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. 2025 Nov 21;60(5):4213–4227. doi: 10.1021/acs.est.5c10008

Biosynthesis and Export of Membrane-Enveloped Selenium Nanoparticles by Escherichia coli

Anna Ochi †, Kano Shibamoto †, Yosuke Toyotake †, Daiki Fujioka †,‡, Fumiaki Yokoyama §, Hiroki Okanishi ∥, Takeshi Imai ⊥, Daiki Fujita †, Riku Aono †, Masao Inoue †,#, Masaru Takizawa ∇, Ryuta Tobe †, Yoshikatsu Kanai ∥,○,◆,, Tomoya Imai ▶, Hisaaki Mihara †,*
PMCID: PMC12895521  PMID: 41271200

Abstract

Bacteria reduce toxic selenium oxyanions, such as selenite, to elemental selenium (Se0), forming selenium nanoparticles (SeNPs) either intracellularly or extracellularly. However, the mechanism through which extracellular SeNPs (Ex-SeNPs) are exported remains unclear. In this study, we characterized Ex-SeNPs biosynthesized by Escherichia coli during the aerobic reduction of selenite. The SeNPs appeared within 2 h of exposure, remained extracellular, and displayed a consistent spherical morphology (∼100 nm). Purified Ex-SeNPs consisted of an Se0 core enveloped by a membrane-like layer containing lipids, proteins, carbohydrates, peptidoglycan, and lipopolysaccharides. Fluorescence microscopy and gas chromatography–mass spectrometry indicated that the encapsulated membrane originates from the E. coli cell membrane. Notably, mutants deficient in the outer membrane proteins OmpC or TolA failed to excrete SeNPs, resulting in intracellular accumulation despite efficient Se0 synthesis. Our findings suggest that E. coli forms SeNPs intracellularly and exports them via an envelope-dependent process, during which the particles may become encapsulated in membrane-like structures. These findings help clarify the mechanism underlying a membrane-dependent pathway for SeNP detoxification and export that had been suggested but not directly demonstrated.

Keywords: selenium nanoparticles, Escherichia coli, nanoparticle export, cell envelope, selenite reduction

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Introduction

Selenium nanoparticles (SeNPs) have garnered increasing attention due to their broad-spectrum antimicrobial and anticancer properties, as well as relatively low cytotoxicity toward nontarget cells. − When biosynthesized by bacteria, SeNPs acquire a natural coating composed of proteins, lipids, and polysaccharides, which contributes to enhanced colloidal stability and aqueous dispersibility. , These features make biogenic SeNPs an attractive alternative to chemically synthesized counterparts, particularly in the context of environmentally sustainable applications such as biomedicine, food preservation, and nutraceutical development. ,

In microbial systems, SeNP biosynthesis is initiated by the uptake of selenite (SeO3 2−), primarily via phosphate and sulfate transporters. , Intracellularly, selenite is reduced to insoluble elemental selenium (Se0) through a network of redox pathways that include glutathione- and thioredoxin-dependent systems, as well as various respiratory and detoxification enzymes, such as fumarate, nitrate, nitrite, sulfite, and selenite reductases, whose involvement varies depending on the bacterial species. − The resulting Se0 are proposed to nucleate into small clusters, which may subsequently grow into nanometer-scale particles via Ostwald ripening. , These particles are subsequently stabilized by adsorption of endogenous biomacromolecules, conferring physicochemical properties distinct from chemically synthesized SeNPs. −

Despite recent advances in understanding SeNP biosynthesis, , the mechanism by which Se0 or SeNPs are exported from the cytoplasm to the extracellular space remains poorly characterized. While membrane vesicles (MVs) have been proposed as potential mediators of nanoparticle transport, direct ultrastructural evidence remains limited. − Moreover, the extent to which cell-derived membranes contribute to the formation and stabilization of extracellular SeNPs (Ex-SeNPs) remains unclear. This knowledge gap poses a barrier to the rational design of microbial systems for scalable and controllable SeNP production.

Although Escherichia coli has not been widely utilized in SeNP-related research, recent observations suggest that it is capable of producing Ex-SeNPs under aerobic conditions, ,, providing a valuable system to explore the molecular basis of SeNP formation and export. In E. coli, selenite reduction is facilitated by thiol-dependent redox systems, often referred to as “painter-type” reactions. Although specific intracellular proteins, such as AdhP, have been identified on SeNP surfaces and implicated in modulating particle formation, the cellular machinery responsible for exporting SeNPs from the cytoplasm to the extracellular space has not been elucidated.

In this study, we investigated the molecular and structural features of Ex-SeNPs produced by E. coli BW25113, to uncover the mechanism underlying their secretion. Purified Ex-SeNPs were found to possess an Se0 core surrounded by a membrane-like structure containing lipids, proteins, carbohydrates, peptidoglycan, and lipopolysaccharides, consistent with a cell-derived origin. Furthermore, genetic analyses revealed that mutants lacking key components of envelope integrity, such as OmpC or TolA, failed to secrete SeNPs despite unaltered Se0 production, resulting in pronounced intracellular accumulation. These findings support the existence of a membrane-dependent export pathway for SeNPs in E. coli, providing new insights into cellular strategies for managing selenium toxicity through nanoparticle encapsulation and excretion.

Materials and Methods

Bacterial Strains and Culture Conditions

E. coli K-12 BW25113 (wild type) and its single-gene deletion mutants, JW2203 (ΔompC), JW0940 (ΔompA), JW3132 (ΔnlpI), and JW0729 (ΔtolA), were obtained from the National BioResource Project (National Institute of Genetics, Shizuoka, Japan) and precultured overnight in lysogeny broth (LB) containing 5 g/L yeast extract, 10 g/L tryptone, and 10 g/L sodium chloride at 37 °C. The precultures were then inoculated in 5 mL tryptic soy broth (TSB; 17 g/L pancreatic digest of casein, 3 g/L papaic digest of soybean, 2.5 g/L dextrose, 5 g/L sodium chloride, and 2.5 g/L dipotassium phosphate) at an initial OD600 of 0.01 and incubated at 37 °C with shaking at 220 rpm under aerobic conditions. At the exponential phase (18 h), 1 mM sodium selenite was added, and the culture was continued for 2–24 h. For the mutant strains, 50 mg/L kanamycin (Nacalai Tesque, Kyoto, Japan) was added to the preculture medium to maintain the kanamycin resistance marker.

Selenite Determination by Hydride Generation Atomic Fluorescence Spectrometry

Selenite levels in the bacterial culture supernatants (obtained by centrifugation at 15,000g for 15 min) were determined using a hydride generation atomic fluorescence spectrometer (HG–AFS; Millenium Excalibur, PSA, Orpington, UK) equipped with a selenium PS analytical lamp (P849SF Photoron Pty Ltd., Victoria, Australia), as previously described. The HG–AFS operating conditions were as follows: injection volume, 100 μL; acid carrier, 0.5% (w/v) KBr in 6 M HCl; reductant, 0.7% (w/v) NaBH4 in 0.1 M NaOH.

SeNP Determination by Lead Selenide Formation

The amount of pellet-associated SeNPs was quantified by converting elemental selenium (Se0) to selenide and measuring the formation of lead selenide (PbSe) turbidity at 400 nm. The pellets (cells plus cell-associated material) were resuspended in phosphate-buffered saline (PBS; 8 g/L NaCl, 0.2 g/L KCl, 1.4 g/L Na2HPO4, 0.2 g/L KH2PO4; pH 7.4), followed by the addition of an equal volume of 250 mM dithiothreitol (DTT) to reduce Se0 to selenide chemically. The resulting mixture was centrifuged at 15,000g for 2.5 min. Subsequently, 100 μL supernatant was mixed with 700 μL of 5 mM lead acetate in 0.1 M HCl, and A 400 was recorded. This assay quantifies pellet-associated SeNPs (intracellular plus tightly cell-associated). A calibration curve was generated using sodium selenide (Na2Se) (Figure S1). Na2Se solutions at 0.04, 0.08, 0.16, 0.31, 0.63, and 1.25 mM (50 μL each) were mixed with 50 μL of 250 mM DTT; then 100 μL of this mixture was added to 700 μL of 5 mM lead acetate in 0.1 M HCl, and A 400 was measured. Reagent blanks (without Na2Se) were subtracted. Sample concentrations were interpolated from the standard curve.

Ex-SeNP Purification

An E. coli culture incubated with 1 mM sodium selenite in TSB for 6 h was centrifuged at 4000g for 5 min to remove the cells. The resulting supernatant was filtered through a 0.45 μm membrane filter (CA Syringe Filter, Membrane Solutions, Texas, USA) and centrifuged at 11,000g for 1 h. The pellet was washed twice with PBS (pH 7.4) and resuspended in the same buffer.

Chemically Synthesized SeNP Preparation

Chemically synthesized SeNPs (Chem-SeNPs) were prepared as previously described by Lin and Chris Wang. Briefly, 5.2 mM selenium dioxide and 5.2 mM sodium thiosulfate were mixed in a 0.01 M sodium dodecyl sulfate (SDS) solution and incubated at 25 °C. This procedure yielded Chem-SeNPs with an average diameter of approximately 90 nm, similar to those biosynthesized by E. coli (Figure S2).

Phase-Contrast Microscopy

Bacterial cells were observed under a CX43 phase-contrast microscope (Olympus, Osaka, Japan) at 1000× magnification. After 24 h of incubation with selenite under the conditions described above, the cells were harvested by centrifugation at 4000g for 1 min, washed twice with PBS, and resuspended in PBS for microscopic observation.

Transmission Electron Microscopy Analysis

Bacterial cells were fixed with 4% (w/v) paraformaldehyde for 2 h at room temperature, harvested, and washed in HEPES-buffered saline (HBS; 40 mM HEPES-NaOH, pH 7.4, 150 mM NaCl) containing 4% (w/v) paraformaldehyde and 0.1% (v/v) glutaraldehyde. Postfixation was performed using 1% (v/v) osmium tetroxide in HBS for 2 h at room temperature. The cells were washed twice with HBS and resuspended in the same buffer. A 2.5 μL aliquot of each fixed sample was applied to hydrophilized carbon-coated copper grids. The grids were washed with HBS, stained twice for 1 min using an EM Stainer (Nisshin-EM, Tokyo, Japan), and then air-dried for negative staining observation. Ex-SeNP samples were examined without chemical fixation. For ultrathin sectioning, the cells fixed with glutaraldehyde and postfixed with osmium tetroxide were resuspended in 2% agarose L (NIPPON GENE Co. Ltd., Tokyo, Japan) and solidified on ice. The embedded samples were dehydrated using a graded ethanol series (50%, 70%, 90%, 99.5%, and absolute ethanol) and then finally substituted with propylene oxide. The dehydrated gel-embedded sample was embedded in low-viscosity Spurr resin (Merck KGaA, Darmstadt, Germany) according to the manufacturer’s instructions. Following heat curing, ultrathin sections (70 nm) were prepared using a diamond knife (Drukker Ultramicrotome knife, element six, Luxembourg, Luxembourg) on an ultramicrotome EM UC7 (Leica Inc., Wetzlar, Germany) and stained with EM Stainer. Transmission electron microscopy (TEM) was performed using a JEM-2100Plus transmission electron microscope (JEOL Ltd., Tokyo, Japan), operated at an accelerating voltage of 120 kV. The images were captured using a charge-coupled device (CCD) camera.

High-Angle Annular Dark-Field Scanning TEM and Energy-Dispersive X-ray Spectrometry Analysis

The samples prepared as described above were analyzed using high-angle annular dark-field scanning TEM (HAADF–STEM) combined with energy-dispersive X-ray spectrometry (EDS). The structural characterization of the nanomaterials was carried out by acquiring selected area electron diffraction (SAED) patterns.

X-ray Absorption Fine Structure Analysis of SeNPs

The Se L-edge was used for X-ray absorption fine structure (XAFS) analysis because it provides higher sensitivity to the Se oxidation state and bonding environment, allowing clearer discrimination of chemical states compared with the Se K-edge.

Freeze-dried Ex-SeNPs and standard selenium compounds, including elemental selenium (Nacalai Tesque, Kyoto, Japan) and selenomethionine (Tokyo Chemical Industry Co., Ltd., Tokyo, Japan), were prepared for XAFS analysis. The measurements were performed at beamline BL-13 of the Ritsumeikan SR Center (Shiga, Japan). XAFS spectra were collected using total electron yield (TEY) and partial fluorescence yield (PFY) modes, which provide surface (∼50 nm) and bulk (∼1 μm) information, respectively. All measurements were performed at room temperature under an ultrahigh vacuum of approximately 10–7 Pa.

X-ray Photoelectron Spectroscopy Measurement of SeNPs

X-ray photoelectron spectroscopy (XPS) measurements were carried out using a hemispherical energy analyzer (SCIENTA SES2002) using Al Kα (1486.6 eV) at BL-7 of Ritsumeikan SR Center. The measurements were performed at room temperature under an ultrahigh vacuum of approximately 10–8 Pa.

Fourier Transform Infrared Spectroscopy Analysis of SeNPs

The Fourier transform infrared (FT-IR) spectra of SeNPs were recorded using a Nicolet iS50 FT-IR spectrometer (Thermo Fisher Scientific, MA, USA) equipped with an attenuated total reflectance (ATR) attachment. Spectra were recorded from 1750 to 750 cm–1. Prior to the analysis, the samples were dried at room temperature for 30 min without additional treatment. For comparison, the FT-IR spectra of the Chem-SeNP were obtained under the same conditions.

Proteomic Analysis of SeNP-Bound Proteins

Purified Ex-SeNPs and negative control (NC) samples (precipitates from selenite-free E. coli cultures) were reduced with 5 mM DTT and heated at 95 °C for 30 min to denature the bound proteins. Each sample was separated by 1 cm using a 12% SDS–PAGE gel and the proteins were visualized using Coomassie Brilliant Blue staining. The stained gel regions were excised, cut into 1 mm3 blocks, and destained. In-gel protein digestion and subsequent liquid chromatography–tandem mass spectrometry (LC–MS/MS) analysis were performed as described previously. The MS and MS/MS spectral data were analyzed using Proteome Discoverer version 2.3 (Thermo Fisher Scientific, Waltham, MA, USA). Peptide-spectrum matches were assigned using product-ion search algorithms against the E. coli K-12 protein database (4,360 entries). The precursor and fragment mass tolerances were set at 10 ppm and 0.02 Da, respectively. Carbamidomethylation of cysteine residues was specified as a static modification, whereas methionine and N-terminal acetylation oxidation were set as dynamic modifications. The search results were filtered using a q-value threshold of 0.01 to ensure high-confidence protein identification. Based on the resulting protein lists, in silico analysis of subcellular localization was performed using DeepLocPro (https://biolib.com/KU/DeepLocPro/), a tool for predicting protein subcellular localization.

Fluorescence Imaging of Membrane-Stained SeNPs

Freeze-dried Ex-SeNPs and Chem-SeNPs were resuspended in 100 μL Milli-Q water. Then, 1 μL suspension was mixed with 10 μL Milli-Q water with or without 10 μg/mL FM4-64 (Thermo Fisher Scientific), and 2 μL mixture was loaded on glass slides (0.13–0.17 μm thickness, Matsunami Glass, Osaka, Japan) and air-dried at 25 °C for ∼30 min to fix these small nanoparticles on a focus plane for imaging. Bright-field and FM4-64 fluorescence images were obtained using a Leica SP8 confocal microscope with a 100× oil-immersion objective (HC PL APO C52, N.A. 1.40, #11506372, Leica, Wetzlar, Germany) under a Leica LasX operation at 400 Hz scan speed and 1 Airy unit pinhole. Images (1024 × 1024 pixels with pixel size ≈ 0.12 μm) were obtained using a 552 nm laser (1% intensity) and 570–650 nm emission window (gain: 100, 4× average scanning). Bright-field images were captured simultaneously (gain: 350). Five fields were imaged per sample, with three independent replicates. FM4-64 fluorescence intensity and SeNP areas were quantified using Fiji/ImageJ v2.16.0/1.54p. SeNPs were segmented from bright-field images using the Otsu method and their areas were quantified using the Analyze Particle function. The binary masks were prepared by dividing the pixel values of the segmented masks by 255 (8-bit maximum). These masks were applied to the fluorescence images to quantify the FM4-64 signals from the SeNPs. The background signals and areas were extracted from the inverted bright-field images using the same procedure.

Lipid Extraction and Lipid Composition Analysis by Gas Chromatography–Mass Spectrometry

Lipids from the Ex-SeNPs and whole E. coli cells were extracted using a modified Bligh and Dyer method. The samples were resuspended in 0.4 mL distilled water, followed by the sequential addition of 0.5 mL methanol and 1 mL chloroform. The tubes were then flushed with nitrogen gas and sonicated at 40 kHz for 4 min at room temperature. Subsequently, 0.5 mL distilled water and 0.5 mL chloroform were added, and the tubes were reflushed with nitrogen gas. After vortex mixing and centrifugation, the chloroform phase was collected and evaporated under nitrogen gas. The dried lipid extracts were then dissolved in 10% (w/v) HCl in methanol for acid methanolysis reaction. The resulting fatty acid methyl esters were extracted with dichloromethane and n-hexane and analyzed using a gas chromatography–mass spectrometry (GC/MS) system (Clarus 680 gas chromatograph interfaced with a Clarus SQ 8C mass spectrometer, PerkinElmer, Wellesley, MA, USA) equipped with an Agilent J and W GC column DB-1 (Agilent Technologies Inc., Santa Clara, CA, USA). The compounds were identified based on their mass spectra.

MV Isolation from Ex-SeNPs

The Ex-SeNPs were resuspended in PBS containing 20 mM DTT and gently reduced. Once the red color of the SeNPs disappeared, the sample was ultracentrifuged at 145,650g for 1 h. The resulting MV pellet was washed twice with PBS, resuspended in PBS, and analyzed using TEM.

Peptidoglycan Detection by Silkworm Larval Plasma Assay

Peptidoglycan detection was performed on MVs isolated from Ex-SeNPs using the Silkworm Larval Plasma (SLP) reagent (Fujifilm Wako Pure Chemical Industries, Ltd., Osaka, Japan), which employs a pro-phenol oxidase cascade system to induce melanin pigmentation in response to trace amounts of peptidoglycan.

Lipopolysaccharides Detection by Limulus Amebocyte Lysate Assay

Lipopolysaccharides detection was performed on Ex-SeNPs suspended in Milli-Q water using a Limulus Amebocyte Lysate (LAL) gel-clot assay (Fujifilm Wako Pure Chemical Industries, Ltd., Osaka, Japan), which utilizes the horseshoe crab coagulation cascade to form a clot in response to trace amounts of endotoxin.

Gene Complementation Analysis

The ompC and tolA genes were PCR-amplified from E. coli BW25113 genomic DNA using the primer sets OmpC-f/OmpC-r and TolA-f/TolA-r, respectively (Table S1). The amplified fragments were digested with SacI and SalI (New England Biolabs, MA, USA) and ligated into the corresponding sites of pBAD-Cm (American Type Culture Collection, Manassas, VA, USA) to generate pBAD-OmpC and pBAD-TolA plasmids, which were introduced into ΔompC and ΔtolA strains, respectively. The transformed cells were precultured overnight at 37 °C in LB medium containing 30 μg/mL chloramphenicol (Nacalai Tesque). The cultures were then inoculated into 5 mL TSB medium at an initial OD600 of 0.01, supplemented with 0.0002% (w/v) l-arabinose, and cultivated under the conditions described above. This low concentration of l-arabinose was used to avoid toxicity associated with overexpression of these membrane-associated proteins.

Minimal Inhibitory Concentrations of Sodium Selenite

To evaluate selenite tolerance, an initial broad-range screening was performed to roughly determine inhibitory concentrations (MICs). ΔompC, and ΔtolA strains were tested at 0–50 mM sodium selenite, whereas the WT strain was tested up to 100 mM. Based on these results, MICs were determined more precisely by inoculating log-phase cultures (approximately 1 × 105 CFU/mL) into 5 mL of TSB containing narrower concentration ranges near the predicted MICs (ΔompC/ΔtolA: 0, 15, 20, 25, 30, 35, 40, 45, 50 mM; WT: 0, 50, 55, 60, 65, 70, 75, 80, 85, 90, 95, 100 mM). Cultures were incubated at 37 °C with shaking, and cell growth was monitored at 18 h and again at 44 h to account for delayed outgrowth. For concentrations showing low OD600, cultures were diluted 10–1000× and plated on TSB agar to determine CFU/mL. MIC was defined as the lowest concentration at which CFU/mL fell below 103. All experiments were performed in triplicate.

Statistical Analysis

The Kruskal–Wallis test was used to compare SeNP particle sizes among the three groups. Pairwise comparisons were conducted using the Wilcoxon rank-sum test with continuity correction where appropriate, and p-values were adjusted for multiple comparisons using the Bonferroni method. Fluorescence intensity data were log10-transformed prior to analysis to reduce skewness and stabilize variance, assuming a log-normal distribution. Group differences in fluorescence intensity were assessed using Welch’s t-test, with Bonferroni correction applied to account for multiple comparisons. All statistical analyses were performed using R software, and differences with p-value <0.01 were considered statistically significant.

Results

Selenite Reduction and SeNP Formation by E. coli BW25113

E. coli BW25113 reduces selenite to form Ex-SeNPs under aerobic conditions. , To better understand when and how SeNPs emerge during selenite reduction, we monitored the time course and spatial distribution of SeNP formation under aerobic conditions. When 1 mM selenite was added to E. coli cultures grown in TSB medium for 18 h, 86% of the selenite was reduced to red Se0 within 24 h (Figure a, b). The TEM images obtained at 2, 4, and 6 h after selenite addition revealed the emergence of electron-dense extracellular particles, likely corresponding to SeNPs (Figure c). These SeNPs were detected 2 h postincubation and predominantly localized in the extracellular space. They exhibited a consistent spherical shape, with particle diameter sizes remaining stable regardless of the incubation period (98 ± 20, 96 ± 32, and 96 ± 29 nm at 2, 4, and 6 h, respectively; Figure d). No evidence of cell lysis or morphological deterioration was observed in E. coli cultures for at least 6 h, suggesting that SeNP release occurs without damaging the cell envelope.

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Selenite reduction and SeNP formation by E. coli. (a) Photographic images showing red SeNP formation in test tubes after incubating E. coli cells with 1 mM selenite for varying periods. (b) Time-dependent changes in selenite consumption (blue) and Se0 production (red) by E. coli. Data represent the mean ± standard deviation of three replicates. (c) TEM images of negatively stained E. coli cells cultured with 1 mM sodium selenite for 2, 4, and 6 h. Scale bar: 1 μm. (d) Size distribution of SeNPs in E. coli cells after 2, 4, and 6 h of incubation with 1 mM sodium selenite. Particle diameters were measured from TEM images using ImageJ software, based on 68, 185, and 218 particles at 2, 4, and 6 h, respectively. Statistical significance was assessed using the Kruskal–Wallis test, and no significant differences were observed among the three groups.

Physicochemical Analysis of Ex-SeNPs Produced by E. coli

To investigate the physicochemical properties of the SeNPs released into the extracellular space, we purified SeNPs from the culture supernatant ofE. coli exposed to selenite. The purified SeNPs were analyzed using TEM and HAADF-STEM-EDS. No contamination by bacterial cells or MVs was observed in the purified SeNP samples (Figure S3). Negative staining, followed by TEM analysis, revealed electron-dense spherical particles (Figure a). EDS spectra of these particles indicated the presence of selenium, with characteristic absorption peaks at 1.4, 11.2, and 12.5 keV corresponding to the SeLα, SeKα, and SeKβ transitions, respectively (Figure b–d). The average size of the SeNPs, as measured using ImageJ software, was 93 ± 23 nm (Figure e). The SAED pattern exhibits a diffuse ring, indicating the amorphous nature of bacteriogenic SeNPs (Figure f). These characteristics are consistent with those of SeNPs produced by phylogenetically diverse bacteria, including another E. coli strain, Bacillus paramycoides, Stenotrophomonas maltophilia, and Comamonas testosterone, suggesting that bacterially synthesized SeNPs share conserved physicochemical features across species.

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Physicochemical analysis of Ex-SeNPs purified from E. coli culture supernatant. (a) TEM image showing the spherical morphology of Ex-SeNPs. Scale bar: 200 nm. (b) HAADF–STEM image highlighting electron-dense regions of SeNPs as bright white areas. (c) EDS elemental mapping indicating the presence of selenium in the particles. (d) EDS spectra of the nanospheres showing characteristic peaks of selenium. (e) Histogram of particle size distribution based on measurements of 190 particles from TEM images. (f) SAED pattern demonstrating the amorphous nature of SeNPs.

Analysis of SeNP Core and Surface Composition by XAFS and XPS

The Se L-edge PFY and TEY XAFS spectra of black elemental selenium (Se–Se bond), selenomethionine (Se–C bond), and Ex-SeNP are shown in Figure a. The peak position of the normalized XAFS spectra of Ex-SeNP was 1434 eV, which aligned with the characteristic absorption peak position of Se0 (black selenium) but differed from that of selenomethionine (1,436 eV, Se–C bond). These XAFS results indicate that the SeNP consist mainly of Se0 with Se–Se bonds.

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Se L-edge PFY and TEY XAFS spectra (a), normalized Se 3d XPS spectra (b), and XPS survey scans (c) of Ex-SeNPs (red), black selenium (blue), and selenomethionine (green). (d) Proposed particle composition model based on the spectral data in (a)–(c).

XPS analysis was performed to explore the surface-sensitive chemical properties of the SeNPs (∼5 nm from the surface). The Se 3d spectrum of Ex-SeNPs shifted to a higher binding energy compared to that of black selenium and was closer to that of selenomethionine (Figure b), indicating the prevalence of Se–C bonds on the SeNP surface. Additionally, increased carbon (C 1s) and oxygen (O 1s) levels were detected in SeNPs compared to that in black Se, suggesting that the outer surface of SeNPs was primarily composed of carbon and oxygen (Figure c). These results suggest that the core of the Ex-SeNPs produced by E. coli consists of Se0 with Se–Se bonds, whereas the surface, extending several nm, is enriched with carbon and oxygen, with features consistent with Se–C bonds (Figure d).

FT-IR Spectroscopy Characterization of Biomolecules Coating Ex-SeNPs

FT-IR spectroscopy was employed to characterize the biomolecular composition of the surface coating of Ex-SeNP. The absorption spectrum of Ex-SeNPs exhibited a peak at 1646 cm– 1 (Amide I), corresponding to CO and C–O stretching vibrations in proteins, and a peak at 1540 cm– 1 (Amide II), indicative of N–H and C–N vibrations in peptide bonds (Figure ). Absorption bands between 1450 and 1400 cm– 1 were attributed to CH2/CH3 and C­(CH3)2 bending vibrations from lipids and proteins. A peak at 1234 cm– 1 (Amide III) likely reflects C–N stretching and N–H bending vibrations, though it may also suggest PO vibrations associated with DNA, RNA, or phospholipids. Additional peaks at 1150 and 1026 cm– 1 correspond to C–O and C–OH stretching vibrations from alcohols, and C–O–C and C–H stretching vibrations from polysaccharides, respectively. In contrast, the absorption spectrum of Chem-SeNPs was markedly different (Figure ). A small peak at 1646 cm– 1 (Amide I) was likely derived from SDS, which was used as a capping agent during synthesis. These results indicate that the surface of Ex-SeNPs is coated with a complex layer of biomolecules, including proteins, carbohydrates, and lipids.

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FT-IR analysis of Ex-SeNPs. FT-IR spectra of Ex-SeNPs (red line) and chemically synthesized SeNPs (Chem-SeNPs, black line), showing differences in surface functional groups.

Proteomic Analysis of SeNP-Bound Proteins

Proteomic analysis was conducted to identify proteins associated with Ex-SeNP by comparing protein profiles from purified Ex-SeNPs with those from control precipitates obtained from an E. coli culture grown without selenite (NC) (Figure , Data set S1). SDS-PAGE analysis revealed that numerous proteins were bound to Ex-SeNPs, whereas only a few were detected in the NC (Figure S4). A total of 1034 proteins were identified in the Ex-SeNP sample, compared to 203 in the NC, with 190 proteins shared between the two samples. Subcellular localization analysis using DeepLocPro (https://biolib.com/KU/DeepLocPro/) indicated that Ex-SeNP-associated proteins originated not only from the outer membrane and extracellular space but also from the cytoplasm, cytoplasmic membrane, and periplasm. In contrast, proteins identified in the NC were predominantly derived from the cytoplasm, outer membrane, and periplasm, likely reflecting the protein composition of OMVs. Notably, a high degree of overlap was observed between Ex-SeNPs and NC for the outer membrane (75%) and periplasmic (51%) proteins. However, the overlap was markedly lower for cytoplasmic (10%) and cytoplasmic membrane (4%) proteins, suggesting that Ex-SeNPs associate with a broader spectrum of proteins than those typically found in OMVs. This broader association may reflect intrinsic properties of SeNPs that enable nonspecific adsorption of various proteins.

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Proteomic analysis of Ex-SeNP-bound proteins. Subcellular localization of proteins associated with Ex-SeNPs and NC samples (precipitates from selenite-free E. coli cultures), as identified by proteomic analysis. Proteins enriched in Ex-SeNPs are shown in light blue, those in NC are shown in green, and proteins detected in both are shown in magenta.

Characterization of Lipid Membrane Surrounding Ex-SeNPs

To confirm the presence of a lipid membrane surrounding the Ex-SeNP, the particles were stained with the lipid-staining fluorescent dye FM4-64 and observed using fluorescence microscopy. The fluorescence images revealed clear signals for FM4-64-stained Ex-SeNPs (Figure a, middle row). In contrast, no fluorescence was detected in the unstained Ex-SeNPs, which served as a NC (Figure a, top row). Quantitative analysis indicated that the fluorescence intensity per unit area of the FM4-64-stained Ex-SeNPs was significantly higher than that of the unstained controls (Figure b). Furthermore, the fluorescence intensity of FM4-64 was markedly lower than that of Ex-SeNPs/FM4-64 (Figure b). In comparison, Chem-SeNPs stained with FM4-64 exhibited fluorescence signals both at the particle sites and in the surrounding area, likely owing to nonspecific binding and residual background fluorescence (Figure a, bottom row, Chem-SeNPs/FM4-64). A comparative analysis of the ratios of fluorescence intensity per area (SeNPs vs background, meaning the outside of the particles) revealed significantly greater FM4-64 accumulation on Ex-SeNPs than on Chem-SeNPs (Figure c). These results indicate that lipids are present in regions accessible to FM4-64, supporting the presence of a lipid membrane surrounding the Ex-SeNPs.

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Lipid membrane components associated with Ex-SeNPs. (a) Bright-field and fluorescence microscopic images of SeNPs. The top, middle, and bottom rows show Ex-SeNPs without FM4-64 staining (labeled “Ex-SeNPs”), FM4-64-stained Ex-SeNPs (Ex-SeNPs/FM4-64), and FM4-64-stained chemically synthesized SeNPs (Chem-SeNPs/FM4-64), respectively. Each row presents, from left to right: bright-field image, fluorescence image, and merged image. Scale bar: 10 μm. (b) Fluorescence intensity per area for Ex-SeNPs and FM4-64 alone. Each gray dot represents the mean value from five fields of view in one independent experiment. Horizontal black bars indicate the average of the three independent experiments. Fluorescence intensity data were log10-transformed prior to analysis to reduce skewness and stabilize variance. Statistical comparisons were performed using Welch’s t test. **p-value <0.005 compared with Ex-SeNPs/FM4-64. (c) Fluorescence intensity ratios (SeNPs vs background) for Ex-SeNPs and Chem-SeNPs. Background refers to regions excluding SeNPs. Data were log10-transformed prior to analysis, and Welch’s t test was used for statistical comparison. **p-value <0.01. (d) Gas chromatography (GC) chromatograms of fatty acids extracted from E. coli cells (upper panel) and Ex-SeNPs (lower panel). Identified fatty acid derivatives include myristoyl (14:0), palmitoleoyl (16:1), palmitoyl (16:0), stearoyl (18:0), and cis-vaccenoyl (18:1). Cyclopropane fatty acid derivatives from 16:1 and 18:1 are denoted as 17:0c and 19:0c, respectively.

To further characterize the lipid membrane surrounding the Ex-SeNPs, the fatty acid compositions were analyzed using GC/MS after lipid extraction from Ex-SeNPs and E. coli cells cultured without selenite. The analysis revealed distinct fatty acid peaks, confirming the presence of lipids associated with the Ex-SeNPs (Figure d). The fatty acid profile of Ex-SeNP resembled that of phospholipids from E. coli cell membranes, although the peaks for stearic acid (18:0) and cis-vaccenic acid (18:1) were more prominent than those in E. coli cells. These results provide additional evidence that Ex-SeNPs are enveloped by E. coli cell membrane-derived components.

MV-like Structures Encapsulating Ex-SeNPs Identified via Reducing Reagent Treatment

To examine the morphological characteristics of the lipid membrane surrounding Ex-SeNPs, purified Ex-SeNPs were resuspended in PBS (0.8 mM Se0 equiv) and treated for 1 min at room temperature with increasing DTT concentrations. The characteristic red color of the Ex-SeNPs suspension gradually faded to near transparency with higher DTT levels (Figure a), indicating reduction and solubilization of the Se0 core. TEM imaging (Figure b) showed that Ex-SeNPs were mostly spherical at 0 mM DTT, became slightly deformed at 2 mM (selenium signals confirmed by HAADF-STEM-EDS; Figure S5), and appeared smaller above 5 mM. Concomitantly, MV-like structures increased with DTT concentration and dominated at 100 mM (Figure b,c and Table S2). Many of them displayed bilayer membranes typical of OMVs, , whereas some exhibited double-lamellar envelopes resembling outer-inner MVs (O-IMVs) (Figure b). ,

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Formation of MV-like structures encapsulating Ex-SeNPs upon DTT treatment. (a) Photograph of SeNPs after incubation with varying concentrations of DTT solution. (b) Representative TEM images (two per condition) of Ex-SeNPs after incubation with varying concentrations of DTT solution. White and red arrows indicate OMVs and O-IMVs, respectively. Scale bar: 100 nm. (c) Relative abundance of spherical SeNPs (gray), deformed SeNPs (striped), and MV-like structures (black), as observed by TEM after incubation with varying concentrations of DTT. (d) Size distribution of MV-like structures, based on diameter measurements of 54 particles using ImageJ software. (e) Two representative TEM images of ultrathin sections of MV-like structures. Scale bar: 50 nm.

The MV-like structures generated at 100 mM DTT were isolated by ultracentrifugation. TEM analysis showed a mean diameter of 79 ± 26 nm (Figure d), comparable to Ex-SeNPs (93 ± 23 nm; Figure e). Consistent with the observations in Figure b, the isolated MVs included both OMV- and O-IMV-like morphologies (Figure S6). Ultrathin sections of the sample revealed membrane layers ∼5 nm thick (Figure e), typical of bacterial cell envelopes.

To further characterize their composition, we analyzed cell-envelope markers in both isolated MV fractions and whole Ex-SeNP preparations. Peptidoglycan was detected in the MV fractions obtained after DTT treatment (Figure S7), consistent with previous findings of peptidoglycan-containing O-IMVs in Acinetobacter baumannii. To assess the outermost surface, whole Ex-SeNP preparations were subjected to a Limulus amebocyte lysate assay, which sensitively detects the Lipid A moiety of lipopolysaccharides. The assay confirmed the presence of lipopolysaccharides in the whole Ex-SeNP preparations (Figure S8). These results indicate that the Ex-SeNPs are associated with both peptidoglycan and outer membrane components, further supporting their encapsulation by bacterial envelope-derived bilayers. While this is consistent with envelope-dependent release, it does not by itself distinguish between vesiculation driven by specific envelope components and release emerging from nanoparticle–membrane physicochemical interactions.

Role of Cell Envelope Integrity in Ex-SeNP Secretion

To further investigate the link between SeNP extracellular secretion and the cell membrane, we analyzed SeNP formation in envelope integrity-related gene (ompC, ompA, tolA, and nlpI) deletion mutants. − Growth in the absence of selenite and phase-contrast microscopy revealed that the fraction of ghost cells, an indicator of compromised cell integrity, was higher in the mutants than in the wild-type (WT) (Figure S9a,b). All mutants reduced selenite and produced red SeNPs, with a slight variation in efficiency compared to the WT (Figures S9c and 8 a). Furthermore, optical density measured 24 h after selenite addition did not show drastic changes in either WT or mutant strains (Figure S9d), arguing against large-scale lysis as the primary explanation for reduced extracellular SeNP detection. After centrifugation, the WT supernatant exhibited a faint orange-red color, whereas the mutant supernatant remained clear and yellowish, indicating decreased Ex-SeNP secretion (Figure a). Conversely, mutant resuspended pellets (especially those of ΔompC and ΔtolA) retained a stronger red color than WT, indicating that more SeNPs were associated with mutant cells. Phase-contrast microscopy revealed frequent ghost cells and crystal-like deposits (most likely SeNPs) in these mutants, particularly ΔompC and ΔtolA (Figure b,c). These phenotypes of ΔompC and ΔtolA were restored to WT-like levels by complementation with plasmids expressing the corresponding WT genes (Figure S10). Collectively, the data indicate that the decreased abundance of extracellular SeNPs in ΔompC and ΔtolA is unlikely to result from impaired selenite reduction, gross growth defects, or large-scale lysis.

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Comparison of Ex-SeNP production in WT and mutant strains. (a) Visual comparison of whole cultures, culture supernatants, culture pellets, and resuspended pellets from WT and mutant strains. (b) Phase-contrast microscopic images of resuspended pellets from WT and mutant strains with (left panels) or without (right panels) selenite. Scale bar: 5 μm. White and red arrows indicate intracellular particle accumulation and ghost cell-like structures, respectively. (c) Percentage of bacterial cells containing visible particles, based on phase-contrast microscopy. Quantification was performed on 117, 347, 303, 430, and 455 cells for WT, ΔompC, ΔompA, ΔnlpI, and ΔtolA strains, respectively.

To determine whether the SeNPs were localized intracellularly or on the cell surface, we performed TEM, HAADF–STEM, and EDS analyses. These observations revealed marked intracellular accumulation of SeNPs in both ΔompC and ΔtolA strains (Figure ). TEM images of ultrathin sections clearly showed electron-dense SeNPs within the cytoplasmic compartment of these mutants (Figure h,l). In contrast, WT exhibited only Ex-SeNPs (Figure a–d). Field-wide quantification of SeNP localization from negative-stain images showed that particles were predominantly extracellular in WT cultures (Figure S11a), in agreement with ultrathin-section data. Notably, intracellular SeNPs in ΔompC and ΔtolA were larger and more heterogeneous in size compared to the Ex-SeNPs produced by WT (Figure S11b). These findings indicate that the integrity of the cell envelope is crucial for the proper export of SeNPs, and its disruption leads to intracellular retention and abnormal SeNP accumulation.

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Subcellular localization of selenium in (a–d) WT, (e–h) ΔompC, and (i–l) ΔtolA strains cultured with sodium selenite. (a, e, i) TEM images of negatively stained cells, (b, f, j) HAADF–STEM images highlighting electron-dense regions as bright white areas, (c, g, k) EDS elemental maps indicating selenium distribution, and (d, h, l) TEM images of ultrathin sections. White arrows indicate SeNPs. Scale bar: 500 nm.

To assess whether this defect also affects cellular tolerance, we determined the MICs of sodium selenite for WT, ΔompC, and ΔtolA strains at 18 and 44 h. ΔompC showed MICs of 30 and 40 mM, whereas ΔtolA showed 20 and 25 mM at the respective time points. In contrast, the WT strain grew at concentrations up to 100 mM, indicating an MIC > 100 mM. These results demonstrate that both mutants have markedly reduced tolerance under selenite exposure, which may reflect the additional stress imposed by intracellular SeNP accumulation when export is impaired.

Discussion

From an applied perspective, SeNPs have attracted growing interest as functional biomaterials due to their antimicrobial and anticancer properties, biocompatibility, and potential use in drug delivery. − However, the biological mechanisms underlying their secretion and membrane encapsulation remain poorly understood. In this study, we observed membrane-like structure-enveloped Ex-SeNPs during selenite reduction in E. coli. Our results suggest that these Ex-SeNPs are not merely inorganic particles, but are released along with cell-derived lipid, protein, polysaccharide, peptidoglycan, and lipopolysaccharides components. These findings provide a novel biological framework for understanding the mechanisms underlying the export of SeNP.

The intact morphology of WT cells during SeNP release (Figure ) suggests that the export process does not involve cell lysis, but may instead occur through a membrane-mediated process. This finding contrasts with previous reports in which SeNP release was associated with cell rupture in other bacterial systems. ,, In E. coli, the size distribution of Ex-SeNPs in the WT strain remained constant over time (Figures and ). Ultrathin sections of WT cells revealed no detectable intracellular SeNPs (Figure d), likely due to rapid export immediately after particle maturation. These findings suggest that SeNP size may be tightly regulated during both formation and secretion. Supporting this notion, the mutant strains defective in SeNP export accumulated larger and more heterogeneous intracellular SeNPs (Figures , , and S11b), consistent with a particle size control mechanism linked to the release pathway. In contrast, broader SeNP size distributions have been reported in other bacterial species, such as Bacillus mycoides SelTE01 and S. maltophilia SeITE02. , Notably, in E. coli, no specific proteins were found to be enriched on the surface of Ex-SeNPs (Figure ). A similar lack of specificity in protein adsorption has also been reported for C. testosteroni. In contrast, Thauera selenatis utilizes a dedicated protein, SefA, which has been shown to control SeNP formation even in vitro.

Ex-SeNPs exhibited a three-layered structure: a selenium core, an inner layer of biomolecules, and an encapsulated membrane layer (Figures , , and ). This architecture supports a model where the selenium core is initially capped nonspecifically by cytoplasmic proteins, followed by a budding-like process that incorporates both the outer and inner membranes along with periplasmic components, ultimately forming a vesicle that possibly resembles O-IMV, , which is secreted extracellularly (Figure ). Although similar vesicle-like structures associated with SeNPs have been proposed in particular bacterial species, − direct ultrastructural evidence for such a three-layered SeNP architecture has not been previously reported. Whether this structure is unique to E. coli or has simply been overlooked in other organisms remains unclear. Nonetheless, our study provides a structural and mechanistic basis for further comparative studies of biogenic SeNPs across different bacterial species.

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Schematic model for Ex-SeNP formation and excretion in E. coli. Selenite is transported into the cytoplasm and reduced to Se0, primarily via the Trx and GSH systems. The resulting Se0 species nucleate, aggregate, and are nonspecifically capped by intracellular proteins to form SeNPs with an average diameter of approximately 100 nm. These protein-capped SeNPs are then secreted from the cell, encapsulated within a membrane-derived structure enriched in phospholipids with cis-vaccenic acid and stearic acid. The encapsulating membrane surface also contains polysaccharides, membrane proteins, peptidoglycan, and lipopolysaccharides. This Ex-SeNP excretion pathway is dependent on bacterial cell envelope integrity, which is maintained by proteins such as OmpC and TolA.

We demonstrate that the mutant strains ΔompC and ΔtolA, both deficient in genes essential for maintaining cell envelope integrity, fail to release SeNPs extracellularly. TolA is a key component of the Tol-Pal complex that spans the periplasm and physically links the inner and outer membranes. During cell division, TolA helps pull the OM toward the division septum, thereby preventing detachment of the OM and contributing to envelope stability. , Recent studies also suggest that TolA contributes to maintaining lipid asymmetry in the OM. OmpC, in turn, interacts with MlaA to form a channel that removes phospholipids mislocalized to the outer leaflet, thereby preserving lipid asymmetry. , Disruption of lipid asymmetry has been shown to promote OMV formation by altering membrane curvature. These findings suggest that proper lipid distribution and coordinated envelope remodeling may be a critical factor in SeNP export. Consistent with this interpretation, both ΔompC and ΔtolA mutants exhibited markedly lower minimal inhibitory concentrations for sodium selenite compared with the WT strain. Selenite is known to trigger oxidative stress by depleting glutathione and generating reactive oxygen species, and Ex-SeNP release may serve as a detoxification route that alleviates this burden. The reduced tolerance of the mutants therefore likely reflects not only their impaired envelope maintenance but also the additional cellular stress imposed by intracellular SeNP accumulation when export is blocked.

Although previous studies have reported increased OMV production in ΔompC and ΔtolA mutants, this does not necessarily correlate with enhanced export of particulate cargo such as SeNPs. OMV biogenesis typically involves localized budding of the outer membrane, whereas SeNP release may require a more elaborate vesiculation process that engages both the inner and outer membranes, reminiscent of the formation of O-IMVs. , The inability of these mutants to export SeNPs extracellularly may therefore reflect a failure in the coordinated remodeling of the entire cell envelope, rather than a simple defect in outer membrane vesiculation. This distinction raises the possibility that SeNP export may involve vesiculation events engaging both membranes, which could reflect a specialized pathway or, alternatively, more general remodeling processes. Although our observations argue against large-scale cell lysis as the main source of Ex-SeNPs, we cannot fully exclude that some membrane-like structures arise from spontaneous vesiculation or limited cell rupture.

In addition to these pathway-level considerations, physicochemical factors are also likely to contribute. The permeation of nanoparticles across membranes is governed by properties including size, shape, charge, hydrophobicity, and electrophilicity, as well as the biophysical characteristics of the membrane itself. Previous studies have shown that nanoparticle penetration can occur when the adhesive interaction between a particle and the membrane is sufficient to overcome the energetic cost of membrane bending. Consistent with this, our GC/MS analysis (Figure ) revealed an enrichment of unsaturated fatty acids, such as cis-vaccenic acid, in the membrane encapsulating Ex-SeNPs. These lipids are known to enhance membrane fluidity , and promote curvature, , potentially facilitating SeNP budding and export.

Membrane encapsulation and extracellular SeNP release may represent adaptive strategies for mitigating selenium toxicity in E. coli. By packaging SeNPs within MVs prior to export, cells may reduce intracellular oxidative stress caused by excessive selenium accumulation. OMVs and other extracellular vesicles are known to participate in diverse bacterial processes, including stress responses, , virulence factor transport, , and intercellular communication. − Our findings expand the functional repertoire of bacterial vesiculation, suggesting that membrane-mediated nanoparticle export may constitute a previously underappreciated mechanism of environmental detoxification.

Supplementary Material

es5c10008_si_001.pdf (1,004.5KB, pdf)
es5c10008_si_002.xlsx (127KB, xlsx)

Acknowledgments

We thank Prof. Tatsuo Kurihara of Kyoto University, Japan, for his valuable assistance with the lipid composition analysis using GC/MS. The E. coli K-12 BW25113 strain and the gene-disrupted derivatives used in this study were provided by the National BioResource Project (NBRP), Japan. This study was supported by KAKENHI grants from JSPS (JP20J40237 to A.O. and JP22H04823 to H.M.), a research grant from the Noda Institute for Scientific Research (to A.O.; Chiba, Japan), the Ritsumeikan Global Innovation Research Organization, the Program for the Fourth-Phase R-GIRO Research (to H.M.), and the Analysis and Development System for Advanced Materials (ADAM) of RISH, Kyoto University as a collaborative program.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.est.5c10008.

  • Table S1: Primers used in this study. Table S2: Counts of SeNPs, deformed SeNPs, and MV-like structures in TEM Images of DTT-treated samples. Table S3: MICs of sodium selenite for WT, ΔompC, and ΔtolA strains. Figure S1: Standard curve for SeNP quantification. Figure S2: Preparation and characterization of Chem-SeNPs. Figure S3: Representative TEM image of purified Ex-SeNPs. Figure S4: SDS–PAGE analysis of Ex-SeNPs and NC to assess protein composition. Figure S5: TEM image, HAADF–STEM image, and EDS map of deformed particles obtained after reduction with 2 mM DTT. Figure S6: TEM image of MV-like structures. Figure S7: Peptidoglycan detection using SLP reagent. Figure S8: LAL gel-clot assay showing the presence of lipopolysaccharides in Ex-SeNP preparations. Figure S9: Growth, ghost cell fraction, time course of selenite reduction, and optical density changes in WT and mutant E. coli. Figure S10: Complementation of Ex-SeNP formation deficiency phenotypes in ΔompC and ΔtolA strains by exogenous gene expression. Figure S11: Localization and size distribution of SeNPs in WT, ΔompC, and ΔtolA strains cultured with sodium selenite for 6 h. (PDF)

  • Data set S1: List of identified proteins associated with Ex-SeNPs and NC samples from proteomic analysis (XLSX)

⋈.

Graduate School of Agricultural Science, Tohoku University, 468–1 Aramakiaza-aoba, Aoba-ku, Sendai, Miyagi 980–8572, Japan

The authors declare no competing financial interest.

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