# tRNA-dependent addition of amino acids to cell wall and membrane components

Guillaume Grob, Marine Hemmerle, Nathaniel Yakobov, Nassira Mahmoudi,
Frédéric Fischer, Bruno Senger, Hubert Becker

► To cite this version:

Guillaume Grob, Marine Hemmerle, Nathaniel Yakobov, Nassira Mahmoudi, Frédéric Fischer, et al. tRNA-dependent addition of amino acids to cell wall and membrane components. Biochimie, 2022, 203, pp.93-105. 10.1016/j.biochi.2022.09.017. hal-04235462

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Submitted on 2 Jul 2025

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1 tRNA-dependent addition of amino acids to cell wall and membrane components

2

3 Guillaume Grob<sup>a*</sup>, Marine Hemmerle<sup>b</sup>, Nathaniel Yakobov<sup>c</sup>, Nassira Mahmoudi<sup>a</sup>, Frederic

4 Fischer<sup>a</sup>, Bruno Senger<sup>a</sup>, Hubert Dominique Becker<sup>a*</sup>

5

6 <sup>a</sup> Génétique Moléculaire, Génomique, Microbiologie UMR7156, University of Strasbourg, 4

7 allée Konrad Roentgen, Strasbourg 6700, France.

8 <sup>b</sup> The Hospital for Sick Children, 555 University Avenue, Toronto, ON M5G 1X8, Canada

9 <sup>c</sup> University of Geneva, Department of Molecular Biology, Sc3 2045b, 30, quai Ernest

10 Ansermet, CH-1211, Geneva 4, Switzerland.

11

12 * Corresponding authors: Guillaume Grob: g.grob@unistra.fr and Prof. H. D. Becker: hbecker@unistra.fr

13

14

15 Abstract

16 The objective of the present review is to provide an insight into modifications of microbial
17 cell walls and membrane constituents by using the aminoacyl-tRNA as amino acid donor. In
18 bacteria, phospholipids are modified by Multiple peptide resistance Factor enzymes and
19 peptidoglycan precursors by so called fem ligases. Although these modifications were
20 thought to be restricted to procaryotes, we discovered enzymes that modify ergosterol (the
21 main component of fungal membrane) with glycine and aspartate.

22

23 The focus of this review is to present the molecular mechanisms underlying all these
24 processes together with the structure of the enzymes and their substrates. This article also
25 reviews how substrates are recognized and modified and how the products are subsequently
26 exported in various organisms. Finally, the physiological outcome and the discoveries of each
27 family of enzymes is also discussed.

28

29
## 1. Introduction

### 1.1. Cell membranes and their modification

The cell is the smallest unit of all living organisms, from unicellular bacteria to multicellular eukaryotes. It represents a protected microenvironment in which biochemical processes required to sustain life occur. All cells possess a plasma membrane (PM) composed of various types of lipids [1-4] as well as integral membrane and/or membrane-associated proteins. The PM acts as a selective barrier, controlling the flow of various chemicals that need to enter or exit the cell, as well as electrochemical gradients [5]. The PM is always composed of lipids, the nature of which changes from one organism to another according to the growth state or depending on stress responses. Besides the plasma membrane that separates the cytoplasm from the environment, eukaryotic cells also harbor several intracellular membrane-delimited compartments often organized in networks, such as the nucleus, the endoplasmic reticulum, the chloroplasts, the mitochondria, etc... Each of these compartments has its own lipid repertoire [6].

In bacteria, one single plasma membrane (PM) exists in monoderm, whereas diderm display a second (outer) membrane (OM) which further encloses the cell, creating the periplasmic compartment, delimited by both membranes [7]. Bacteria also always possess peptidoglycan (PGN) [8,9], which forms a complex molecular mesh that sits outside the plasma membrane or within the periplasm. It is not a lipidic component, but it highly contributes to maintaining cell shape and integrity. In the case of diderm, the 2 membranes have contrasting compositions and properties [10], with the OM mainly displaying a complex and highly decorated glycolipid, the lipopolysaccharide (LPS). It is composed of 2 glucosamines, on which 4 to 6 fatty acid chains are linked (i.e., the lipid A component) and one of the glucosamine molecule can be decorated with a chain of oligosaccharide (i.e. the core oligosaccharide), further linked to a O-antigen polysaccharide chain. Membranes and cell envelopes are at the interface with the environment and their composition and lipid modification status is highly regulated in response to environmental changes. For example, LPS can be highly modified in its lipid A part, as well as on its oligosaccharide conjugate in response to environmental stress [10]. Lipid components of membranes are also modified. Among these modifications, transfer RNA (tRNA)-dependent aminoacylation of glycerolipids (GLs), i.e., the addition of amino acids (aa) on hydroxyl groups of lipids, is one of the main subject of this review.

In both mono- and diderm, glycerolipids (GLs) can be conjugated to aa by enzymes that use aminoacyl-transfer RNAs (aa-tRNAs) as substrates, i.e., aminoacyl-tRNA transferases (ATTs). Among them, aminoacyl-glycerolipid synthases (aaGLSs) transfer the aa moiety of alanyl- and/or lysyl-tRNAs onto phosphatidylglycerol (PG), cardiolipin (CL) or diacylglycerol (DAG). The amino groups of the aa modifiers change the net charge of glycerolipids and enable bacteria to resist cationic anti-microbial peptides (CAMP), one of the first lines of defense of innate immunity in animals [4,11] (Fig. 1B). Likewise, other ATTs mainly found in monoderm, the Fem-ligases (FemA, FemB and FemX), were shown to transfer aa from aa- tRNAs onto the stem peptide of PGN subunits. This enables the synthesis of interpeptide cross-links that modify the meshing of PGN, but also of peptide tails that serve to anchor extracellular proteins to the PGN (Fig. 1A).

In fungi, apart from classical glycerol(phospho)lipids, the membrane is composed of eukaryotic specific lipids thereby increasing its complexity [12]. These lipids include sterols
such as ergosterol [13]. In fungi, the latter can be modified, on its 3β-OH group by acetylation [14], fatty acylation [15], glycosylation [16], and as discovered recently, by aminoacylation with aspartic acid (Asp) and/or glycine (Gly) [17,18]. Ergosterol aminoacylations have been shown to involve two ATTs that are homologs of the bacterial aaGLSs, that were named ergosteryl-amino acid synthases (ErxS) (Fig1. C).

In plants and animals, there are so far no known tRNA-dependent aa modifications of lipids achieving similar roles.

## 1.2. Amino acid conjugating enzymes, a structurally conserved family of protein

All aa-conjugating enzymes mentioned thus far share at least one common substrate: the aa-tRNA (the "first substrate") which is a source of activated aa for subsequent transfer onto a cell wall component or a lipid substrate, referred to as the "second" substrate. These ATTs share a common and characteristic structural domain which corresponds to a double General Control Non derepressible 5 related N-acetyltransferase (GNAT) domain in which the two GNAT submodules are separated by a small, positively charged, alpha helix ($\alpha^{(+)}$) (Fig. 2). We name this architecture the double GNAT$\alpha^{(+)}$ fold, referred to as the DUF2156/LPG_synthase_C (PF09924) domain in the Protein FAMily Database [11,17–21]. This fold belongs to the N-acetyltransferase-like clan (Pfam 0257) which, itself, includes all families that will be described in the present review: FemX, FemA, and FemB of the FemAB family (PF02388); aaGLSs and ErxS. Note that other types of ATTs, with different and unrelated folds exist, but are not included in this review; they have been extensively described elsewhere [22]. Although they belong to different families in PFAM, Fem-ligases (FemA, B and X) on the one hand and aaGLSs and ErxS on the other hand share the same double GNAT$\alpha^{(+)}$ fold, and highly similar structure, regarding the GNAT II plus $\alpha^{(+)}$ helix domain, but do not present significant sequence similarities, a common feature of GNAT-related proteins [19](Fig 2.).

This review will cover most aa-conjugating enzymes that modify membrane or membrane-related constituents in bacteria and fungi, with a particular focus on the molecular and biochemical players in place.

# 2. Amino acid phosphatidyl-glycerol synthases modify the charge of phospholipids

## 2.1. History and discovery

The existence of aminoacyl-glycerolipids (aaGL) in bacteria was discovered in the early 1960s with the identification of alanyl-phosphatidylglycerol (AlaPG) in *Clostridium perfringens* and lysyl-phosphatidylglycerol (LysPG) in *Staphylococcus aureus* [23,24]. AaGL are commonly found in Gram-positive (monoderm) bacteria including most Firmicutes and several *Actinobacteria* but are also present in Gram-negative (diderm) bacteria [11,25]. In the staphylococcal membrane, LysPG represents 20 to 40 % of all phospholipids. AlaPG, however, is found less often and is only a minor component of bacterial membrane (up to 5 % of total phospholipids) in many Gram-positive and in some Gram-negative bacteria. In addition, PG can also rarely be aminoacylated with arginine (ArgPG). For example, only traces of ArgPG are found in enterococci [4]. All these aminoacylated lipids are synthesized by tRNA-dependent aaGLSs [11]. In addition to PG, cardiolipin (CL) can also be tRNA-dependently aminoacylated with lysine or alanine by aaGLSs. Lysyl-cardiolipin (LysCL) is found in *Listeria* and *Enterococcus* species and represents up to 30 % of total phospholipids in *Listeria* membranes. In *Anoxybacillus rupiensis* and *Vagococcus fluvialis*, both LysCL and
119 AlaCL were detected [4]. The aminoacyl-phosphatidylglycerol synthetase (aaPGS)
120 responsible for LysPG production was discovered almost 40 years after LysPG in one S.
121 *aureus* transposon mutant that displayed increased susceptibility to a wide range of CAMP
122 produced by eukaryotic hosts (host defense peptides) or by prokaryotic competitors
123 (bacteriocins) [26]. These enzymes were thus named Multiple peptide resistance Factors
124 (MprF). In 2015, one MprF homolog found in Corynebacterium glutamicum (Actinobacteria)
125 was shown to use Ala-tRNA and DAG to produce AlaDAG. In addition, an esterase of the α/β-
126 hydrolase family (PesT), found in the same bacterium, seemingly uses AlaDAG to
127 transesterify Ala onto PG, yielding AlaPG, which the AlaDAG synthase cannot produce by
128 itself [25].

129 To date, hundreds of MprF homologs, distributed into 7 architectural types, have been
130 identified and are detected across 93 bacterial genera (mostly Firmicutes, Actinobacteria,
131 and Proteobacteria) and in several archaeal species [25]. These homologous enzymes
132 present variable aa-tRNA specificity for Lys-, Ala- and/or Arg-tRNA but also use different GL
133 such as PG, CL, or DAG, depending on the organism in which it is present.

## 2.2. Role and mechanism

135 The precursor molecules PG and CL possess negatively charged phosphate head groups (PG
136 (-1) and CL (-2)). Aminoacylation of PG and CL with L-lysine (two α-amino groups) or L-
137 alanine (one α-amino group) thus creates cationic or neutral zwitterionic molecules (LysPG
138 (+1), AlaPG (0), LysCL (0)). These modifications change the surface properties of membranes
139 and reduce its affinity for and permeability to CAMPs. Most prominent human pathogens
140 achieve CAMP resistance using aaPGS-dependent lipid lysylation and/or alanylation.
141 Contrary to other GNAT-type acyl- or acetyl-transferases that use acyl-/acetyl-Coenzyme A
142 as acyl donors [19], double GNAT α(+) enzymes, such as aaGLSs, use aa-tRNAs as acyl (here
143 aminoacyl) donors. The aa-tRNA substrates are produced by dedicated aminoacyl-tRNA
144 synthetases (lysyl-tRNA synthetase for Lys-tRNA, alanyl-tRNA synthetase for Ala-tRNA) and
145 deviated from protein synthesis [27,28]. In Lys- and Ala-tRNA, the α-carboxyl groups of the
146 aa moieties are engaged in an ester bond with the 3' hydroxyl group of the terminal
147 adenosine of their corresponding tRNAs [29]. In aaGL, the same α-carboxyl is used to create
148 an ester bond with a hydroxyl group of the acceptor lipid. The aaPGS are thus aa-tRNA-
149 dependent transesterases. In aaPG, for example, the α-carboxyl group of the aa modifier is
150 esterified with the last hydroxyl- group of the terminal glycerol moiety of PG [28,30]. In
151 *Pseudomonas aeruginosa* and *Enterococcus faecium*, specific extracytoplasmic esterases that
152 hydrolyze the aa moiety from PG have been identified and are involved in maintaining a
153 specific level of aaPG which is important for antimicrobial resistance and cell growth [31,32].

## 2.3. aaPGS modular organization

155 AaPGS are integral membrane proteins, usually constituted of two domains with distinct
156 enzymatic functions: i) the C-terminal DUF2156/LPG_synthase domain, which transfers the
157 aa from aa-tRNA to the lipid substrate and ii) the N-terminal integral membrane anchoring
158 domain with an aaGL flippase activity. There is limited sequence conservation between N-
159 terminal domain of aaPGS, contrasting with the well-conserved C-terminal synthase domain.
160 The N-terminal hydrophobic domain contains 2 to 14 transmembrane helices (TMHs) [25]
161 responsible for anchoring the aaPGS proteins to the PM. The largest N-ter domains (12-14
162 TMH) display a flippase activity responsible for translocating aaPG from the inner leaflet of
163 the PM (where it is produced) towards the outer leaflet (where it is exposed) [33]. Smaller N-
ter domains (< 12 TMH) usually lack this activity [34]. For instance, *C. perfringens* possesses two aaPGSs, a LysPGS with a 12-14 TMH N-ter domain that functions as a flippase and a 6 TMH-containing AlaPGS that lacks the flippase activity. The former, which can use both Lys- and AlaPG, complements the lack of flippase activity of the second [35]. The LysPGS of *M. tuberculosis* and related actinomycetes present reduced size N-terminal domains, with no demonstrated flippase activity, but additional, separated proteins with similarities to the aaPGS flippase domain are present. Of note, many uncharacterized aaPGS homologs lack the N-terminal extension [25] and possess only the synthase domain. In this case, separate, *trans*-acting, LysPG and/or AlaPG flippases, either from other aaPGSs or from free-standing flippases, may complement the lacking flippase activity. *Myxococcus xanthus* and related species illustrate this case, in which the flippase and synthase domains of aaPGS are separated (encoded by two separate genes) and cooperate. This suggests that aaPGSs might have evolved from fusion of two separate proteins [4,36]. Other modular aaPGS forms exist, such as in *M. tuberculosis*, where the LysPGS is fused to a lysyl-tRNA synthetase paralog that synthesizes and directly delivers the Lys-tRNA<sup>Lys</sup> substrates for LysPG synthesis [37].

In *S. aureus*, the LysPGS is an integral membrane protein that presents 14 TMH connected by 2- to 56-aa-long loops (predicted lengths) [3], followed by the synthase domain. Even if the synthase and flippase domains have distinct roles, they display multiple inter-domain interactions. Two TMHs in the synthase domain are for instance required for optimal LysPG flippase activity [3]. Topology of the flippase domain has also been studied. While loops 1, 9, and 13 are located in the extracytoplasmic space, localization of loop 7 (between TMS 7 and 8) is ambiguous [38]. Some algorithms predict that loop 7 is extracytoplasmic, whereas experimental topology investigations revealed a preferential intra-cytoplasmic localization [3]. Slavetinsky et al. developed monoclonal antibodies (mAbs) targeting extracellular loops of the N-ter flippase module to inhibit the LysPG flippase domain (of which topology is described in Fig. 3). They discovered that mAbs targeting loop 7 do not impact the LysPG synthase activity but rather inhibits the flippase function, thereby making sensitizing *S. aureus* to antimicrobial host peptides, daptomycin, and CAMP-produced by human polymorphonuclear leukocytes. Loop 7, at the junction between the LysPG synthase and flippase domain, has therefore been postulated to be alternatively exposed at both sides of the PM, playing a role in LysPG translocation [38]. In addition, the two conserved residues D254 (TMS 7) and R279 (TMS 8), which flank loop 7, could be part of a binding site for LysPG translocation [3,38]. However, all these results needed to be confirmed by solving the 3D structure of the protein, to decipher the molecular mechanism of LysPG translocation.

This structure was obtained in 2021 by Song and colleagues [39]. When expressed in *E. coli*, LysPGS from *Rhizobium tropici* homodimerizes within the PM, mainly through interactions between membrane domains [39]. Four lipid molecules located at the dimer interface contribute to dimerization by simultaneously forming polar and hydrophobic interactions with two adjacent monomers [39]. The N-terminal flippase domain has 12 long TMHs (TM1-2, TM4-6, TM8-14) and two pairs of short α-helices (TM3a – TM3b and TM7a-TM7b) spanning halfway in the membrane [39]. The cytoplasmic side of the flippase domain appears positively charged, whereas the periplasmic one is mostly negatively charged, as expected from the “positive inside rule” in membrane protein topogenesis [40].

The C-terminal synthase domain contains both the binding site for aa-tRNA and that for PG and displays a structure similar to that of AlaPGS and LysPGS homologs from *P. aeruginosa* and *Bacillus licheniformis*, respectively, e.g. tandem repeats of the GNAT folds (GNAT folds 1
and 2) [3,39,41]. The GNAT 1 domain of the synthase module is attached to TM14 of the
210
211 flippase domain by a flexible loop and folds back onto the membrane domain through non-
212 covalent interactions. The latter interactions contribute to forming the so-called Interfaces 1
213 and 2 between the synthase and the flippase domains. Residue Arg456, involved in the
214 formation of Interface 1, is highly conserved in aaPGS homologs [39]. The synthase domain
215 of aaPGS from *P. aeruginosa*, *R. tropicii* and *B. licheniformis* are all structurally similar to the
216 FemX alanyl-transferase involved in peptidoglycan interpeptide bridge formation [41]. The
217 structural superposition of LysPGS from *B. licheniformis* or AlaPGS from *P. aeruginosa* with
218 FemX peptidyl-RNA complex shows that the terminal tRNA nucleotides are accommodated
219 in a defined cavity located in the GNAT 2 domain of both aaPGS. This cavity is delineated
220 mainly by secondary structure elements comprising a series of fully conserved aa residues
221 [41].

222 **2.4. Aa-tRNA and phospholipid substrate recognition and enzymatic mechanism**

While some aaPGSs are highly specific, some others possess relaxed substrate specificity. For
223 example, *S. aureus* and *P. aeruginosa* aaPGS only synthesize LysPG and AlaPG respectively,
224 while *Enterococcus faecalis* MprF2, using 3 different aa-tRNAs, synthesizes Ala- and LysPG
225 along with small amounts of ArgPG [42–44]. Moreover, the acceptor substrate specificity can
226 also be relaxed as exemplified by *Listeria monocytogenes* aaPGS, which produces both LysPG
227 and LysCL [45]. Concerning the flippase domain, *S. aureus* and *C. perfringens* LysPGSs can
228 both translocate AlaPG and LysPG, indicating that this domain has relaxed specificity for the
229 aa modifier [35].
230

Structural analysis of the aminoacyl-binding cavity of *P. aeruginosa* AlaPGS and *B. licheniformis* LysPGS indicates that the specific recognition or steric exclusion of the lysyl-versus alanyl-substrate moiety is not mediated by any conserved or supplementary aa residues [41]. However, structural analyses and biochemical experiments demonstrated that specific recognition of the aa donor relies on the tRNA acceptor stem, the aminoacyl-moiety, and that lipid recognition, in another pocket adjacent to that for aa-tRNA, depends on its polar head group of PG [41]. In addition, both proteins contain a 23-Å tunnel with a diameter of 4-8 Å, that is directly connected to the aa-tRNA-binding cavity. The tunnel is delineated by residues of GNAT domains 1 and 2 and could facilitate binding of the polar aa-tRNA opposite to the hydrophobic lipid and thus be the place of aaPGS catalysis at the water-lipid interface [41].
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241

242 **3. FemAB enzymes are peptidoglycan remodelers**

3.1. Peptidoglycan structure and function

Bacterial peptidoglycan (PGN) is composed of repeated disaccharides consisting of *N*-acetyl
244 glucosamine (GlcNAc) linked to *N*-acetyl muramic acid (MurNAc) [46]. The pathway of PGN
245 synthesis (and therefore its structure) can slightly vary across species. Synthesis is sequential
246 and starts in the cytoplasm with (Fig. 3) an activated sugar precursor, i.e., UDP-GlcNAc
247 (Uridine-5'-diphospho-1'-glucosamine) that is converted to UDP-MurNAc by the MurA and
248 MurB enzymes through attachment of pyruvate (from phosphoenolpyruvate) and its
249 reduction to a 3'-*O*-lactoyl group. Then, UDP-MurNAc serves for the synthesis, by 4 enzymes
250 (MurC, D, E and F), of a 5 aa stem peptide. In Gram-negative bacteria, the resulting
251 pentapeptide has a conserved structure and is composed of L-*Ala*₁-*γ*-D-*Glu*₂-*mDAP*₃-D-*Ala*₄-
252 D-*Ala*₅ (L- and D- denoting the aa isomer and mDAP, meso-diaminopimelic acid and numbers
253 the position of the aa relative to the lactoyl group). The resulting pentapeptide-
254
monosaccharide then becomes conjugated by the MraY transferase to an undecaprenyl-
phosphate membrane anchor, resulting in the undecaprenyl-diphospho-MurNAc-
pentapeptide (lipid I) product facing the cytoplasm. Another UDP-GlcNAc unit is used by the
MurG protein and GlcNAc conjugated to the MurNAc moiety of lipid I, resulting in the
formation of lipid II. Lipid II is then translocated to the extracytoplasmic space by a flippase,
where can be used as the building block for PGN synthesis. After translocation, MurNAc and
GlcNAc moieties from adjacent Lipid II molecules become linked through trans-glycosylation
reactions catalyzed by dedicated transglycosylases. Transpeptidases then catalyze the
formation of a peptide bond between the α-carboxyl group of the D-Ala₄ residue (linked to
D-Ala₅) of one stem peptide and the amino group of the mDAP₃'s side chain of the adjacent
stem peptide, which eliminates the D-Ala₅ residue. The stem-peptides of two adjacent units
are therefore directly cross-linked [47–49]. Consequently, D-Ala₅ is released from the mature
stem-peptide, and the level of stem-peptide cross-linking determines the rigidity of the PGN
[46]. In gram-negative bacteria, this rate is low (25-50 %) in comparison to Gram-positive
PGN (70-90 %). Even though the above-mentioned stem-peptide composition is found in
most Gram-negative bacteria, variations were reported at various positions, especially mDAP
that is replaced by L-Lys in Gram-positive bacteria [46]. The PGN building block (Lipid II) of
Gram-positive bacteria is, for example, composed of L-Ala₁-γ-D-Gln₂-L-Lys₃-D-Ala₄-D-Ala₅. As
mentioned above, structure variations of the stem-peptide have been described, such as the
presence of D-Lac in position 5 (D-Lac₅), replacing D-Ala, which increases resistance of
Enterococci to vancomycin. In contrast to Gram-negative bacteria, where residues from two
stem-peptides are directly cross-linked, Gram-positive stem-peptides are generally
interspaced by an interpeptide-bridge whose size and composition is variable across species
[50].

279 **3.2. Cross-linking of peptidoglycan by Fem-ligases in S. aureus**

280 Early discovery of PGN pentaglycine cross-linking

281 Investigation of PGN synthesis started as early as in the 1960s [51] and it is in 1961 that the
282 then "atypical" pentaglycine bridge happened to be discovered in *S. aureus* [52]. In 1965, a
283 study suggested that enzymatic formation of pentaglycine (interpeptide bridges in *S. aureus*
284 required the presence of so-called soluble RNA (sRNA, which is now known to include tRNA
285 amongst other soluble RNAs) [53]. Studies on the incorporation in PGN of glycine in
286 *Micrococcus lysodeikticus* (1967) [54], of alanine in *Arthrobacter crystallopoides* (1968) [55]
287 and threonine in *Micrococcus roseus* (1968) [56] evidenced the same RNA-dependency. It is
288 in 1968 that the same group unambiguously demonstrated the use of Gly-tRNA by dedicated
289 Gly-tRNA transferases to add the five Gly residues on lipid II in *S. aureus* [57,58]. The use of
290 seryl-tRNA to add Ser residues in the PGN of *S. aureus* was also described in the same
291 studies, which generalized the idea that these modifications of the PGN were tRNA-
292 dependent.

293 Synthesis of the pentaglycine bridge was shown to rely on the expression of 3 genes, 2 of
294 which had been functionally linked to methicillin resistance [59]. These two genes were
295 ascribed the names *femA* and *femB* and further studies soon showed that the two genes
296 were expressed as a single transcriptional unit [60]. The *ΔfemAB* deletion leads to a drastic
297 slow growth phenotype and hypersensitivity to methicillin [61]. In 1997, a study showed that
298 in this mutant, the interpeptide bridge was reduced down to one Gly unit (monoglycine
299 bridge), indicating that 1) they were likely Gly-tRNA transferases but that 2) another enzyme
300 was responsible for the addition of the first Gly. Complementation of the *ΔfemAB* mutant with
301 only the *femA* gene led to the synthesis of a truncated triglycine bridge, showing that the
302 FemA Gly-tRNA transferase could elongate the preexisting monoglycine chain of 2 more Gly
303 residues. FemB alone could not elongate the monopeptide bridge. Complementation with
304 both *femA* and *femB* restored synthesis of the full-length pentapeptide bridge [61], revealing
305 that FemB acted after FemA by completing the synthesis through addition of 2 other
306 residues on the triglycine primer. However, the first enzyme required for the addition of the
307 first Gly residue remained unknown.

308 In 1999, Rohrer and col. analyzed the PGN of a modified *S. aureus* strain in which the
309 expression of the essential gene *fmhB* was put under the control of an inducible xylose
310 promoter[62]. In absence of xylose (depletion of the FmhB protein), cross-links in the PGN
311 were dramatically reduced, while lipid II was found to lack any Gly residue, thereby proving
312 that this protein was likely responsible for the addition the first Gly on lipid II. The absence
313 of this first glycine upon *fmhB* downregulation leads to tremendous growth defects and
314 complete instability of the PGN. Actually, cross-linking with pentapeptide bridges is essential
315 for cell integrity in *S. aureus*, while it is not the case in other Gram-positive bacteria [63]. The
316 whole pathway was then fully reconstituted in vitro in 2004 [64](Fig. 3).

317 **3.3. Fem ligases are aa-tRNA transferases**

318 Following the seminal studies that identified Gly-tRNA transferases in the 60s, biochemical
319 characterization of the whole pathway using purified FemA, FemB and FemX and substrates
320 (lipid II and lipid II intermediates), was reached in 2004. It confirmed that FemX acts first,
321 using one Gly-tRNA<sup>Gly</sup> to transfer Gly onto lipid II —more specifically on the ε-amino group of
322 the Lys<sub>3</sub> residue of the stem peptide. Then, this Lipid II-Gly intermediate serves as a
323 substrate for FemA that seems to sequentially use 2 Gly-tRNA<sup>Gly</sup> to elongate the Gly modifier
324 by 2 more Gly residues. The Lipid II-Gly<sub>3</sub> product is the substrate of FemB that terminates the
325 synthesis by adding two more Gly residues in a Gly-tRNA<sup>Gly</sup>-dependent manner [64]. In 1968,
326 seminal studies showed that in *S. aureus*, three tRNA<sup>Gly</sup> isoacceptors participate in the
327 interpeptide bridge synthesis, two of which being also substrates for ribosomal protein
328 synthesis, whereas one was seemingly not usable in translation [58]. First reinforced by
329 bioinformatic studies and Elongation factor-Tu (EF-Tu) binding experiments [65], this fact
330 was later confirmed with experiments showing that a pool of non-proteinogenic Gly-tRNA<sup>Gly</sup>
331 actually exists, which are specifically dedicated to Fem transferases; this pool is actually
332 more abundant (63 %) than the proteinogenic Gly-tRNA<sup>Gly</sup> [66]. Non proteinogenic Gly-
333 tRNA<sup>Gly</sup> present negligible affinity of EF-Tu [65], when compared to proteinogenic Gly-
334 tRNA<sup>Gly</sup>. Such a partition in binding features may represent an evolutionary advantage, since
335 a Gly-tRNA pool dedicated to Fem ligases prevents depletion of that necessary to sustain
336 protein synthesis and ensures that the essential pentaglycine biosynthesis pathway remains
337 active even under higher Gly-tRNA demands in protein synthesis.

338 Note that in *S. aureus*, other Fem-like enzymes exist [67, 68] that can replace or pair with
339 FemA and/or FemB to catalyze the transfer of Ser residues from Ser-tRNA<sup>Ser</sup> onto the lipid II-
340 Gly precursor, resulting in interpeptide bridges with Ser-Gly dipeptides following the first Gly
341 residue. The FmhA Ser-tRNA transferase can, for example, add Ser residues at positions 3
342 and 5, whereas the FmhC Ser-tRNA transferase only elongates tetra-aa chains by one Ser
343 residue. These variations increase resistance of *S. aureus* to lysostaphin, a secreted
344 bacteriocin enzyme, secreted by competing bacteria, which cleaves the pentaglycine cross-
345 bridge [69].
346 Enzymatic characterization and molecular modeling of FemX suggest that the enzyme, at 347 least in the case of FemX from *Streptococcus pneumonia* (named murM, see below) acquires 348 its full activity when bound onto lipid II in membrane regions enriched in CL that help it 349 correctly orientate onto its membrane-anchored substrate with a cooperative effect [70]. 350 Authors suggest that, on grounds of this membrane interaction, FemX/MurM likely acquires 351 its Ala-tRNAAla substrate in the cytoplasm before interacting in a second step, once loaded 352 with this substrate, with the stem peptide of lipid II, aided by the CL-rich environment. 353 Interestingly, in *Weissella viridescens*, FemX is an Ala-tRNA transferase that binds to the 354 UDP-MurNAc-pentapeptide cytoplasmic precursor rather than to lipid II [71].

355 **3.4. Structural studies of Fem transferases from S. aureus**

356 Structure of *S. aureus* FemA was obtained in 2002 [72] and displays, like aaPGS synthase 357 domains, a double GNAT fold [72]. FemA harbors, in GNAT subdomain 2, an additional 358 helical arm, absent in aaPGSs, similar to the N-terminal domain of bacterial seryl-tRNA 359 synthetases (involved in the variable loop binding of tRNA<sup>Ser</sup>) [73], that likely serves in Gly- 360 tRNA<sup>Gly</sup> binding. No FemB structure has been reported. In the case of FemX, early studies 361 conducted in 2003-2004 reported the structure of FemX from *W. viridescens* [74,75], an Ala- 362 tRNA transferase that adds Ala onto the soluble UDP-MurNAc-pentapeptide rather than on 363 lipid II (see below). This enzyme shares the classical double GNAT α<sup>(+)</sup> fold that is found in 364 aaPGSs, with no helical arm.

365 Last year, the structure of FemX from *S. aureus* was resolved and used as a template for the 366 modeling of the structure of the homologous MurM from *S. pneumoniae* [70]. Just like 367 FemA, *S. aureus* FemX displays a double GNAT α<sup>(+)</sup> fold with a long helical arm emerging 368 from within subdomain GNAT 2. This arm is absent in *W. viridescens* FemX [75] highlighting 369 structural variations between these transferases that may account for differences observed 370 in their substrates specificity.

371 **3.5. Regulation of peptidoglycan remodeling**

372 Modifications of the interpeptide bridge levels seem to be tightly regulated since it has been 373 reported that the Stk kinase [76] of *S. aureus* can phosphorylate FemX *in vitro* and may 374 represent a way to regulate the level of pentaglycine cross-linking in bacteria.

375 **3.6. MurM/N enzymes from S. pneumoniae are Fem ligases homologs**

376 In the late 1980s, it was shown in *Streptococcus pneumoniae* that abnormal cross-linking of 377 the peptide moieties in PGN biosynthesis leads to increased resistance level to the penicillin 378 family of antibiotics. These different interpeptide bridges contain either L-Ala-L-Ala or L-Ala- 379 L-Ser and change the PGN structure [47,77]. The genetic determinants for this resistance to 380 penicillin were in the 2000s in studies showing that an operon coding for two genes that 381 were named *murM* and *murN* was responsible for the addition of the first and the second aa 382 of the interpeptide bridge, respectively. Like in *S. aureus*, aa are added by the MurM and 383 MurN ATTs to lipid II before its export on the outer layer of the plasma membrane. Despite 384 being functional homologs, MurM and MurN are different from *S. aureus* Fem transferases 385 in that they each catalyze the transfer of only one aa instead of 2. Molecular determinants of 386 these differences and of substrate specificity are still unknown. Recently, a mechanism has 387 been proposed for MurM enzymatic activity (see 3.2.2).
388

**3.7. *Weissella viridescens* displays an alternative modification pathway**

389
390
391
392
393
394
395
396
397
398
399

Finally, another system of Fem transferases has been described in *W. viridescens* (formerly
classified as Lactobacillus species). This organism's genome encodes one homolog of *S.
aureus* FemX, the activity of which had been known since the 1970s. As noted above, *W.
viridescens* FemX transfers Ala from Ala-tRNAAla onto the stem peptide. Properties of *W.
viridescens* FemX change in comparison with that of *S. aureus* or *S. pneumoniae*: i) the
enzyme is found in the soluble fraction (unlike the "particulate fraction" in the case of *S.
aureus*) and ii) it conjugates Ala before MurNAc is transferred onto the undecaprenyl-
phosphate anchor, i.e. it uses UDP-MurNAc-pentapeptide as an L-Ala acceptor substrate
[78](Fig. 3). FemX was crystalized with both its UDP-MurNAc substrate [71,79] and a
peptidyl-RNA conjugate that mimics the alanylated CCA-end of tRNAAla and UDP-MurNAc
[80].

400

**3.8. Structural information about Fem ligases**

401
402
403
404
405
406
407

Fem transferases are of interest since they can serve as targets of choice for antimicrobial
therapy, especially in organisms, like *S. aureus*, in which they are essential (FemX).
Structures of FemA and FemX from *S. aureus* were determined while those of FemB, MurM,
and MurN have yet to be solved. However, the growing amount of protein structures
available as well as the release of models using structure prediction tools like AlphaFold2
enables structural comparisons with a good degree of confidence. Such approaches could
also be used as a basis for the design of antimicrobial drugs.

408

**4. Ergosteryl-amino acid synthases**
409 **4.1. Homologs of bacterial aaPGS**

410 Although aminoacylated phospholipids had been discovered as early as the 60s in bacteria,
411 no equivalent molecules seemed to exist in eukaryotes. Moreover, whereas aaPGSs are
412 widely distributed in bacteria and present in some archaea [24], sequence searches failed to
413 detect homologs in eukaryotes, suggesting that no tRNA-dependent lipid aminoacylation
414 pathway would exist in this domain of life. This fact changed, starting in 2020, with the
415 discovery that Dikarya — a sub-kingdom of fungi [81], also referred to as “higher fungi”—
416 were equipped with one, sometimes two, distant homologs of bacterial aaPGSs. The first to
417 be discovered corresponded to a DUF2156/LPG_synthase domain, homologous to the
418 synthase domain of aaPGSs fused at the C-terminus of a eukaryotic-type aspartyl-tRNA
419 synthetase (AspRS), i.e., the enzyme that produces Asp-tRNA<sup>Asp</sup> from free L-Asp, ATP and
420 tRNA<sup>Asp</sup>. This enzyme, named ErdS, is present in most Dikarya including prominent human
421 pathogens such as *Aspergillus fumigatus* or *Cryptococcus neoformans* but also in model
422 organisms and species of biotechnological interest like *Neurospora crassa* or *Aspergillus
423 oryzae* [17]. The second type of fungal aaPGS-like enzyme, named ErgS, was described in
424 2022 [18]. It corresponds to a freestanding DUF2156/LGP_synthase domain, also distantly
425 related to bacterial aaPGSs. Distribution of ErgS is narrower than for ErdS, and restricted to
426 the division Ascomycota within Dikarya, while ErdS is present in both Ascomycota and
427 Basidiomycota. Interestingly, *Saccharomycetes*, that comprises the baker's yeast
428 *Saccharomyces cerevisiae*, possess none of these enzymes. Neither ErdS, nor ErgS present
429 membrane domains.
430

430 **4.2. Aminoacylations of ergosterol**

431 ErdS is an AspRS-DUF2156 fusion. Biochemical characterization of the enzyme coupled with
432 *in vivo* investigations on *A. fumigatus*, *A. oryzae* and *N. crassa* mutants deleted from the
433 corresponding *erdS* gene demonstrated that it is a bifunctional enzyme. The AspRS domain
434 uses L-Asp, ATP and tRNA<sup>Asp</sup> to produce Asp-tRNA<sup>Asp</sup>, like a canonical AspRS, but this aa-tRNA
435 is then used as a substrate by the appended DUF2156/LGP_synthase module that transfers
436 Asp onto a lipid acceptor. Contrary to bacterial aaPGS that use GLs, ErdS was found to use
437 ergosterol, the main sterol of fungal membrane, functionally equivalent to the cholesterol
438 found in animal membranes [13] L-Asp is transferred on the β-hydroxyl group at position 3
439 of the A ring of the sterol, yielding ergosteryl-3β-*O*-L-aspartate (Erg-Asp), reason for which
440 the enzyme was named ergosteryl-3β-*O*-L-aspartate synthase (Fig. 5)(KEGG orthology:
441 K24278), shortened ErdS, with ‘Er’ for ergosterol, ‘d’ the one letter code for L-Asp and ‘S’ for
442 synthase [17]. Although the whole enzyme was termed ErdS, it is the C-terminal DUF2156
443 module that carries the Asp-tRNA:ergosterol transferase activity to which the name ErdS
444 refers to. Structure predictions and sequence analyses show that the transferase domain
445 presents the same double GNAT α<sup>(+)</sup> topology as the Fem transferases and the synthase
446 domain of aaPGSs. It is therefore believed to function in a similar fashion, a hypothesis
447 which biochemical experiments tend to advocate. It is of note that, despite a large
448 distribution of ErdS across Dikarya, Erg-Asp had never been detected before, although its
449 synthesis has now been confirmed in several fungi from different lineages [17]. In *A. oryzae*,
450 Erg-Asp represents ~7 % of total ergosterol content, which is not negligible. Of note is that
451 ErdS, despite its AspRS domain, likely does not participate in protein synthesis, since all fungi
452 that possess this enzyme always present an independent AspRS that can ensure this
453 essential function. Only tRNA<sup>Asp</sup> must be partitioned between the two pathways, because
454
455

the AspRS domain of ErdS ensures Asp-tRNA<sup>Asp</sup> synthesis to fuel the transferase/ErdS domain.

456
457
458
459
460
461
462
463
464
465
466
467
468
469
470
471
472
473

The second enzyme was found to be an ergosteryl-3β-*O*-glycine synthase, shortened ErgS (using the same nomenclature as for ErdS, with ‘Er’ for ergosterol, ‘g’ the one letter code for glycine, and ‘S’ for synthase) [18]. This enzyme is a freestanding DUF2156/LGP_synthase domain, which sequence analyses and structural predictions showed to harbor the classical double GNAT α<sup>(+)</sup> topology. Experiments that characterized its enzymatic activity *in vitro* also showed that ErgS from *A. fumigatus* and *Yarrowia lipolytica* lose activity when the α<sup>(+)</sup> helix that connects subdomains GNAT 1 and 2 is mutated as to become negatively charged or neutral. This advocates for a role of this helix in tRNA binding, as suggested for aaPGSs and Fem transferases [20], but this awaits further investigations. ErgS is not fused to a glycyl-tRNA synthetase (GlyRS) and must deviate Gly-tRNA<sup>Gly</sup> from the cytoplasmic pool before transferring its Gly moiety onto the 3β-hydroxyl of ergosterol, yielding ergosteryl-3β-*O*-glycine (Erg-Gly) (Fig. 6). How ErgS competes with elongation factor 1A (eEF1A) to gain access to Gly-tRNA<sup>Gly</sup> has not been investigated, but the enzyme can use its substrate when heterologously expressed in the yeast *S. cerevisiae*, suggesting that it does not necessarily require specific accessory proteins, competition likely occurring at the binding level. Erg-Gly could be detected in several Ascomycota using mass spectrometry, including in *A. fumigatus*, *A. oryzae* and *Y. lipolytica*, expanding the known repertoire of this newly discovered class of ergosteryl-3β-*O*-(*L*)-amino acids (Erg-aa).

474
475
476
477
478
479
480

Of note, several ascomycota possess both ErdS and ErgS, consequently synthesizing Erg-Asp and Erg-Gly. This resembles the case of some bacteria that possess multiple aaPGS paralogs which together produce multiple aminoacylated GLs. These two enzymes represent a novel fungal-specific class of aa-tRNA:lipid transferases, namely ergosteryl-3β-*O*-amino acid synthases (ErxS, with ‘x’ denoting any aa). More investigations are needed to understand whether more of these exist in fungi, that would remain undetected due to high sequence divergence from bacterial aaPGSs.

481
482
483
484
485
486

Finally, further investigations on the ErdS system showed that a dedicated Erg-Asp α/β-hydrolase (named ErdH) exists in fungi [17]. Its gene lies in a two-gene locus together with the *erdS* gene, that seems conserved across species. This suggests that aminoacylated sterols must be tightly regulated *in vivo*. This fact parallels the existence of extracytoplasmic hydrolases that remove the aa modifiers of GLs in bacteria (see 2.2). Up to date, no Erg-Gly specific hydrolase has been discovered.

487

**4.3. Role of sterols and ergosterol in eukaryotes and fungi**

488
489
490
491
492
493
494
495
496
497

The function(s) of Erg-aa is currently unknown. However, sterols are important components of eukaryotic cell membranes that are also known to act as signaling molecules. In fungi, ergosterol is essential for cell viability because it regulates membrane fluidity, stability and permeability, including to osmolites [82–86]. Consequently, ergosterol biosynthesis defects lead to pleiotropic phenotypes including osmotic disturbance, disruption of cell growth, sensitivity to antimicrobial, and decreased proliferation [87–90]. Fungal cells lacking normal ergosterol biosynthesis also show defects in endocytosis [91,92], vacuolar fusion [93], and mitochondrial respiration [94]. Because of its importance, ergosterol represents a good therapeutic target to fight pathogenic fungi. Ergosterol biosynthesis enzymes and/or ergosterol itself are targeted, by most of common antifungal drugs when infecting humans.
498 Sterols, including ergosterol, can be modified *in vivo*, which changes their functions, 499 localization and/or influence molecular pathways. For example, the 3β-hydroxyl of either 500 ergosterol in fungi or cholesterol in mammals can be fatty acylated, as to serve as a lipid and 501 sterol store, which translocates within lipid droplets, and that can be mobilized back through 502 deacylation [15]. In *S. cerevisiae*, sterols, including ergosterol, can additionally be acetylated 503 on the 3β-hydroxyl by the Atf2 acetyltransferase using acetyl-coenzyme A [14]. Only 504 ergosterol, not other sterols, can be deacetylated by the dedicated Say1 deacetylase, which 505 participates in membrane sterols homeostasis and antifungal resistance. In many organisms, 506 sterols can also be glycosylated on the 3β-hydroxyl position [16,95,96], which is the case of 507 ergosterol with glucose, by the Atg26 UDP-glucose utilizing enzyme. This modification 508 participates in the onset of macroautophagy in fungi. The Egh1 glucosidase removes the Glc 509 unit off ergosterol. In mammals, sterols and steroids can be sulfated on the same position, 510 as to regulate gene expression or sterols/steroids turnover [97]. In all these cases, at least 511 one modification enzyme exists, that use the appropriate activated modifier (fatty acyl- or 512 acetyl-CoA, UDP-glucose, 3'-phosphoadenylyl-sulfate) to add one acyl- or acetyl- group, a 513 sugar unit, or a sulfate group on the 3β-hydroxyl, whereas de-modification enzymes can be 514 mobilized to cleave it off ergosterol. The ErdS/ErdH system, which relies on a tRNA- 515 dependent aspartylation (Asp-tRNA<sup>Asp</sup> is the activated form of Asp) of ergosterol to yield Erg- 516 Asp and its hydrolysis by the ErdH Erg-Asp hydrolase to recover free ergosterol, is 517 reminiscent of these systems. In the case of Erg glycylation by ErgS (Gly-tRNA<sup>Gly</sup> represents 518 the activated Gly), no dedicated Erg-Gly hydrolase has been found yet, but comparison of 519 these modification cycles suggests that it might also be present in Erg-Gly-producing fungi. 520

4.4. Role of aminoacylated ergosterol

521 In fungi, the role of aminoacylated ergosterol remains to be discovered. By analogy with the 522 lipid aminoacylation systems of bacteria, aminoacylated sterols could play a role in 523 antimicrobial resistance, membrane remodeling, host/pathogen interaction, virulence 524 and/or immune escape. This awaits further studies. In filamentous fungi, Erg-aa could also 525 be involved in processes such as membrane trafficking and/or signaling as described for 526 other modified sterols or modified lipids. The example of ergosteryl-3β-*O*-glucoside involved 527 in the onset of autophagy in several yeasts and in *A. oryzae* [95,98] [99] illustrates this 528 possibility. Aminoacylation of ergosterol could similarly serve in recruiting protein factors on 529 membranes and influence regulatory or trafficking pathways. 530

530 Several types of ergosterol modifications are now known. As described, they can be 531 important for central homeostasis processes in fungi. Such modifications (acetylation, 532 glycosylation, aminoacylation) have been identified in pathogenic fungi, making the enzymes 533 responsible for ergosterol modification a good target for the development of antifungal 534 drugs (as exemplified by ergosteryl-glucoside [16]), which is a critical public health issue. 535

5. Conclusion

536 ATT are widely distributed in both bacteria and fungi. Despite being identified for the first 537 time almost 60 years ago, new ones are still being discovered nowadays. The physiological 538 roles of their molecular targets are still under investigation. 539

539 In bacteria, ATT can achieve a wide range of functions ranging from aaPGSs which are tuning 540 the charge of the plasma membrane to Fem transferases that are adapting crosslinking level 541 of the PGN. A large diversity of aa transferring enzymes exists and only a few were 542 exhaustively studied. 543
543 For a long period of time, membrane modification by ATT was thought to be restricted to
544 bacteria until recently when a similar family of proteins was described in fungi: the EraS
545 protein family. Similarly, to aaPGS or Fem transferases, they can transfer different aa
546 originating from aminoacyl-tRNAs onto various substrates.

547 Finally, ATT are a highly diverse family of proteins and all of those which were described
548 here are modifying membrane related compounds. However, other families are involved in
549 the N-terminal rule pathway for protein degradation or in antibiotics and metabolite [100].
550 Thus, the flexibility of the double-GNAT fold makes it is possible to accommodate a variety
551 of substrates while conserving a similar 3D structure.

552

553

554 Acknowledgment

555 This work was supported by the "N-FLAMS" project by the Agence Nationale de la
556 Recherche (ANR-20-CE44-0002), by the "MitoCross" Laboratory of Excellence funding
557 (Labex, ANR-10-IDEX-0002-02), by the University of Strasbourg and by the CNRS (H. D. B., F.
558 F., N. M., N. Y., G. G., B. S.). N. Y., M. H. and G.G. were supported by a fellowship from the
559 French Ministère de l'Enseignement Supérieur et de la Recherche, N. M. by a postdoctoral
560 fellowship from ANR (ANR-20-CE44-0002).

561

562
563 **Bibliography**

564 [1] T. Harayama, H. Riezman, Understanding the diversity of membrane lipid
565 composition, Nat. Rev. Mol. Cell Biol. 19 (2018) 281–296.
566 https://doi.org/10.1038/nrm.2017.138.

567 [2] I.M. López-Lara, O. Geiger, Bacterial lipid diversity, Biochim. Biophys. Acta - Mol. Cell
568 Biol. Lipids. 1862 (2017) 1287–1299. https://doi.org/10.1016/j.bbalip.2016.10.007.

569 [3] C.M. Ernst, S. Kuhn, C.J. Slavetinsky, B. Krismer, S. Heilbronner, C. Gekeler, D. Kraus, S.
570 Wagner, A. Peschel, The lipid-modifying multiple peptide resistance factor is an
571 oligomer consisting of distinct interacting synthase and flippase subunits, MBio. 6
572 (2015). https://doi.org/10.1128/MBIO.02340-
573 14/SUPPL_FILE/MBO004142134ST2.DOCX.

574 [4] C. Slavetinsky, S. Kuhn, A. Peschel, Bacterial aminoacyl phospholipids – Biosynthesis
575 and role in basic cellular processes and pathogenicity, Biochim. Biophys. Acta - Mol.
576 Cell Biol. Lipids. 1862 (2017) 1310–1318.
577 https://doi.org/10.1016/j.bbalip.2016.11.013.

578 [5] N.J. Yang, H. Marlon J., Getting Across the Cell Membrane: An Overview for Small
579 Molecules, Peptides, and Proteins, Site-Specific Protein Labeling Methods Protoc.
580 1266 (2015) 1–267. https://doi.org/10.1007/978-1-4939-2272-7.

581 [6] G. Van Meer, D.R. Voelker, G.W. Feigenson, Membrane lipids: Where they are and
582 how they behave, Nat. Rev. Mol. Cell Biol. 9 (2008) 112–124.
583 https://doi.org/10.1038/nrm2330.

584 [7] D. Megrian, N. Taib, J. Witwinowski, C. Beloin, S. Gribaldo, One or two membranes?
585 Diderm Firmicutes challenge the Gram-positive/Gram-negative divide, Mol. Microbiol.
586 113 (2020) 659–671. https://doi.org/10.1111/mmi.14469.

587 [8] A.J.F. Egan, J. Errington, W. Vollmer, Regulation of peptidoglycan synthesis and
588 remodelling, Nat. Rev. Microbiol. 18 (2020) 446–460. https://doi.org/10.1038/s41579-
589 020-0366-3.

590 [9] R. Mithila, W. Suzanne, Envelope Structures of Gram-Positive Bacteria, Assess. Eval.
591 High. Educ. 37 (2012) 435. https://doi.org/10.1007/82_2015_5021.

592 [10] J.C. Henderson, S.M. Zimmerman, A.A. Crofts, J.M. Boll, L.G. Kuhns, C.M. Herrera, M.S.
593 Trent, The Power of Asymmetry: Architecture and Assembly of the Gram-Negative
594 Outer Membrane Lipid Bilayer, Annu. Rev. Microbiol. 70 (2016) 255–278.
595 https://doi.org/10.1146/annurev-micro-102215-095308.

596 [11] R.N. Fields, H. Roy, Deciphering the tRNA-dependent lipid aminoacylation systems in
597 bacteria: Novel components and structural advances, RNA Biol. 15 (2018) 480–491.
598 https://doi.org/10.1080/15476286.2017.1356980.

599 [12] A. Rella, A.M. Farnoud, M. Del Poeta, Plasma membrane lipids and their role in fungal
600 virulence, Prog. Lipid Res. 61 (2016) 63–72.
601 https://doi.org/10.1016/j.plipres.2015.11.003.

602 [13] R. Marcio L., The multifunctional fungal ergosterol, MBio. 9 (2018) 1–5.

603 [14] R. Tiwari, R. Köffel, R. Schneiter, An acetylation/deacetylation cycle controls the
604 export of sterols and steroids from S. cerevisiae, EMBO J. 26 (2007) 5109–5119.
605 https://doi.org/10.1038/sj.emboj.7601924.

606 [15] N. Jacquier, R. Schneiter, Mechanisms of sterol uptake and transport in yeast, J. 607 Steroid Biochem. Mol. Biol. 129 (2012) 70-78. 608 https://doi.org/10.1016/j.jsbmb.2010.11.014.

609 [16] T.G. Normile, K. Mcevoy, M. Del Poeta, Steryl Glycosides in Fungal Pathogenesis: An 610 Understudied Immunomodulatory Adjuvant, 2019. www.mdpi.com/journal/jof.

611 [17] N. Yakobov, F. Fischer, N. Mahmoudi, Y. Saga, C.D. Grube, H. Roy, B. Senger, G. Grob, 612 S. Tatematsu, D. Yokokawa, I. Mouyna, J.P. Latgé, H. Nakajima, T. Kushiro, H.D. 613 Becker, RNA-dependent sterol aspartylation in fungi, Proc. Natl. Acad. Sci. (2020). 614 https://doi.org/10.1073/pnas.2003266117.

615 [18] N. Yakobov, N. Mahmoudi, G. Grob, Y. Saga, H. Roy, B. Senger, S. Tatematsu, D. 616 Yokokawa, T. Kushiro, H.D. Becker, F. Fischer, RNA-dependent synthesis of ergosteryl- 617 3β-O-glycine expands the diversity of steryl-amino acid conjugates in ascomycota, J. 618 Biol. Chem. (2021).

619 [19] L. Favrot, J.S. Blanchard, O. Vergnolle, Bacterial GCN5-Related N-Acetyltransferases: 620 From Resistance to Regulation, Biochemistry. 55 (2016) 989-1002. 621 https://doi.org/10.1021/acs.biochem.5b01269.

622 [20] S. Hebecker, J. Krausze, T. Hasenkampf, J. Schneider, M. Groenewold, J. Reichelt, D. 623 Jahn, D.W. Heinz, J. Moser, Structures of two bacterial resistance factors mediating 624 tRNA-dependent aminoacylation of phosphatidylglycerol with lysine or alanine, Proc. 625 Natl. Acad. Sci. U. S. A. 112 (2015) 10691-10696. 626 https://doi.org/10.1073/pnas.1511167112.

627 [21] S. El-Gebali, J. Mistry, A. Bateman, S.R. Eddy, A. Luciani, S.C. Potter, M. Qureshi, L.J. 628 Richardson, G.A. Salazar, A. Smart, E.L.L. Sonnhammer, L. Hirsh, L. Paladin, D. 629 Piovesan, S.C.E. Tosatto, R.D. Finn, The Pfam protein families database in 2019, 630 Nucleic Acids Res. 47 (2019) D427-D432. https://doi.org/10.1093/nar/gky995.

631 [22] M. Moutiez, P. Belin, M. Gondry, Aminoacyl-tRNA-Utilizing Enzymes in Natural 632 Product Biosynthesis, Chem. Rev. 117 (2017) 5578-5618. 633 https://doi.org/10.1021/acs.chemrev.6b00523.

634 [23] E.F. Gale, J.P. Folkes, the Incorporation of Glycerol and Lysine Into the Lipid Fraction of 635 Staphylococcus Aureus., Biochem. J. 94 (1965) 390-400. 636 https://doi.org/10.1042/bj0940390.

637 [24] B.M.G. Macfarlane, Characterization of lipoamino-acids as O-amino-acids, 1 (1962) 638 136-138.

639 [25] A.M. Smith, J.S. Harrison, C.D. Grube, A.E.F. Sheppe, N. Sahara, R. Ishii, O. Nureki, H. 640 Roy, tRNA-dependent alanylation of diacylglycerol and phosphatidylglycerol in 641 Corynebacterium glutamicum, Mol. Microbiol. 98 (2015) 681-693. 642 https://doi.org/10.1111/mmi.13150.

643 [26] A. Peschel, R.W. Jack, M. Otto, L.V. Collins, P. Staubitz, G. Nicholson, H. Kalbacher, 644 W.F. Nieuwenhuizen, G. Jung, A. Tarkowski, K.P.M. Van Kessel, J.A.G. Van Strijp, 645 Staphylococcus aureus resistance to human defensins and evasion of neutrophil killing 646 via the novel virulence factor MprF is based on modification of membrane lipids with
647 L-lysine, J. Exp. Med. 193 (2001) 1067-1076. https://doi.org/10.1084/jem.193.9.1067.

648 [27] J.A. Nesbitt, W.J. Lennarz, Participation of aminoacyl transfer ribonucleic acid in aminoacyl phosphatidylglycerol synthesis. I. Specificity of lysyl phosphatidylglycerol synthetase., J. Biol. Chem. 243 (1968) 3088-3095. https://doi.org/10.1016/S0021-9258(18)93381-8.

652 [28] W.J. Lennarz, J.A. Nesbitt, J. Reiss, The participation of sRNA in the enzymatic synthesis of O-L-lysyl phosphatidylglycerol in Staphylococcus aureus., Proc. Natl. Acad. Sci. U. S. A. 55 (1966) 934-941. https://doi.org/10.1073/pnas.55.4.934.

655 [29] M. Ibba, D. Soll, Aminoacyl-tRNA synthesis, Annu. Rev. Biochem. 69 (2000) 1064-1068.

657 [30] R.M. Gould, W.J. Lennarz, Biosynthesis of aminoacyl derivatives of phosphatidylglycerol, Biochem. Biophys. Res. Commun. 26 (1967) 510-515. https://doi.org/10.1016/0006-291X(67)90578-5.

660 [31] W. Arendt, M.K. Groenewold, S. Hebecker, J.S. Dickschat, J. Moser, Identification and characterization of a periplasmic aminoacyl-phosphatidylglycerol hydrolase responsible for Pseudomonas aeruginosa lipid homeostasis, J. Biol. Chem. 288 (2013) 24717-24730. https://doi.org/10.1074/jbc.M113.482935.

664 [32] A.M. Smith, J.S. Harrison, K.M. Sprague, H. Roy, A conserved hydrolase responsible for the cleavage of aminoacylphosphatidylglycerol in the membrane of Enterococcus faecium, J. Biol. Chem. 288 (2013) 22768-22776. https://doi.org/10.1074/jbc.M113.484402.

668 [33] C.M. Ernst, P. Staubitz, N.N. Mishra, S.J.S.-J. Yang, G. Hornig, H. Kalbacher, A.S. Bayer, D. Kraus, A. Peschel, The bacterial defensin resistance protein MprF consists of separable domains for lipid lysinylation and antimicrobial peptide repulsion, PLoS Pathog. 5 (2009) 1-9. https://doi.org/10.1371/journal.ppat.1000660.

672 [34] R.N. Fields, H. Roy, Deciphering the tRNA-dependent lipid aminoacylation systems in bacteria: Novel components and structural advances, RNA Biol. 15 (2018) 480-491. https://doi.org/10.1080/15476286.2017.1356980.

675 [35] H. Roy, M. Ibba, Monitoring Lys-tRNALys phosphatidylglycerol transferase activity, Methods. 44 (2008) 164-169. https://doi.org/10.1016/j.ymeth.2007.09.002.

677 [36] A. Day, D. Wall, A Genetic Screen in Myxococcus xanthus Identifies Mutants That Uncouple Outer Membrane Exchange from a Downstream Cellular Response, J. Bacteriol. 196 (2014) 4324-4332. https://doi.org/10.1128/JB.02217-14.

680 [37] E. Maloney, D. Stankowska, J. Zhang, M. Fol, Q.-J.J. Cheng, S. Lun, W.R. Bishai, M. Rajagopalan, D. Chatterjee, M. V. Madiraju, The two-domain LysX protein of Mycobacterium tuberculosis is required for production of lysinylated phosphatidylglycerol and resistance to cationic antimicrobial peptides, PLoS Pathog. 5 (2009) e1000534. https://doi.org/10.1371/journal.ppat.1000534.

685 [38] C.J. Slavetinsky, J.N. Hauser, C. Gekeler, J. Slavetinsky, A. Geyer, A. Kraus, D. Heilingbrunner, S. Wagner, M. Tesar, B. Krismer, S. Kuhn, C.M. Ernst, A. Peschel, Sensitizing Staphylococcus aureus to antibacterial agents by decoding and blocking the lipid flippase MprF, Elife. 11 (2022) 1-20. https://doi.org/10.7554/elife.66376.
689 [39] D. Song, H. Jiao, Z. Liu, Phospholipid translocation captured in a bifunctional
690 membrane protein MprF, Nat. Commun. 12 (2021) 1–16.
691 https://doi.org/10.1038/s41467-021-23248-z.

692 [40] W. Dowhan, H. Vitrac, M. Bogdanov, Lipid-Assisted Membrane Protein Folding and
693 Topogenesis, Protein J. (2019). https://doi.org/10.1007/s10930-019-09826-7.

694 [41] S. Hebecker, J. Krausze, T. Hasenkampf, J. Schneider, M. Groenewold, J. Reichelt, D.
695 Jahn, D.W. Heinz, J. Moser, Structures of two bacterial resistance factors mediating
696 tRNA-dependent aminoacylation of phosphatidylglycerol with lysine or alanine, Proc.
697 Natl. Acad. Sci. U. S. A. 112 (2015) 10691–10696.
698 https://doi.org/10.1073/pnas.1511167112.

699 [42] P. Staubitz, H. Neumann, T. Schneider, I. Wiedemann, A. Peschel, MprF-mediated
700 biosynthesis of lysylphosphatidylglycerol, an important determinant in staphylococcal
701 defensin resistance, FEMS Microbiol. Lett. 231 (2004) 67–71.
702 https://doi.org/10.1016/S0378-1097(03)00921-2.

703 [43] S. Klein, C. Lorenzo, S. Hoffmann, J.M. Walther, S. Storbeck, T. Piekarski, B.J. Tindall, V.
704 Wray, M. Nimtz, J. Moser, Adaptation of Pseudomonas aeruginosa to various
705 conditions includes tRNA-dependent formation of alanyl-phosphatidylglycerol, Mol.
706 Microbiol. 71 (2009) 551–565. https://doi.org/10.1111/j.1365-2958.2008.06562.X.

707 [44] H. Roy, M. Ibba, Broad range amino acid specificity of RNA-dependent lipid
708 remodeling by multiple peptide resistance factors, J. Biol. Chem. 284 (2009) 29677–
709 29683. https://doi.org/10.1074/jbc.M109.046367.

710 [45] K. Thedieck, T. Hain, W. Mohamed, B.J. Tindall, M. Nimtz, T. Chakraborty, J. Wehland,
711 L. Jänsch, The MprF protein is required for lysinylation of phospholipids in listerial
712 membranes and confers resistance to cationic antimicrobial peptides (CAMPs) on
713 Listeria monocytogenes, Mol. Microbiol. 62 (2006) 1325–1339.
714 https://doi.org/10.1111/j.1365-2958.2006.05452.x.

715 [46] W. Vollmer, D. Blanot, M.A. De Pedro, Peptidoglycan structure and architecture, FEMS
716 Microbiol. Rev. 32 (2008) 149–167. https://doi.org/10.1111/j.1574-
717 6976.2007.00094.x.

718 [47] J. Garcia-Bustos, A. Tomasz, A biological price of antibiotic resistance: Major changes
719 in the peptidoglycan structure of penicillin-resistant pneumococci, Proc. Natl. Acad.
720 Sci. U. S. A. 87 (1990) 5415–5419. https://doi.org/10.1073/pnas.87.14.5415.

721 [48] T.D.H. Bugg, Bacterial Peptidoglycan Biosynthesis and its Inhibition, Compr. Nat. Prod.
722 Chem. (1999) 241–294. https://doi.org/10.1016/b978-0-08-091283-7.00080-1.

723 [49] A. Typas, M. Banzhaf, C.A. Gross, W. Vollmer, From the regulation of peptidoglycan
724 synthesis to bacterial growth and morphology, Nat. Rev. Microbiol. 10 (2012) 123–
725 136. https://doi.org/10.1038/nrmicro2677.

726 [50] K. Dare, M. Ibba, Roles of tRNA in cell wall biosynthesis, Wiley Interdiscip. Rev. RNA. 3
727 (2012) 247–264. https://doi.org/10.1002/wrna.1108.

728 [51] L.F. Leloir, Two Decades of Research on the Biosynthesis of Saccharides, 172 (1971)
729 1299–1303.

730 [52] M.H. Mandelstam, J.L. Strominger, On The Structure Of The Cell Wall Of
731 Staphylococcus Aureus, Biochem. Biophys. Res. Commun. 5 (1961) 466-471.

732 [53] M. Matsuhashi, C.P. Dietrich, J.L. Strominger, Incorporation of glycine into the cell wall glycopeptide in Staphylococcus aureus: role of sRNA and lipid intermediates., Proc.
733
734 Natl. Acad. Sci. U. S. A. 54 (1965) 587-594. https://doi.org/10.1073/pnas.54.2.587.

735 [54] W. Katz, M. Matsuhashi, C.P. Dietrich, J.L. Strominger, Biosynthesis of the Peptidoglycan of Bacterial Cell Walls, J. Biol. Chem. 242 (1967) 3207-3217.
736
737 https://doi.org/10.1016/s0021-9258(18)95951-x.

738 [55] W.S.L. Roberts, J.F. Petit, J.L. Strominger, Biosynthesis of the Peptidoglycan of Bacterial Cell Walls, J. Biol. Chem. 243 (1968) 768-772.
739
740 https://doi.org/10.1016/s0021-9258(19)81731-3.

741 [56] W.S.L. Roberts, J.L. Strominger, D. Söll, Biosynthesis of the Peptidoglycan of Bacterial Cell Walls, J. Biol. Chem. 243 (1968) 749-756. https://doi.org/10.1016/s0021-9258(19)81729-5.

744 [57] J.F. Petit, J.L. Strominger, D. Söll, Biosynthesis of the Peptidoglycan of Bacterial Cell Walls, J. Biol. Chem. 243 (1968) 757-767. https://doi.org/10.1016/s0021-9258(19)81730-1.

747 [58] R.M. Bumsted, J.L. Dahl, D. Söll, J.L. Strominger, Biosynthesis of the Peptidoglycan of Bacterial Cell Walls, J. Biol. Chem. 243 (1968) 779-782.
748
749 https://doi.org/10.1016/s0021-9258(19)81733-7.

750 [59] B. Berger Bachi, Insertional inactivation of staphylococcal methicillin resistance by Tn551, J. Bacteriol. 154 (1983) 479-487. https://doi.org/10.1128/jb.154.1.479-487.1983.

753 [60] B. Berger-Bächi, L. Barberis-Maino, A. Strässle, F.H. Kayser, FemA, a host-mediated factor essential for methicillin resistance in Staphylococcus aureus: Molecular cloning and characterization, MGG Mol. Gen. Genet. 219 (1989) 263-269.
754
755
756 https://doi.org/10.1007/BF00261186.

757 [61] A.M. Strandén, K. Ehlert, H. Labischinski, B. Berger-BÄCHI, Cell wall monoglycine cross-bridges and methicillin hypersusceptibility in a femAB null mutant of methicillin-resistant Staphylococcus aureus, J. Bacteriol. 179 (1997) 9-16.
758
759
760 https://doi.org/10.1128/jb.179.1.9-16.1997.

761 [62] S. Rohrer, K. Ehlert, M. Tschierske, H. Labischinski, B. Berger-Bachi, The essential Staphylococcus aureus gene fmhB is involved in the first step of peptidoglycan pentaglycine interpeptide formation, Proc. Natl. Acad. Sci. U. S. A. 96 (1999) 9351-9356. https://doi.org/10.1073/pnas.96.16.9351.

765 [63] J.M. Monteiro, G. Covas, D. Rausch, S.R. Filipe, T. Schneider, H.G. Sahl, M.G. Pinho, The pentaglycine bridges of Staphylococcus aureus peptidoglycan are essential for cell integrity, Sci. Rep. 9 (2019) 1-10. https://doi.org/10.1038/s41598-019-41461-1.

768 [64] T. Schneider, M.M. Senn, B. Berger-Bächi, A. Tossi, H.G. Sahl, I. Wiedemann, In vitro assembly of a complete, pentaglycine interpeptide bridge containing cell wall precursor (lipid II-Gly5) of Staphylococcus aureus, Mol. Microbiol. 53 (2004) 675-685.
769
770
771 https://doi.org/10.1111/j.1365-2958.2004.04149.x.

772 [65] S. Giannouli, A. Kyritsis, N. Malissovas, H.D. Becker, C. Stathopoulos, On the role of an
773 unusual tRNAGly isoacceptor in Staphylococcus aureus, Biochimie. 91 (2009) 344-351.
774 https://doi.org/10.1016/j.biochi.2008.10.009.

775 [66] L. Rietmeyer, N. Fix-Boulier, C. Le Fournis, L. Iannazzo, C. Kitoun, D. Patin, D. Mengin-Lecreulx, M. Ethève-Quelquejeu, M. Arthur, M. Fonvielle, Partition of tRNAGly isoacceptors between protein and cell-wall peptidoglycan synthesis in Staphylococcus aureus, Nucleic Acids Res. 49 (2021) 684-699. https://doi.org/10.1093/nar/gkaa1242.

779 [67] M. Tschierske, C. Mori, S. Rohrer, K. Ehlert, K.J. Shaw, B. Berger-Bächi, Identification of three additional femAB-like open reading frames in Staphylococcus aureus, FEMS Microbiol. Lett. 171 (1999) 97-102. https://doi.org/10.1016/S0378-1097(98)00586-2.

782 [68] M. Sugai, T. Fujiwara, K. Ohta, H. Komatsuzawa, M. Ohara, H. Suginaka, epr, which encodes glycylglycine endopeptidase resistance, is homologous to femAB and affects serine content of peptidoglycan cross bridges in Staphylococcus capitis and Staphylococcus aureus, J. Bacteriol. 179 (1997) 4311-4318.
786 https://doi.org/10.1128/jb.179.13.4311-4318.1997.

787 [69] S. Willing, E. Dyer, O. Schneewind, D. Missiakas, FmHA and FmHC of Staphylococcus aureus incorporate serine residues into peptidoglycan cross-bridges, J. Biol. Chem. 295 (2020) 13664-13676. https://doi.org/10.1074/jbc.ra120.014371.

790 [70] A. York, A.J. Lloyd, C.I. del Genio, J. Shearer, K.J. Hinxman, K. Fritz, V. Fulop, C.G. Dowson, S. Khalid, D.I. Roper, Structure-based modeling and dynamics of MurM, a Streptococcus pneumoniae penicillin resistance determinant present at the cytoplasmic membrane, Structure. 29 (2021) 731-742.e6.
794 https://doi.org/10.1016/j.str.2021.03.001.

795 [71] S. Biarrotte-Sorin, A.P. Maillard, J. Delettré, W. Sougakoff, M. Arthur, C. Mayer, Crystal Structures of Weissella viridescens FemX and Its Complex with UDP-MurNAc-Pentapeptide: Insights into FemABX Family Substrates Recognition, Structure. 12 (2004) 257-267. https://doi.org/10.1016/j.str.2004.01.006.

799 [72] T.E. Benson, D.B. Prince, V.T. Mutchler, K.A. Curry, A.M. Ho, R.W. Sarver, J.C. Hagadorn, G.H. Choi, R.L. Garlick, X-ray crystal structure of Staphylococcus aureus FemA, Structure. 10 (2002) 1107-1115. https://doi.org/10.1016/S0969-2126(02)00807-9.

803 [73] V. Biou, A. Yaremchuk, M. Tukalo, S. Cusack, The 2.9 Å crystal structure of T. thermophilus Seryl-tRNA Synthetase Complexed with tRNASer, Science (80-.). 263 (1994) 1404-1410.

806 [74] S. Biarrotte-Sorin, A.P. Maillard, J. Delettré, W. Sougakoff, M. Arthur, C. Mayer, Crystal Structures of Weissella viridescens FemX and Its Complex with UDP-MurNAc-Pentapeptide: Insights into FemABX Family Substrates Recognition, Structure. 12 (2004) 257-267. https://doi.org/10.1016/J.STR.2004.01.006.

810 [75] S. Biarrotte-sorin, A.P. Maillard, crystallization papers Crystallization and preliminary X-ray analysis of Weissella viridescens FemX UDP-MurNAc-pentapeptide : L -alanine ligase crystallization papers, (2003) 1055-1057.

813 [76] M. Jarick, U. Bertsche, M. Stahl, D. Schultz, K. Methling, M. Lalk, C. Stigloher, M. Steger, A. Schlosser, K. Ohlsen, The serine/threonine kinase Stk and the phosphatase Stp regulate cell wall synthesis in Staphylococcus aureus, Sci. Rep. 8 (2018) 1-13.
816 https://doi.org/10.1038/s41598-018-32109-7.

817 [77] J.F. Garcia-Bustos, B.T. Chait, A. Tomasz, Altered peptidoglycan structure in a pneumococcal transformant resistant to penicillin, J. Bacteriol. 170 (1988) 2136-2147.
818
819 https://doi.org/10.1128/jb.170.5.2143-2147.1988.

820 [78] S.S. Hegde, T.E. Shrader, FemABX Family Members Are Novel Nonribosomal Peptidyltransferases and Important Pathogen-specific Drug Targets, J. Biol. Chem. 276 (2001) 6998-7003. https://doi.org/10.1074/JBC.M008591200.

823 [79] A.P. Maillard, S. Biarrotte-Sorin, R. Villet, S. Mesnage, A. Bouhss, W. Sougakoff, C. Mayer, M. Arthur, Structure-based site-directed mutagenesis of the UDP-MurNAc-pentapeptide- binding cavity of the FemX alanyl transferase from Weissella viridescens, J. Bacteriol. 187 (2005) 3833-3838.
826
827 https://doi.org/10.1128/JB.187.11.3833-3838.2005.

828 [80] M. Fonvielle, I. Li De La Sierra-Gallay, A.H. El-Sagheer, M. Lecerf, D. Patin, D. Mellal, C. Mayer, D. Blanot, N. Gale, T. Brown, H. Van Tilbeurgh, M. Ethève-Quelquejeu, M. Arthur, The structure of FemXWv in complex with a peptidyl-RNA conjugate: Mechanism of aminoacyl transfer from Ala-tRNAAla to peptidoglycan precursors, Angew. Chemie - Int. Ed. 52 (2013) 7278-7281.
833
834
835
836 https://doi.org/10.1002/anie.201301411.

834 [81] J.W. Spatafora, M.C. Aime, I. V. Gregoriev, F. Martin, J.E. Stajich, M. Blackwell, The fungal tree of life: From molecular systematics to genome-scale phylogenies, The Fungal Kingdom. (2017) 3-34. https://doi.org/10.1128/9781555819583.ch1.

837 [82] C. Askew, A. Sellam, E. Epp, H. Hogues, A. Mullick, A. Nantel, M. Whiteway, Transcriptional regulation of carbohydrate metabolism in the human pathogen Candida albicans, PLoS Pathog. 5 (2009).
839
840 https://doi.org/10.1371/journal.ppat.1000612.

841 [83] B.M. Barker, K. Kroll, M. Vödisch, A. Mazurie, O. Kniemeyer, R.A. Cramer, Transcriptomic and proteomic analyses of the Aspergillus fumigatus hypoxia response using an oxygen-controlled fermenter, BMC Genomics. 13 (2012) 62.
842
843
844 https://doi.org/10.1186/1471-2164-13-62.

845 [84] C.D. Chun, O.W. Liu, H.D. Madhani, A link between virulence and homeostatic responses to hypoxia during infection by the human fungal pathogen Cryptococcus neoformans, PLoS Pathog. 3 (2007) 0225-0238.
847
848 https://doi.org/10.1371/journal.ppat.0030022.

849 [85] E.R. Setiadi, T. Doedt, F. Cottier, C. Noffz, J.F. Ernst, Transcriptional Response of Candida albicans to Hypoxia: Linkage of Oxygen Sensing and Efg1p-regulatory Networks, J. Mol. Biol. 361 (2006) 399-411.
851
852 https://doi.org/10.1016/j.jmb.2006.06.040.

853 [86] K. Shimizu, M. Osada, K. Takemoto, Y. Yamamoto, T. Asai, N. Oku, Temperature-dependent transfer of amphotericin B from liposomal membrane of AmBisome to fungal cell membrane, J. Control. Release. 141 (2010) 208-215.
854
855
856 https://doi.org/10.1016/j.jconrel.2009.09.019.

857 [87] N. Tao, Q. OuYang, L. Jia, Citral inhibits mycelial growth of Penicillium italicum by a membrane damage mechanism, Food Control. 41 (2014) 116-121.
858
859 https://doi.org/10.1016/j.foodcont.2014.01.010.

860 [88] A. Khan, A. Ahmad, F. Akhtar, S. Yousuf, I. Xess, L.A. Khan, N. Manzoor, Ocimum sanctum essential oil and its active principles exert their antifungal activity by disrupting ergosterol biosynthesis and membrane integrity, Res. Microbiol. 161 (2010) 816-823. https://doi.org/10.1016/j.resmic.2010.09.008.

864 [89] C. Onyewu, J.R. Blankenship, M. Del Poeta, J. Heitman, Ergosterol biosynthesis inhibitors become fungicidal when combined with calcineurin inhibitors against Candida albicans, Candida glabrata, and Candida krusei, Antimicrob. Agents Chemother. 47 (2003) 956-964. https://doi.org/10.1128/AAC.47.3.956-964.2003.

868 [90] X. Sun, J. Wang, D. Feng, Z. Ma, H. Li, PdCYP51B, a new putative sterol 14α-demethylase gene of Penicillium digitatum involved in resistance to imazalil and other fungicides inhibiting ergosterol synthesis, Appl. Microbiol. Biotechnol. 91 (2011) 1107-1119. https://doi.org/10.1007/s00253-011-3355-7.

872 [91] A.L. Munn, A. Heese-Peck, B.J. Stevenson, H. Pichler, H. Riezman, Specific sterols required for the internalization step of endocytosis in yeast, Mol. Biol. Cell. 10 (1999) 3943-3957. https://doi.org/10.1091/mbc.10.11.3943.

875 [92] A. Heese-Peck, H. Pichler, B. Zanolari, R. Watanabe, G. Daum, H. Riezman, Multiple Functions of Sterols in Yeast Endocytosis Antje, Mol. Biol. Cell. 13 (2002) 2664-2680. https://doi.org/10.1091/mbc.E02.

878 [93] M. Kato, W. Wickner, Ergosterol is required for the Sec18/ATP-dependent priming step of homotypic vacuole fusion, EMBO J. 20 (2001) 4035-4040. https://doi.org/10.1093/emboj/20.15.4035.

881 [94] L.W. Parks, W.M. Casey, Physiological implications of sterol biosynthesis in yeast, Annu. Rev. Microbiol. 49 (1995) 95-116. https://doi.org/10.1146/annurev.mi.49.100195.000523.

884 [95] S. Grille, A. Zaslawski, S. Thiele, J. Plat, D. Warnecke, The functions of steryl glycosides come to those who wait: Recent advances in plants, fungi, bacteria and animals, Prog. Lipid Res. 49 (2010) 262-288. https://doi.org/10.1016/j.plipres.2010.02.001.

887 [96] M. Shimamura, Structure, metabolism and biological functions of steryl glycosides in mammals, Biochem. J. 477 (2020) 4243-4261. https://doi.org/10.1042/BCJ20200532.

889 [97] J.W. Mueller, L.C. Gilligan, J. Idkowiak, W. Arlt, P.A. Foster, The regulation of steroid action by sulfation and desulfation, Endocr. Rev. 36 (2015) 526-563. https://doi.org/10.1210/er.2015-1036.

892 [98] T. Kikuma, T. Tadokoro, J.I. Maruyama, K. Kitamoto, AoAtg26, a putative sterol glucosyltransferase, is required for autophagic degradation of peroxisomes, mitochondria, and nuclei in the filamentous fungus Aspergillus oryzae, Biosci. Biotechnol. Biochem. 81 (2017) 384-395. https://doi.org/10.1080/09168451.2016.1240603.

897 [99] S.I. Yamashita, M. Oku, Y. Sakai, Functions of PI4P and sterol glucoside are necessary for the synthesis of a nascent membrane structure during pexophagy, Autophagy. 3 (2007) 35-37. https://doi.org/10.4161/auto.3311.

900 [100] M. Hemmerle, M. Wendenbaum, G. Grob, N. Yakobov, N. Mahmoudi, B. Senger, S.
901 Debard, F. Fischer, H.D. Becker, Noncanonical inputs and outputs of tRNA
902 aminoacylation, Enzymes. 48 (2020) 117-147.
903 https://doi.org/10.1016/bs.enz.2020.04.003.

904
Figure 1: mains families of aminoacyl-tRNA transferases (ATT) that transfer amino acid on a membrane related compound. These are so-called ATT because they use an aminoacyl-tRNA (aa-tRNA) synthetized by the cognate aminoacyl-tRNA synthetase as a substrate and transfer the amino acid moiety on a wide range of "acceptor" substrate. Three families of AAT are represented: (A) the family of Fem transferases which adds an interpeptide bridge between stem peptides of peptidoglycan. (B) the EraS family, found in fungi which add an aspartate or a glycine onto ergosterol and the Multiple peptide resistance Factor (MprF) which transfer amino acids on phospholipids or diacyl-glycerol (here only phosphatidyl-glycerol, a major phospholipid is depicted).

Figure 2: Comparison of the crystallographic 3D structures of 3 different Aminoacyl-tRNA transferases, GNAT I, GNAT II and α(+) helix are depicted in blue, green, and red respectively. Despite having different activities and encoded by the genome of 3 different organisms, the structure of the GNAT II and the α(+) helix is widely similar, which reflect the common aa-tRNA substrate for those domains. GNAT I structure on the other hand, is much more divergent, which also reflects the necessity for each enzyme to recognize and accommodate a structurally different substrate through this domain.

Figure 3: Topology of the N-terminal flippase domain from the MprF protein from *Staphylococcus aureus*. Transmembrane helices are depicted as light blue box and extra- or intra-cellular loops are depicted as string. The conformation of helices 7 and 8 is shown only in 1 conformation but is believed to change upon flippase activity. Aminoacyl-tRNA transferase domain is depicted in C-terminus and is responsible for the synthesis of the lysyl-phosphatidyl-glycerol.

Figure 4: overview of peptidoglycan interpeptide bridge synthesis by Fem ligase proteins. Peptidoglycan is made of glycans strands of N-acetyl-muramic acid (MurNac) and N-acetyl-glucosamine (GlcNAc) which are linked together by pentapeptide bridges. Bridges are linked together to form a net-like structure. However, peptide bridges can be indirectly crosslinked by several amino acids that are added in an aminoacyl-tRNA dependent manner by fem ligases. This indirect crosslinking can be performed either after synthesis of the lipid II stem peptide, on the inner leaflet and before its export in outer leaflet of the plasma membrane. This pathway was the first described in *S. aureus*, but another pathway was identified in *W. viridescens* where the crosslinking bridge is added on UDP-MurNac-pentapeptide precursor, which is then used to synthetize lipid II stem-peptide. The following of the synthesis is similar in both organisms.

Figure 5: Ergosteryl-aspartate synthase uses Aspartate and tRNA to yield Asp-tRNA through its synthetase domain. Aspartate can then be transferred onto the 3-b-hydroxyle group of ergosterol. In addition to this synthesis process, a hydrolysis process catalyzed by ergosteryl-aspartate hydrolase can take place, possibly leading to a tight regulation of the amount of Ergosteryl-aspartate.

Figure 6: Ergosteryl-glycine synthase uses directly Gly-tRNA, produced by Glycyl-tRNA synthetase, to transfer glycine to the 3-b-hydroxyle group of ergosterol, leading to the production of an ester product, namely ergosteryl-glycine.
Ergosterol aminotransferase

Phosphatidyl glycerol

C

Amino acid phosphatase
(Ps6024)

Phosphatidyl inositol
(Ps6024)

Phosphatidyl inositol
(Ps6024)

B

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

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

<figure><img src="image_4.png" /></figure>

<figure><img src="image_5.png" /></figure>

C

<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /></figure>
α' helix

<figure><img src="image_8.png" /><figcaption>GNAT I</figcaption></figure>

<caption>GNAT II</caption>

* LPGS L Lanthipeptide (transmembrane domain; PDB 4J6)

* FasA A surface protein (PDB 4J6)

* Farn W protein (PDB 4J6)
<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>
<figure><img src="image_11.png" /></figure>

The diagram shows a network of interconnected cells. Each cell is represented by a circle, and the connections between cells are shown by lines. The lines represent the presence of chemical or physical bonds between the cells. The size of the circles is proportional to the number of connections each cell has. The color of the circles represents the type of cell: red forSupport cells, blue for Epithelial cells, green for Endothelial cells, and purple for myeloid cells.

<figure><img src="image_12.png" /></figure>

The diagram also shows a network of blood vessels (red), capillaries (blue), blood vessels (green), and myeloid cells (purple). The size of the circles is proportional to the number of connections each cell has. The color of the circles represents the type of cell: red for myeloid cells, blue for macrophages, green for neutrophils, and purple for eosinophils.
<figure><img src="image_13.png" /></figure>
<figure><img src="image_14.png" /></figure>