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J Bacteriol. Oct 2012; 194(19): 5294–5304.
PMCID: PMC3457211

Membrane Disruption by Antimicrobial Fatty Acids Releases Low-Molecular-Weight Proteins from Staphylococcus aureus


The skin represents an important barrier for pathogens and is known to produce fatty acids that are toxic toward Gram-positive bacteria. A screen of fatty acids as growth inhibitors of Staphylococcus aureus revealed structure-specific antibacterial activity. Fatty acids like oleate (18:1Δ9) were nontoxic, whereas palmitoleate (16:1Δ9) was a potent growth inhibitor. Cells treated with 16:1Δ9 exhibited rapid membrane depolarization, the disruption of all major branches of macromolecular synthesis, and the release of solutes and low-molecular-weight proteins into the medium. Other cytotoxic lipids, such as glycerol ethers, sphingosine, and acyl-amines blocked growth by the same mechanisms. Nontoxic 18:1Δ9 was used for phospholipid synthesis, whereas toxic 16:1Δ9 was not and required elongation to 18:1Δ11 prior to incorporation. However, blocking fatty acid metabolism using inhibitors to prevent acyl-acyl carrier protein formation or glycerol-phosphate acyltransferase activity did not increase the toxicity of 18:1Δ9, indicating that inefficient metabolism did not play a determinant role in fatty acid toxicity. Nontoxic 18:1Δ9 was as toxic as 16:1Δ9 in a strain lacking wall teichoic acids and led to growth arrest and enhanced release of intracellular contents. Thus, wall teichoic acids contribute to the structure-specific antimicrobial effects of unsaturated fatty acids. The ability of poorly metabolized 16:1 isomers to penetrate the cell wall defenses is a weakness that has been exploited by the innate immune system to combat S. aureus.


Staphylococcus aureus is a common cutaneous pathogen responsible for serious infections that are becoming increasingly dangerous due to the prevalence of antibiotic-resistant organisms (8). Lipids have an important role in innate immunity. Human skin deploys a variety of innate defenses against S. aureus colonization that include antimicrobial peptides and fatty acids (9, 18, 47, 49, 51). In mice, 16:1Δ9 is the most potent antibacterial fatty acid, whereas humans synthesize a different isomer, 16:1Δ6 (49). It has been known for decades that these skin fatty acids block the growth of S. aureus (21, 27, 44). Humans (49) and mice (18) deficient in the production of these 16-carbon monounsaturated fatty acids are more susceptible to S. aureus skin infections. However, it is much less clear how these specific fatty acids produce their antibacterial effect. Ideas include the destabilization of the bacterial membrane due to their surfactant properties (19), uncoupling of ATP synthesis (17), the formation of fatty acid hydroperoxides that elicit oxidative stress (29), increased membrane fluidity due to the incorporation in unsaturated fatty acids in phospholipid (5, 7), and the inhibition of de novo fatty acid synthesis at the FabI step (44, 54). These toxic properties of fatty acids stand in contrast to the observations that S. aureus readily incorporates exogenous fatty acids into membrane phospholipids (1, 3, 4, 41) and that acetyl-coenzyme A (CoA) carboxylase knockout mutants can be isolated as fatty acid auxotrophs (41). Taken together, this body of work paints a rather confusing picture of the impact of extracellular fatty acids on S. aureus physiology.

The pathways for the incorporation of exogenous fatty acids are established in Escherichia coli (53). Exogenous fatty acids traverse the outer membrane via the FadL porin and are activated on the cytoplasmic aspect of the inner membrane by acyl-CoA synthetase (13). Acyl-CoAs are substrates for the fatty acid β-oxidation system in E. coli (13) and are substrates for both the glycerol-phosphate (PlsB) and acyl-glycerol-phosphate (PlsC) acyltransferases (53). Gram-positive bacteria, like S. aureus, initiate phospholipid biosynthesis using a different mechanism than E. coli. In place of PlsB, S. aureus expresses the PlsY glycerol-phosphate acyltransferase that utilizes only the unique acyl-PO4 intermediate produced by PlsX from acyl-acyl carrier protein (ACP) end products of type II fatty acid synthesis (FASII) (36). The second acylation is catalyzed by a PlsC that utilizes only acyl-ACP. There is no role for acyl-CoA in S. aureus phospholipid synthesis, and the bacterium does not degrade fatty acids. The only activity that is known to be required for fatty acid incorporation is acyl-ACP synthetase (41). Although the gene encoding acyl-ACP synthetase activity remains to be identified in S. aureus, extracts of S. aureus ligate fatty acids to ACP, which can then be utilized for phospholipid synthesis by either PlsX/PlsY and/or PlsC or elongated by the FabF condensing enzyme (41).

The goal of this study is to systematically study the effect of exogenous fatty acids on growth and membrane integrity of S. aureus to address the underlying mechanism for fatty acid intoxication and the apparent contradiction between the toxicity of exogenous fatty acids and their utilization for membrane formation in S. aureus. We find that only specific fatty acid structures are capable of blocking S. aureus growth and do so by triggering the dissolution of the cytoplasmic membrane, allowing the leakage of metabolites and low-molecular-weight proteins from the cell.


Bacterial strains and growth.

S. aureus strain RN4220 was obtained from Richard Novick (32). Other S. aureus strains, Streptococcus pneumoniae R6, and Bacillus subtilis 168 were obtained from ATCC. Strain EBII53 (tarO::Spec/pG164-tarO) was obtained from Eric Brown (11). Strains PDJ28 (ΔgpsA) and PDJ29 (ΔSA2339) were constructed by the insertion of a group II intron into the respective genes using the primer design software and plasmid system provided by Sigma-Aldrich (Targetron system) (55). The introns were inserted at bp 216 of the SA2339 gene and bp 42 of the gpsA gene. The presence of the insertions was verified by PCR using primers outside the intron insertion site. Growth medium for B. subtilis and S. aureus was either 1% tryptone broth (TB) or Luria broth (LB). Growth medium for S. pneumoniae was C+Y (casein-based medium with yeast extract [CY]) medium (34). RN medium consisted of M9 salts, 1 mM MgSO4, 10 mM CaCl2, 15 μM vitamin B1, 32 μM vitamin B3, 0.1% casein hydrolysate, 0.4% glucose. 0.1 mg/liter biotin, 2 mg/liter pantothenic acid, 10 μM FeCl2, 6 mg/liter citrate, 10 mg/liter MnCl2, 4 μg/liter l-tryptophan, and 0.1 mg/liter lipoic acid. The MICs of the various lipids against S. aureus, B. subtilis, and S. pneumoniae were determined using a broth microdilution method. The strains were grown to an optical density at 600 nm (OD600) of 1.0 and diluted 30,000-fold in TB or CY medium. A 10-μl aliquot of diluted cells was added to each well of a U-bottom 96-well plate containing 100 μl of medium with the appropriate concentration of compound. The plate was incubated at 37°C for 20 h and read using a Fusion plate reader at 600 nm. Cells grown in medium containing dimethyl sulfoxide (DMSO; 1%) were used as 100% growth. Fatty acids and other lipids were purchased from Sigma, Matreya, or Larodan Fine Chemicals.

Cell permeability assays.

Membrane potential was determined by flow cytometry using a BacLight Bacterial Membrane Potential Kit (Life Technologies/Molecular Probes). S. aureus (RN4220) was grown to OD600 of 0.4 in TB, and treated with DMSO (1%), 100 μM 16:1Δ9 or 18:1Δ9, or 5 μM carbonyl cyanide m-chlorophenylhydrazone (CCCP) for 15 min at 37°C. After treatment, cultures were incubated with 30 μM DiOC2(3) (3,3-diethyloxacarbocyanine iodide) for 5 min at room temperature. Cell permeability was determined by flow cytometry using TO-PRO-3 iodide (Life Technologies/Molecular Probes). S. aureus strain RN4220 was grown to an OD600 of 0.4 in TB and treated with DMSO (1%), 100 μM a15:0, 16:1Δ9, a17:0 or 18:1Δ9, 5 μM CCCP, or 80 μg/ml nisin for 15 min at 37°C. After treatment, cultures were incubated with 100 nM TO-PRO-3 iodide at room temperature for 5 min. Flow cytometry was performed using a FACSCalibur (BD Biosciences) running CellQuestPro software (BD Biosciences) on 20,000 S. aureus cells per run. DiOC2(3) was excited with the 488-nm argon laser, and emission was detected as follows: green fluorescence signal was detected using a 530/30 filter, and red fluorescence was detected with a 585/42 filter. TO-PRO-3 iodide was excited with a 635-nm red diode laser, and a 661/16 emission filter was used to detect fluorescence.

Metabolic activity was determined using a Vybrant Cell Metabolic Assay Kit (Life Technologies/Molecular Probes). S. aureus strain RN4220 was grown to an OD600 of 0.4 in TB and treated with DMSO (1%), 100 μM 16:1Δ9 or 18:1Δ9, or 5 μM CCCP for 15 min at 37°C. Cells were diluted back to an OD600 of 0.4 with TB containing the appropriate compound. In a 96-well plate, 200 μl of culture along with 5 μM C12-resazurin was incubated at 37°C for 15 min. Fluorescence was measured using a Fusion Reader set with an excitation of 550 nm (slit width, 20) and an emission of 600 nm (slit width, 10 nm).

The release of ATP from cells was determined after growing strain RN4220 to an OD600 of 0.5. The cells were treated with either fatty acids or DMSO (1%) for 30 min at 37°C with moderate shaking (225 rpm). Strains RN4220 and PDJ28 were grown in LB and RN medium supplemented with 0.1% glycerol, respectively, to an OD600 of 0.5. Strain RN4220 (10 ml) was treated with 400 ng/ml AFN-1252 for 1 h. PDJ28 cells were centrifuged, washed with 50 ml of RN medium without glycerol, and used to inoculate 10 ml of glycerol-free RN medium. Intracellular glycerol-3-phosphate was depleted by incubation of culture for 1 h at 37°C. A 1-ml aliquot of the treated cells was centrifuged at 14,000 × g for 5 min. The supernatant was separated from the cell pellet, and the cell pellet was resuspended in 1 ml of phosphate-buffered saline. A 100-μl aliquot of BacTiter-Glo reagent (BacTiter-Glo Microbial Cell Viability Assay Kit; Promega) with 500 μg of lysostaphin was added to 100 μl of either the supernatant or resuspended cell pellet in an OptiPlate-96F solid-bottom black plate. The mixture was shaken for 2 min at 250 rpm, and luminescence was determined on a Packard Fusion plate reader. The concentration of ATP was determined by comparison to the luminescence obtained from the addition of the BacTiter-Glo reagent to an ATP standard curve. Protein determination was performed using a Bio-Rad protein determination reagent.

Metabolic labeling.

Macromolecular pathway labeling was performed using [1-14C]acetate to indicate lipid biosynthesis, 3H-labeled amino acid mixture for protein synthesis, [3H]thymidine for DNA synthesis, and [3H]uracil for RNA as described previously (41). Fatty acid metabolism was measured in cells grown in TB and labeled with the concentration of [1-14C]palmitoleate (specific activity, 56 mCi/mmol) or [1-14C]oleate (specific activity, 55 mCi/mmol) (American Radiolabeled Chemicals) indicated in the text and figure legends. Cells were harvested by centrifugation and washed two times with TB and two times with phosphate-buffered saline. The lipids were extracted (2), and the total radioactivity incorporated was determined by liquid scintillation counting. Incorporation of fatty acids into the phospholipid fraction was determined by thin-layer chromatography on Silica Gel H layers developed with chloroform-methanol-acetic acid (55:20:5, vol/vol/vol). Fatty acid labeling of the neutral lipid fraction was determined by thin-layer chromatography on Silica Gel G layers developed with chloroform-methanol-acetic acid (98/2/1, vol/vol/vol). The 14C-labeled lipids were visualized and quantified using a Bioscan Imaging detector.

The ability of fatty acids to extract lipids from cells was measured in strain RN4220 or EBII53 grown overnight with or without 1 mM isopropyl-β-d-thiogalactopyranoside (IPTG). Overnight cultures were used to inoculate 10 ml of LB containing 10 μCi/ml [1-14C]acetate (specific activity, 55 Ci/mmole) to label endogenous lipids. Cultures were grown to an OD600 of 1.0 before centrifugation at 4,000 × g for 10 min. [1-14C]acetate was removed by washing with 50 ml of LB medium. Cells were resuspended in 1 ml of medium and used to inoculate 10 ml of LB containing 100 μM 16:1Δ9 with or without 1 mM IPTG in a 50-ml Falcon tube to an OD600 of 0.5. Cells were incubated with shaking at 37°C for 2 h. Subsequently, cultures were centrifuged at 4,000 × g for 10 min, and the supernatant was removed. Cell pellets were resuspended in 1 ml of H2O, and 1 ml of the medium from each culture was aliquoted for lipid extraction. Lipids were extracted by the addition of 3.6 ml of CHCl3-methanol-HCl (1:2:0.02) and incubation at room temperature for 20 min. Next, 1.2 ml of chloroform and 1.2 ml of KCl were added for phase separation, and tubes were centrifuged for 10 min at low speed. The lower phase was removed, and 100 μl was analyzed by scintillation counting.

Lipid structural methods.

Fatty acid composition was determined using cultures (10 ml) of S. aureus RN4220 grown to mid-log phase in TB medium with or without the specified fatty acid. Cells were harvested by centrifugation and washed two times with TB and two times with phosphate-buffered saline. The lipids were extracted (2), and methyl esters were prepared and quantified using a Hewlett-Packard 5890 gas chromatograph as described previously (52). Mass spectrometry of phosphatidylglycerol (PtdGro) was performed using a Finnigan TSQ Quantum (Thermo Electron, San Jose, CA) triple quadrupole mass spectrometer. The instrument was operated in the negative ion mode using parent ion scanning corresponding to the loss of phosphoryl glycerol-H2O. Ion source parameters were as follows: spray voltage of 2,000 V, capillary temperature of 270°C, and capillary offset of −35 V. In addition the tube lens offset was set by infusion of polytyrosine tuning and calibration solution (Thermo Electron, San Jose, CA) in the electrospray mode. Acquisition parameters for parent ion scanning were as follows: scan range, 600 to 900 m/z; scan time, 0.4 s; product mass, 153.0 m/z; collision energy, 45 V; peak width of the first and third quadrupoles (Q1 and Q3, respectively), 0.7 full-width half-maximum (FWHM); and Q2 collision-induced dissociation (CID) gas, 0.5 mTorr. Instrument control and data acquisition were performed with Finnigan Xcalibur (version 1.4 SR1) software.

Gel electrophoresis and ACP immunoblotting.

S. aureus strain RN4220 was grown to an OD600 of 0.4 in TB. Cells were lysed using lysostaphin and Triton X-100 and centrifuged at 80, 000 × g to remove cell debris. The lysates were separated using a 13% polyacrylamide gel containing 1 M urea. The separated proteins were transferred to polyvinylidene difluoride membrane by electroblotting. ACP was detected using an enhanced chemifluorescence (ECF) detection kit (GE Healthcare). The primary ACP-specific antibody was used at a 1:500 dilution, followed by a secondary anti-rabbit IgG conjugated with alkaline phosphatase at a 1:5,000 dilution. Affinity-purified anti-rabbit ACP antibody was described previously (22). The blot was developed using the ECF substrate, and the fluorescent signal was recorded using the Typhoon 9200 PhosphoIMager.

ACP was isolated from the medium of strain RN4220 grown to an OD600 of 0.5 in LB medium. The culture was divided into 100-ml aliquots and treated with the lipid indicated in the text, tables, or figures or an equal volume of DMSO (1% final) as a control. Cells were incubated at 37°C for 30 min, and the cells were pelleted by centrifugation. The medium was removed and applied to a 2.5-ml DE-52 column equilibrated with 20 mM bis-Tris, pH 6.5. The column was washed with 50 ml of equilibration buffer and 50 ml of equilibration buffer containing 0.15 M LiCl and eluted with 10 ml of buffer containing 0.45 M LiCl. The elution fractions were concentrated to 1.5 ml using a 3,000-molecular-weight-cutoff (MWCO) spin filter. The samples were separated by urea gel electrophoresis and immunoblotted for ACP as described above.

Protein assay and identification.

S. aureus strain RN4220 was grown to an OD600 of 0.5 in LB medium. The cells were divided into 100-ml aliquots, centrifuged, and resuspended in an equal volume of M9 minimal medium supplemented with 1 mM MgSO4, 0.1 mM CaCl2, and protease inhibitor cocktail (Roche). The cells were treated with either 200 μM 16:1Δ9 or an equal volume of DMSO (1%) for 30 min. The cultures were centrifuged and the medium was removed and passed through a 0.2-μm-pore-size filter. Proteins in the filtrate were precipitated overnight, and the protein pellet was sedimented by centrifugation at 10,000 × g for 1 h, washed twice with cold acetone, and resuspended in 50 μl of SDS sample buffer (6). Proteins were separated by SDS-PAGE and identified by peptide mass fingerprinting. The proteins were reduced and alkylated with iodoacetamide, and a tryptic digest was prepared. Mass spectrometric analysis was performed using a Model 4700 Proteomics Analyzer from Applied Biosystems (Foster City, CA). This instrument employs matrix-assisted laser desorption/ionization in conjunction with tandem time-of-flight mass analyzers. The digest was introduced into the instrument in a crystalline matrix of α-cyano-4-hydroxycinnamic acid also containing 2 mM ammonium citrate to suppress ionization of matrix clusters. Database searches were performed with Applied Biosystem's GPS explorer software, which uses the Mascot search engine. Swissprot (release date, 22 January 2012) was used for protein identification.

Protein released into the medium was assayed after strain EBII53 was grown overnight with or without 1 mM IPTG. Overnight cultures of strain RN4220 or EBII53 were used to inoculate 20 ml of LB and grown to an OD600 of 2.0. Cultures were centrifuged at 4,000 × g for 10 min and resuspended in 10 ml of RN medium to minimize interference in the protein estimation assay. The OD600 was adjusted to 2.0, and 200 μM 16:1Δ9 was added to each tube. Cultures were incubated for 2 h at 37°C with shaking in a 50-ml Falcon tube. Cells were removed by centrifugation at 4,000 × g for 20 min, and proteins in the medium were quantified by a Bradford protein assay.

Affymetrix arrays and qRT-PCR.

The abundance of gene transcripts was analyzed using the S. aureus Affymetrix array technology. Strain RN4220 was grown in tryptone broth or tryptone broth plus 500 μM 18:1 at 37°C to an OD600 of 0.5. Cells were harvested by centrifugation and washed with 1 ml of RNAlater solution from Ambion. Total RNA was then immediately isolated from the bacterial cells using an RNAqueous Kit (Ambion) per the manufacturer's protocol, including the treatment with lysostaphin and LiCl precipitation. The pelleted RNA was resuspended in nuclease-free water, and a 5-μl aliquot was used to verify RNA quality with an Agilent Technologies 2100 Bioanalyzer Laboratory-on-a-chip before proceeding with the microarray. Synthesis of cDNA and its subsequent fragmentation and labeling with biotin were performed following the protocol from Affymetrix for GeneChip expression analysis. Hybridization, washing, and scanning of the cDNA to the GeneChip S. aureus Genome Array was performed according to the manufacturer's instructions (Affymetrix). Triplicate arrays from three independent samples were analyzed with GeneChip Operating Software, and global scaling was used to normalize the data from different arrays. Spotfire DecisionSite, version 9.1.1, was used to analyze the data. The effects of exogenous 18:1Δ9 on the expression of fab and virulence factor genes were corroborated by quantitative reverse transcription-PCR (qRT-PCR) using the primers listed in Table S2 in the supplemental material. RNA abundance was normalized to the amount of gmk transcript present in the sample, and the amounts present in cells grown with and without 18:1Δ9 were compared.

Electron microscopy.

RN4220 was grown in LB medium to an OD600 of 0.5. A 50-ml culture was treated with either 50 μl of DMSO, 200 μM 16:1Δ9, or 100 μg/ml nisin for 30 min. Cells were centrifuged at 4,000 × g for 10 min and washed twice with 5 ml of PBS. The samples were fixed in 2% paraformaldehyde and 2.5% glutaraldehyde and postfixed in 2% osmium tetroxide in 0.1 M sodium cacodylate buffer with 0.3% potassium ferrocyanide for 2 h. After cells were rinsed in same buffer, they were dehydrated through a series of graded ethanol to propylene oxide washes. The samples were infiltrated and embedded in epoxy resin and polymerized at 70°C overnight. Semithin sections (0.5 μm) were stained with toluidine blue for light microscope examination. Ultrathin sections (80 nm) were cut and imaged using a JEOL 1200 electron microscope with an AMT XR 111 camera.

Microarray data accession number.

The complete data set was deposited under accession number GSE36231 in the NCBI Gene Expression Omnibus (GEO) database (www.ncbi.nlm.nih.gov/geo).


Structure-specific inhibition of S. aureus growth by exogenous fatty acids.

A comparative study of the MICs of different fatty acid structures against S. aureus using a broth microdilution experiment is shown in Fig. 1. Fatty acids were not uniformly toxic toward S. aureus. The most toxic fatty acids all had an MIC of 31 μM, indicating the minimum concentration that was required for growth arrest. A key finding from this survey was the markedly different effects of closely related fatty acid structures. The 16:1 isomers had low MICs, whereas the 18:1 isomers were not toxic at all. Similarly, a15:0 was toxic, whereas a17:0 was not. The three 16:1 isomers tested (cis Δ6, Δ9, and Δ11) were approximately equally effective, but the trans isomer of 16:1Δ9 was considerably less potent. In the 18-carbon series, fatty acid toxicity increased with increasing unsaturation. The only saturated fatty acids that exhibited toxicity were 12:0 and 14:0. This structure-specific pattern of fatty acid toxicity was recapitulated in three other S. aureus strains and Bacillus subtilis (see Fig. S1 in the supplemental material). The pattern of fatty acid toxicity was completely different in Streptococcus pneumoniae (see Fig. S1), which was more sensitive to saturated than unsaturated fatty acids. This result was important because it illustrated that our findings with S. aureus may apply to closely related species but cannot be considered typical for Gram-positive pathogens. The striking differences in toxicity between closely related fatty acids were recapitulated in growing cultures. Cell growth was immediately arrested by 50 μM 16:1Δ9, whereas exposure to 4 mM 18:1Δ9 had no effect on cell growth (Fig. 2A). Growth was impacted in the same way when the toxic a15:0 (50 μM) was compared to a17:0 (4 mM) (data not shown). Examining the time course of growth inhibition by 16:1Δ9 at shorter time intervals showed that 16:1Δ9 stopped growth within minutes of fatty acid addition (Fig. 2A, inset). The abrupt cessation of cell growth elicited by 16:1Δ9 most closely resembled the effect of the protonophore carbonyl cyanide-m-chlorophenylhydrazone (CCCP) rather than effects of the pathway-selective inhibitors mupirocin, a protein synthesis inhibitor, and AFN-1252, a fatty acid synthesis inhibitor (Fig. 2B). Exposure to 16:1Δ9 significantly inhibited fatty acid synthesis from acetate, RNA synthesis from uracil, protein synthesis from amino acids, and DNA synthesis from thymidine (Fig. 2C). The nontoxic 18:1Δ9 did not affect protein, DNA, or RNA synthesis; however, it did reduce acetate incorporation into fatty acids by about 50% (Fig. 2C). This was expected because when strain RN4220 was grown with exogenous 18:1Δ9, ~50% of the phospholipid fatty acids were derived from 18:1Δ9 (41). The rapid onset of growth inhibition by toxic fatty acids coupled with the inhibition of all major pathways for macromolecular biosynthesis suggested that they acted as a general metabolic poison rather than having a specific effect on one branch of metabolism.

Fig 1
Structure-specific fatty acid toxicity in S. aureus. The MICs of a series of fatty acid structures were determined using the broth microdilution assay described in Materials and Methods. Data shown are for the laboratory S. aureus strain RN4220. MIC data ...
Fig 2
Rapid S. aureus growth arrest and membrane depolarization by 16:1Δ9. (A) S. aureus strain RN4220 was grown to mid-log phase in LB–0.1% Brij 58 and treated with either 4 mM 18:1Δ9 (○) or 100 μM 16:1Δ9 (●) ...

The addition of either detergents or bovine serum albumin to the cultures significantly reduces the potency of the fatty acids. These agents bind fatty acids and effectively lower the concentration of free fatty acids in the medium. The inclusion of albumin or detergent in the medium prevents toxicity and allows S. aureus fatty acid auxotrophs to be supplemented with exogenous fatty acids (41).

Effects of toxic fatty acids on cell physiology.

Fatty acids are known to facilitate proton movement across biological membranes (23), and the similarity of the growth arrest by the protonophore CCCP and the toxic fatty acids suggested that fatty acid intoxication was also associated with compromised cytoplasmic membrane function. Growing S. aureus strain RN4220 was treated with diethyloxacarbocyanine, a validated indicator of the proton motive force in S. aureus and other bacteria (37, 46). This two-color flow cytofluorometry assay showed the expected shift in the red-green fluorescence ratio following the addition of CCCP to growing cells (Fig. 2D). Treatment of the cells with 18:1Δ9 yielded a color ratio similar to that of untreated cells, whereas treatment of the cells with 16:1Δ9 showed a ratio shift similar to CCCP treatment, indicating the collapse of the proton gradient (Fig. 2E). Another approach to examining cell function was to use C12-resazurin to indicate the reducing environment within the cell. When the oxidized form of the reagent was added to living cells, it was taken up and converted to the reduced form that exhibits a characteristic fluorescent signal (39). The ability of strain RN4220 to reduce the dye in control and 18:1Δ9- and CCCP-treated cells was clearly evident, but 16:1Δ9-treated cells were unable to reduce the dye (Fig. 2F). Thus, 16:1Δ9-treated cells failed to maintain an intracellular reducing environment, showing that fatty acid intoxication was more deleterious to metabolism than CCCP. These data were consistent with the growth phenotype of 16:1Δ9-treated cells and the ability of 16:1Δ9 to block all major branches of macromolecular synthesis (Fig. 2).

These data indicated that toxic fatty acids may depolarize cells by creating pores in the membrane rather that acting as protonophores. The extent of membrane permeabilization by 16:1Δ9 was assessed by measuring the fluorescence intensity of TO-PRO-3 iodide, which increases significantly when it interacts with DNA. This dye is unable to penetrate cell membranes and was used to distinguish depolarized cells from cells with porous membranes (40). The mean fluorescence of untreated strain RN4220 was low and comparable to that of CCCP-, 18:1Δ9-, and a17:0-treated cells (Fig. 3A). In contrast, treatment of the cells with nisin, an antibiotic that interacts with lipid II to form pores in cytoplasmic membranes (20, 50), resulted in a significant increase in fluorescence, indicating the permeabilization of the cells. The 16:1Δ9-treated cells had an increase in the mean fluorescence that was similar to that with nisin treatment and distinct from that with CCCP treatment, indicating that 16:1Δ9 also permeabilized the cells. The idea from these data that fatty acid intoxication triggered the release of solutes from the cytoplasm was corroborated by measuring the release of ATP from the cells (Fig. 3B). Like nisin treatment, 16:1Δ9- or a15:0-treated cells released virtually all of their intracellular ATP into the medium, whereas CCCP-, 18:1Δ9-, or a17:0-treated cells did not. Transmission electron microscopy of RN4220 cells treated with either 16:1Δ9 or nisin showed no significant morphological difference compared to DMSO-treated cells (see Fig. S2 in the supplemental material). These data indicated that the fatty acid intoxication introduced pores in the cytoplasmic membrane that allowed the passage of solutes out of the cell but did not cause a catastrophic dissolution of the cytoplasmic membrane.

Fig 3
Release of solutes and ACP from S. aureus by toxic fatty acids. (A and B) Two assays were used to determine the permeability properties of cells treated with toxic fatty acids. Controls contained 1% DMSO; fatty acids were added to a final concentration ...

The pores created by toxic fatty acids were large enough to allow the escape of low-molecular-weight proteins into the medium. The phenomenon was clearly illustrated by following the fate of ACP using an anti-ACP antibody to localize the protein. Treatment of strain RN4220 with 16:1Δ9, a15:0, or 18:2 resulted in the disappearance of ACP from the cells, whereas treatment with nontoxic fatty acids did not (Fig. 3C). The ACP was recovered from the medium (Fig. 3C). The time course for cellular ACP depletion indicated that it took >20 min for cells to become completely depleted of ACP (Fig. 3D), which was considerably slower than growth arrest or depolarization (Fig. 2F). Treatment of S. aureus with nisin also released ACP from the cells (Fig. 3E), indicating that pores the size of the nisin-lipid II pores were sufficient for ACP release. The extent of general protein release following fatty acid intoxication was determined by gel electrophoresis of cell supernatants following treatment with 16:1Δ9. Staining of the gel revealed a smear of low-molecular-mass proteins in the <20-kDa region of the gel. Mass spectrometry was used to identify some of the released proteins by mass spectrometry. The goal was not to perform an extensive proteomic analysis of the released proteins but, rather, to positively identify representative proteins that would indicate the scope of protein release by fatty acid intoxication. Proteins positively identified were ACP, (9 kDa), GcvH (14 kDa), CspA (7 kDa), and thioredoxin (13 kDa) and hypothetical protein SA1443 (12 kDa) and TrxA (18 kDa). These data showed that fatty acid intoxication was associated with the permeabilization of the cytoplasmic membrane and the release of low-molecular-mass proteins (<20 kDa) into the medium.

Relationship between fatty acid intoxication and metabolism.

The conclusion that fatty acids disrupt S. aureus membranes due to their surfactant properties fits the data obtained with 16:1Δ9 but does not explain why 18:1Δ9 is not toxic. Metabolic labeling with the toxic 14C-labeled 16:1Δ9 ([14C]16:1Δ9) was compared to that with the nontoxic [14C]18:1Δ9 to determine if differences in the cellular metabolism contributes to the toxicity of specific fatty acid structures. [14C]18:1Δ9-labeled S. aureus primarily synthesized phospholipids from the exogenous fatty acid, but there was always a level of diacylglycerol in the neutral lipid fraction arising from phosphatidylglycerol (PtdGro) turnover for lipoteichoic acid biosynthesis (Fig. 4A). This distribution of label was the same as the distribution of [14C]acetate in the lipid fraction, which tracks the fate of fatty acids produced de novo (41). Metabolic labeling at a nontoxic concentration of [14C]16:1Δ9 (25 μM) did not give rise to free fatty acid accumulation (Fig. 4B). In contrast, labeling of strain RN4220 with 100 μM [14C]16:1Δ9 showed much reduced phospholipid labeling and the appearance of a major new species, free fatty acid (Fig. 4C). These data led to the idea that the structure-specific toxicity of fatty acids occurs when the influx of fatty acids exceeds the ability of the cells to metabolize the fatty acid.

Fig 4
16:1Δ9 is a poor substrate for phospholipid synthesis. Following labeling of strain RN4220 with exogenous fatty acids, the lipids were extracted and separated by thin-layer chromatography to identify the neutral lipids present. PL, phospholipid; ...

The metabolic labeling experiments suggested that the 16:1 isomers were toxic because they were the poor substrates for phospholipid synthesis in S. aureus. Exogenous fatty acids are not degraded by S. aureus (41), and their only metabolic destination is membrane phospholipids and their derivatives (lipoteichoic acids, lipoproteins, etc.). a15:0 and a17:0 are the major fatty acids of S. aureus giving rise to PtdGro molecular species of 32:0 consisting of a17:0 in 1-position and a15:0 in the 2-position (41) (Fig. 4D) (see Table S1 in the supplemental material). Growth of cells with 1 mM 18:1Δ9 showed that 18:1Δ9 was incorporated into the 1-position of PtdGro, the principal phospholipid of S. aureus, and was also elongated to 20:1Δ11 giving rise to 33:1 and 35:1 PtdGro molecular species (41) (Fig. 4E) (see Table S1). These data showed that 18:1Δ9 was a substrate for the PlsX/PlsY acyltransferase system and was elongated by FabF to 20:1Δ11 that was also incorporated into the 1-position. Neither 18:1Δ9 nor 20:1Δ11 was a suitable substrate for PlsC and was found only in the 1-position paired with a15:0 in the 2-position (Fig. 4E). Exposure of strain RN4220 to subtoxic levels of 16:1Δ9 (25 μM) and analysis of the fatty acid composition showed <1% 16:1Δ9 present in the membrane phospholipids (see Table S1). Instead, 16:1Δ9 was elongated to 18:Δ11 and 20:1Δ13 prior to its incorporation into the 1-position to produce 33:1 and 35:1 PtdGro molecular species (Fig. 4F). These data suggested that 16:1Δ9 entered the cell and was converted to 16:1Δ9-ACP, but this acyl-ACP is not an efficient substrate for either the PlsX/PlsY or PlsC acyltransferase systems. Instead, 16:1Δ9 must be elongated by FabF in order to be efficiently utilized by the PlsX/PlsY acyltransferase system that inserts the longer chain unsaturated fatty acids into the 1-position of the glycerol backbone. This idea was further tested by growth of S. aureus with 12:1Δ5 (100 μM) and determining the phospholipid fatty acid composition. Only 18:1Δ11 (7%) and 20:1Δ13 (4%) unsaturated fatty acids were detected in the phospholipids. This experiment showed that 12:1Δ5 entered the cells and was elongated by FabF, but none of the elongation intermediates were incorporated into phospholipid until they reached 18 and 20 carbons in length. This stringent discrimination against 16:1 utilization for phospholipid synthesis correlated with the toxicity of the fatty acid.

Effects of 18:1Δ9 on gene expression.

E. coli and B. subtilis both have specific transcriptional responses to the presence of exogenous fatty acids governed by repressors that sense the influx of fatty acids (16). The effect of exogenous fatty acids on S. aureus gene expression was investigated using Affymetrix arrays to profile mRNA levels in strain RN4220 grown to mid-log phase in the presence or absence of 500 μM 18:1Δ9. The complete triplicate data sets for control and 18:1Δ9-treated cells were deposited in the GEO database under accession number GSE36231. The goal was to identify oleate-regulated genes and determine if they were involved in fatty acid toxicity. A list of genes that were regulated ±2-fold with a P value of <0.05 and with a statistically significant signal in all six samples is provided as a searchable Excel spreadsheet as Table S3 in the supplemental material. The most notable downregulated genes were a group of virulence factors that are controlled by the SaeRS two-component regulator (38). A selection of these virulence genes was analyzed by quantitative real-time PCR to verify their strong downregulation by growth in 18:1Δ9 (see Fig. S3 in the supplemental material). The SaeRS system autoregulates the saePQRS operon, which also had significantly reduced expression in 18:1Δ9-treated cells, as indicated by the strong reduction in the level of saeP mRNA (see Fig. S3). Surprisingly, fabH and plsX were upregulated by growth in 18:1Δ9 (see Table S3), and these effects of exogenous 18:1Δ9 on gene expression were also confirmed by qRT-PCR (see Fig. S3). The observation that fatty acid biosynthetic genes were upregulated by exogenous 18:1Δ9 seems counterintuitive because fatty acid synthesis from acetate was repressed in 18:1Δ9-treated cells (Fig. 2B) (41).

The most highly upregulated gene in strain RN4220 grown in 18:1Δ9 was SA2339, a hypothetical protein belonging to the MmpL family off lipid transporters (14, 48). This intriguing connection prompted testing the potential role for SA2339 in exogenous fatty acid metabolism. The SA2339 gene was inactivated in strain PDJ29 by inserting a type II intron sequence at nucleotide 216 of the SA2339 gene. Metabolic labeling of strain PDJ29 with [14C]18:1Δ9 did not reveal any differences in the uptake and incorporation of exogenous fatty acids between strains RN4220 and PDJ29 (ΔSA2339) (data not shown). Kenny et al. (27) screened a transposon library of S. aureus mutants and reported that an insertion in gene SA2339 (SAR2632) had a 4-fold higher plating efficiency on 18:2 agar plates than the wild type. However, our MIC and growth experiments failed to show altered sensitivity to 16:1Δ9 toxicity in strain PDJ29 (ΔSA2339) compared to the parent strain RN4220 (data not shown). The transcriptional profiling experiments did not identify genes that were clearly involved in modulating fatty acid intoxication or exogenous fatty acid metabolism.

Other antibacterial lipids have the same mechanism of action as the toxic fatty acids.

Our results suggested that the surfactant properties of fatty acids were responsible for cell permeabilization although their effects are modulated by the cell wall and their utilization for membrane formation. Therefore, other lipids reported to kill S. aureus were tested to determine if these molecules functioned by triggering the leakage of cellular components. 1-Laurylglycerol (12-MG) has antibacterial activity against S. aureus (35, 45) and is regarded as a safe food preservative. Because lauric acid (12:0) was a moderately toxic fatty acid (Fig. 1), the antibacterial activity of 12-MG may arise from esterase cleavage of the monoglyceride. This idea was supported by the equal toxicity for 12:0 and 12-MG (Table 1) (43). The ether lipid analog of 12-MG (12-GE) was more potent than the corresponding monoglyceride (Table 1) (35), indicating that the cleavage of the fatty acid from 12-MG and its incorporation into phospholipids protected the cells from toxicity. Long-chain amines, like laurylamine (12-NH2), were also more potent antibacterials than the corresponding fatty acids (Table 1) (28). Finally, sphingoid bases have antibacterial activity (15), and we confirmed that sphingosine (Sph) was toxic to S. aureus (Table 1). The ACP release assay was used as a diagnostic experiment to determine if these potent antibacterial lipids act by the same permeabilization mechanism as the toxic fatty acids. All of the potent antibacterial lipids (long-chain amines, ether lipids, and sphingosine) released ACP into the medium, indicating that they disrupted the membrane to the same extent as 16:1Δ9 (Fig. 3E). Thus, toxic lipids share the common property of rapid cell growth arrest associated with the release of solutes and low-molecular-weight proteins from the cell.

Table 1
MICs of a series of antibacterial lipids against S. aureus strain RN4220

The role of fatty acid metabolism in determining the toxicity profile was tested using two systems. First, strain RN4220 was treated with the enoyl-ACP reductase inhibitor AFN-1252, which was established to sequester the cellular ACP as an elongation pathway intermediate, thus preventing the utilization of exogenous fatty acids for phospholipid synthesis (41). Second, we constructed strain PDJ28 (ΔgpsA). GpsA is responsible for supplying glycerol-3-phosphate for phospholipid synthesis, and strain PDJ28 was a glycerol auxotroph. As reported previously in another S. aureus glycerol auxotroph (42), the removal of glycerol from the medium led to an abrupt cessation of phospholipid synthesis based on [14C]acetate incorporation (data not shown). If incorporation into phospholipid was an important detoxification mechanism for 18:1Δ9, then 18:1Δ9 should release solutes from strain RN4220 treated with AFN-1252 or in strain PDJ28 following the removal of glycerol. Toxicity was assessed by determining the release of ATP from the AFN-1252-treated or glycerol-depleted cells, and in both cases 16:1Δ9 caused the release of ATP, but 18:1Δ9 did not (Fig. 5A). These data argued against a dominant role for metabolism in modulating fatty acid intoxication.

Fig 5
Role of metabolism and wall teichoic acids in fatty acid intoxication. (A) Strain RN4220 or PDJ28 (ΔgpsA) was grown to mid-log phase and either treated with AFN-1252 to arrest fatty acid synthesis (RN4220) or deprived of glycerol for 1 h (PDJ28) ...

Role of wall teichoic acids in selective resistance to fatty acids.

The Gram-positive cell wall presents a barrier that confers resistance to surfactants like Triton X-100 (31) and the antimicrobial fatty acid 16:1Δ6 (30). Our goal was to determine if wall teichoic acids were involved in conferring the structure specificity of unsaturated fatty acid toxicity (Fig. 1). This question was addressed by examining the fatty acid sensitivity of strain EBII53 (ΔtarO, also known as tagO) that harbors an IPTG-inducible plasmid expressing a complementing tarO gene (11). TarO catalyzes the first step in wall teichoic acid synthesis, and cells lacking tarO expression lack wall teichoic acids (11). The sensitivity of strain EBII53 to 16:1Δ9 and 18:1Δ9 was tested in a broth microdilution assay in the presence and absence of IPTG (Fig. 5B). These experiments showed that 18:1Δ9 was just as toxic as 16:1Δ9 in the TarO-deficient cells, but it was not toxic when tarO was complemented by induction with IPTG. These data indicated that wall teichoic acids were a major determinant of the structural specificity of antimicrobial unsaturated fatty acids. Next, a series of experiments were performed to determine if the wall teichoic acids were involved in establishing the pore size for the leakage of components from cells treated with antimicrobial fatty acids. Cells were incubated with [1-14C]acetate to label the phospholipids. Treatment with 16:1Δ9 extracted a modest amount of labeled phospholipid into the medium in either strain RN4220 or EBII53 grown with IPTG. However, about 50% of the labeled cellular lipid was released by 16:1Δ9 treatment in the absence of wall teichoic acids (Fig. 5C). Consistent with this observation, significantly more protein was released from the TarO-deficient cells than from the complemented strain, which had the same amount of protein release as strain RN4220 (Fig. 5D). The proteins released by 16:1Δ9 from strain EBII53 in the presence and absence of IPTG were analyzed by SDS-gel electrophoresis. The increase in total protein released in the absence of IPTG-induced complementation showed an increase in proteins with molecular masses of >25 kDa (data not shown). These data support the idea that the cell wall allows the selective leakage of solutes and low-molecular-weight proteins when the cytoplasmic membrane was compromised by 16:1Δ9, but in the absence of teichoic acids, fatty acids cause a more complete release of cell components. These results showed that wall teichoic acids contributed to the selective toxicity of fatty acid structures and contribute to the retention of cellular phospholipids and proteins in 16:1Δ9-treated cells.


The systematic examination of the permeability properties of S. aureus following treatment with a prototypical toxic fatty acid, 16:1Δ9, shows the accumulation of cell-associated free fatty acids that correlates with the disruption of the cytoplasmic membrane and leakage of solutes and low-molecular-weight proteins into the medium. This conclusion is consistent with the general concept proposed in the 1970s that exogenous fatty acids trigger membrane damage that interferes with energy metabolism (17, 19). However, other explanations that were subsequently proposed for fatty acid intoxication, like formation of hydroperoxides (29), increased membrane fluidity (5, 7), and the inhibition of fatty acid synthesis at the FabI step (44, 54), are not consistent with the data. The 16:1Δ9-dependent release of solutes and low-molecular-mass proteins (<25 kDa) into the medium, coupled with the retention of macromolecules and phospholipids within the cell, suggests the creation of pores with a defined size. Nisin is an antibiotic that forms a complex with lipid II to form structured pores (~2 nm) in the cytoplasmic membrane (20, 50), consistent with the observed nisin-dependent release of solutes and ACP from the cell (Fig. 3E). Specific interactions between 16:1Δ9 and other membrane components to assemble pores appears unlikely because structurally unrelated surfactants like alkylglycerol, sphingosine, and alkylamines have the same effect on protein release as toxic fatty acids (Fig. 3). A more tenable explanation is that the lipid surfactants accumulate in the cytoplasmic membrane, disrupting the phospholipid bilayer. In this scenario, the apparent pore size would correspond to the size of the channels through the cell wall, which would not be dissolved by the lipids. The Gram-positive cell wall acts as a barrier to large molecules but contains openings of about ~2 nm in size that allow the diffusion of solutes and proteins up to ~25 kDa (12). Thus, we suggest that the cell wall acts as a dialysis membrane, following the dissolution of the cytoplasmic membrane by toxic lipids, that immediately collapses the proton gradient and rapidly releases solutes like ATP into the medium. Proteins are released more slowly, depending on their size. This role for the cell wall is consistent with the enhanced release of proteins and phospholipids by fatty acids in an S. aureus strain lacking wall teichoic acids. This model provides a framework for understanding the rapid cessation of growth, the inhibition of all major branches of macromolecular biosynthesis, and the eventual loss of cell viability associated with toxic lipids.

The data reveal a key role for wall teichoic acids in determining the structural specificity of fatty acid toxicity as well as determining the extent of cellular damage by antimicrobial fatty acids. There are a number of reports that link cell wall components to the sensitivity of S. aureus to antimicrobial compounds produced by the innate immune system. The absence of wall teichoic acids (30), a reduction in the level of d-alanine modification (10), and the absence of major surface protein IsdA (9) all increase the sensitivity of cells to antimicrobial host factors. Our work extends the role of the cell wall by showing that wall teichoic acids contribute to the structure-specific toxicity of fatty acids. This finding explains why closely related fatty acid structures with nearly the same surfactant properties have markedly different effects on cell physiology (Fig. 1). The biochemical mechanism that underlies the wall teichoic acid contribution to unsaturated fatty acid action on cells is unknown but may involve the wall teichoic acid network itself or may reflect the absence of cell components that require the wall teichoic acids for proper assembly. Wall teichoic acids are also responsible for mitigating the damage caused by antimicrobial fatty acids. Cells with an intact cell wall leak solutes and low-molecular-weight proteins from the cell, but in the absence of the cell wall, the release of cellular components is more complete. Thus, the mammalian innate skin defense system produces poorly metabolized 16:1 isomers to exploit the inability of the cell wall to defend S. aureus against these surfactants.

There remain a number of unknowns about exogenous fatty acid metabolism in S. aureus. In E. coli, the FadL porin is required for fatty acids to transit of the outer membrane (33), but there is no experimental evidence that a specific protein is required for fatty acid penetration of the Gram-positive cell wall. The second step in fatty acid translocation is the movement of the fatty acid from the outside to the inner aspect of the cytoplasmic membrane where it can be accessed by acyl-ACP (or acyl-CoA) synthetases. Such transporters have not been found in bacteria despite extensive mutagenesis studies in E. coli to identify the components of the fatty acid uptake process. A protein transporter may not be required because the ΔpH would drive the protonation of the fatty acid intercalated into the outer aspect of the cytoplasmic membrane, which allows the fatty acid to rapidly and spontaneously flip to the inner leaflet, where it is accessible to metabolic enzymes (26). Thus, it seems certain that acyl-ACP synthetase may be the only essential activity for the utilization of fatty acids for phospholipid synthesis in S. aureus. Acyl-ACP synthetases are known (24, 25), but the gene responsible for encoding this activity in S. aureus remains to be identified.

Supplementary Material

Supplemental material:


This research was supported by National Institutes of Health grant GM034496, Cancer Center (CORE) Support Grant CA21765, and the American Lebanese Syrian Associated Charities.


Published ahead of print 27 July 2012

Supplemental material for this article may be found at http://jb.asm.org/.


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