhibited impaired swarming and twitching motilities, leading to a reduced level of virulence in the *Caenorhabditis elegans* (*C. elegans*) infection model [34,98]. A recent study showed that the regulator FabR (also known as DesT), which is involved in UFA biosynthesis, can reduce the robustness of *Klebsiella pneumoniae* (*K. pneumoniae*) by altering its FA composition. It increases the susceptibility of the biofilm to detachment under dynamic conditions by weakening it and making it more rigid and compact, which is likely to facilitate the biofilm dispersion process [99]. A conserved desaturase, DesC, in *A. baumannii*, which is regulated by DesR and plays a critical role in converting C18:0 to C18:1n-9, supporting the pathogen's adaptation to host niches [21]. Mutants lacking DesC showed significant attenuation in systemic infections, particularly in the blood and pleural cavity, highlighting its importance in virulence during host adaptation. These findings suggest that FA desaturation is a critical factor in bacterial pathogenicity, making it a promising new target for the development of antibacterial treatments.
8. Conclusions and future perspectives

This review provides a comprehensive overview of the FADs found in non-photosynthetic bacteria. Desaturases are integral members of a diverse family of enzymes that modify FAs by the insertion of double bonds. These enzymes play critical roles in essential physiological functions, including cellular homeostasis and stress response. While some soluble bacterial FADs have been successfully purified and studied, the study of membrane-bound FADs presents significant challenges. This difficulty limits our ability to accurately characterize their physical and chemical properties, as well as their reaction mechanisms. However, recent advances in deep learning-based protein folding prediction models, such as AlphaFold2 and RoseTTAFold, offer promising avenues for improving our understanding of these complex enzymes in the future [100,101]. In addition, the emergence of innovative biotechnologies such as cryo-correlative light and electron microscopy (cryo-CLEM), cryo-electron tomography (cryo-ET), and cryo-X-ray tomography may help overcome existing research limitations. These advanced techniques have the potential to revolutionize lipidomic and proteomic studies, paving the way for the exploration of alternative sources of unsaturated fatty acids (UFAs) and novel methods derived from the study of membrane proteins. By integrating these cutting-edge approaches, researchers can gain deeper insights into the functional roles of desaturases and their importance in microbial metabolism and adaptability.
References

1. Renne, M.F.; Ernst, R. Membrane homeostasis beyond fluidity: control of membrane compressibility. *Trends Biochem. Sci.* **2023**, *48*, 963–977, doi:10.1016/j.tibs.2023.08.004.

2. Tamby, A.; Sinninghe Damsté, J.S.; Villanueva, L. Microbial membrane lipid adaptations to high hydrostatic pressure in the marine environment. *Front. Mol. Biosci.* **2023**, *9*, 1058381-1058392, doi: 10.3389/fmolb.2022.1058381.

3. Sohlenkamp, C.; Geiger, O. Bacterial membrane lipids: Diversity in structures and pathways. *FEMS Microbiol. Rev.* **2015**, *40*, 133–159, doi:10.1093/femsre/fuv008.

4. Oursel, D.; Loutelier-Bourhis, C.; Orange, N.; Chevalier, S.; Norris, V.; Lange, C.M. Identification and relative quantification of fatty acids in *Escherichia coli* membranes by gas chromatography/mass spectrometry. *Rapid Commun. Mass Spectrom.* **2007**, *21*, 3229–3233, doi:10.1002/rcm.3177.

5. Ishida, Y.; Kitagawa, K.; Nakayama, A.; Ohtani, H. Complementary analysis of lipids in whole bacteria cells by thermally assisted hydrolysis and methylation-GC and MALDI-MS combined with on-probe sample pretreatment. *J. Anal. Appl. Pyrolysis* **2006**, *77*, 116–120, doi:10.1016/j.jaap.2006.02.006.

6. Gozdzik, P.; Magkos, F.; Sledzinski, T.; Mika, A. Monomethyl branched-chain fatty acids: health effects and biological mechanisms. *Prog. Lipid Res.* **2023**, *90*, 101226, doi:10.1016/j.plipres.2023.101226.

7. Melchionna, M. V.; Gullett, J.M.; Bouveret, E.; Shrestha, H.K.; Abraham, P.E.; Hettich, R.L.; Alexandre, G. Bacterial homologs of progestin and AdipoQ receptors (PAQRs) affect membrane energetics homeostasis but not fluidity. *J. Bacteriol.* **2022**, *204*, e00583-21. doi:10.1128/jb.00583-21.

8. Diomandé, S.E.; Nguyen-The, C.; Guinebretière, M.H.; Broussolle, V.; Brillard, J. Role of fatty acids in *Bacillus* environmental adaptation. *Front. Microbiol.* **2015**, *6*, 1–20, doi:10.3389/fmicb.2015.00813.

9. Passos, L.S.; de Freitas, P.N.N.; Menezes, R.B.; de Souza, A.O.; Silva, M.F. da; Converti, A.; Pinto, E. Content of lipids, fatty acids, carbohydrates, and proteins in continental cyanobacteria: A systematic analysis and database application. *Appl. Sci.* **2023**, *13*, 3162-3190, doi:10.3390/app13053162.

10. Okuyama, H.; Orikasa, Y.; Nishida, T.; Watanabe, K.; Morita, N. Bacterial genes responsible for the biosynthesis of eicosapentaenoic and docosahexaenoic acids and their heterologous expression. *Appl. Environ. Microbiol.* **2007**, *73*, 665–670, doi:10.1128/AEM.02270-06.

11. Nachtschatt, M.; Okada, S.; Speight, R. Integral membrane fatty acid desaturases: a review of biochemical, structural, and biotechnological advances. *Eur. J. Lipid Sci. Technol.* **2020**, *122*, 2000181. doi: 10.1002/ejlt.202000181.

12. Adams, F.G.; Pokhrel, A.; Brazel, E.B.; Semenec, L.; Li, L.; Trappetti, C.; Paton, J.C.; Cain, A.K.; Paulsen, I.T.; Eijkelkamp, B.A. *Acinetobacter baumannii* fatty acid desaturases facilitate survival in distinct environments. *ACS Infect. Dis.* **2021**, *7*, 2221–2228. doi: 10.1021/acsinfecdis.1c00192.

13. Vergoz, D.; Schaumann, A.; Schmitz, I.; Afonso, C.; Dé, E.; Loutelier-Bourhis, C.; Alexandre, S. Lipidome of *Acinetobacter baumannii* antibiotic persister cells. *Biochim.*
*Biophys. Acta - Mol. Cell Biol. Lipids* **2024**, *1869*, 159539.
doi:10.1016/j.bbalip.2024.159539.

14. De Carvalho, C.C.C.R.; Marques, M.P.C.; Hachicho, N.; Heipieper, H.J. Rapid adaptation of *Rhodococcus erythropolis* cells to salt stress by synthesizing polyunsaturated fatty acids. *Appl. Microbiol. Biotechnol.* **2014**, *98*, 5599–5606, doi:10.1007/s00253-014-5549-2.

15. Moi, I.M.; Thean, A.; Leow, C.; Shukuri, M.; Ali, M.; Noor, R.; Raja, Z.; Rahman, A.; Salleh, A.B.; Sabri, S. Polyunsaturated fatty acids in marine bacteria and strategies to enhance their production. *Appl. Microbiol. Biotechnol.* **2018**, *102*, 5811–5826. doi:10.1007/s00253-018-9063-9.

16. Holthuis, J.C.M.; Menon, A.K. Lipid landscapes and pipelines in membrane homeostasis. *Nature* **2014**, *510*, 48–57, doi:10.1038/nature13474.

17. Cronan, J.E. Unsaturated fatty acid synthesis in bacteria: mechanisms and regulation of canonical and remarkably noncanonical pathways. *Biochimie* **2024**, *218*, 137-151. doi:10.1016/j.biochi.2023.09.007

18. Bloch, K. Enzymatic synthesis of monounsaturated fatty acids. *Acc. Chem. Res.* **1969**, *2*, 193–202, doi:10.1021/ar50019a001.

19. Spiering, M.J. The work of Konrad Bloch's laboratory on unsaturated fatty acid biosynthesis in bacteria. *J. Biol. Chem.* **2019**, *294*, 14876–14878, doi:10.1074/jbc.CL119.010972.

20. Cronan, J.E.; Birge, C.H.; Vagelos, P.R. Evidence for two genes specifically involved in unsaturated fatty acid biosynthesis in *Escherichia coli*. *J. Bacteriol.* **1969**, *100*, 601–604, doi:10.1128/JB.100.2.601-604.1969.

21. Adams, F.G.; Alquethamy, S.; Waters, J.K.; Davies, B.J.; Haracic, E.; Nanson, J.D.; Paton, J.C.; Forwood, J.K.; Hassan, K.A.; Brazel, E.B.; et al. *Acinetobacter baumannii* employs a rare fatty acid desaturase for niche-specific host adaptation. *ACS Infect. Dis.* **2025**, doi:10.1021/acsinfecdis.4c00765.

22. Dessenne, C.; Ménart, B.; Acket, S.; Dewulf, G.; Guerardel, Y.; Vidal, O.; Rossez, Y. Lipidomic analyses reveal distinctive variations in homeoviscous adaptation among clinical strains of *Acinetobacter baumannii*, providing insights from an environmental adaptation perspective. *Microbiol. Spectr.* **2024**, *12*, 1–19, doi:10.1128/spectrum.00757-24.

23. Dyer, D.H.; Lyle, K.S.; Rayment, I.; Fox, B.G. X-ray structure of putative acyl-ACP desaturase DesA2 from *Mycobacterium tuberculosis* H37Rv. *Protein Sci.* **2009**, *14*, 1508–1517. doi:10.1110/ps.041288005.

24. Bailo, R.; Radhakrishnan, A.; Singh, A.; Nakaya, M.; Fujiwara, N.; Bhatt, A. The mycobacterial desaturase DesA2 is associated with mycolic acid biosynthesis. *Sci. Rep.* **2022**, *12*, 1–10, doi:10.1038/s41598-022-10589-y.

25. Tocher, D.R.; Leaver, M.J.; Hodgson, P.A. Recent advances in the biochemistry and molecular biology of fatty acyl desaturases. *Prog. Lipid Res.* **1998**, *37*, 73–117, doi:10.1016/S0163-7827(98)00005-8.

26. Shanklin, J.; Cahoon, E.B. Desaturation and related modifications of fatty acids. *Annu. Rev. Plant Physiol. Plant Mol. Biol.* **1998**, *49*, 611–641, doi:10.1146/annurev.arplant.49.1.611.
27. Los, D.; Mironov, K. Modes of fatty acid desaturation in *Cyanobacteria: an update*. *Life* **2015**, *5*, 554–567, doi:10.3390/life5010554.

28. Gohrbandt, M.; Lipski, A.; Grimshaw, J.W.; Buttress, J.A.; Baig, Z.; Herkenhoff, B.; Walter, S.; Kurre, R.; Deckers-Hebestreit, G.; Strahl, H. Low membrane fluidity triggers lipid phase separation and protein segregation in living bacteria. *EMBO J.* **2022**, *41*, 1–21, doi:10.15252/embj.2021109800.

29. Budin, I.; Rond, T. de; Chen, Y.; Chan, L.J.G.; Petzold, C.J.; Keasling, J.D. Viscous control of cellular respiration by membrane lipid composition. *Science*. **2018**, *362*, 1186–1189, doi:10.1126/science.aat7925.

30. Cronan, J.E.; Thomas, J. Chapter 17 Bacterial fatty acid synthesis and its relationships with polyketide synthetic pathways. *Methods Enzymol.* **2009**, *459*, 395–433. doi: 10.1016/S0076-6879(09)04617-5.

31. Lee, H.; Park, W.J. Unsaturated fatty acids, desaturases, and human health. *J. Med. Food* **2014**, *17*, 189–197. doi: 10.1089/jmf.2013.2917

32. Yoon, Y.; Lee, H.; Lee, S.; Kim, S.; Choi, K.H. Membrane fluidity-related adaptive response mechanisms of foodborne bacterial pathogens under environmental stresses. *Food Res. Int.* **2015**, *72*, 25–36. doi:10.1016/j.foodres.2015.03.016.

33. de Mendoza, D.; Pilon, M. Control of membrane lipid homeostasis by lipid-bilayer associated sensors: a mechanism conserved from bacteria to humans. *Prog. Lipid Res.* **2019**, *76*, 100996. doi: 10.1016/j.plipres.2019.100996.

34. Schweizer, H.P.; Choi, K.-H. *Pseudomonas aeruginosa* aerobic fatty acid desaturase DesB is important for virulence factor production. *Arch. Microbiol.* **2011**, *193*, 227–234, doi:10.1007/s00203-010-0665-6.

35. Sakamoto, T.; Murata, N. Regulation of the desaturation of fatty acids and its role in tolerance to cold and salt stress. *Curr. Opin. Microbiol.* **2002**, *5*, 206–210. doi:10.1016/S1369-5274(02)00306-5.

36. Ishii, E.; Eguchi, Y. Diversity in sensing and signaling of bacterial sensor histidine kinases. *Biomolecules* **2021**, *11*, 1524-1551. doi:10.3390/biom11101524.

37. Chwastek, G.; Surma, M.A.; Rizk, S.; Grosser, D.; Lavrynenko, O.; Rucińska, M.; Jambor, H.; Sáenz, J. Principles of membrane adaptation revealed through environmentally induced bacterial lipidome remodeling. *Cell Rep.* **2020**, *32*, 108165, doi:10.1016/j.celrep.2020.108165.

38. Tao, Y.; Acket, S.; Beaumont, E.; Galez, H.; Duma, L.; Rossez, Y. Colistin treatment affects lipid composition of *Acinetobacter baumannii*. *Antibiotics* **2021**, *10*, 528 doi:10.3390/antibiotics10050528.

39. Justice, I.; Saenz, J.P. A tuneable minimal cell membrane reveals that two lipids suffice for life. *bioRxiv* **2023**, *10*, 563757, doi:10.1038/s41467-024-53975-y.

40. Heipieper, H.J.; Meinhardt, F.; Segura, A. The *cis-trans* isomerase of unsaturated fatty acids in *Pseudomonas* and *Vibrio*: Biochemistry, molecular biology and physiological function of a unique stress adaptive mechanism. *FEMS Microbiol. Lett.* **2003**, *229*, 1–7, doi:10.1016/S0378-1097(03)00792-4.

41. Mauger, M.; Ferreri, C.; Chatgilialoglu, C.; Seemann, M. The bacterial protective armor
against stress: The *cis-trans* isomerase of unsaturated fatty acids, a cytochrome-c type enzyme. *J. Inorg. Biochem.* **2021**, *224*, doi:10.1016/j.jinorgbio.2021.111564.

42. Raza, S.; Sievertsen, T.; Jafari, M.; Vermaas, J. V. BPS2025 - Dynamics and structural changes associated with *cis-trans* isomerization in bacterial lipid bilayers. *Biophys. J.* **2025**, *124*, 257a, doi: 10.26434/chemrxiv-2025-jm423.

43. Cerone, M.; Smith, T.K. Desaturases: structural and mechanistic insights into the biosynthesis of unsaturated fatty acids. *IUBMB Life* **2022**, *74*, 1036-1051, doi:10.1002/iub.2671.

44. Lee, J.M.; Lee, H.; Kang, S.B.; Park, W.J. Fatty acid desaturases, polyunsaturated fatty acid regulation, and biotechnological advances. *Nutrients* **2016**, *8*, 23-36. doi:10.3390/nu8010023.

45. Wang, M.; Chen, H.; Gu, Z.; Zhang, H.; Chen, W.; Chen, Y.Q. ω3 fatty acid desaturases from microorganisms: structure, function, evolution, and biotechnological use. *Appl. Microbiol. Biotechnol.* **2013**, *97*, 10255-10262. doi:10.1007/s00253-013-5336-5.

46. Zhu, K.; Choi, K.H.; Schweizer, H.P.; Rock, C.O.; Zhang, Y.M. Two aerobic pathways for the formation of unsaturated fatty acids in *Pseudomonas aeruginosa*. *Mol. Microbiol.* **2006**, *60*, 260-273. doi:10.1111/j.1365-2958.2006.05088.x

47. Gallego-García, A.; Monera-Girona, A.J.; Pajares-Martínez, E.; Bastida-Martínez, E.; Pérez-Castaño, R.; Iniesta, A.A.; Fontes, M.; Padmanabhan, S.; Elías-Arnanz, M. A bacterial light response reveals an orphan desaturase for human plasmalogen synthesis. *Science.* **2019**, *366*, 128-132, doi:10.1126/science.aay1436.

48. Koelmel, J.P.; Ulmer, C.Z.; Jones, C.M.; Yost, R.A.; Bowden, J.A. Common cases of improper lipid annotation using high-resolution tandem mass spectrometry data and corresponding limitations in biological interpretation. *Biochim. Biophys. Acta - Mol. Cell Biol. Lipids* **2017**, *1862*, 766-770, doi:10.1016/j.bbalip.2017.02.016.

49. Jackson, D.R.; Cassilly, C.D.; Plichta, D.R.; Vlamakis, H.; Liu, H.; Melville, S.B.; Xavier, R.J.; Clardy, J. Plasmalogen biosynthesis by anaerobic bbacteria: identification of a two-gene operon responsible for plasmalogen production in *Clostridium perfringens*. *ACS Chem. Biol.* **2021**, *16*, 6-13, doi:10.1021/acscchembio.0c00673.

50. Lemke, R.A.S.; Olson, S.M.; Morse, K.; Karlen, S.D.; Higbee, A.; Beebe, E.T.; Ralph, J.; Coon, J.J.; Fox, B.G.; Donohue, T.J. A bacterial biosynthetic pathway for methylated furan fatty acids. *J. Biol. Chem.* **2020**, *295*, 9786-9801, doi:10.1074/jbc.RA120.013697.

51. Ross, C.; Scherlach, K.; Kloss, F.; Hertweck, C. The molecular basis of conjugated polyyne biosynthesis in phytopathogenic bacteria. *Angew. Chemie - Int. Ed.* **2014**, *53*, 7794-7798, doi:10.1002/anie.201403344.

52. Kim, H.J.; Ishida, K.; Hertweck, C. Thiotemplated biosynthesis of bacterial polyyne fatty acids by a designated desaturase triad. *ChemBioChem* **2022**, *23*, 7-11, doi:10.1002/cbic.202200430.

53. Suenaga, M.; Katayama, N.; Kitamura, K.; Kai, K. Structures and biosynthesis of caryoynencins, unstable bacterial polyynes from *Pseudomonas protegens* recombinant expressing the *cayG* gene. *J. Org. Chem.* **2023**, *88*, 16280-16291, doi:10.1021/acs.joc.3c01789.
54. Kermani, A.A. A guide to membrane protein X-ray crystallography. *FEBS J.* **2021**, *288*, 5788-5804, doi:10.1111/febs.15676.

55. Bai, Y.; McCoy, J.G.; Levin, E.J.; Sobrado, P.; Rajashankar, K.R.; Fox, B.G.; Zhou, M. X-ray structure of a mammalian stearoyl-CoA desaturase. *Nature* **2015**, *524*, 252–256, doi:10.1038/nature14549.

56. Wang, H.; Klein, M.G.; Zou, H.; Lane, W.; Snell, G.; Levin, I.; Li, K.; Sang, B.C. Crystal structure of human stearoyl-coenzyme A desaturase in complex with substrate. *Nat. Struct. Mol. Biol.* **2015**, *22*, 581–585, doi:10.1038/nsmb.3049.

57. Sastre, D.E.; Saita, E.; Uttaro, A.D.; De Mendoza, D.; Altabe, S.G. Structural determinant of functionality in acyl lipid desaturases. *J. Lipid Res.* **2018**, *59*, 1871–1879. doi:10.1194/jlr.M085258.

58. Li, D.; Moorman, R.; Vanhercke, T.; Petrie, J.; Singh, S.; Jackson, C.J. Classification and substrate head-group specificity of membrane fatty acid desaturases. *Comput. Struct. Biotechnol. J.* **2016**, *14*, 341–349, doi:10.1016/j.csbj.2016.08.003.

59. Halim, N.F.A.A.; Ali, M.S.M.; Leow, A.T.C.; Rahman, R.N.Z.R.A. Membrane fatty acid desaturase: biosynthesis, mechanism, and architecture. *Appl. Microbiol. Biotechnol.* **2022**, *106*, 5957–5972. doi:10.1007/s00253-022-12142-3.

60. Birch, J.; Axford, D.; Foadi, J.; Meyer, A.; Eckhardt, A.; Thielmann, Y.; Moraes, I. The fine art of integral membrane protein crystallisation. *Methods* **2018**, *147*, 150–162, doi:10.1016/j.ymeth.2018.05.014.

61. Mansilla, M.C.; De Mendoza, D. The *Bacillus subtilis* desaturase: a model to understand phospholipid modification and temperature sensing. *Arch. Microbiol.* **2005**, *183*, 229–235. doi:10.1007/s00203-005-0759-8.

62. Olenic, S.; Buchanan, F.; VanPortfliet, J.; Parrell, D.; Kroos, L. Conserved proline residues of *Bacillus subtilis* intramembrane metalloprotease SpoIVFB are important for substrate interaction and cleavage. *J. Bacteriol.* **2022**, *204*, e00386-21. doi:10.1128/jb.00386-21.

63. Diaz, A.R.; Mansilla, M.C.; Vila, A.J.; De Mendoza, D. Membrane topology of the acyl-lipid desaturase from *Bacillus subtilis*. *J. Biol. Chem.* **2002**, *277*, 48099–48106. doi:10.1074/jbc.M208960200

64. Shanklin, J.; Guy, J.E.; Mishra, G.; Lindqvist, Y. Desaturases: emerging models for understanding functional diversification of diiron-containing enzymes. *J. Biol. Chem.* **2009**, *284*, 18559–18563. doi:10.1074/jbc.R900009200.

65. Liang, J.; Adamian, L.; Jackups, R. The membrane-water interface region of membrane proteins: structural bias and the anti-snorkeling effect. *Trends Biochem. Sci.* **2005**, *30*, 355–357, doi:10.1016/j.tibs.2005.05.003.

66. Chang, Y.; Fox, B.G. Identification of Rv3230c as the NADPH oxidoreductase of a two-protein DesA3 acyl-CoA desaturase in *Mycobacterium tuberculosis* H37Rv. *Biochemistry* **2006**, *45*, 13476–13486, doi:10.1021/bi0615285.

67. Di Capua, C.B.; Doprado, M.; Belardinelli, J.M.; Morbidoni, H.R. Complete auxotrophy for unsaturated fatty acids requires deletion of two sets of genes in *Mycobacterium smegmatis*. *Mol. Microbiol. 2017*, *106*, 93–108, doi:10.1111/mmi.13753.

68. Buist, P.H. Fatty acid desaturases: selecting the dehydrogenation channel. *Nat. Prod.*
Rep. **2004**, *21*, 249–262. doi:10.1039/b302094k.

69. Chazarreta-Cifre, L.; Martiarena, L.; de Mendoza, D.; Altabe, S.G. Role of ferredoxin and flavodoxins in *Bacillus subtilis* fatty acid desaturation. *J. Bacteriol.* **2011**, *193*, 4043–4048, doi:10.1128/JB.05103-11.

70. Mutter, A.C.; Tyrishkin, A.M.; Campbell, I.J.; Poudel, S.; Bennett, G.N.; Silberg, J.J.; Nanda, V.; Falkowski, P.G. De novo design of symmetric ferredoxins that shuttle electrons in vivo. *Proc. Natl. Acad. Sci. U. S. A.* **2019**, *116*, 14557–14562, doi:10.1073/pnas.1905643116.

71. Lawson, R.J.; Von Wachenfeldt, C.; Haq, I.; Perkins, J.; Munro, A.W. Expression and characterization of the two flavodoxin proteins of *Bacillus subtilis*, YkuN and YkuP: biophysical properties and interactions with cytochrome P450 biol. *Biochemistry* **2004**, *43*, 12390–12409, doi:10.1021/bi049131t.

72. Shanklin, J.; Cahoon, E.B. Desaturation and related modifications of fatty acids. *Annu. Rev. Plant Biol.* **1998**, *49*, 611–641. doi:10.1146/annurev.arplant.49.1.611.

73. Trajtenberg, F.; Graña, M.; Ruétalo, N.; Botti, H.; Buschiazzo, A. Structural and enzymatic insights into the ATP binding and autophosphorylation mechanism of a sensor histidine kinase. *J. Biol. Chem.* **2010**, *285*, 24892–24903. doi:10.1074/jbc.M110.147843.

74. Di Capua, C.B.; Doprado, M.; Belardinelli, J.M.; Morbidoni, H.R. Complete auxotrophy for unsaturated fatty acids requires deletion of two sets of genes in *Mycobacterium smegmatis*. *Mol. Microbiol.* **2017**, *106*, 93–108, doi:doi.org/10.1111/mmi.1375.

75. Garba, L.; Mohamad Yussoff, M.A.; Abd Halim, K.B.; Ishak, S.N.H.; Mohamad Ali, M.S.; Oslan, S.N.; Raja Abd Rahman, R.N.Z. Homology modeling and docking studies of δ19-fatty acid desaturase from a cold-tolerant *Pseudomonas* sp. AMS8. *PeerJ* **2018**, *6*, 4347–4348. doi: 10.7717/peerj.4347.

76. Garba, L.; Ali, M.S.M.; Oslan, S.N.; Rahman, R.N.Z.R.A. Heterologous expression of PA8FAD9 and functional characterization of a Δ9-fatty acid desaturase from a cold-tolerant *Pseudomonas* sp. A8. *Mol. Biotechnol.* **2016**, *58*, 718–728. doi:10.1007/s12033-016-9971-9.

77. Soares-Castro, P.; Montenegro-Silva, P.; Heipieper, H.J.; Santos, P.M. Functional characterization of a 28-kilobase catabolic island from *Pseudomonas* sp. strain M1 involved in biotransformation of β-myrcene and related plant-derived volatiles. *Appl. Environ. Microbiol.* **2017**, *83*, e03112-16. doi:10.1128/AEM.03112-16.

78. Li, Y.; Xu, X.; Dietrich, M.; Urlacher, V.B.; Schmid, R.D.; Ouyang, P.; He, B. Identification and functional expression of a Δ9 fatty acid desaturase from the marine bacterium *Pseudoalteromonas* sp. MLY15. *J. Mol. Catal. B Enzym.* **2009**, *56*, 96–101, doi:10.1016/j.molcatb.2008.07.012.

79. Li, Y.; Dietrich, M.; Schmid, R.D.; He, B.; Ouyang, P.; Urlacher, V.B. Identification and functional expression of a Δ9-fatty acid desaturase from *Psychrobacter urativorans* in *Escherichia coli*. *Lipids* **2008**, *43*, 207–213, doi:10.1007/s11745-007-3150-5.

80. Araki, H.; Hagihara, H.; Takigawa, H.; Tsujino, Y.; Ozaki, K. Novel genes encoding hexadecanoic acid Δ6-desaturase activity in a *Rhodococcus* sp. *Curr. Microbiol.* **2016**, *73*, 646–651, doi:10.1007/s00284-016-1106-9.

81. Lemke, R.A.S.; Olson, S.M.; Morse, K.; Karlen, S.D.; Higbee, A.; Beebe, E.T.; Ralph, J.;
Coon, J.J.; Fox, B.G.; Donohue, T.J. A bacterial biosynthetic pathway for methylated furan fatty acids. *J. Biol. Chem.* **2020**, *295*, 9786-9801, doi:10.1074/jbc.RA120.013697.

82. Dyall, S.C.; Balas, L.; Bazan, N.G.; Brenna, J.T.; Chiang, N.; da Costa Souza, F.; Dalli, J.; Durand, T.; Galano, J.M.; Lein, P.J.; et al. Polyunsaturated fatty acids and fatty acid-derived lipid mediators: recent advances in the understanding of their biosynthesis, structures, and functions. *Prog. Lipid Res.* **2022**, *86*, 101165, doi:10.1016/j.plipres.2022.101165.

83. Woodward, X.; Kelly, C. V. Single-lipid dynamics in phase-separated supported lipid bilayers. *Chem. Phys. Lipids* **2020**, *233*, 104991-105012. doi:10.1016/j.chemphyslip.2020.104991.

84. Mansilla, M.C.; Cybulski, L.E.; Albanesi, D.; De Mendoza, D. Control of membrane lipid fluidity by molecular thermosensors. *J. Bacteriol.* **2004**, *186*, 6681-6688. doi:10.1128/JB.186.20.6681-6688.2004.

85. Parsons, J.B.; Rock, C.O. Bacterial lipids: metabolism and membrane homeostasis. *Prog. Lipid Res.* **2013**, *52*, 249-276. doi:10.1016/j.plipres.2013.02.002.

86. Aguilar, P.S.; De Mendoza, D. Control of fatty acid desaturation: a mechanism conserved from bacteria to humans. *Mol. Microbiol.* **2006**, *62*, 1507-1514. doi:10.1111/j.1365-2958.2006.05484.x.

87. Altabe, S.G.; Mansilla, M.C.; De Mendoza, D. Remodeling of membrane phospholipids by bacterial desaturases. *Stearoyl-CoA Desaturase Genes Lipid Metab.* **2013**, *209*, 209-231. doi:10.1007/978-1-4614-7969-7_15.

88. Diomandé, S.E.; Nguyen-the, C.; Abee, T.; Tempelaars, M.H.; Broussolle, V.; Brillard, J. Involvement of the CasK/R two-component system in optimal unsaturation of the *Bacillus cereus* fatty acids during low-temperature growth. *Int. J. Food Microbiol.* **2015**, *213*, 110-117. doi:10.1016/j.ijfoodmicro.2015.04.043.

89. Fernández, P.; Porrini, L.; Pereyra, J.I.; Albanesi, D.; Mansilla, M.C. Unveiling the coordinated action of DesK/DesR and YvfT/YvfU to control the expression of an ABC transporter in *Bacillus subtilis*. *Mol. Microbiol.* **2024**, 730-742, doi:10.1111/mmi.15320.

90. Miller, D.J.; Zhang, Y.M.; Subramanian, C.; Rock, C.O.; White, S.W. Structural basis for the transcriptional regulation of membrane lipid homeostasis. *Nat. Struct. Mol. Biol.* **2010**, *17*, 971-975, doi:10.1038/nsmb.1847.

91. Cooper, C.; Peterson, E.J.R.; Bailo, R.; Pan, M.; Singh, A.; Moynihan, P.; Nakaya, M.; Fujiwara, N.; Baliga, N.; Bhatt, A. MadR mediates acyl CoA-dependent regulation of mycolic acid desaturation in mycobacteria. *Proc. Natl. Acad. Sci. U. S. A.* **2022**, *119*, 1-10, doi:10.1073/pnas.2111059119.

92. Thomas, K.; Rutter, A.; Suller, M.; Harwood, J.; Lloyd, D. Oxygen induces fatty acid (n-6)-desaturation independently of temperature in *Acanthamoeba castellanii*. *FEBS Lett.* **1998**, *425*, 171-174, doi:10.1016/S0014-5793(98)00218-X.

93. Nakazawa, M.S.; Keith, B.; Simon, M.C. Oxygen availability and metabolic adaptations. *Nat. Rev. Cancer* **2016**, *16*, 663-673, doi:10.1038/nrc.2016.84.

94. Panconi, L.; Lorenz, C.D.; May, R.C.; Owen, D.M.; Makarova, M. Phospholipid tail asymmetry allows cellular adaptation to anoxic environments. *J. Biol. Chem.* **2023**, *299*, 105134, doi:10.1016/j.jbc.2023.105134.
95. Guerzoni, M.E.; Lanciotti, R.; Cocconcelli, P.S. Alteration in cellular fatty acid composition as a response to salt, acid, oxidative and thermal stresses in *Lactobacillus helveticus*. *Microbiology* **2001**, *147*, 2255–2264. doi:10.1099/00221287-147-8-2255.

96. Bayer, A.S.; Kupferwasser, L.I.; Brown, M.H.; Skurray, R.A.; Grkovic, S.; Jones, T.; Mukhopadhay, K.; Yeaman, M.R. Low-level resistance of *Staphylococcus aureus* to thrombin-induced platelet microbicidal protein 1 in vitro associated with *qacA* gene carriage is independent of multidrug efflux pump activity. *Antimicrob. Agents Chemother.* **2006**, *50*, 2448–2454, doi:10.1128/AAC.00028-06.

97. Vadyvaloo, V.; Hastings, J.W.; Van der Merwe, M.J.; Rautenbach, M. Membranes of class IIa bacteriocin-resistant *Listeria monocytogenes* cells contain increased levels of desaturated and short-acyl-chain phosphatidylglycerols. *Appl. Environ. Microbiol.* **2002**, *68*, 5223–5230, doi:10.1128/AEM.68.11.5223-5230.2002.

98. Gao, P.; Davies, J.; Kao, R.Y.T. Dehydrosqualene desaturase as a novel target for anti-virulence therapy against *Staphylococcus aureus*. *MBio* **2017**, *8*, e01224-17. doi: 10.1128/mBio.01224-17.

99. Dramé, I.; Rossez, Y.; Krzewinski, F.; Charbonnel, N.; Ollivier-Nakusi, L.; Briandet, R.; Dague, E.; Forestier, C.; Balestrino, D. FabR, a regulator of membrane lipid homeostasis, is involved in *Klebsiella pneumoniae* biofilm robustness. *MBio* **2024**, *0*, 1–20, doi:10.1128/mbio.01317-24.

100. Versini, R.; Sritharan, S.; Aykac Fas, B.; Tubiana, T.; Aimeur, S.Z.; Henri, J.; Erard, M.; Nüsse, O.; Andreani, J.; Baaden, M.; et al. A perspective on the prospective use of Al in protein structure prediction. *J. Chem. Inf. Model.* **2024**, *64*, 26–41, doi:10.1021/acs.jcim.3c01361.

101. Callaway, E. Chemistry Nobel goes to developers of AlphaFold Al that predicts protein structures. *Nature* **2024**, *634*, 525–526, doi:10.1038/d41586-024-03214-7.
Declaration of competing interest

I have no conflicts of interest to declare, and the same applies to all associated authors.
Highlights

* Structural and functional diversity of FADs in non-photosynthetic bacteria

* Role of fatty acid desaturases in bacterial pathogenesis

* Standardizing bacterial desaturase nomenclature

* FADs that modify FAs to produce specialized lipids beyond membrane flexibility

* Identification of desaturases in bacteria