Leuconostoc mesenteroides AA001: A High-Efficiency Nitrite Degrader Facilitating Controlled and Safe Traditional Vegetable Fermentation

Abstract

In traditional vegetable fermentation, the lack of effective microbial community control often leads to excessive nitrite accumulation, a major food safety concern. To address this challenge, this study aimed to isolate and characterize a high-performance starter culture strain capable of simultaneously degrading nitrite and guiding safe, controlled fermentation. A strain of L. mesenteroides AA001 was isolated from traditionally fermented Yingcai. It exhibited strong nitrite-degrading activity, with degradation rates consistently exceeding 90% under various environmental conditions, and demonstrated robust environmental adaptability. When used as a starter culture in vegetable fermentation, L. mesenteroides AA001 significantly accelerated nitrite degradation and consistently maintained peak nitrite concentrations below the Chinese national standard limit (20 mg/kg), while also shortening the fermentation period. Moreover, inoculation with L. mesenteroides AA001 had no significant impact on most nutrients in fermented vegetables across 8 crucial nutritional indicators (comprising 22 specific parameters), and only β-carotene content shows differences. The sensory attributes of the inoculated samples are basically similar to those of the naturally fermented samples, except that they are significantly brighter in color. No harmful substances were detected among 16 tested safety indicators, and the profile of 46 major volatile flavor compounds showed no significant difference compared to the spontaneously fermented control. Microbial community profiling throughout fermentation revealed that early dominance by this strain rapidly established a lactic acid–driven anaerobic environment, effectively suppressing nitrite-accumulating microorganisms and steering the process toward a stable, safe, and flavor-consistent trajectory. Thus, L. mesenteroides AA001 is a safe starter strain that combines potent nitrite-degrading capacity with fermentation-guiding functionality, effectively ensuring safety and process controllability in traditional vegetable fermentation.

1. Introduction

Fermented vegetables represent a traditional preservation method with a history spanning thousands of years [1,2,3]. The process was not scientifically understood until the 19th century, when Pasteur elucidated the role of microorganisms in fermentation [4]. Today, fermented vegetables are widely consumed as functional foods prized for their distinctive flavor, low caloric content, and rich probiotic properties [5,6,7,8]. Research suggests that consuming fermented vegetables can improve gut health, enhance immunity, and offer antioxidant benefits. Lactic acid bacteria (LAB) are the dominant functional microorganisms in vegetable fermentation [9,10]. This group is taxonomically diverse, encompassing several genera such as Lactobacillus (e.g., L. plantarum), Leuconostoc (e.g., L. mesenteroides), Pediococcus (e.g., P. acidilactici), and Streptococcus (e.g., S. thermophilus) [11,12]. By using selected, beneficial strains as starter cultures, manufacturers can exercise greater control over the fermentation process, thereby improving product quality and safety [13,14]. For example, L. mesenteroides B1, a strain isolated by Jung et al., has been shown to act as an effective starter for kimchi fermentation. It accelerates the process, optimizes the microbial community structure and metabolite profile, and thus enhances flavor development and product stability [15].

L. mesenteroides, a lactic acid bacterium commonly associated with various plant surfaces, is a critical microorganism in spontaneous fermentations [15,16]. Compared to other lactic acid bacteria, this species initiates fermentation more rapidly and efficiently converts fermentable sugars into metabolites such as lactic acid, acetic acid, ethanol, and carbon dioxide via the heterolactic pathway [17,18]. Furthermore, metabolic activity produces a diverse array of compounds, including flavor precursors (e.g., diacetyl [19], acetaldehyde [20]), functional oligosaccharides (e.g., dextran [21], levan [21]), and bacteriocins (e.g., leucocin K7 [22], leucocin C-607 [23]). These metabolites contribute to the unique sensory characteristics, probiotic properties, and biopreservation capabilities of fermented products. Consequently, L. mesenteroides is a pivotal starter culture in food fermentation and exhibits significant potential for industrial applications.

During vegetable fermentation, nitrite is generated as a potentially harmful byproduct, typically peaking early in the process [24,25]. This poses significant challenges to both food safety and industrial-scale production. The toxicological risks of nitrite are twofold: oxidation of ferrous iron (Fe²⁺) in hemoglobin to ferric iron (Fe³⁺), forming methemoglobin and causing tissue hypoxia and neurological damage; and reaction with secondary amines under acidic gastric conditions to form N-nitroso compounds, many of which are potent carcinogens [26,27,28,29]. Therefore, screening for efficient nitrite-degrading strains is an effective and environmentally friendly strategy to reduce the risk. Bacillus subtilis sp. N4, isolated from natto, efficiently degrades nitrite via simultaneous nitrification and denitrification [30]. Similarly, Lactiplantibacillus plantarum PK25, isolated from traditional Sichuan pickles (China), achieves a nitrite degradation rate exceeding 90% within 24 h by activating pyruvate metabolism, suppressing fatty acid biosynthesis, initiating DNA repair, and accelerating carbon source uptake—processes that promote rapid acidification and drive chemical decomposition of nitrite [31]. The application of such strains can significantly lower peak nitrite concentrations, thereby enhancing the safety and quality of fermented vegetables. To develop a microbiome-compatible starter culture specifically suited for traditional vegetable fermentation, this study aimed to: isolate a highly efficient nitrite-degrading strain from naturally fermented vegetables; characterize its functional and ecological properties, and evaluate its efficacy in controlling nitrite accumulation during fermentation. In line with these objectives, A highly efficient nitrite-degrading strain was isolated from traditional fermented vegetables and identified as L. mesenteroides AA001. After the characterization of its biological properties and evaluation of its nitrite-degrading capability, the strain was applied as a starter culture for vegetable fermentation and compared with spontaneous fermentation. Furthermore, by analyzing the microbial community during the fermentation process, this study further clarified the intrinsic mechanism by which this strain mitigates nitrite accumulation.

2. Materials and Methods

2.1. Bacteria, Reagents, and Raw Materials

Chemical reagents were purchased from Macklin (Shanghai, China). MRS and Mueller–Hinton (MH) media were purchased from Qingdao Hopebio (Qingdao, China). The bacterial strains were sourced from the Microbiology Laboratory, Center for Livestock Product Quality and Safety Technology, Jilin Academy of Agricultural Sciences (Gongzhuling, China). LMM solid medium was prepared with the following composition (per liter): sucrose, 101.0 g; dipotassium phosphate, 4.0 g; yeast extract, 2.5 g; ammonium sulfate, 0.2 g; manganese sulfate, 0.2 g; sodium chloride, 0.6 g; and agar, 14.0 g. The mixture was dissolved in distilled water to a final volume of 1 L and sterilized by autoclaving at 121 °C for 16 min. The raw vegetables and fermentation auxiliaries were purchased from Ouya Supermarket, Linhe Street Store (Changchun, China). Three traditional fermented vegetable models were prepared using the following raw substrates:

Fermented Yingcai: prepared from leaves of Lepidium sp. (family Brassicaceae);

Dongbei suancai: prepared from heads of Chinese cabbage (Brassica rapa subsp. pekinensis, family Brassicaceae);

Radish kimchi: prepared from roots of white radish (Raphanus sativus, family Brassicaceae).

2.2. Isolation and Purification of Strains

A total of 32 naturally fermented vegetable samples—produced by local households using traditional spontaneous fermentation methods—were purchased from farmers’ markets in five cities of Jilin Province, China: Yanji (n = 8), Changchun (n = 6), Jilin City (n = 6), Baishan (n = 5), and Changbai Korean Autonomous County (n = 7). The samples included fermented Yingcai, cabbage kimchi, radish kimchi, pickled radish, and Dongbei suancai. From these samples, the target strains were isolated as follows: 1 mL of homogenate from each fermented vegetable sample was spread onto LMM agar plates under aseptic conditions. Plates were inverted and incubated aerobically at 28 °C for 48–72 h. Colonies surrounded by droplet-shaped viscous polysaccharides were selected and streaked onto MRS agar supplemented with 30 μg/mL vancomycin for isolation and purification. This process was repeated 2–3 times to ensure purity. Single colonies were then subjected to microscopic examination to confirm the acquisition of pure bacterial cultures.

2.3. Screening of Nitrite-Degrading Bacterial Strains

The isolated strains were inoculated at 3% inoculation volume into MRS liquid medium containing 150 μg/mL sodium nitrite, with plain MRS liquid medium serving as the blank control. The cultures were incubated at 30 °C with constant shaking at 140 rpm for 24 h. The nitrite content in the bacterial suspension was determined using the N-(1-naphthyl) ethylenediamine dihydrochloride method [32], and strains exhibiting a nitrite degradation rate exceeding 50% were selected. The calculation formula is as follows:

$$X={\displaystyle \frac{Y-Y1}{Y}}\times 100\%$$

X = the nitrite degradation rate, %; Y = the initial nitrite sodium content, mg/kg; Y₁ = the nitrite sodium content after fermentation, mg/kg.

2.4. 16S rRNA Gene Sequencing

Total DNA of the screened L. mesenteroides AA001 was extracted using a bacterial DNA extraction kit (Sangon Biotech, Shanghai, China). The extracted DNA served as a template for PCR amplification with universal primers targeting the 16S rRNA gene. The PCR products were analyzed via agarose gel electrophoresis, and a single, appropriately intense band of approximately 1500 bp was selected and subsequently sent to Sangon Biotech for sequencing.

2.5. Growth Curve

L. mesenteroides AA001 was cultured in MRS liquid medium at 30 °C and 140 rpm under constant-temperature shaking. Samples were collected every 4 h. Optical density at 600 nm was measured using a microplate reader, and after appropriate dilution, samples were spread onto MRS agar plates for colony counting following incubation [33].

2.6. Temperature Tolerance Test

L. mesenteroides AA001 was cultured in MRS liquid medium for 24 h, with samples collected at predetermined time points. Optical density was measured at 600 nm using a microplate reader (Multiskan SkyHigh, Thermo Fisher Scientific, Waltham, MA, USA).

2.7. Acid-Base Tolerance Test

L. mesenteroides AA001 was inoculated into a series of MRS liquid media pre-adjusted to different pH levels. The cultures were incubated at 30 °C with constant shaking at 140 rpm for 24 h. After incubation, samples were collected and the optical density of each suspension was measured at 600 nm using a microplate reader (Multiskan SkyHigh, Thermo Fisher Scientific, Waltham, MA, USA).

2.8. Salt Tolerance Test

L. mesenteroides AA001 was inoculated into MRS broth supplemented with 0%, 4%, 6%, 8%, or 10% (w/v) NaCl. The cultures were incubated at 30 °C with constant shaking at 140 r/min for 24 h. Absorbance at 600 nm was measured using a microplate reader(Multiskan SkyHigh, Thermo Fisher Scientific, Waltham, MA, USA).

2.9. Acid Production Test

L. mesenteroides AA001 was inoculated into MRS liquid medium and incubated at 30 °C with shaking at 140 rpm. Samples were collected every 4 h to measure pH. Uninoculated MRS served as the blank control.

2.10. Antagonistic Activity Assay

The antagonistic activity of L. mesenteroides AA001 was evaluated against six pathogenic indicator strains using the agar well diffusion method. Sterile culture medium was used as the negative control, and ciprofloxacin (25 μg/mL) served as the positive control. After incubation, inhibition zones were measured and interpreted in accordance with the CLSI M100 standard as follows: resistant (≤10 mm), intermediate (11–17 mm), and susceptible (≥18 mm) [34]. The indicator strains used were: Escherichia coli ATCC 25922, Salmonella enteritidis ATCC 13076, Pseudomonas aeruginosa ATCC 27853, Staphylococcus aureus ATCC 25923, Acinetobacter baumannii ATCC 19606, and Shigella dysenteriae ATCC 13313. All strains were obtained from the Jilin Academy of Agricultural Sciences.

2.11. Preparation of Fermented Vegetables

Fresh vegetable raw materials (Yingcai, Chinese cabbage, and white radish) were thoroughly washed and then air-dried in a clean environment to remove surface moisture. Radish was cut into uniform cubes of approximately 2 cm³, while Yingcai and Chinese cabbage were kept intact as whole plants. During fermentation, Yingcai and radish kimchi were mixed with traditional seasonings (e.g., salt, garlic, ginger, and chili peppers) before being packed into fermentation vessels; Dongbei suancai was placed directly into the vessels without any added seasonings. All fermentation containers and associated utensils were sterilized by autoclaving at 121 °C for 20 min to minimize exogenous microbial contamination.

The experiment included two treatment groups: (1) natural fermentation (NF), relying solely on the indigenous microbial community naturally present on the vegetable surfaces; and (2) inoculated fermentation, in which L. mesenteroides AA001 was added at an inoculation level of 1.0 × 10⁶ CFU per gram of fresh weight. Each group was fermented according to the respective local traditional practices for the specific vegetable type.

Strict anaerobic conditions were not required during fermentation; instead, vessels were sealed moderately (breathable sealing films) to allow carbon dioxide produced during fermentation to escape while limiting oxygen ingress and preventing surface mold growth or spoilage. All treatments were statically fermented in the dark at ambient temperature (25 ± 3 °C) for 30 days.

2.12. Nitrite Degradation Capability Test

2.12.1. Evaluation of Nitrite Degradation Capability in Culture Medium

L. mesenteroides AA001 was inoculated into culture media containing sodium nitrite at final concentrations of 50, 100, 150, 200, and 250 μg/mL, and incubated for 24 h under varying conditions (temperature: 5–30 °C; pH: 4.0–8.0; NaCl concentration: 2–6%). During incubation, samples were collected at predetermined time points, and the nitrite degradation rate was determined using the N-(1-naphthyl)ethylenediamine dihydrochloride (NED) method.

2.12.2. Nitrite Degradation by L. mesenteroides AA001 in Fermented Vegetables

Accurately weighed 5.0 g portions of homogenized fermented vegetables, including fermented Yingcai, Dongbei suancai, and radish kimchi, were each mixed with saturated borax solution and hot water at 70 °C, followed by heating in a boiling water bath for 15 min. After cooling, the mixture was adjusted to volume, and potassium ferrocyanide and zinc acetate solutions were sequentially added to precipitate proteins. The suspension was allowed to stand for 30 min, defatted, and filtered, with the initial filtrate discarded. Nitrite content in the resulting filtrate was determined using the NED method.

2.13. Sensory Analysis

A structured descriptive sensory analysis was employed to determine the sensory maturation period of fermented samples and to compare the key sensory attributes between two fermentation methods: inoculation with L. mesenteroides AA001 (LmAA001) and natural fermentation (NF). The analysis was conducted following the general framework of Quantitative Descriptive Analysis (QDA). Detailed procedures, including the definition of sensory attributes, a 9-point intensity scale, evaluation criteria (Supplementary Tables S1–S5), environmental conditions conforming to ISO 8589 standards [35], and sample presentation order and coding methods, are provided in the Supplementary Materials. The sensory panel consisted of 10 trained assessors who demonstrated good inter-rater reliability (ICC ≥ 0.80). The study protocol was approved by the Institutional Review Board of Jilin University (Approval No. JNK20240316-1) and was conducted in accordance with the Declaration of Helsinki. Written informed consent was obtained from all panelists prior to participation.

2.14. Physicochemical Characterization

The samples (5 g) were accurately weighed and then homogenized.

(a) The pH was measured directly using a pH meter (PHS-3C, INESA Scientific Instrument Co., Ltd., Shanghai, China) in accordance with the National Food Safety Standard for Determination of pH in Foods (GB 5009.237–2016). The English version of the standard was referenced from ChineseStandard.net.

(b) The total acidity (TA) was determined by titration in accordance with the National Food Safety Standard for Determination of Total Acidity in Foods (GB 12456–2021). A homogeneous mixture was prepared from the sample. Subsequently, 10 mL of the sample supernatant (or an appropriate aliquot) was transferred to a titration vessel, and 2–3 drops of phenolphthalein indicator were added. The mixture was titrated with standardized 0.1 M NaOH solution until a persistent pale pink endpoint was observed. The volume of NaOH consumed (V, in mL) was recorded, and the TA value was calculated using the following formula:

$$TA(g/100g)=(V\times M\times 90)/(m\times 10)$$

V = Volume of NaOH solution consumed (mL); M = Molarity of NaOH solution (mol/L); 90 = Molecular weight of lactic acid (g/mol); m = Mass of sample (g); 10 = Conversion factor for 100 g basis.

The results were expressed as grams of lactic acid equivalent per 100 g of sample (g/100 g).

(c) Lactic acid content was determined by HPLC (Waters e2695, Waters Corporation, Milford, MA, USA) after sample preparation (10.0000 g sample centrifuged, supernatant diluted to 50 mL, and filtered). Separation used a Venusil MP C18 column (4.6 × 250 mm, 5 μm) with isocratic elution (0.1% H₃PO₄-methanol, 97.5:2.5) at 40 °C, 1 mL/min, 210 nm detection, and 2 μL injection. The method was based on GB 12456-2021 (National Food Safety Standard for the Determination of Total Acidity in Food).

(d) Reducing sugar content was measured by Fehling titration. The method was based on GB 5009.7-2016 (National Food Safety Standard for the Determination of Reducing Sugars in Food). A 20 g sample was extracted with 50 mL water, clarified with 5 mL zinc acetate and 5 mL potassium ferrocyanide, diluted to 250 mL, and filtered after 30 min standing. The filtrate was titrated with Fehling’s solutions A/B (5 mL each) at boiling point (1 drop/2 s) until decolorization, recording the volume used.

2.15. Vitamin Analysis

(a) Vitamin C was determined by semi-micro titration. Sample (100 g) was homogenized with oxalic acid (100 g), and 20 g homogenate was diluted to 100 mL and filtered. Filtrate (10 mL) was titrated with 2,6-dichloroindophenol to a persistent pink endpoint (15 min).

$$VitaminC(mg/100g)=(V\times T\times D\times 100)/W$$

V = titrant volume (mL), T = titer (mg/mL), D = dilution factor, W = sample weight (g).

(b) Vitamin B1 was determined by HPLC (Waters e2695, Waters Corporation, Milford, MA, USA). Sample (10 g) was hydrolyzed in 0.1 mol/L HCl (121 °C, 30 min), adjusted to pH 4.0, and digested with enzymes (37 °C, 16 h). The digest was diluted to 100 mL, centrifuged, and 2 mL supernatant was derivatized with alkaline K₃Fe(CN)₆ and n-butanol. The butanol phase was filtered (0.22 μm) and analyzed by HPLC: C18 column (250 × 4.6 mm, 5 μm), sodium acetate-methanol (65:35), 0.8 mL/min, 30 °C, fluorescence detection (Ex 375/Em 435 nm), 20 μL injection. The method was based on GB 5009.84-2016 (National Food Safety Standard for the Determination of Vitamin B1).

(c) Vitamin B2 was determined by HPLC (Waters e2695, Waters Corporation, Milford, MA, USA). Sample (10 g) was hydrolyzed in 0.1 mol/L HCl (121 °C, 30 min), adjusted to pH 6.5, and digested with enzymes (37 °C, overnight). The digest was diluted to 100 mL, centrifuged, and the supernatant was filtered (0.22 μm). HPLC conditions: C18 column (150 × 4.6 mm, 5 μm), sodium acetate-methanol (65:35), 1 mL/min, 30 °C, fluorescence detection (Ex 462/Em 522 nm), 20 μL injection. The method was based on GB 5009.85-2016 (National Food Safety Standard for the Determination of Vitamin B2).

(d) Vitamin B6 was determined by HPLC-MS/MS (Triple Quad 5500+, Sciex, Redwood City, CA, USA). Sample (5 g) was hydrolyzed in 0.1 mol/L HCl (100 °C, 30 min) with internal standard (500 μL), adjusted to pH 4.8, and digested with enzymes under nitrogen (37 °C, 18 h). The digest was diluted to 100 mL, and 1 mL filtrate was diluted to 10 mL and filtered (0.22 μm). HPLC-MS/MS: PFP column, mobile phase A (ammonium formate-formic acid in water) and B (formic acid in methanol), 0.4 mL/min, 30 °C, 10 μL injection. MS: ESI+, MRM, 5500 V, 550 °C. The method was based on GB 5009.154-2016 (National Food Safety Standard for the Determination of Vitamin B6).

(e) Vitamin A was determined by HPLC (Waters e2695, Waters Corporation, Milford, MA, USA). Sample (5.0 g) was saponified (80 °C, 30 min) with ascorbic acid, BHT, ethanol, and KOH, then extracted twice. The combined ether layer was washed to neutrality, dehydrated (Na₂SO₄), concentrated (40 °C), dried under N₂, and reconstituted in methanol (10 mL). After filtration (0.22 μm), analysis used a C30 column with gradient elution (water/methanol), 20 °C, 0.8 mL/min, UV detection at 325 nm, 10 μL injection. The method was based on GB 5009.82-2016 (National Food Safety Standard for the Determination of Vitamins A, D, and E in Food).

(f) Folic acid was determined by UV spectrophotometry (UNICO). Sample (5.0 g) was hydrolyzed in phosphate buffer (121 °C, 15 min), digested with enzymes (36 °C, 16 h), and filtered. The culture medium was sterilized, inoculated, and incubated (36 °C, 20 h). Absorbance was measured at 540 nm with a spectrophotometer, using the blank control to set 100% transmittance. The method was based on GB 5009.211-2022 (National Food Safety Standard for the Determination of Folic Acid).

(g) β-carotene was determined by HPLC (Waters e2695, Waters Corporation, Milford, MA, USA). Sample (5.0 g) was saponified (53 °C) with ascorbic acid/ethanol (60 °C, oscillation), extracted twice with petroleum ether, and the combined organic phase was washed, dehydrated, and concentrated. The residue was dissolved in dichloromethane, filtered, and analyzed by HPLC-C30 with gradient elution (methanol-acetonitrile-water/MTBE), detection at 450 nm. The method was based on GB 5009.83-2016 (National Food Safety Standard for the Determination of Carotene).

2.16. Analysis of Trace Elements and Heavy Metals

Sample analysis was performed by MD-ICP-MS (PerkinElmer NexION® 2000G, PerkinElmer, Inc., Waltham, MA, USA). Briefly, 1.0 g of sample was digested in 5 mL of HNO₃ overnight, followed by a three-stage microwave program (120, 150, 190 °C). The cooled digest was degassed by ultrasound for 5 min and diluted to 50 mL. Analysis was performed on a PerkinElmer NexION 2000G ICP-MS in KED mode (RF power: 1600 W; plasma gas: 15 L/min; carrier gas: 0.80 L/min; auxiliary gas: 0.40 L/min; He: 4 mL/min). The method was based on GB 5009.268-2016 (National Food Safety Standard for the Determination of Multi-elements).

2.17. Dietary Fiber Analysis

Weigh 0.25 g of the sample and subject it to enzymatic hydrolysis using amylase and protease. Precipitate the hydrolysate with ethanol, then filter and wash the residue under suction. Dry the residue at 105 °C to constant weight and record the mass. Determine the protein and ash content. Soluble dietary fiber is quantified using a HPLC (Waters e2695 Separations Module and Waters 2489 UV/Vis Detector, Waters Corporation, Milford, MA, USA) equipped with an aqueous size-exclusion gel chromatography column. The method was based on GB 5009.88-2014 (National Food Safety Standard for the Determination of Dietary Fiber).

2.18. Crude Fat Analysis

Sample (2.0 g) was placed in a filter paper thimble and extracted with petroleum ether in Automatic Fat Extractor (SOX406, Xinzhi Instrument Co., Ltd., Shanghai, China) (6–10 h, until no oil spot). The solvent was recovered, evaporated to dryness, and the residue was dried (100 °C ± 5 °C, 1 h), cooled, and weighed to constant weight. Crude fat content was calculated. The method was based on GB 5009.6-2016 (National Food Safety Standard for the Determination of Fat).

2.19. Crude Protein Analysis

Sample (0.5000 g) was digested with CuSO₄ (0.4 g), K₂SO₄ (6 g), and H₂SO₄ (20 mL) at 420 °C (graphite digester) until clear green. After cooling, diluted to 100 mL. A 10 mL aliquot was distilled with NaOH (40%, 30 mL) and boric acid (2%, 25 mL) using Kjeldahl system (FOSS 8400, FOSS Analytical A/S, Hillerød, Denmark) (5 min), then titrated with 0.1 mol/L HCl to grayish-red endpoint. Crude protein content was calculated. The method was based on GB 5009.5-2016 (National Food Safety Standard for the Determination of Protein).

2.20. Flavor Compounds Analysis

Pickle flavor compounds were analyzed by HS-GC-MS (Agilent 7000D GC/TQ, 8890GC, 7697A HS, Agilent Technologies, Inc., Santa Clara, CA, USA) [36]. Sample (5.0000 g) was equilibrated at 70 °C (30 min) in a headspace vial. GC conditions: DB-FastFAME column (30 m × 0.25 mm × 0.25 μm), flow 1 mL/min, split 10:1. Oven: 40 °C (1 min) → ramp to 220 °C. HS syringe: 130 °C; transfer line: 150 °C.

2.21. Aflatoxin B1 Analysis

Sample (5.0000 g) was spiked with isotopic internal standard, oscillated, and extracted with acetonitrile-water via ultrasonication. After centrifugation, the supernatant was PBS-diluted, purified by immunoaffinity column (water wash → methanol elution). The eluate was nitrogen-evaporated, reconstituted, filtered, and analyzed for AFB1 by LC-MS/MS (Triple Quad 5500+, Sciex, Redwood City, CA, USA). The method was based on GB 5009.22-2016 (National Food Safety Standard for the Determination of Aflatoxin B Group).

2.22. Analysis of Pathogenic Microorganisms

(a) Clostridium botulinum was detected using an ELISA kit (Shanghai Xuanya Biotechnology Co., Ltd., Shanghai, China), with homogenized and centrifuged samples measured for OD values at 450 nm.

(b) Enterobacteriaceae, Salmonella, Staphylococcus aureus, Listeria monocytogenes, and Pseudomonas aeruginosa were identified using specific chromogenic or selective agar plates as follows:

Enterobacteriaceae: Chromogenic Enterobacteriaceae Agar (HB7018), supplied by Qingdao Hopebio (Qingdao, China);

Salmonella: Chromogenic Salmonella Agar (CRM004B), supplied by Huankai Microbial (Guangzhou, China);

Staphylococcus aureus: Baird-Parker Agar (HB7025), supplied by Qingdao Hopebio;

Listeria monocytogenes: Chromogenic Listeria Agar (CRM014P1), supplied by Huankai Microbial;

Pseudomonas aeruginosa: CHROMagar™ P. aeruginosa Agar (CHROMagar, Paris, France), following enrichment in mPA broth at 42 °C for 24 h.

Positive identification was based on the following criteria: blue colonies (Enterobacteriaceae); bright red colonies (Salmonella); magenta colonies with clear zones (S. aureus); bluish-green colonies (L. monocytogenes); and blue-green or blue-gray colonies exhibiting yellow-green fluorescence under 365 nm UV light (P. aeruginosa).

(c) Mold was cultured on Rose Bengal Agar plates (Huankai Microbial, Guangzhou), presenting red or pink colonies with filamentous margins (preliminarily distinguishable from the smooth, cream-colored yeast colonies).

2.23. Biogenic Amine Analysis

Biogenic amines were analyzed by HPLC according to GB 5009.208–2016 with minor modifications. Briefly, 1.000 g of frozen fermented vegetable sample was homogenized with 10 mL of 5% (w/v) trichloroacetic acid at 16,000 rpm for 4 min, followed by centrifugation at 12,900 rpm for 20 min at 4 °C. The supernatant was filtered (0.45 μm), and 2.0 mL was derivatized with 1.0 mL each of 2 mol/L NaOH and dansyl chloride (10 mg/mL in acetone) at 60 °C for 30 min in the dark. The reaction was quenched with 100 μL of 10% ammonia, followed by double extraction with 5 mL hexane. The aqueous layer was filtered (0.22 μm) and analyzed by HPLC-UV (254 nm) on a Waters e2695 system (Waters Corporation, Milford, MA, USA) using a C18 column (250 mm × 4.6 mm, 5 μm). The mobile phase (acetonitrile and 0.1 mol/L ammonium acetate, pH 4.0) was applied under gradient elution at 0.8 mL/min. Total biogenic amines were quantified via a mixed standard calibration curve; tyramine and histamine were quantified using their individual standards.

2.24. Microbial Analysis

Fermented vegetable samples were collected at 0, 1, 2, 3, 4, 5, 6, 7, 14, 21, and 28 days, immediately frozen in liquid nitrogen, and stored at −80 °C until further analysis. Total genomic DNA was extracted from each sample using the CTAB method. DNA integrity and purity were assessed by agarose gel electrophoresis and spectrophotometry (NanoDrop; A260/A280 ratio ≈ 1.8). The V3–V4 hypervariable region of the bacterial 16S rRNA gene was amplified by PCR using universal primers appended with unique barcodes. PCR products were purified, quantified, and pooled in equimolar amounts to construct an Illumina MiSeq™ PE300 sequencing library. Raw paired-end reads were first processed with Cutadapt v1.2.1 to remove adapter and primer sequences. Paired reads were then merged into full-length amplicons using PEAR v0.9.6, and demultiplexed according to their barcodes. Subsequently, PRINSEQ v0.20.4 was employed for quality filtering: low-quality bases (Q < 20) were trimmed, sequences containing ambiguous bases (N) were discarded, and reads shorter than 200 bp or exhibiting low complexity were excluded. Chimeric sequences were detected and removed using UCHIME v4.2.40 (implemented in USEARCH v5.2.236), and non-target amplification products were filtered out by BLAST v2.28 alignment against the RDP database. High-quality sequences were clustered into operational taxonomic units (OTUs) at a 97% similarity threshold using USEARCH. The most abundant sequence within each OTU was selected as the representative sequence and taxonomically classified using the RDP Classifier v2.12 with a confidence threshold of ≥80%. Based on the resulting OTU abundance table, alpha diversity indices were calculated, and beta diversity was assessed using Bray–Curtis dissimilarity. Community succession was visualized via UPGMA clustering and non-metric multidimensional scaling (NMDS). Additionally, the functional potential of the microbial community was predicted using PICRUSt v1.0.0.

2.25. Statistical Analysis

All experimental data were analyzed using one-way analysis of variance (ANOVA) with SPSS software (version 21.0; IBM Corp., Armonk, NY, USA). Post hoc comparisons between groups were performed using Tukey’s test or Student’s t-test, as appropriate. Results are expressed as mean ± standard deviation (SD). A p value < 0.05 was considered statistically significant.

3. Results

3.1. Screening and Characterization of Nitrite-Degrading L. mesenteroides AA001

From 32 fermented vegetable samples, 22 colonies exhibiting a viscous, droplet-like morphology were isolated on Leuconostoc mesenteroides Medium (LMM)—a morphology typical of Leuconostoc species. Taking advantage of the intrinsic vancomycin resistance of Leuconostoc, 15 suspected colonies were further screened on de Man, Rogosa and Sharpe medium (MRS) supplemented with 30 μg/mL vancomycin. These isolates were then inoculated into MRS liquid medium containing 150 μg/mL sodium nitrite, a concentration that mimics nitrite levels commonly observed during the early stage of natural vegetable fermentation. Four strains exhibited nitrite degradation rates exceeding 50%: AA001 (97.06%), AA007 (89.77%), AA002 (58.73%), and AA011 (58.66%) (Figure 1A). All four strains were Gram-positive, as confirmed by Gram staining. The cells were predominantly spherical or lenticular, arranged in pairs or short chains, and non-motile—morphological features consistent with the genus Leuconostoc (Figure 1B). 16S rRNA gene sequencing was performed on the four isolates, and a phylogenetic tree was constructed to determine their taxonomic affiliation (Figure 1C). All were classified within the genus Leuconostoc: strain AA001 as Leuconostoc mesenteroides, strain AA011 as Leuconostoc carnosum, and strains AA002 and AA007 as Leuconostoc gelidum.

The metabolic profiles of the strains were assessed using three biochemical tests: arginine hydrolysis, citrate utilization, and dextran production [37,38,39]. Among the four strains, only AA001 tested positive in all three assays (

Table 1. Metabolic profiles of the four strains based on three biochemical tests.

Metabolic Characterization Test	Strain AA001	Strain AA002	Strain AA007	Strain AA011
Arginine Hydrolysis	+	+	+	+
Citrate Utilization	+	+	−	−
Dextran Formation	+	−	−	+

Note: “+” indicates a positive result; “−” indicates a negative result.

). Given that carbon source utilization is crucial for microbial growth and competitiveness, the ability of each strain to utilize a panel of 20 carbon sources was evaluated (

Table 2. Carbon source utilization profiles of the four strains.

Carbon Sources	Strain AA001	Strain AA002	Strain AA007	Strain AA011
Galactose	+	+	−	+
Erythritol	−	+	+	−
Sorbitol	+	−	+	−
Cellobiose	+	−	+	+
Melibiose	+	+	−	−
Lactose	+	+	+	+
Maltose	+	−	+	−
Sucrose	+	+	+	+
Fructose	+	+	+	+
L-Arabinose	−	−	+	−
Arbutin	+	−	+	+
Trehalose	+	+	+	+
Mannose	+	+	+	+
D-Xylose	+	+	−	−
Raffinose	+	−	−	−
L-Sorbose	+	−	+	+
L-Rhamnose	−	−	−	−
Salicin	+	+	+	+
Amygdalin	−	+	−	−
Ribose	+	−	−	+

Note: “+” indicates a positive result; “−” indicates a negative result.

). Strain AA001 exhibited the broadest utilization profile, utilizing 16 out of 20 substrates (80%), compared to AA007 (65%) and AA002 and AA011 (55% each).

In summary, strain AA001 demonstrated superior performance in nitrite degradation, biochemical activity, and carbon source utilization, suggesting strong ecological competitiveness and high potential as a starter culture. Furthermore, L. mesenteroides is widely recognized for its rapid acidification, high lactic acid production, ability to generate favorable flavor compounds, and broad environmental adaptability. These characteristics render it particularly well-suited for vegetable fermentation compared to L. carnosum and L. gelidum. Based on these comprehensive advantages, this study selected strain AA001 as the research subject.

3.2. Environmental Tolerance and Growth Characteristics of L. mesenteroides AA001

Achieving efficient and safe vegetable fermentation requires microbial strains with high tolerance to harsh environmental conditions, such as high salinity, low pH, and elevated temperature. Accordingly, to evaluate the fermentation potential of L. mesenteroides AA001, we assessed the growth under these stress conditions, as well as its lactic acid production and ability to degrade nitrite.

3.2.1. Growth Curve of L. mesenteroides AA001

Growth dynamics serve as a core indicator of a strain’s physiological activity, with the growth curve providing a quantitative profile of its cell proliferation [40]. The strain entered the logarithmic growth phase between 4 and 12 h, marked by exponential growth. Subsequently, from 12 to 24 h, it transitioned into the stationary phase, with viable cell counts plateauing (Figure 2A).

3.2.2. Environmental Tolerance of L. mesenteroides AA001

L. mesenteroides AA001 exhibited a broad pH tolerance range, growing between pH 3.5 and 8.0, with optimal growth observed at pH 5.5–6.5 (Figure 2B). Growth was considered positive when OD 600 nm ≥ 0.2; this criterion was applied consistently across all subsequent growth assessments. The strain also demonstrated wide temperature tolerance, growing from 4 °C to 46 °C, with an optimum at 30 °C (Figure 2C). Regarding salinity, strain AA001 grew in NaCl concentrations up to 7% (w/v), exceeding the typical range used in vegetable fermentation (2–5%) and indicating strong potential to inhibit spoilage microorganisms (Figure 2D). Collectively, these results demonstrate that L. mesenteroides AA001 possesses robust adaptability to key fermentation parameters.

3.2.3. Acidification Profile of L. mesenteroides AA001

The acidifying capability of lactic acid bacteria plays a key role in shaping the flavor and preservation of fermented foods [14,41]. L. mesenteroides AA001 demonstrated rapid acidification, with a significant drop in pH during the initial 8 h of fermentation. The pH decreased to approximately 3.9 by 24 h and remained stable thereafter, indicating that the acidogenesis process had reached a steady state (Figure 2E).

3.2.4. Inhibition of Common Pathogens by L. mesenteroides AA001

Inhibiting pathogenic bacteria is an important function of fermentative strains, reducing contamination risk in fermented vegetables [25,42,43]. The inhibitory capability of the cell-free culture supernatant from L. mesenteroides AA001 was evaluated in vitro using the agar well diffusion method, performed in accordance with general antimicrobial susceptibility testing guidelines (CLSI M100). The supernatant produced significant inhibitory zones against all six tested pathogens: Escherichia coli ATCC 25922, Salmonella enteritidis ATCC 13076, Pseudomonas aeruginosa ATCC 27853, Staphylococcus aureus ATCC 25923, Acinetobacter baumannii ATCC 19606, and Shigella dysenteriae ATCC 13313 (Figure 2F). The inhibition zone diameters ranged from 14 to 21 mm, indicating strong antimicrobial activity. A positive control (e.g., nisin or lactic acid) showed larger inhibition zones, confirming the effectiveness of the assay.

3.2.5. Nitrite Degradation by L. mesenteroides AA001

Nitrite is a common byproduct of food fermentation and poses a significant threat to food safety because of its associated health risks [44,45]. As fermentation conditions are complex and variable, we evaluated the nitrite-degrading capability of L. mesenteroides AA001 across a range of temperatures (5–30 °C), pH (4.0–8.0), and NaCl concentrations (2–6%). Within 24 h, the strain degraded over 90% of sodium nitrite (50–250 μg/mL) under all tested conditions, with the exception of the 6% NaCl environment, where the rate remained above 80% (Figure 2G). The degradation kinetics revealed a rapid increase in activity, particularly after 16 h, leading to these high final rates (Figure 2H). In short, L. mesenteroides AA001 provided a stable and efficient nitrite-degrading capability for complex fermentation applications.

3.3. L. mesenteroides AA001 Effectively Degrades Nitrite in Vegetable Fermentation

We evaluated the ability of L. mesenteroides AA001 to control nitrite accumulation during vegetable fermentation when used as a starter culture. Using three model systems—fermented Yingcai (raw material: Lepidium sp., family Brassicaceae), Dongbei suancai (raw material: Chinese cabbage, genus Brassica, family Brassicaceae), and radish kimchi (raw material: white radish, genus Raphanus, family Brassicaceae)—we compared nitrite accumulation profiles between natural and inoculated fermentations.

The nitrite peak is a critical indicator of fermentation safety, where a lower value and earlier onset are desirable [25]. In all three tested vegetable fermentations, inoculation with L. mesenteroides AA001 accelerated the onset and reduced the magnitude of the nitrite peak compared to natural fermentation (Figure 3A–C). Specifically, in radish kimchi, the AA001 group reached a nitrite peak of ≤10 mg/kg on days 2–3, while in fermented Yingcai and Dongbei suancai, the peak occurred on days 2–3 as well. In contrast, the natural fermentation groups peaked later (days 3–6) with significantly higher nitrite levels ranging from 20 to 46 mg/kg. As a result, the use of L. mesenteroides AA001 as a starter culture resulted in nitrite peaks consistently below the Chinese National Standard limit of 20 mg/kg, thereby enhancing fermentation safety.

3.4. L. mesenteroides AA001 as a Starter Accelerates the Vegetable Fermentation

Fermentation speed is a critical determinant of the production cycle, as accelerating the process shortens production time, improves equipment utilization, and reduces overall costs. We compared the fermentation speed of natural fermentation versus inoculation with L. mesenteroides AA001 using fermented Yingcai as a model. The comparison was based on a comprehensive set of sensory (color, flavor, aroma), physicochemical (total acid, lactic acid, reducing sugar), and microbiological (lactic acid bacteria count) indicators.

3.4.1. Sensory Evaluation

Sensory evaluation serves as an important indicator for determining the optimal fermentation duration [46]. The sensory criteria for mature fermented Yingcai include a yellowish-green color, a characteristic fermented aroma, and a balanced sweet-and-sour taste without bitterness. Based on these criteria, the fermentation period required to reach maturity was significantly shorter in samples inoculated with L. mesenteroides AA001 (5–7 days) compared to those undergoing natural fermentation (11–14 days) (Figure 4A–C), indicating that this strain accelerates the fermentation process.

3.4.2. Physicochemical and Microbiological Analysis

An objective evaluation of the fermentation process enables a precise and quantitative presentation of the dynamics. To this end, the crucial parameters were dynamically monitored throughout fermentation: total acidity, pH, lactic acid concentration, reducing sugar content, and total lactic acid bacteria (LAB) count. Fermentation maturity was defined by the following criteria: pH ≤ 4.0, total acidity ≥ 6 g/kg, LAB count reaching 10⁹ CFU/g, and peak concentrations of lactic acid and reducing sugar [47,48,49,50,51]. Compared to the natural fermentation (NF) group (9–14 days), the group inoculated with L. mesenteroides AA001 (LmAA001) significantly accelerated the fermentation process, achieving maturity within 4–7 days. This reduced the fermentation period by 5–7 days (Figure 5A–F).

3.5. Comparison of L. mesenteroides AA001-Inoculated and Natural Fermentation

Fermented Yingcai was used as a model food to compare nutritional composition, flavor compounds, and sensory quality between starter culture-based fermentation with L. mesenteroides AA001 and natural fermentation.

3.5.1. No Significant Difference in Nutritional Composition Between L. mesenteroides AA001-Inoculated and Natural Fermentation

Different fermentation processes may lead to significant variations in nutritional composition. However, an ideal starter culture–based fermentation should achieve high consistency with natural fermentation in terms of nutritional profile. Nutritional equivalence is an important indicator of starter culture performance and its ability to mimic natural fermentation. In fermented Yingcai, there were no significant differences between natural fermentation and fermentation with L. mesenteroides AA001 as a starter culture across eight crucial nutritional indicators (comprising 22 specific parameters): vitamins (excluding β-carotene), trace elements, minerals, antioxidants, moisture content, dietary fiber, crude protein, crude fat, and amino nitrogen (Figure 6). Additionally, the two fermentation methods showed similar levels of free amino acids and comparable proportions of essential amino acids for humans (Figure 7B). In conclusion, the two fermentation methods were nutritionally equivalent, with the only difference being β-carotene content, which was higher in the L. mesenteroides AA001–inoculated fermented Yingcai.

3.5.2. No Significant Difference in Major Flavor Compounds Between L. mesenteroides AA001-Inoculated and Natural Fermentation

Flavor compounds in fermented vegetables are generated through microbial metabolism and enzymatic catalysis [46,51,52]. These compounds not only impart complex flavor profiles, including sourness, sweetness, umami, and aroma to fermented vegetables but also enhance the sensory quality of the product by modulating overall flavor balance [46]. Common flavor compounds include acids [53,54], alcohols [51], esters [55], aldehydes [56], as well as free amino acids that influence flavor [53]. Here, we compared the two fermentation methods and found no significant differences in the contents of 46 major volatile flavor compounds (including alcohols, esters, aldehydes, ketones, terpenes, sulfides, and isothiocyanates), seven major organic acids, and flavor-related free amino acids between natural and AA001-inoculated fermentation (Figure 7A–C).

3.5.3. No Significant Difference in Sensory Attributes Between L. mesenteroides AA001-Inoculated and Natural Fermentation

Sensory evaluation is an intuitive method for assessing the quality of fermented vegetables. We employed a structured descriptive sensory analysis based on the general framework of Quantitative Descriptive Analysis (QDA) to compare the two fermentation methods. No significant differences were observed between natural fermentation and AA001-inoculated fermentation in appearance, aroma, flavor, or texture, with the only exception being color (Figure 8). The sample fermented with L. mesenteroides AA001 exhibited a brighter color compared to that of natural fermentation. This starter culture can preserve the sensory characteristics of natural fermentation to the greatest extent.

3.6. Absence of Detectable Harmful Substances in L. mesenteroides AA001 Fermentation

Vegetable fermentation can lead to the formation of potentially harmful substances, including nitrites, biogenic amines, mycotoxins, and pathogenic bacteria [9,57,58]. Excessive intake of these compounds may pose significant health risks, making their monitoring essential for ensuring product safety and quality. In this study, heavy metal levels in all samples of L. mesenteroides AA001-inoculated fermented Yingcai complied with the limits set by Chinese national standards. No pathogenic bacteria or aflatoxin B₁ were detected, and biogenic amine levels, including histamine and tyramine, were all below established safety thresholds, confirming that the fermented products meet safety requirements for consumption (

Table 3. Detection of harmful substances (heavy metals, pathogenic bacteria, aflatoxin B₁ and Biogenic Amines) in fermented Yingcai inoculated with L. mesenteroides AA001.

Substances Hazardous		Limit (LOD)	Regulatory Maximum Level (ML)	A1	A2	A3	A4	A5
Heavy Metals	Pb	0.001	≤0.5 a	0.003	<0.001	<0.001	0.004	<0.001
(mg/kg)	Hg	0.001	≤0.05 a	<0.001	<0.001	0.001	<0.001	<0.001
	As	0.001	≤0.5 a	<0.001	0.001	<0.001	<0.001	<0.001
	Cr	0.001	≤1.0 a	0.002	<0.001	<0.001	<0.001	0.003
	Cd	0.001	≤0.1 a	0.001	<0.001	<0.001	0.002	<0.001
Pathogenic Bacteria	Enterobacteriaceae	—	≤100 CFU/g b	ND	ND	ND	ND	ND
(CFU/g)	Clostridium botulinum	—	Absence required c	ND	ND	ND	ND	ND
	Salmonella	—	Absence required c	ND	ND	ND	ND	ND
	Staphylococcus aureus	—	Absence required c	ND	ND	ND	ND	ND
	Listeria monocytogenes	—	Absence required c	ND	ND	ND	ND	ND
	Pseudomonas aeruginosa	—	Absence recommended d	ND	ND	ND	ND	ND
	Molds	10 CFU/g	≤103 CFU/g e	ND	ND	ND	ND	ND
Toxins (µg/kg)	Aflatoxin B1	0.1	≤5.0 f	ND	ND	ND	ND	ND
Biogenic Amines	Total biogenic amines	—	<100 g	79.27 ± 3.50	88.33 ± 6.58	71.87 ± 2.37	69.63 ± 8.81	70.18 ± 2.31
(mg/kg)	Histamine	—	≤50 g	24.12 ± 3.29	31.05 ± 3.91	24.71 ± 5.73	21.14 ± 3.24	17.60 ± 2.97
	Tyramine	—	≤50 g	27.31 ± 3.53	24.78 ± 3.65	18.33 ± 3.66	26.36 ± 3.40	21.91 ± 2.86

Notes: A1–A5 represent five independent production batches, with each batch analyzed in triplicate; ND: Not detected (below LOD); Regulatory and guidance references: ^a GB 2762–2022 National Food Safety Standard—Maximum Levels of Contaminants in Foods; ^b Industry hygiene criterion for fermented vegetables; ^c GB 29921–2021 National Food Safety Standard—Microbiological Criteria for Pathogenic Bacteria in Prepackaged Foods; ^d Recommended absence due to its role as an indicator of poor hygiene (not regulated in GB 29921 for vegetable products); ^e Molds ≤ 10³ CFU/g, as specified in DB51/T 978–2023 Sichuan Paocai; ^f GB 2761–2017 National Food Safety Standard—Maximum Levels of Mycotoxins in Foods (Aflatoxin B₁ ≤ 5.0 µg/kg for relevant plant-based products); ^g Histamine ≤ 50 mg/kg: the U.S. FDA action level for scombroid toxin prevention; tyramine ≤ 50 mg/kg: the EFSA-recommended threshold to prevent hypertensive crisis in individuals taking monoamine oxidase inhibitors (MAOIs); total biogenic amines < 100 mg/kg: this threshold is considered indicative of controlled fermentation in vegetables, as levels above this value may reflect poor hygiene (EFSA, 2011).

).

3.7. Microbial Community Succession

This study systematically investigated the microbial community dynamics at 11 time points (0–28 days) spanning the entire traditional Yingcai fermentation process, encompassing the initiation, primary fermentation, and maturation stages, through high-throughput sequencing of the 16S rRNA gene. The aim was to elucidate microbial succession patterns and their correlations with product safety and flavor development.

3.7.1. Microbial Community Composition Analysis

At day 0 (initial colonization phase), the dominance of Proteobacteria was primarily driven by Pseudomonas and Acinetobacter, both of which possess broad substrate utilization capabilities and rapid aerobic respiration kinetics, enabling them to rapidly colonize vacant ecological niches. The secondary abundance of Actinobacteria was attributed to early colonizers such as Arthrobacter and Gaiella. During days 1–6 (rapid succession phase), the sharp increase in Firmicutes was mainly due to the proliferation of Lactobacillus, Leuconostoc, and Weissella, whose relative abundances rose markedly. This period also saw a concurrent increase in Barnesiella. Notably, the explosive growth of Lactobacillus (days 1–4) and Leuconostoc (days 5–6) led to a rapid decline in dissolved oxygen and pH, indicative of homolactic fermentation that established an anaerobic, acidic microenvironment. This shift effectively suppressed obligate aerobes such as Pseudomonas and Flavobacterium, thereby explaining the concomitant decline of Bacteroidetes during this stage. From days 7–14 (stabilization and transition phase), the abundances of Lactobacillus and Leuconostoc peaked and subsequently declined, whereas Weissella remained relatively stable. Concurrently, Akkermansia, a genus within Verrucomicrobia known for its capacity to degrade mucin and peptidoglycan, increased in abundance. This suggests that Akkermansia utilized cell wall fragments released by lactic acid bacteria during their growth and lysis, marking the onset of a cross-feeding interaction within the community. During days 21–28 (maturation and stabilization phase), although the abundances of Lactobacillus, Leuconostoc, and Weissella decreased, they retained a substantial presence within the community (Figure 9A,B). Notably, facultative anaerobes such as Pseudomonas exhibited a modest increase in relative abundance during late fermentation, likely reflecting their ability to tolerate the low-pH, organic acid–rich environment. However, Pseudomonas remained a minor component of the microbial community and did not become dominant. Consistent with this, culturable Pseudomonas counts were extremely low (<10² CFU/g; Supplementary Table S6), and no pathogenic species—such as P. aeruginosa—were detected (Table 3). These results suggest that the observed increase in relative abundance likely stems from residual DNA, non-viable cells, or benign environmental strains, rather than active proliferation of viable populations. Consequently, the presence of Pseudomonas poses no food safety risk.

3.7.2. Functional Profiling of Bacterial Metabolic Potential

The enrichment of oxidative phosphorylation and the tricarboxylic acid (TCA) cycle (KEGG—Energy Metabolism) during the early fermentation stage confirmed the rapid aerobic colonization by Proteobacteria. From day 1 to day 6, functional signals associated with carbohydrate metabolism (COG category G and KEGG—Carbohydrate Metabolism) sharply increased, consistent with homolactic fermentation dominated by Lactobacillus, which drove environmental acidification and oxygen depletion. Between days 7 and 14, pathways related to cell wall biosynthesis (COG category M) and the metabolism of amino acids and glycans (KEGG—Amino Acid and Glycan Metabolism) were upregulated, corresponding to mucin degradation and cross-feeding activities mediated by Akkermansia. From day 21 to day 28, genes involved in inorganic ion transport (COG category P) and xenobiotic biodegradation (KEG—Xenobiotics Biodegradation and Metabolism) showed increased abundance, reflecting the utilization of trace elements and recalcitrant organic compounds by oligotrophic taxa such as Acidobacteria. This conferred greater metabolic redundancy and ecological stability to the mature microbial community. Collectively, the dynamic shifts in functional gene profiles recapitulated a clear ecological succession trajectory: aerobic respiration, fermentative acid production, polymer degradation, recalcitrant compound turnover. Furthermore, all samples exhibited strong enrichment in genes linked to carbohydrate metabolism [G] and energy metabolism [C], supporting efficient glycolysis-driven energy generation. In certain samples, functions related to amino acid metabolism [E] and secondary metabolite biosynthesis [Q] were notably enhanced, suggesting a potential role in flavor development. Overall, the functional profile was highly congruent with taxonomic composition, indicating that the structure of the microbial community directly governs the functional output of the fermentation process (Figure 10A,B).

4. Discussion

Fermented vegetables are rich in probiotics but may accumulate nitrite, a potential carcinogen, during production, posing a significant challenge for the industry [9]. Microbial degradation offers an efficient and eco-friendly solution that relies on screening for high-performance strains. A starter culture comprising a mixed culture of Lactobacillus and Staphylococcus xylosus, isolated by Hu et al. from traditional fermented foods, reduced nitrite residues by 54–68% in fermented sausage, demonstrating the practical value of this approach [59]. In this study, L. mesenteroides AA001, isolated from traditionally fermented Yingcai, exhibits a nitrite degradation rate exceeding 90%. When applied to vegetable fermentation, L. mesenteroides AA001 effectively reduced nitrite, preserved flavor and nutrition, and accelerated the fermentation process, thereby enhancing both product safety and quality. Notably, L. mesenteroides is inherently well-suited for vegetable fermentation. Reduced-salt kimchi inoculated with L. mesenteroides LA81 maintained excellent texture and flavor, further supporting the application potential of this species [60].

The fermentation process is often accompanied by harsh environmental conditions, such as high salinity and high acidity, which pose a severe challenge to the tolerance of starter cultures. Therefore, we further evaluated the environmental adaptability of L. mesenteroides AA001. This strain exhibited robust environmental adaptability, characterized by a broad growth temperature range (4–46 °C), wide pH tolerance (3.5–8.0), and the ability to proliferate in NaCl concentrations up to 7%. Notably, the typical NaCl concentration in fermented vegetables (2–5%) falls well within this growth range. Furthermore, this strain produces acid rapidly, reducing the pH value to below 4.0 within 24 h, which is crucial for the flavor of fermented foods and the inhibition of harmful bacteria. This performance is comparable to the ability of Lactiplantibacillus plantarum LP1–3 and Lacticaseibacillus paracasei LC1 to reduce the pH to below 3.9 within 24 h [61]. Crucially, its nitrite degradation efficiency is highly stable, remaining above 90% across various environmental conditions. Even under high-salinity stress (6% NaCl), the degradation rate remained above 80%. These findings collectively demonstrate the strong potential of L. mesenteroides AA001 as an excellent fermentation strain, offering a viable technical solution to mitigate nitrite accumulation in fermented vegetables.

To validate the efficacy of L. mesenteroides AA001 as a starter culture, it was applied to the fermentation of fermented Yingcai, Dongbei suancai, and radish kimchi. Compared with natural fermentation, inoculation with this strain exhibited a marked ability to suppress nitrite accumulation and accelerate its degradation. This effect likely stems from the strain’s rapid acidification and its potential enzymatic nitrite-reducing activity, which creates an unfavorable environment for nitrate-reducing bacteria while simultaneously promoting nitrite breakdown [31,62]. Consequently, AA001 inoculation not only reduced the nitrite peak—keeping maximum concentrations well below the Chinese National Standard limit of ≤20 mg/kg (specifically, below 10 mg/kg)—but also advanced the timing of the peak by 2–3 days. Moreover, the overall fermentation duration was significantly shortened: AA001-fermented products reached completion in 4–7 days, in stark contrast to the 9–14 days required for natural fermentations. This early and effective control of nitrite, coupled with accelerated fermentation kinetics, substantially enhances product safety and may shorten the window before safe consumption—a practical advantage for both household and industrial production. In terms of overall product quality, the AA001-fermented samples were comparable to naturally fermented ones in most nutritional and sensory aspects (appearance, aroma, flavor, and texture). Notably, they exhibited significantly higher β-carotenoid content and more vibrant coloration. This may be attributed to two factors: strain L. mesenteroides AA001 potentially secretes plant cell wall-degrading enzymes that facilitate carotenoid release from vegetable matrices, and the rapid acidification during early fermentation inhibits oxidative enzymes such as polyphenol oxidase, thereby reducing carotenoid degradation and browning [63,64]. However, direct evidence for enzyme production by AA001 remains to be investigated in future work. Safety assessments confirmed that none of the targeted harmful substances—including heavy metals, mycotoxins, pathogenic microorganisms, and biogenic amines—were detected above the method’s limit of detection, and all results complied with Chinese national food safety standards.

Microbial community succession and functional profiling revealed the intrinsic mechanism behind efficient nitrite degradation. During the early fermentation stage, aerobic bacteria, represented by Proteobacteria, were the primary source of nitrite. Acting as an ecological engineer, L. mesenteroides AA001 rapidly colonized and dominated the early fermentation. Its drastically enhanced carbohydrate metabolism rapidly lowered the pH and consumed oxygen, thereby promoting nitrite degradation and fundamentally inhibiting the growth of nitrite-producing bacteria. This driving force accelerated the community’s transition from an initial aerobic state to a Lactobacillus-dominated anaerobic ecosystem. Correspondingly, the functional gene profile shifted from aerobic respiration to metabolism centered on lactic acid fermentation, ultimately forming a structurally simplified but functionally robust mature community. Notably, although the relative abundance of Pseudomonas slightly increased in the late fermentation stage, it never became a dominant member of the microbial community. This observation does not indicate active proliferation or spoilage potential; rather, it likely represents a molecular imprint of community succession—either transient signal remnants from nutrients released during lactic acid bacteria lysis or background DNA from non-viable cells [65,66]. From the perspective of environmental control and microbial antagonism, the AA001 strain rapidly established a typical fermented environment characterized by low pH (3.0–4.0) and 2–5% NaCl, and likely produced antimicrobial compounds that exerted antagonistic effects against Gram-negative pathogens such as P. aeruginosa. This multi-layered inhibitory barrier, arising not only from the inherent antagonistic properties of Leuconostoc against harmful microorganisms—including pathogenic species—but also from the low-pH and moderately saline environment it established, effectively suppressed the proliferation of Pseudomonas and its potential spoilage-related metabolic activities [67,68]. Moreover, the spoilage capacity of Pseudomonas depends on the production of heat-stable extracellular enzymes, whose activity was likely impaired under the acidic conditions and by antimicrobial metabolites present in the fermented matrix [69]. This interpretation is further supported by safety assessments: all final products complied with national food safety standards for key parameters, including biogenic amines, heavy metals, mycotoxins, and pathogenic microorganisms such as P. aeruginosa. Therefore, L. mesenteroides AA001 likely established a “structure–function–safety” triad regulatory mechanism that not only blocked nitrite accumulation at its source by shaping microbial community succession and optimizing metabolic functionality, but also ensured the stability of the fermentation ecosystem and the safety of the final product through the establishment of a multi-layered inhibitory barrier, thereby achieving efficient detoxification and quality assurance throughout the fermentation process.

5. Conclusions

In conclusion, L. mesenteroides AA001, isolated from traditionally fermented Yingcai, demonstrates exceptional nitrite-degrading capacity coupled with broad environmental adaptability and rapid acidification. When used as a starter, it effectively suppresses nitrite accumulation below regulatory limits while preserving key nutritional and sensory attributes comparable to spontaneous fermentation. These features address a critical safety bottleneck in traditional vegetable fermentation without compromising authenticity. Although further scale-up and comparative studies are warranted, L. mesenteroides AA001 offers a promising microbial tool for enhancing the safety and standardization of naturally fermented foods.