Skip to content
naulon
Get started

No retraction, correction or concern found when last checked on Oct 27, 2026.

Mini Review: Bacterial Membrane Composition and Its Modulation in Response to Stress

Jessica R. Willdigg, John D. Helmann

Frontiers in Molecular Biosciences · 2021 · Review · CC BY

doi:10.3389/fmolb.2021.634438

Abstract

Antibiotics and other agents that perturb the synthesis or integrity of the bacterial cell envelope trigger compensatory stress responses. Focusing on Bacillus subtilis as a model system, this mini-review summarizes current views of membrane structure and insights into how cell envelope stress responses remodel and protect the membrane. Altering the composition and properties of the membrane and its associated proteome can protect cells against detergents, antimicrobial peptides, and pore-forming compounds while also, indirectly, contributing to resistance against compounds that affect cell wall synthesis. Many of these regulatory responses are broadly conserved, even where the details of regulation may differ, and can be important in the emergence of antibiotic resistance in clinical settings.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 27, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction: Membrane Homeostasis and Its Modulation in Response to Stress

The cell envelope is a multilayered outer barrier that protects the cell from a changing environment. Cell envelope stress responses (CESRs) are regulatory pathways that sense threats and mount a protective response, often involving modification of lipopolysaccharides (in Gram-negative bacteria), teichoic acids (Gram-positive bacteria), peptidoglycan, and the inner membrane. Here, we focus on Bacillus subtilis as a Gram-positive model for the role of CESRs in membrane homeostasis.

The cell membrane is a dynamic, fluid mosaic comprising a lipid bilayer and associated proteins (Figure 1). In B. subtilis, the major lipid species are phospholipids, glucolipids, and the lipoteichoic acids (LTA). The membrane proteome includes proteins for transport and signaling, as well as membrane synthesis, remodeling, and protection. As the innermost and last line of defense, the cell membrane is critical for viability. In B. subtilis, for example, collapsing the proton motive force activates autolysins resulting in rapid cell lysis. Membrane-active compounds such as detergents, antimicrobial peptides, and pore-forming compounds often trigger stress responses that modify the lipidome and membrane proteome to confer resistance. Membrane stress responses can modify the cell membrane, by (i) modulating the length, branching, and saturation of the fatty acid (FA) acyl chains, (ii) altering membrane lipid composition, or (iii) synthesizing proteins that modify or protect the membrane (Table 1).

The Regulation of FA Synthesis During Growth

Most bacteria utilize a type II FA synthase that catalyzes repeated cycles of acyl chain elongation. The committed step, catalyzed by acetyl-CoA carboxylase (ACC), generates malonyl-CoA and then malonyl-ACP to serve in FA chain initiation by FabH and elongation by FabF. B. subtilis has two isoforms of FabH, and both preferentially synthesize branched chain FAs (BCFAs). Acylation of glycerol-3-phosphate by the PlsX/PlsY/PlsC acyltransferase system with long chain FAs generates phosphatidic acid, the precursor to all other phospholipids.

FapR is the key transcriptional regulator of membrane lipid synthesis in B. subtilis and clinically relevant pathogens such as Staphylococcus aureus, Bacillus anthracis, and Listeria monocytogenes, and modulates the overall rate of membrane synthesis in response to precursor availability. B. subtilis FapR represses genes for FA and phospholipid synthesis, and this repression is relieved by allosteric interactions with malonyl-CoA or malonyl-ACP.

As a branchpoint enzyme, ACC is often under complex regulation. In B. subtilis, ACC is regulated in part by YqhY, a conserved DUF322/Asp23 protein which is highly expressed and often encoded together with ACC subunits as part of an accB-accC-yqhY operon. The namesake, S. aureus Asp23, is a membrane-associated protein originally linked to alkaline shock. Loss of Asp23/YqhY causes cell wall stress and poor growth. In B. subtilis, yqhY null mutants acquire suppressors that decrease ACC activity, but this selective pressure is alleviated in medium supplemented with acetate. We suggest that ACC-dependent depletion of acetyl-CoA may contribute to wall stress by negatively affecting synthesis of UDP-N-acetylglucosamine needed for peptidoglycan synthesis. A key challenge for future research will be to understand the precise role of YqhY/Asp23 proteins and how they control ACC activity to balance FA synthesis with other cellular needs.

Modulating FA Composition for Homeoviscous Adaptation

Tuning of FA composition provides one way in which the cell can optimize membrane properties in response to a changing environment. Even under non-stressed conditions, B. subtilis membranes contain ∼7 distinct FAs varying in length from C14 to C18 (indicating the number of carbon atoms) and include both branched (∼24% iso and 66% anteiso) and straight chain (∼10%) FAs. Since membrane phospholipids and glucolipids each contain 2 FA chains, the lipidome contains a complex mix of species (Figure 1B), with a preponderance containing one C15 and one C17 FA chain.

Modifications of FAs are important for regulating membrane fluidity in a process known as homeoviscous adaptation. In B. subtilis, temperature downshift induces a FA desaturase (Des) controlled by the DesKR two-component system (TCS). Des modifies existing membrane lipids, and is thereby suited for rapid adaptation. DesK is one of the better understood TCS sensors, with both kinase and phosphatase activity. DesK lacks an extracellular sensor domain, but has multiple transmembrane segments that sense changes in the membrane physical state. DesK phosphorylates the DesR response regulator, which induces des, encoding a FA Δ5 desaturase. The resultant unsaturated FAs increase bilayer fluidity, which restores DesK phosphatase activity in a negative feedback loop. Longer term adaptation to low temperatures relies on an isoleucine-dependent switch to primarily anteiso-FAs. Since anteiso-FAs perturb the lateral interactions between adjacent lipids to a greater extent than iso-FAs (Figure 1B), this shift increases membrane fluidity. This shift may result from a cold-dependent change in FabH activity.

Membranes must also adapt to conditions that increase fluidity. In B. subtilis, the ECF σ factor σW is activated by detergents, antibiotics, and bacteriocins active on the membrane. A σW promoter within the fabHA-fabF operon plays a major role in homeoviscous adaptation. Activation of σW leads to a decrease in FabHA levels, resulting in increased reliance on FabHB and an increase in straight chain FAs (from ∼10 to 30%). Elevated expression of the FabF elongation enzyme leads to increased FA chain length. The combined effect is a membrane with longer acyl chains and less BCFA. This increased membrane rigidity serves to protect cells against detergents and antimicrobial peptides. The activation of σW is controlled by regulated proteolysis of its membrane-bound anti-σW factor (RsiW). However, the mechanisms by which membrane stressors trigger σW activation remain unclear.

To better understand the role of FA heterogeneity in controlling membrane properties, it would be desirable to study bacteria with chemically simple membranes. This has been achieved in B. subtilis by feeding exogenous FAs to cells with de novo FA synthesis blocked by cerulenin and a mutation to inhibit FA degradation. Growth can be rescued with only two FA species: a straight-chain C16 FA (high melting) and an anteiso C15 FA (low melting). Even with only these two FA species, four distinct arrangements are possible upon acylation of glycerol-3-phosphate to generate phosphatidic acid. Cells compensate for this reduced FA complexity by altering the distribution of phospholipid headgroups, a modest induction of the DesRK system, apparent downregulation of the σW stress response, and an increase in isoprenoid lipids. These results highlight the remarkable adaptability of bacterial membranes, and the interconnection between diverse stress responses.

Overview of Membrane Lipid Composition and Synthesis

One of the persistent challenges in membrane biology is to define the roles of the diverse constituent lipids. Although membranes have a complex and adaptable composition (the lipidome), cells are remarkably resilient to genetic alterations that remove lipid species. Because of its single membrane and ease of genetic manipulation, B. subtilis presents an attractive model system. The B. subtilis lipidome comprises ∼70% phospholipids and ∼30% neutral glucolipids. The major phospholipids are phosphatidylglycerol (PhG) and phosphatidylethanolamine (PE), with minor contributions from cardiolipin and lysylphosphatidylglycerol (LPG). Variations in phospholipid headgroup size and charge modulate membrane properties (Figure 1B). Membranes also contain LTA anchored to neutral glucolipids, which together with peptidoglycan-linked wall teichoic acid (WTA) can account for up to 60% of the dry weight of the cell wall. However, LTA fractionates with the wall during membrane lipid extraction, and is often not considered in lipidome measurements.

The only essential phospholipid in B. subtilis is PhG. Remarkably, the membrane can be simplified to contain close to 100% PhG with no glucolipids. Despite a greatly simplified membrane, such mutants can grow rapidly, albeit with a highly abnormal coiled filament morphology. Genetic perturbations of membrane composition can lead to resistance to cationic antimicrobial peptides (CAMP). For example, gain-of-function mutations in mprF, encoding the LPG synthase/flippase, can confer daptomycin resistance possibly by reducing surface charge. Consistently, mprF null mutants have increased daptomycin sensitivity and overexpression decreases sensitivity in B. subtilis. Daptomycin resistance also results from pgsA mutations that decrease PhG levels.

In addition to the dominant phospholipids and glucolipids, membranes contain numerous other lipid species. Most prominent are the isoprenoid lipids synthesized by polymerization of C5 isoprene units (Figure 1C). The key intermediate farnesyl-PP (C15) can be joined (head-to-head) to generate squalene (C30), a precursor of cholesterol and other sterols in eukaryotes and of structurally related hopanoid lipids in many bacteria. One major hopanoid is diploterol (Figure 1C), with five fused rings that can be further modified in a variety of ways. Farnesyl-PP can also be extended by UppS, which sequentially adds eight isopentenyl units to generate undecaprenyl-PP, the C55 carrier lipid that supports cell wall synthesis (Figure 1C). Alternatively, the HepST complex can extend farnesyl-PP to generate heptaprenyl (C35)-PP, an isoprenoid used as a lipid anchor for menaquinone (MK-7), the electron carrier for respiration. In B. subtilis, this same precursor can be processed to polycyclic C35-sesquarterpenoids, which may be functionally similar to C30 hopanoids. This process is initiated by YtpB, which generates tetraprenyl-β-curcumene, and then SqhC (a homolog of squalene-hopene cyclases) to generate the C35 tetracyclic product known as baciterpenol A. Although initially described in spores, and named “sporulenes”, these sesquarterpenoids are found in vegetative cells. Finally, heptaprenyl-PP can be coupled to glycerol-1-phosphate by PcrB, and then further processed by an unidentified phosphatase and the YvoF acetyltranferase to generate an ether linked lipid of unknown function.

Lateral Heterogeneity and Functional Membrane Microdomains

In eukaryotes, cholesterol is associated with the generation of functional membrane microdomains (FMM), also called lipid rafts. These regions have relatively low membrane fluidity (a liquid-ordered, or Lo phase) and are associated with flotillins. B. subtilis also encodes flotillin homologs, regulated by σW. These proteins, subsequently renamed FloA and FloT, are implicated in FMM formation (Figure 1A). The notion of FMMs in bacteria received strong impetus from the finding that yisP mutants, lacking a putative squalene synthase, was defective in biofilm formation. Together with the finding of a punctate localization for FloT, and chemical inhibition studies with compounds that affect sterol synthesis, this led to the proposal that bacteria harbor FMMs. However, subsequent work revealed that YisP is a farnesyl-PP phosphatase that generates farnesol (Figure 1C), rather than squalene. Moreover, farnesol itself complements the biofilm defect of the yisP mutant, suggesting that this long chain alcohol may have an ordering effect on FMMs provided in other systems by hopanoids or carotenoids. B. subtilis FMMs are enriched in flotillins (FloA and FloT) and their associated signaling complexes, with FMM formation apparently stabilized by farnesol (YisP product). No role for the C35 isoprenoid lipids has yet been demonstrated in biofilm formation or in FMM formation or function.

Lateral heterogeneity, including FMMs, is likely a feature of most bacterial membranes. However, the lipid species that are required to form FMMs are still poorly understood, but likely include carotenoids, hopanoids, and other polycyclic isoprenoid lipids. Hopanoids are structurally diverse and fulfill a broad range of functions in bacterial membranes. The hopanoid diplopterol (Figure 1C) orders saturated lipids and glycolipids in the outer membrane of Methylobacterium extorquens, and deficient mutants are impaired in multidrug transport. Hopanoids and other polycyclic isoprenoids are present in many Gram-positive bacteria as well, suggestive of a role in the plasma membrane. In methicillin-resistant S. aureus, the carotenoid staphyloxanthin (Figure 1C) colocalizes in FMMs with FloA, and disruption of these domains with isoprenoid synthesis inhibitors interferes with the function of the penicillin-binding protein required for β-lactam resistance (PBP2a). The formation and function of FMMs, in both the inner (plasma) and outer membrane, remains an important area for future research.

Cell Envelope Stress Responses That Modulate Lipid Composition

Bacteria generally have a negatively charged membrane, which contributes to their susceptibility to CAMPs, bacteriocins, and antimicrobials. In B. subtilis, membrane composition and properties are regulated by ECF σ factors. Because of their overlapping activation and promoter recognition properties, these CESRs are intertwined and referred to as an σECF stress response. In B. subtilis, activation of σX reduces the net negative charge of the membrane by increasing zwitterionic PE levels. The net negative charge of the cell wall can be further reduced by D-alanylation of teichoic acids, activated by σX and σV, a lysozyme-responsive CESR. In S. aureus, surface membrane charge is modified by the induction of mprF by the GraRS TCS, thereby increasing LPG levels. In B. anthracis, the membrane-active compound targocil activates the EdsRS TCS, which induces expression of a cardiolipin synthase. Thus, many different stimuli can trigger changes in the membrane lipidome.

Bacillus subtilis σECF factors also control other membrane-related functions, although the effects are not yet understood. For example, σM activates the ytpAB operon. The YtpA lysophospholipase cleaves FAs from the 2 position of phospholipids resulting in a lysophospholipid (bacilysocin) suggested to function as an antibiotic. However, it is unclear if bacilysocin is ever released at levels sufficient to serve as an antibiotic, and it may instead modify membrane properties or be an intermediate in lipid remodeling. As noted above, YtpB initiates synthesis of baciterpenol (Figure 1C;;; ). Genetic studies have revealed only modest phenotypes for ytpAB mutants, including effects on antibiotic sensitivity, sporulation, and germination. In the case of ytpB, the observed phenotype (bacitracin sensitivity) was due to the accumulation of the substrate (heptaprenyl-PP) rather than a loss of baciterpenol.

Genetic perturbations of membrane composition can also trigger CESRs. For example, deletion of LTA synthases induces σECF factors. An ltaS mutation upregulates σM, which then activates expression of the alternate LTA synthase LtaSa. The absence of both ltaS and ltaSa leads to activation of additional σECF factors. The depletion of PhG, a building block of LTA, also activates σM and to a lesser extent σV. The effects of mutations that affect glucolipids have been particularly challenging to understand. Glucolipids produced by UgtP are important membrane lipids and also serve as the lipid anchor of LTA. ugtP mutants lacking glucolipids are shorter and rounder, have abnormal localization of MreB, and altered assembly of FtsZ. Whether this abnormal morphology is due, in part, to the loss of glucolipids is unclear. Mutation of ugtP activates a σECF stress response and can be suppressed by production of monoglycosyldiacylglycerol (MGlcDG) using a heterologous synthase. Since this product does not function as an LTA anchor lipid, this suggests that it is the loss of glucolipids that induces the σECF response. The mechanistic basis for activation of σECF factors in the absence of glucolipids is unclear, but at least for σV does not require intramembrane proteolysis of the anti-σ factor. One hypothesis is that glucolipids might regulate folding and function of intramembrane proteins.

Cell Envelope Stress Responses That Function Through Membrane Proteins

In addition to modulating lipid composition, CESRs also induce proteins that function in membrane protection and remodeling. In B. subtilis, these proteins include two flotillin homologs (FloA, FloT), two members of the phage shock protein family (LiaH, PspA), as well as antibiotic specific detoxification modules. The roles of these proteins in stabilizing and repairing the membrane are increasingly appreciated, although the precise mechanisms remain controversial.

Flotillins and Modulation of Membrane Fluidity

Flotillins are members of the widely conserved stomatin, prohibitin, flotillin, and HflK/C (SPFH) domain proteins. Flotillins localize to FMMs and are thought to be required for FMM function. In S. aureus, FloA colocalizes with staphyloxanthin in FMMs. In other systems, flotillins and FMMs are associated with flagellar function and chemotaxis, type VII secretion, signaling, and interaction with the host during infection. Ongoing efforts strive to track the mobility, oligomerization state, and interaction partners of flotillins in living cells.

Bacillus subtilis FloA and FloT are oligomeric, integral membrane proteins implicated in the formation of FMMs, and regulated by σW. FloA and FloT are thought to help partition the membrane into low fluidity FMM regions that are spatially distinct from more fluid regions. A direct role for flotillins in FMM formation has been challenged, however, since B. subtilis FloA and FloT do not always colocalize, and form separated foci of ∼100 nm in diameter that appear spatially distinct from FMMs. Counter-intuitively, flotillins appear to be required for regions of increased fluidity (RIFs), which are the counterpart to the FMMs. A lack of flotillins leads to a decrease in membrane fluidity and a concomitant reduction in activity of the MreB-directed elongasome complex that synthesizes peptidoglycan. This loss of membrane fluidity can be chemically complemented with fluidizing agents such as benzoyl alcohol.

Flotillins also functionally interact with DynA, a constitutively expressed dynamin homolog. Dynamins are membrane-associated GTPases implicated in membrane remodeling, fusion and fission, and lipid mixing. DynA may help repair damaged membrane regions, and contribute to resistance against antibiotics that bind membrane components, including nisin, bacitracin, and daptomycin. Our understanding of flotillins and dynamins, and their roles in bacterial physiology is still incomplete and rapidly evolving.

Phage-Shock Proteins Protect Membrane Integrity

Cell envelope stress responses also support membrane stability through induction of PspA proteins, including two paralogs in B. subtilis: PspA an LiaH. Originally defined as part of the phage-shock protein response in Escherichia coli, PspA proteins comprise a conserved family including the vesicle-inducing protein in plastids (VIPP1/IM30) and mammalian ESCRT III. PspA proteins have a conserved N-terminal amphipathic helix required for membrane binding, which seems to depend on anionic lipid content and regions with unfavorable packing geometries creating stored curvature elastic stress. Structural studies reveal that VIPP1 forms oligomeric rings of various symmetries that stack together to form domes. These rings are dynamic, and are thought to stabilize membranes during budding, tubulation, and fusion. However, the role of these oligomeric structures has been questioned. An alternative model suggests that these rings dissociate, and the resultant intrinsically disordered monomers interact with the membrane surface to form a protective protein “carpet” to stabilize the membrane and suppress proton leakage.

Although PspA proteins are assumed to function in membrane protection and repair, their regulation differs markedly. B. subtilis PspA is regulated by σW, whereas the paralog LiaH is regulated by the LiaRS TCS. Both paralogs localize to the membrane in response to stress and protect against membrane-damaging antibiotics. In the case of LiaH, membrane association is mediated by interaction with the integral membrane protein LiaI. LiaH may also protect the membrane against proton leakage during the export of proteins through the twin-arginine translocation (TAT) system. While the B. subtilis LiaRS regulon is rather limited in scope, LiaRS orthologs (e.g., S. aureus VraRS) play an important role in stress resistance in many Gram-positive pathogens, and mutations in these regulators are associated with clinical antibiotic resistance. In Mycobacterium tuberculosis, the PspA ortholog is also under control of the ECF σ factor σE, whereas E. coli pspA requires the σ54 RNAP and PspF activator. A common theme in these systems is that PspA-like proteins are often regulated by a specific CESR; they can accumulate to high levels in stressed cells, and they seem to protect the membrane against disruptions that can dissipate the proton gradient.

Antibiotic Specific Detoxification Modules

Bacillus subtilis, like many soil bacteria, can synthesize a wide range of antimicrobial compounds and also encodes diverse resistance mechanisms. Many antimicrobial peptides induce the B. subtilis LiaRS stress response that protects cells through induction of LiaH. Induction of σW also leads to expression of the SppA membrane-localized protease and its regulatory protein SppI, which function to clear the membrane of embedded peptides to protect against lantibiotics. Other antimicrobial peptides induce specific detoxification machinery, often including ABC transporters that either export the peptide antibiotic or disassemble membrane-bound peptide complexes.

A prototype for such systems is the BceRS TCS, which regulates the bacitracin-specific induction of the BceAB ABC transporter. Bacitracin is a peptide antibiotic made by Bacillus spp. that inhibits cell wall synthesis by binding to undecaprenyl-PP. The BceAB system appears to act in disassembly of bacitracin complexes to confer resistance. In addition, BceAB interacts with the BceRS TCS to allow sensing of bacitracin. The BceRS-AB system provides a first line of defense against bacitracin, with higher levels of antibiotic activating the protective responses mediated by the LiaRS and σECF regulons. The detailed study of the B. subtilis bacitracin stress response has provided lessons relevant to the understanding of other antibiotic detoxification modules. Similar genetic modules, encoding both TCS and ABC transporter/sensors have been described for several other antimicrobial peptides. Since induction can be quite specific, these systems provide a basis for antibiotic-inducible gene expression systems.

Bacillus subtilis also encodes and responds to many other secondary metabolites that can induce membrane stress. For example, the toxic peptide YydF∗ is encoded by the yydFGHIJ operon, together with a radical-SAM epimerase (YydG), protease (YydH), and ABC transporter (YydIJ). Transposon insertions in the presumptive efflux pump lead to the upregulation of the LiaRS stress system. Subsequent studies revealed that YydF is post-translationally processed to convert two L-amino acids to D-amino acids. The resulting epipeptide, YydF∗, induces LiaRS-regulated LiaH and the FloT flotillin. The modified YydF∗ peptide kills B. subtilis cells by dissipating the membrane potential via membrane permeabilization. The associated concomitant decrease in membrane fluidity together with increased membrane permeabilization induces liaIH. YydF∗ peptides are likely synthesized by a variety of Gram-positive organisms including Enterococcus, Staphylococcus, and Streptococcus spp. as well as members of the human microbiome.

Bacillus subtilis also has CESRs induced by polyketide and polyene-type antimicrobials. For example, Streptomyces spp. produce linear polyketides (linearmycins) that depolarize the membrane. Linearmycins strongly activate the LnrJK TCS that regulates an ABC transporter, LnrLMN. This ABC transporter also provides resistance against other polyenes, including the anti-fungal amphotericin.

Outlook

Here we provide a brief overview of the diverse ways in which CESRs help modify and protect the membrane in response to environmental threats (Table 1). This is a rapidly evolving field, and the impact of membrane composition on cell physiology is still mysterious. We have much to learn about the synthesis and roles of minor lipids (sesquarterpenes, ether lipids, lysophospholipids). There is a growing need to reconcile current models of lipid rafts, and the role that isoprenoid lipids and flotillins play in their formation. The activities of the VIPP1/IM30/PspA family of proteins in membrane repair and protection, and in particular the specific role of different oligomeric states, are still debated. Finally, the mechanisms by which diverse CESRs sense membrane perturbations are largely unknown, although considerable progress has been made in the specific cases of the DesK sensor kinase, flux-sensing by peptide detoxification modules, and the lysozyme-mediated induction of the σV protein. The overall picture is of the cell membrane as a complex and adaptable assemblage of many different lipid and protein species that still has many secrets to reveal.

Author Contributions

Both authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

References 145

  1. 1.

    Abriata L. A. Albanesi D. Dal Peraro M. de Mendoza D. (2017 ). Signal sensing and transduction by histidine kinases as unveiled through studies on a temperature sensor. Acc. Chem. Res. 50 1359–1366.

    Not in our library10.1021/acs.accounts.6b00593

  2. 2.

    Albanesi D. Reh G. Guerin M. E. Schaeffer F. Debarbouille M. Buschiazzo A. (2013 ). Structural basis for feed-forward transcriptional regulation of membrane lipid homeostasis in Staphylococcus aureus. PLoS Pathog. 9:e1003108.

    Not in our library10.1371/journal.ppat.1003108

  3. 3.

    Altabe S. G. Aguilar P. Caballero G. M. de Mendoza D. (2003 ). The Bacillus subtilis acyl lipid desaturase is a delta5 desaturase. J. Bacteriol. 185 3228–3231.

    Not in our library10.1128/jb.185.10.3228-3231.2003

  4. 4.

    Bach J. N. Bramkamp M. (2013 ). Flotillins functionally organize the bacterial membrane. Mol. Microbiol. 88 1205–1217.

    Not in our library10.1111/mmi.12252

  5. 5.

    Belin B. J. Busset N. Giraud E. Molinaro A. Silipo A. Newman D. K. (2018 ). Hopanoid lipids: from membranes to plant-bacteria interactions. Nat. Rev. Microbiol. 16 304–315.

    Not in our library10.1038/nrmicro.2017.173

  6. 6.

    Bell S. A. Chappell J. (2014 ). Now playing: farnesol in the biofilm. Chem. Biol. 21 1421–1422.

    Not in our library10.1016/j.chembiol.2014.11.001

  7. 7.

    Benjdia A. Guillot A. Ruffie P. Leprince J. Berteau O. (2017 ). Post-translational modification of ribosomally synthesized peptides by a radical SAM epimerase in Bacillus subtilis. Nat. Chem. 9 698–707.

    Not in our library10.1038/nchem.2714

  8. 8.

    Beranova J. Jemiola-Rzeminska M. Elhottova D. Strzalka K. Konopasek I. (2008 ). Metabolic control of the membrane fluidity in Bacillus subtilis during cold adaptation. Biochim. Biophys. Acta 1778 445–453.

    Not in our library10.1016/j.bbamem.2007.11.012

  9. 9.

    Bernal-Cabas M. Miethke M. Antelo-Varela M. Aguilar Suarez R. Neef J. Schon L. (2020 ). Functional association of the stress-responsive LiaH protein and the minimal TatAyCy protein translocase in Bacillus subtilis. Biochim. Biophys. Acta Mol. Cell Res. 1867:118719.

    Not in our library10.1016/j.bbamcr.2020.118719

  10. 10.

    Bosak T. Losick R. M. Pearson A. (2008 ). A polycyclic terpenoid that alleviates oxidative stress. Proc. Natl. Acad. Sci. U.S.A. 105 6725–6729.

    Not in our library10.1073/pnas.0800199105

  11. 11.

    Bramkamp M. Lopez D. (2015 ). Exploring the existence of lipid rafts in bacteria. Microbiol. Mol. Biol. Rev. 79 81–100.

    Not in our library10.1128/mmbr.00036-14

  12. 12.

    Butcher B. G. Helmann J. D. (2006 ). Identification of Bacillus subtilis σW-dependent genes that provide intrinsic resistance to antimicrobial compounds produced by Bacilli. Mol. Microbiol. 60 765–782.

    Not in our library10.1111/j.1365-2958.2006.05131.x

  13. 13.

    Butcher B. G. Lin Y. P. Helmann J. D. (2007 ). The yydFGHIJ operon of Bacillus subtilis encodes a peptide that induces the LiaRS two-component system. J. Bacteriol. 189 8616–8625.

    Not in our library10.1128/jb.01181-07

  14. 14.

    Cao M. Helmann J. D. (2004 ). The Bacillus subtilis extracytoplasmic-function σX factor regulates modification of the cell envelope and resistance to cationic antimicrobial peptides. J. Bacteriol. 186 1136–1146.

    Not in our library10.1128/jb.186.4.1136-1146.2004

  15. 15.

    Cao M. Kobel P. A. Morshedi M. M. Wu M. F. Paddon C. Helmann J. D. (2002a ). Defining the Bacillus subtilis σW regulon: a comparative analysis of promoter consensus search, run-off transcription/macroarray analysis (ROMA), and transcriptional profiling approaches. J. Mol. Biol. 316 443–457.

    Not in our library10.1006/jmbi.2001.5372

  16. 16.

    Cao M. Wang T. Ye R. Helmann J. D. (2002b ). Antibiotics that inhibit cell wall biosynthesis induce expression of the Bacillus subtilis σW and σM regulons. Mol. Microbiol. 45 1267–1276.

    Not in our library10.1046/j.1365-2958.2002.03050.x

  17. 17.
    Overview of the Antimicrobial Compounds Produced by Members of the Bacillus subtilis Group · 2019
    No notice
  18. 18.

    Choi K. H. Heath R. J. Rock C. O. (2000 ). beta-ketoacyl-acyl carrier protein synthase III (FabH) is a determining factor in branched-chain fatty acid biosynthesis. J. Bacteriol. 182 365–370.

    Not in our library10.1128/jb.182.2.365-370.2000

  19. 19.

    Chwastek G. Surma M. A. Rizk S. Grosser D. Lavrynenko O. Rucinska M. (2020 ). Principles of membrane adaptation revealed through environmentally induced bacterial lipidome remodeling. Cell Rep. 32:108165.

    Not in our library10.1016/j.celrep.2020.108165

  20. 20.

    Datta P. Ravi J. Guerrini V. Chauhan R. Neiditch M. B. Shell S. S. (2015 ). The Psp system of Mycobacterium tuberculosis integrates envelope stress-sensing and envelope-preserving functions. Mol. Microbiol. 97 408–422.

    Not in our library10.1111/mmi.13037

  21. 21.

    de Mendoza D. (2014 ). Temperature sensing by membranes. Annu. Rev. Microbiol. 68 101–116.

    Not in our library10.1146/annurev-micro-091313-103612

  22. 22.

    Dempwolff F. Graumann P. L. (2014 ). Genetic links between bacterial dynamin and flotillin proteins. Commun. Integr. Biol. 7:e970972.

    Not in our library10.4161/cib.29578

  23. 23.

    Dempwolff F. Schmidt F. K. Hervas A. B. Stroh A. Rosch T. C. Riese C. N. (2016 ). Super resolution fluorescence microscopy and tracking of bacterial flotillin (Reggie) paralogs provide evidence for defined-sized protein microdomains within the bacterial membrane but absence of clusters containing detergent-resistant proteins. PLoS Genet. 12:e1006116.

    Not in our library10.1371/journal.pgen.1006116

  24. 24.

    Dempwolff F. Wischhusen H. M. Specht M. Graumann P. L. (2012 ). The deletion of bacterial dynamin and flotillin genes results in pleiotrophic effects on cell division, cell growth and in cell shape maintenance. BMC Microbiol. 12:298.

    Not in our library10.1186/1471-2180-12-298

  25. 25.

    Devkota S. R. Kwon E. Ha S. C. Chang H. W. Kim D. Y. (2017 ). Structural insights into the regulation of Bacillus subtilis σW activity by anti-sigma RsiW. PLoS One 12:e0174284.

    Not in our library10.1371/journal.pone.0174284

  26. 26.

    Dintner S. Heermann R. Fang C. Jung K. Gebhard S. (2014 ). A sensory complex consisting of an ATP-binding cassette transporter and a two-component regulatory system controls bacitracin resistance in Bacillus subtilis. J. Biol. Chem. 289 27899–27910.

    Not in our library10.1074/jbc.m114.596221

  27. 27.

    Dominguez-Escobar J. Wolf D. Fritz G. Hofler C. Wedlich-Soldner R. Mascher T. (2014 ). Subcellular localization, interactions and dynamics of the phage-shock protein-like Lia response in Bacillus subtilis. Mol. Microbiol. 92 716–732.

    Not in our library10.1111/mmi.12586

  28. 28.

    Dowhan W. Vitrac H. Bogdanov M. (2019 ). Lipid-assisted membrane protein folding and topogenesis. Protein J. 38 274–288.

    Not in our library10.1007/s10930-019-09826-7

  29. 29.

    Eiamphungporn W. Helmann J. D. (2008 ). The Bacillus subtilis σM regulon and its contribution to cell envelope stress responses. Mol. Microbiol. 67 830–848.

    Not in our library10.1111/j.1365-2958.2007.06090.x

  30. 30.

    Ellermeier C. D. Losick R. (2006 ). Evidence for a novel protease governing regulated intramembrane proteolysis and resistance to antimicrobial peptides in Bacillus subtilis. Genes Dev. 20 1911–1922.

    Not in our library10.1101/gad.1440606

  31. 31.

    Ernst C. M. Peschel A. (2019 ). MprF-mediated daptomycin resistance. Int. J. Med. Microbiol. 309 359–363.

    Not in our library10.1016/j.ijmm.2019.05.010

  32. 32.

    Ernst C. M. Slavetinsky C. J. Kuhn S. Hauser J. N. Nega M. Mishra N. N. (2018 ). Gain-of-function mutations in the phospholipid flippase MprF confer specific daptomycin resistance. mBio 9:e01659-18.

    Not in our library10.1128/mbio.01659-18

  33. 33.

    Ernst R. Ejsing C. S. Antonny B. (2016 ). Homeoviscous adaptation and the regulation of membrane lipids. J. Mol. Biol. 428 4776–4791.

    Not in our library10.1016/j.jmb.2016.08.013

  34. 34.
    Investigation of the Staphylococcus aureus GraSR Regulon Reveals Novel Links to Virulence, Stress Response and Cell Wall Signal Transduction Pathways · 2011
    No notice
  35. 35.

    Feng X. Hu Y. Zheng Y. Zhu W. Li K. Huang C. H. (2014 ). Structural and functional analysis of Bacillus subtilis YisP reveals a role of its product in biofilm production. Chem. Biol. 21 1557–1563.

    Not in our library10.1016/j.chembiol.2014.08.018

  36. 36.

    Fernandez P. Porrini L. Albanesi D. Abriata L. A. Dal Peraro M. de Mendoza D. (2019 ). Transmembrane prolines mediate signal sensing and decoding in Bacillus subtilis DesK histidine kinase. mBio 10:e02564-19.

    Not in our library10.1128/mbio.02564-19

  37. 37.

    Flores-Kim J. Darwin A. J. (2016 ). The phage shock protein response. Annu. Rev. Microbiol. 70 83–101.

    Not in our library10.1146/annurev-micro-102215-095359

  38. 38.

    Foster T. J. (2019 ). Can beta-lactam antibiotics be resurrected to combat MRSA? Trends Microbiol. 27 26–38.

    Not in our library10.1016/j.tim.2018.06.005

  39. 39.

    Fritz G. Dintner S. Treichel N. S. Radeck J. Gerland U. Mascher T. (2015 ). A New way of sensing: need-based activation of antibiotic resistance by a flux-sensing mechanism. mBio 6:e00975.

    Not in our library10.1128/mbio.00975-15

  40. 40.

    Fujita Y. Matsuoka H. Hirooka K. (2007 ). Regulation of fatty acid metabolism in bacteria. Mol. Microbiol. 66 829–839.

    Not in our library10.1111/j.1365-2958.2007.05947.x

  41. 41.

    Garcia-Fernandez E. Koch G. Wagner R. M. Fekete A. Stengel S. T. Schneider J. (2017 ). Membrane microdomain disassembly inhibits MRSA antibiotic resistance. Cell 171 1354–1367.e1320.

    Not in our library10.1016/j.cell.2017.10.012

  42. 42.

    Guariglia-Oropeza V. Helmann J. D. (2011 ). Bacillus subtilis σV confers lysozyme resistance by activation of two cell wall modification pathways, peptidoglycan O-acetylation and D-alanylation of teichoic acids. J. Bacteriol. 193 6223–6232.

    Not in our library10.1128/jb.06023-11

  43. 43.

    Guo L. Bramkamp M. (2019 ). Bacterial dynamin-like protein DynA mediates lipid and content mixing. FASEB J. 33 11746–11757.

    Not in our library10.1096/fj.201900844rr

  44. 44.

    Gupta T. K. Klumpe S. Gries K. Heinz S. Wietrzynski W. Ohnishi N. (2020 ). Structural basis for VIPP1 oligomerization and maintenance of thylakoid membrane integrity. BioRxiv [Preprint].

    Not in our library10.1101/2020.08.11.243204

  45. 45.

    Gutu A. Chang F. O’Shea E. K. (2018 ). Dynamical localization of a thylakoid membrane binding protein is required for acquisition of photosynthetic competency. Mol. Microbiol. 108 16–31.

    Not in our library10.1111/mmi.13912

  46. 46.

    Hachmann A. B. Angert E. R. Helmann J. D. (2009 ). Genetic analysis of factors affecting susceptibility of Bacillus subtilis to daptomycin. Antimicrob. Agents Chemother. 53 1598–1609.

    Not in our library10.1128/aac.01329-08

  47. 47.

    Hachmann A. B. Sevim E. Gaballa A. Popham D. L. Antelmann H. Helmann J. D. (2011 ). Reduction in membrane phosphatidylglycerol content leads to daptomycin resistance in Bacillus subtilis. Antimicrob. Agents Chemother. 55 4326–4337.

    Not in our library10.1128/aac.01819-10

  48. 48.

    Hashimoto M. Seki T. Matsuoka S. Hara H. Asai K. Sadaie Y. (2013 ). Induction of extracytoplasmic function sigma factors in Bacillus subtilis cells with defects in lipoteichoic acid synthesis. Microbiology (Reading) 159 23–35.

    Not in our library10.1099/mic.0.063420-0

  49. 49.

    Hashimoto M. Takahashi H. Hara Y. Hara H. Asai K. Sadaie Y. (2009 ). Induction of extracytoplasmic function sigma factors in Bacillus subtilis cells with membranes of reduced phosphatidylglycerol content. Genes Genet. Syst. 84 191–198.

    Not in our library10.1266/ggs.84.191

  50. 50.

    Helmann J. D. (2006 ). Deciphering a complex genetic regulatory network: the Bacillus subtilis σW protein and intrinsic resistance to antimicrobial compounds. Sci. Prog. 89 243–266.

    Not in our library10.3184/003685006783238290

  51. 51.

    Helmann J. D. (2016 ). Bacillus subtilis extracytoplasmic function (ECF) sigma factors and defense of the cell envelope. Curr. Opin. Microbiol. 30 122–132.

    Not in our library10.1016/j.mib.2016.02.002

  52. 52.

    Henriques G. McGovern S. Neef J. Antelo-Varela M. Gotz F. Otto A. (2020 ). SppI forms a membrane protein complex with sppa and inhibits its protease activity in Bacillus subtilis. mSphere 5:e00724-20.

    Not in our library10.1128/msphere.00724-20

  53. 53.

    Ho T. D. Ellermeier C. D. (2019 ). Activation of the extracytoplasmic function sigma factor σV by lysozyme. Mol. Microbiol. 112 410–419.

    Not in our library10.1111/mmi.14348

  54. 54.

    Ho T. D. Hastie J. L. Intile P. J. Ellermeier C. D. (2011 ). The Bacillus subtilis extracytoplasmic function sigma factor σV is induced by lysozyme and provides resistance to lysozyme. J. Bacteriol. 193 6215–6222.

    Not in our library10.1128/jb.05467-11

  55. 55.

    Hofler C. Heckmann J. Fritsch A. Popp P. Gebhard S. Fritz G. (2016 ). Cannibalism stress response in Bacillus subtilis. Microbiology (Reading) 162 164–176.

    Not in our library10.1099/mic.0.000176

  56. 56.

    Hou B. Heidrich E. S. Mehner-Breitfeld D. Bruser T. (2018 ). The TatA component of the twin-arginine translocation system locally weakens the cytoplasmic membrane of Escherichia coli upon protein substrate binding. J. Biol. Chem. 293 7592–7605.

    Not in our library10.1074/jbc.ra118.002205

  57. 57.

    Huang X. Gaballa A. Cao M. Helmann J. D. (1999 ). Identification of target promoters for the Bacillus subtilis extracytoplasmic function sigma factor, σW. Mol. Microbiol. 31 361–371.

    Not in our library10.1046/j.1365-2958.1999.01180.x

  58. 58.

    Hutton M. L. D’Costa K. Rossiter A. E. Wang L. Turner L. Steer D. L. (2017 ). A Helicobacter pylori homolog of eukaryotic flotillin is involved in cholesterol accumulation, epithelial cell responses and host colonization. Front. Cell Infect. Microbiol. 7:219.

    Not in our library10.3389/fcimb.2017.00219

  59. 59.

    Jolliffe L. K. Doyle R. J. Streips U. N. (1981 ). The energized membrane and cellular autolysis in Bacillus subtilis. Cell 25 753–763.

    Not in our library10.1016/0092-8674(81)90183-5

  60. 60.

    Joly N. Engl C. Jovanovic G. Huvet M. Toni T. Sheng X. (2010 ). Managing membrane stress: the phage shock protein (Psp) response, from molecular mechanisms to physiology. FEMS Microbiol. Rev. 34 797–827.

    Not in our library10.1111/j.1574-6976.2010.00240.x

  61. 61.

    Jordan S. Junker A. Helmann J. D. Mascher T. (2006 ). Regulation of LiaRS-dependent gene expression in Bacillus subtilis: identification of inhibitor proteins, regulator binding sites, and target genes of a conserved cell envelope stress-sensing two-component system. J. Bacteriol. 188 5153–5166.

    Not in our library10.1128/jb.00310-06

  62. 62.

    Junglas B. Schneider D. (2018 ). What is Vipp1 good for? Mol. Microbiol. 108 1–5.

    Not in our library10.1111/mmi.13924

  63. 63.

    Junglas B. Orru R. Axt A. Siebenaller C. Steinchen W. Heidrich J. (2020 ). IM30 IDPs form a membrane-protective carpet upon super-complex disassembly. Commun. Biol. 3:595.

    Not in our library10.1038/s42003-020-01314-4

  64. 64.

    Kingston A. W. Liao X. Helmann J. D. (2013 ). Contributions of the σW, σM and σX regulons to the lantibiotic resistome of Bacillus subtilis. Mol. Microbiol. 90 502–518.

    Not in our library10.1111/mmi.12380

  65. 65.

    Kingston A. W. Subramanian C. Rock C. O. Helmann J. D. (2011 ). A σW-dependent stress response in Bacillus subtilis that reduces membrane fluidity. Mol. Microbiol. 81 69–79.

    Not in our library10.1111/j.1365-2958.2011.07679.x

  66. 66.

    Kingston A. W. Zhao H. Cook G. M. Helmann J. D. (2014 ). Accumulation of heptaprenyl diphosphate sensitizes Bacillus subtilis to bacitracin: implications for the mechanism of resistance mediated by the BceAB transporter. Mol. Microbiol. 93 37–49.

    Not in our library10.1111/mmi.12637

  67. 67.

    Kobayashi R. Suzuki T. Yoshida M. (2007 ). Escherichia coli phage-shock protein A (PspA) binds to membrane phospholipids and repairs proton leakage of the damaged membranes. Mol. Microbiol. 66 100–109.

    Not in our library10.1111/j.1365-2958.2007.05893.x

  68. 68.

    Kobras C. M. Piepenbreier H. Emenegger J. Sim A. Fritz G. Gebhard S. (2020 ). BceAB-type antibiotic resistance transporters appear to act by target protection of cell wall synthesis. Antimicrob. Agents Chemother. 64:e02241-19.

    Not in our library10.1128/aac.02241-19

  69. 69.

    Koh A. Gibbon M. J. Van der Kamp M. W. Pudney C. R. Gebhard S. (2020 ). Conformation control of the histidine kinase BceS of Bacillus subtilis by its cognate ABC-transporter facilitates need-based activation of antibiotic resistance. Mol. Microbiol. 115 157–174.

    Not in our library10.1111/mmi.14607

  70. 70.

    Lamsa A. Liu W. T. Dorrestein P. C. Pogliano K. (2012 ). The Bacillus subtilis cannibalism toxin SDP collapses the proton motive force and induces autolysis. Mol. Microbiol. 84 486–500.

    Not in our library10.1111/j.1365-2958.2012.08038.x

  71. 71.

    Laut C. L. Perry W. J. Metzger A. L. Weiss A. Stauff D. L. Walker S. (2020 ). Bacillus anthracis responds to targocil-induced envelope damage through EdsRS activation of cardiolipin synthesis. mBio 11:e03375-19.

    Not in our library10.1128/mbio.03375-19

  72. 72.

    Linde M. Peterhoff D. Sterner R. Babinger P. (2016 ). Identification and characterization of heptaprenylglyceryl phosphate processing enzymes in Bacillus subtilis. J. Biol. Chem. 291 14861–14870.

    Not in our library10.1074/jbc.m115.711994

  73. 73.

    Liu J. Tassinari M. Souza D. P. Naskar S. Noel J. K. Bohuszewicz O. (2020 ). Bacterial Vipp1 and PspA are members of the ancient ESCRT-III membrane-remodelling superfamily. bioRxiv [Preprint].

    Not in our library10.1101/2020.08.13.249979

  74. 74.

    Lopez D. Koch G. (2017 ). Exploring functional membrane microdomains in bacteria: an overview. Curr. Opin. Microbiol. 36 76–84.

    Not in our library10.1016/j.mib.2017.02.001

  75. 75.

    Lopez D. Kolter R. (2010 ). Functional microdomains in bacterial membranes. Genes Dev. 24 1893–1902.

    Not in our library10.1101/gad.1945010

  76. 76.

    Machinandiarena F. Nakamatsu L. Schujman G. E. de Mendoza D. Albanesi D. (2020 ). Revisiting the coupling of fatty acid to phospholipid synthesis in bacteria with FapR regulation. Mol. Microbiol. 114:14574.

    Not in our library10.1111/mmi.14574

  77. 77.

    Manganelli R. Gennaro M. L. (2017 ). Protecting from envelope stress: variations on the phage-shock-protein theme. Trends Microbiol. 25 205–216.

    Not in our library10.1016/j.tim.2016.10.001

  78. 78.

    Martinez M. A. Zaballa M. E. Schaeffer F. Bellinzoni M. Albanesi D. Schujman G. E. (2010 ). A novel role of malonyl-ACP in lipid homeostasis. Biochemistry 49 3161–3167.

    Not in our library10.1021/bi100136n

  79. 79.

    Mascher T. Hachmann A. B. Helmann J. D. (2007 ). Regulatory overlap and functional redundancy among Bacillus subtilis extracytoplasmic function sigma factors. J. Bacteriol. 189 6919–6927.

    Not in our library10.1128/jb.00904-07

  80. 80.

    Mascher T. Zimmer S. L. Smith T. A. Helmann J. D. (2004 ). Antibiotic-inducible promoter regulated by the cell envelope stress-sensing two-component system LiaRS of Bacillus subtilis. Antimicrob. Agents Chemother. 48 2888–2896.

    Not in our library10.1128/aac.48.8.2888-2896.2004

  81. 81.

    Matsuoka S. (2018 ). Biological functions of glucolipids in Bacillus subtilis. Genes Genet. Syst. 92 217–221.

    Not in our library10.1266/ggs.17-00017

  82. 82.

    Matsuoka S. Seki T. Matsumoto K. Hara H. (2016 ). Suppression of abnormal morphology and extracytoplasmic function sigma activity in Bacillus subtilis ugtP mutant cells by expression of heterologous glucolipid synthases from Acholeplasma laidlawii. Biosci. Biotechnol. Biochem. 80 2325–2333.

    Not in our library10.1080/09168451.2016.1217147

  83. 83.

    McDonald C. Jovanovic G. Ces O. Buck M. (2015 ). Membrane stored curvature elastic stress modulates recruitment of maintenance proteins PspA and Vipp1. mBio 6:e001188-15.

    Not in our library10.1128/mbio.01188-15

  84. 84.

    McDonald C. Jovanovic G. Wallace B. A. Ces O. Buck M. (2017 ). Structure and function of PspA and Vipp1 N-terminal peptides: insights into the membrane stress sensing and mitigation. Biochim. Biophys. Acta Biomembr. 1859 28–39.

    Not in our library10.1016/j.bbamem.2016.10.018

  85. 85.

    Mielich-Suss B. Wagner R. M. Mietrach N. Hertlein T. Marincola G. Ohlsen K. (2017 ). Flotillin scaffold activity contributes to type VII secretion system assembly in Staphylococcus aureus. PLoS Pathog. 13:e1006728.

    Not in our library10.1371/journal.ppat.1006728

  86. 86.

    Mitchell A. M. Silhavy T. J. (2019 ). Envelope stress responses: balancing damage repair and toxicity. Nat. Rev. Microbiol. 17 417–428.

    Not in our library10.1038/s41579-019-0199-0

  87. 87.

    Muller M. Reiss S. Schluter R. Mader U. Beyer A. Reiss W. (2014 ). Deletion of membrane-associated Asp23 leads to upregulation of cell wall stress genes in Staphylococcus aureus. Mol. Microbiol. 93 1259–1268.

    Not in our library10.1111/mmi.12733

  88. 88.

    Nickels J. D. Chatterjee S. Mostofian B. Stanley C. B. Ohl M. Zolnierczuk P. (2017 ). Bacillus subtilis lipid extract, a branched-chain fatty acid model membrane. J. Phys. Chem. Lett. 8 4214–4217.

    Not in our library10.1021/acs.jpclett.7b01877

  89. 89.

    Nickels J. D. Poudel S. Chatterjee S. Farmer A. Cordner D. Campagna S. R. (2020 ). Impact of fatty-acid labeling of Bacillus subtilis membranes on the cellular lipidome and proteome. Front. Microbiol. 11:914.

    Not in our library10.3389/fmicb.2020.00914

  90. 90.

    Ohki R. Giyanto K. T. Masuyama W. Moriya S. Kobayashi K. Ogasawara N. (2003 ). The BceRS two-component regulatory system induces expression of the bacitracin transporter, BceAB, in Bacillus subtilis. Mol. Microbiol. 49 1135–1144.

    Not in our library10.1046/j.1365-2958.2003.03653.x

  91. 91.

    Padilla-Vaca F. Vargas-Maya N. I. Elizarraras-Vargas N. U. Rangel-Serrano A. Cardoso-Reyes L. R. Razo-Soria T. (2019 ). Flotillin homologue is involved in the swimming behavior of Escherichia coli. Arch. Microbiol. 201 999–1008.

    Not in our library10.1007/s00203-019-01670-8

  92. 92.

    Pan J. J. Solbiati J. O. Ramamoorthy G. Hillerich B. S. Seidel R. D. Cronan J. E. (2015 ). Biosynthesis of squalene from farnesyl diphosphate in bacteria: three steps catalyzed by three enzymes. ACS Cent. Sci. 1 77–82.

    Not in our library10.1021/acscentsci.5b00115

  93. 93.

    Parsons J. B. Rock C. O. (2013 ). Bacterial lipids: metabolism and membrane homeostasis. Prog. Lipid Res. 52 249–276.

    Not in our library10.1016/j.plipres.2013.02.002

  94. 94.
    Whole Genome Characterization of the Mechanisms of Daptomycin Resistance in Clinical and Laboratory Derived Isolates of Staphylococcus aureus · 2012
    No notice
  95. 95.

    Petersen I. Schluter R. Hoff K. J. Liebscher V. Bange G. Riedel K. (2020 ). Non-invasive and label-free 3D-visualization shows in vivo oligomerization of the staphylococcal alkaline shock protein 23 (Asp23). Sci. Rep. 10:125.

    Not in our library10.1038/s41598-019-56907-9

  96. 96.
    From Modules to Networks: a Systems-Level Analysis of the Bacitracin Stress Response in <named-content content-type="genus-species">Bacillus subtilis</named-content> · 2020
    Not checked
  97. 97.

    Pietiainen M. Gardemeister M. Mecklin M. Leskela S. Sarvas M. Kontinen V. P. (2005 ). Cationic antimicrobial peptides elicit a complex stress response in Bacillus subtilis that involves ECF-type sigma factors and two-component signal transduction systems. Microbiology (Reading) 151 1577–1592.

    Not in our library10.1099/mic.0.27761-0

  98. 98.

    Popp P. F. Benjdia A. Strahl H. Berteau O. Mascher T. (2020 ). The epipeptide YydF intrinsically triggers the cell envelope stress response of bacillus subtilis and causes severe membrane perturbations. Front. Microbiol. 11:151.

    Not in our library10.3389/fmicb.2020.00151

  99. 99.

    Radeck J. Fritz G. Mascher T. (2017 ). The cell envelope stress response of Bacillus subtilis: from static signaling devices to dynamic regulatory network. Curr. Genet. 63 79–90.

    Not in our library10.1007/s00294-016-0624-0

  100. 100.

    Radeck J. Gebhard S. Orchard P. S. Kirchner M. Bauer S. Mascher T. (2016 ). Anatomy of the bacitracin resistance network in Bacillus subtilis. Mol. Microbiol. 100 607–620.

    Not in our library10.1111/mmi.13336

  101. 101.

    Rajagopal M. Walker S. (2017 ). Envelope structures of gram-positive bacteria. Curr. Top. Microbiol. Immunol. 404 1–44.

    Not in our library10.1007/82_2015_5021

  102. 102.
    Amphotericin B Specifically Induces the Two-Component System LnrJK: Development of a Novel Whole-Cell Biosensor for the Detection of Amphotericin-Like Polyenes · 2020
    Not checked
  103. 103.

    Revilla-Guarinos A. Gebhard S. Mascher T. Zuniga M. (2014 ). Defence against antimicrobial peptides: different strategies in Firmicutes. Environ. Microbiol. 16 1225–1237.

    Not in our library10.1111/1462-2920.12400

  104. 104.

    Saenz J. P. Grosser D. Bradley A. S. Lagny T. J. Lavrynenko O. Broda M. (2015 ). Hopanoids as functional analogues of cholesterol in bacterial membranes. Proc. Natl. Acad. Sci. U.S.A. 112 11971–11976.

    Not in our library10.1073/pnas.1515607112

  105. 105.

    Salie M. J. Thelen J. J. (2016 ). Regulation and structure of the heteromeric acetyl-CoA carboxylase. Biochim. Biophys. Acta 1861 1207–1213.

    Not in our library10.1016/j.bbalip.2016.04.004

  106. 106.

    Salzberg L. I. Helmann J. D. (2008 ). Phenotypic and transcriptomic characterization of Bacillus subtilis mutants with grossly altered membrane composition. J. Bacteriol. 190 7797–7807.

    Not in our library10.1128/jb.00720-08

  107. 107.

    Sato T. (2013 ). Unique biosynthesis of sesquarterpenes (C35 terpenes). Biosci. Biotechnol. Biochem. 77 1155–1159.

    Not in our library10.1271/bbb.130180

  108. 108.

    Sato T. Yoshida S. Hoshino H. Tanno M. Nakajima M. Hoshino T. (2011 ). Sesquarterpenes (C35 terpenes) biosynthesized via the cyclization of a linear C35 isoprenoid by a tetraprenyl-beta-curcumene synthase and a tetraprenyl-beta-curcumene cyclase: identification of a new terpene cyclase. J. Am. Chem. Soc. 133 9734–9737.

    Not in our library10.1021/ja203779h

  109. 109.

    Saunders L. P. Sen S. Wilkinson B. J. Gatto C. (2016 ). Insights into the mechanism of homeoviscous adaptation to low temperature in branched-chain fatty acid-containing bacteria through modeling FabH kinetics from the foodborne pathogen Listeria monocytogenes. Front. Microbiol. 7:1386.

    Not in our library10.3389/fmicb.2016.01386

  110. 110.

    Saur M. Hennig R. Young P. Rusitzka K. Hellmann N. Heidrich J. (2017 ). A Janus-faced IM30 ring involved in thylakoid membrane fusion is assembled from IM30 tetramers. Structure 25 1380–1390.e1385.

    Not in our library10.1016/j.str.2017.07.001

  111. 111.

    Sawant P. Eissenberger K. Karier L. Mascher T. Bramkamp M. (2016 ). A dynamin-like protein involved in bacterial cell membrane surveillance under environmental stress. Environ. Microbiol. 18 2705–2720.

    Not in our library10.1111/1462-2920.13110

  112. 112.
    Identification of L-Valine-initiated-germination-active genes in Bacillus subtilis using Tn-seq. · 2019
    Not checked
  113. 113.

    Schobel S. Zellmeier S. Schumann W. Wiegert T. (2004 ). The Bacillus subtilis σW anti-sigma factor RsiW is degraded by intramembrane proteolysis through YluC. Mol. Microbiol. 52 1091–1105.

    Not in our library10.1111/j.1365-2958.2004.04031.x

  114. 114.

    Scholz R. Vater J. Budiharjo A. Wang Z. He Y. Dietel K. (2014 ). Amylocyclicin, a novel circular bacteriocin produced by Bacillus amyloliquefaciens FZB42. J. Bacteriol. 196 1842–1852.

    Not in our library10.1128/jb.01474-14

  115. 115.

    Schujman G. E. Guerin M. Buschiazzo A. Schaeffer F. Llarrull L. I. Reh G. (2006 ). Structural basis of lipid biosynthesis regulation in Gram-positive bacteria. EMBO J. 25 4074–4083.

    Not in our library10.1038/sj.emboj.7601284

  116. 116.

    Schujman G. E. Paoletti L. Grossman A. D. de Mendoza D. (2003 ). FapR, a bacterial transcription factor involved in global regulation of membrane lipid biosynthesis. Dev. Cell 4 663–672.

    Not in our library10.1016/s1534-5807(03)00123-0

  117. 117.

    Seki T. Furumi T. Hashimoto M. Hara H. Matsuoka S. (2019 ). Activation of extracytoplasmic function sigma factors upon removal of glucolipids and reduction of phosphatidylglycerol content in Bacillus subtilis cells lacking lipoteichoic acid. Genes Genet. Syst. 94 71–80.

    Not in our library10.1266/ggs.18-00046

  118. 118.

    Siebenaller C. Junglas B. Schneider D. (2019 ). Functional implications of multiple IM30 oligomeric states. Front. Plant Sci. 10:1500.

    Not in our library10.3389/fpls.2019.01500

  119. 119.

    Sohlenkamp C. Geiger O. (2016 ). Bacterial membrane lipids: diversity in structures and pathways. FEMS Microbiol. Rev. 40 133–159.

    Not in our library10.1093/femsre/fuv008

  120. 120.

    Staron A. Finkeisen D. E. Mascher T. (2011 ). Peptide antibiotic sensing and detoxification modules of Bacillus subtilis. Antimicrob. Agents Chemother. 55 515–525.

    Not in our library10.1128/aac.00352-10

  121. 121.

    Stein T. (2005 ). Bacillus subtilis antibiotics: structures, syntheses and specific functions. Mol. Microbiol. 56 845–857.

    Not in our library10.1111/j.1365-2958.2005.04587.x

  122. 122.

    Stubbendieck R. M. Straight P. D. (2015 ). Escape from lethal bacterial competition through coupled activation of antibiotic resistance and a mobilized subpopulation. PLoS Genet. 11:e1005722.

    Not in our library10.1371/journal.pgen.1005722

  123. 123.

    Stubbendieck R. M. Straight P. D. (2017 ). Linearmycins activate a two-component signaling system involved in bacterial competition and biofilm morphology. J. Bacteriol. 199:JB.00186-17.

    Not in our library10.1128/jb.00186-17

  124. 124.

    Stubbendieck R. M. Brock D. J. Pellois J. P. Gill J. J. Straight P. D. (2018 ). Linearmycins are lytic membrane-targeting antibiotics. J. Antibiot. (Tokyo) 71 372–381.

    Not in our library10.1038/s41429-017-0005-z

  125. 125.

    Sumrall E. T. Keller A. P. Shen Y. Loessner M. J. (2020 ). Structure and function of Listeria teichoic acids and their implications. Mol. Microbiol. 113 627–637.

    Not in our library10.1111/mmi.14472

  126. 126.

    Takekawa N. Isumi M. Terashima H. Zhu S. Nishino Y. Sakuma M. (2019 ). Structure of Vibrio FliL, a new stomatin-like protein that assists the bacterial flagellar motor function. mBio 10:e00292-19.

    Not in our library10.1128/mbio.00292-19

  127. 127.

    Takigawa H. Sugiyama M. Shibuya Y. (2010 ). C35-Terpenes from Bacillus subtilis KSM 6-10. J. Nat. Prod. 73 204–207.

    Not in our library10.1021/np900705q

  128. 128.

    Tamehiro N. Okamoto-Hosoya Y. Okamoto S. Ubukata M. Hamada M. Naganawa H. (2002 ). Bacilysocin, a novel phospholipid antibiotic produced by Bacillus subtilis 168. Antimicrob. Agents Chemother. 46 315–320.

    Not in our library10.1128/aac.46.2.315-320.2002

  129. 129.

    Thurotte A. Brüser T. Mascher T. Schneider D. (2017 ). Membrane chaperoning by members of the PspA/IM30 protein family. Commun. Integrat. Biol. 10:e1264546.

    Not in our library10.1080/19420889.19422016.11264546

  130. 130.
    The Highly Conserved Asp23 Family Protein YqhY Plays a Role in Lipid Biosynthesis in Bacillus subtilis · 2017
    Not checked
  131. 131.

    Tran T. T. Miller W. R. Shamoo Y. Arias C. A. (2016 ). Targeting cell membrane adaptation as a novel antimicrobial strategy. Curr. Opin. Microbiol. 33 91–96.

    Not in our library10.1016/j.mib.2016.07.002

  132. 132.

    van der Donk W. A. (2015 ). Bacteria do it differently: an alternative path to squalene. ACS Cent. Sci. 1 64–65.

    Not in our library10.1021/acscentsci.5b00142

  133. 133.

    Vinayavekhin N. Mahipant G. Vangnai A. S. Sangvanich P. (2015 ). Untargeted metabolomics analysis revealed changes in the composition of glycerolipids and phospholipids in Bacillus subtilis under 1-butanol stress. Appl. Microbiol. Biotechnol. 99 5971–5983.

    Not in our library10.1007/s00253-015-6692-0

  134. 134.

    Wagner R. M. Kricks L. Lopez D. (2017 ). Functional membrane microdomains organize signaling networks in bacteria. J. Membr. Biol. 250 367–378.

    Not in our library10.1007/s00232-016-9923-0

  135. 135.

    Weart R. B. Lee A. H. Chien A. C. Haeusser D. P. Hill N. S. Levin P. A. (2007 ). A metabolic sensor governing cell size in bacteria. Cell 130 335–347.

    Not in our library10.1016/j.cell.2007.05.043

  136. 136.

    Weber M. H. Klein W. Muller L. Niess U. M. Marahiel M. A. (2001 ). Role of the Bacillus subtilis fatty acid desaturase in membrane adaptation during cold shock. Mol. Microbiol. 39 1321–1329.

    Not in our library10.1111/j.1365-2958.2001.02322.x

  137. 137.

    Wiegert T. Homuth G. Versteeg S. Schumann W. (2001 ). Alkaline shock induces the Bacillus subtilis σW regulon. Mol. Microbiol. 41 59–71.

    Not in our library10.1046/j.1365-2958.2001.02489.x

  138. 138.

    Wolf D. Mascher T. (2016 ). The applied side of antimicrobial peptide-inducible promoters from Firmicutes bacteria: expression systems and whole-cell biosensors. Appl. Microbiol. Biotechnol. 100 4817–4829.

    Not in our library10.1007/s00253-016-7519-3

  139. 139.

    Wolf D. Kalamorz F. Wecke T. Juszczak A. Mader U. Homuth G. (2010 ). In-depth profiling of the LiaR response of Bacillus subtilis. J. Bacteriol. 192 4680–4693.

    Not in our library10.1128/jb.00543-10

  140. 140.

    Wormann M. E. Corrigan R. M. Simpson P. J. Matthews S. J. Grundling A. (2011 ). Enzymatic activities and functional interdependencies of Bacillus subtilis lipoteichoic acid synthesis enzymes. Mol. Microbiol. 79 566–583.

    Not in our library10.1111/j.1365-2958.2010.07472.x

  141. 141.

    Yamada Y. Tikhonova E. B. Zgurskaya H. I. (2012 ). YknWXYZ is an unusual four-component transporter with a role in protection against sporulation-delaying-protein-induced killing of Bacillus subtilis. J. Bacteriol. 194 4386–4394.

    Not in our library10.1128/jb.00223-12

  142. 142.

    Yang S. J. Bayer A. S. Mishra N. N. Meehl M. Ledala N. Yeaman M. R. (2012 ). The Staphylococcus aureus two-component regulatory system, GraRS, senses and confers resistance to selected cationic antimicrobial peptides. Infect. Immun. 80 74–81.

    Not in our library10.1128/iai.05669-11

  143. 143.

    Yao J. Rock C. O. (2013 ). Phosphatidic acid synthesis in bacteria. Biochim. Biophys. Acta 1831 495–502.

    Not in our library10.1016/j.bbalip.2012.08.018

  144. 144.

    Zhang Y. M. Rock C. O. (2009 ). Transcriptional regulation in bacterial membrane lipid synthesis. J. Lipid Res. 50(Suppl) S115–S119.

    Not in our library10.1194/jlr.r800046-jlr200

  145. 145.

    Zielinska A. Savietto A. de Sousa Borges A. Martinez D. Berbon M. Roelofsen J. R. (2020 ). Flotillin-mediated membrane fluidity controls peptidoglycan synthesis and MreB movement. Elife 9:e57179.

    Not in our library10.7554/elife.57179

Reads

Read by people on this page

None yet.

Checked or read by AI through naulon

None yet.

A person counts once a day per paper. We keep no cookie and no address: a reader is a hash that changes every day. Any month under five reads shows no number.

Are you an author?

Is this your paper?

Claim it to add your note and be paid when agents cite it. Free.