Preliminary Study on the Inhibitory Effect and Mechanism of Oleic Acid in Cylindrospermopsis raciborskii

Abstract

Cylindrospermopsis raciborskii is a toxin-producing cyanobacterium that is easy to overlook. It has strong environmental adaptability and is currently spreading around the world and gradually dominating to form a persistent bloom, causing ecological and environmental risks and drinking water safety issues. In this study, we systematically investigated the inhibitory effects of oleic acid on C. raciborskii and elucidated the underlying mechanisms through morphological observation, physiological assays, and bioinformatics analysis. Our results demonstrated that oleic acid strongly inhibits the growth of C. raciborskii, with a 72 h half-maximal effective concentration (EC₅₀) of 0.903 mg·L⁻¹. At 1.6 mg·L⁻¹, oleic acid achieved an inhibition rate of 99.5% within 48 h, indicating rapid suppression of cyanobacterial growth. Physiological analyses revealed that oleic acid severely impaired photosynthetic activity, as evidenced by significant reductions in key parameters (rETRmax, α, Fv/Fm, and Fv/Fo) and altered photosynthetic pigment composition, suggesting structural and functional damage to the photosynthetic apparatus. Morphological observations further showed that oleic acid disrupted filament integrity, inducing cell shrinkage, cytoplasmic vacuolation, cell wall detachment, membrane rupture, and eventual cellular disintegration. Kyoto Encyclopedia of Genes and Genomes pathway enrichment analysis indicated that oleic acid interferes with multiple metabolic processes, including nutrient and cofactor synthesis, membrane transport, and signal transduction, ultimately triggering algal cell death. This study highlights oleic acid as a promising eco-friendly agent for mitigating C. raciborskii blooms, offering potential applications in ecological prevention and emergency bloom control.

1. Introduction

Cylindrospermopsis raciborskii is a toxic, cryptically blooming cyanobacterium characterized by high environmental adaptability. Unlike typical bloom-forming species, it is difficult to detect and monitor in time because it does not gather into clusters at the surface of the water during algal outbreaks [1,2,3,4]. Originally endemic to tropical and subtropical regions, C. raciborskii has recently expanded into temperate zones worldwide, where it frequently dominates phytoplankton communities, forming persistent blooms in diverse aquatic systems [5,6,7,8]. The proliferation of C. raciborskii disrupts aquatic ecosystem balance and poses severe risks due to its production of toxic secondary metabolites, including paralytic shellfish poisoning (PSP) and cylindrospermopsin (CYN). These toxins can bioaccumulate in aquatic organisms, potentially threatening human health through the food chain [9,10,11,12,13]. Consequently, there is an urgent need to develop efficient and eco-friendly strategies for controlling C. raciborskii blooms.

Currently, the prevention and control of C. raciborskii blooms face significant challenges. Traditional physicochemical methods, such as coagulation sedimentation [14,15] and membrane filtration [16,17], are effective but often entail high operational costs and risks of secondary pollution. In contrast, algal control through allelochemicals [18,19,20,21,22] combines the advantages of chemical and biological algal control and has the advantages of an obvious algal inhibition effect, a fast algal inhibition rate, low ecological hazards, and easy operation, which shows good prospects for application in the field of water bloom control. Previous studies have demonstrated the potential of chemosensory substances in inhibiting C. raciborskii blooms. Wu et al. (2013) [18] reported that pyrogallic acid effectively suppressed C. raciborskii growth in a dose-dependent manner, primarily through photosynthetic inhibition and oxidative damage. Liu et al. (2015) [19,20] further identified that N-phenyl-2-naphthylamine (EC₅₀-72 h = 1.02 mg·L⁻¹) targeted photosystem II (PSII) of C. raciborskii by disrupting photosynthetic electron transfer and reducing active reaction centers. Xu et al. (2017) [21] observed that linoleic acid (2–4 mg·L⁻¹) strongly inhibited C. raciborskii growth, inducing Q_A⁻ accumulation and blocking electron transfer in PSII, suggesting dual impacts on photosynthetic and oxidative systems. Complementing these findings, Kornelia Duchnik et al. (2021) [22] revealed that prolonged co-cultivation (35 days) with Lemna trisulca reduced both cellular and extracellular CYN concentrations, indicating allelopathic control potential. Collectively, these studies highlight allelochemicals as promising eco-friendly agents for C. raciborskii bloom mitigation.

Among various allelochemicals, aquatic plant-derived fatty acids have emerged as particularly promising algal control agents due to their ecological safety, environmental compatibility, and target specificity [23]. Structure–activity relationship studies reveal that fatty acid efficacy correlates strongly with the carbon chain length and degree of unsaturation. Notably, unsaturated fatty acids demonstrate superior inhibitory effects compared to their saturated counterparts at equivalent chain lengths, with potency increasing proportionally to the number of unsaturated bonds [23,24]. Experimental evidence confirms that representative fatty acids (nonanoic acid, oleic acid, and linoleic acid) exhibit broad-spectrum growth inhibition against multiple algal species, including Microcystis aeruginosa [25,26], Pseudokirchneriella subcapitata [27], and Anabaena sp. [28]. The algicidal mechanism involves multi-target synergistic actions: (1) membrane disruption through altered lipid composition and fluidity [29], specific ion channel interference causing K⁺ efflux [28], and Ca²⁺-mediated membrane damage potentiation by long-chain saturated fatty acids (C16–C22) [30]; (2) oxidative stress induction via reactive oxygen species (ROS) burst accumulation from radical chain reactions [31], leading to membrane lipid peroxidation, increased permeability, and biomolecular damage; (3) photosynthetic inhibition through PSII reaction center targeting (Q_A⁻ accumulation and electron transfer blockade) [21] and photosynthetic electron transport chain disruption (O₂⁻ overproduction) [26]; and (4) metabolic interference via the dysregulation of Ca²⁺-CaM kinase pathway-mediated protein phosphorylation [32]. These concerted actions ultimately culminate in algal cell death.

Previous studies by our research team [33] demonstrated that co-culture with Eichhornia crassipes significantly inhibited the growth of C. raciborskii. While direct use of live water hyacinth for algal control poses ecological invasion risks, extraction of its bioactive components offers a more sustainable alternative for C. raciborskii management. Comprehensive literature reviews [34,35,36,37,38] have identified that Eichhornia crassipes produces various bioactive compounds, including sterols, fatty acids, phenols, terpenoids, and alkaloids. Notably, unsaturated fatty acids consistently show higher relative concentrations and detection frequencies compared to other compound classes.

Building upon existing research and addressing the practical requirements of aquatic ecosystem management, this study focuses on oleic acid—a key bioactive component of Eichhornia crassipes with demonstrated algicidal properties. We systematically evaluated its growth-inhibitory effects on C. raciborskii and elucidated the underlying mechanisms through an integrated multidisciplinary approach, including the following: (1) ultrastructural morphological analysis, (2) physiological and biochemical profiling, and (3) bioinformatics-assisted metabolic pathway interrogation. This work not only advances our understanding of fatty acid-mediated algal inhibition mechanisms but also provides a foundation for developing novel eco-friendly algal control technologies.

2. Materials and Methods

2.1. Experimental Materials

Cylindrospermopsis raciborskii FACHB-1503 in this experiment was purchased from the Freshwater Algae Culture Collection at the Institute of Hydrobiology, Chinese Academy of Sciences (Wuhan, China). The cyanobacterial strain was aseptically cultivated in sterilized BG11 medium and cultivated in an illuminating incubator (LRH-400A-GE, Guangdong Macro-Thermo Scientific Instrument Co., Ltd., Shaoguan, China). Culture conditions were strictly controlled as follows: temperature of 25.0 ± 0.5 °C, light intensity of 4000 ± 200 lux, and a light–dark cycle of 12 h:12 h. The cultures were manually shaken three times daily at regular intervals to prevent algal cell adhesion and sedimentation. The algae were expanded every 15 days, and the cells in the logarithmic growth period were prepared for the experiments to simulate water quality during C. raciborskii blooms.

High-purity oleic acid standard (≥99%, HPLC grade, Shanghai Aladdin Biochemical Technology Co., Ltd., Shanghai, China) was used in this experiment. Preliminary toxicity tests confirmed that 0.1% (v/v) dimethyl sulfoxide (DMSO, AR grade, ≥99.5%, Titan Scientific Co., Ltd., Shanghai, China) showed no significant growth inhibition on C. raciborskii FACHB-1503. Therefore, DMSO was selected as the co-solvent for the preparation of a 1 mg/mL oleic acid working solution, which was freshly prepared under sterile conditions prior to each experiment to ensure compound stability.

2.2. Experimental Design

Log-phase cultures of C. raciborskii were adjusted to an initial density of (2.0 ± 0.1) × 10⁹ cells/L using sterile BG-11 medium to simulate bloom conditions. The algal suspensions were distributed into 500 mL sterile conical flasks (300 mL working volume) and exposed to eight oleic acid concentrations (0, 0.2, 0.4, 0.6, 0.8, 1.0, 1.2, and 1.6 mg·L⁻¹). Freshly prepared oleic acid–DMSO working solution (1 mg/mL) was added, and DMSO was supplemented to ensure consistent final DMSO concentrations across all groups. Three replicates were set for each concentration. During the culture process, the flasks were shaken three times daily at regular intervals and randomly repositioned to ensure uniform light exposure. Samples were collected every 24 h from 0 to 144 h to measure algal density, calculate inhibition rates, and record changes in algal liquid appearance. To investigate the inhibition mechanism, additional experimental groups were set at 0, 0.4, 0.6, 0.8, 1.0, and 1.2 mg·L⁻¹ oleic acid concentrations. Samples were regularly collected for algal filament length statistics, chlorophyll fluorescence parameter measurements, and photosynthetic pigment content determinations. The 1.6 mg·L⁻¹ treatment group underwent intensive sampling within the first 24 h for scanning electron microscopy, transmission electron microscopy observation, and metabolomics analysis. All experimental procedures were conducted under aseptic conditions with appropriate negative controls.

2.3. Experimental Indicator Measurement

2.3.1. Microscopic Observations of the Growth of C. raciborskii

The collected algal liquid samples were fixed with Lugol’s iodine solution, thoroughly mixed, and then left to stand in the dark for 2 h. Then, 0.1 mL of the sample was aspirated into a plankton counting frame (20 × 20 mm) and covered with a cover slip to ensure that the counting area was free of bubbles. It was then observed under an optical microscope (BX41, Olympus Corporation, Tokyo, Japan) and photographed using an algal identification and counting instrument (R100, Hangzhou Shineso Technology Co., Ltd., Hangzhou, China), and the length of the algal filaments was measured using ImageJ software (v1.53k; National Institutes of Health, USA; downloaded from https://imagej.nih.gov/ij/index.html on 12 May 2021)). The algal cell density was calculated using the following formula:

$$\mathrm{N} ={\displaystyle \frac{{\mathrm{C}}_{\mathrm{s}}}{{\mathrm{F}}_{\mathrm{S}}\times {\mathrm{F}}_{\mathrm{n}}\times \mathrm{V}}}\times {\displaystyle \frac{{\mathrm{L}}_{\mathrm{S}}}{{\mathrm{L}}_{\mathrm{a}}}} \times 1000$$

(1)

where N represents the algal cell density (cells/L), C_s represents the counting frame area (mm²), F_s represents the field of view area (mm²), F_n represents the number of fields of view counted for each sample, V represents the counting frame volume (mL), L_s represents the total length of the counted algal filaments (μm), and L_a represents the average algal cell length (μm).

The inhibition ratio is calculated using the following formula:

$$\mathrm{IR}=\left(1-{\displaystyle \frac{{\mathrm{N}}_{\mathrm{i}}}{{\mathrm{N}}_0}}\right)\times 100$$

(2)

where IR represents the inhibition ratio (%); N_i represents the algal cell density of the treatment group; N₀ represents the algal cell density of the control group.

At the same time, the length of the algal filaments was classified into three length categories: short filaments (0–50 μm), medium filaments (50–250 μm), and long filaments (>250 μm). Each sample was counted for 150–200 algal filaments, the average algal filament length was calculated, and the proportion of algal filaments in each length category was analyzed.

2.3.2. Ultrastructural Analysis of C. raciborskii by Electron Microscopy

Scanning electron microscopy (SEM) observation: The algal samples were centrifuged (5000 r/min, 15 min) to collect the algal cell pellets, which were subsequently washed three times with 0.1 mol/L Phosphate-Buffered Saline (PBS). Fixation was performed using 2.5% glutaraldehyde at 4 °C for 24 h under dark conditions, followed by three additional PBS washes. Gradient dehydration was then conducted through an ethanol series (30%, 50%, 70%, 80%, 90%, and 100%), with the absolute ethanol step repeated twice. The samples were pre-cooled at −20 °C for 24 h and then dried in a vacuum freeze-dryer for 24 h. The dried sample was mounted on a copper stage and sprayed with gold using a vacuum spray coater and, finally, observed using a high-resolution field emission scanning electron microscope (Merlin, Zeiss, Jena, Germany).

Transmission electron microscopy (TEM) observation: The algal samples were centrifuged (5000 r/min, 15 min) to collect the algal cell pellets, which were subsequently washed three times with 0.1 mol/L PBS. Fixation was performed using 2.5% glutaraldehyde at 4 °C for 24 h under dark conditions, followed by three additional PBS washes. Subsequently, the samples were fixed with 1% osmium acid solution for 2 h and rinsed with PBS. Gradient dehydration was then conducted through an ethanol series (30%, 50%, 70%, 80%, 90%, 95%, and 100%), after which pure acetone was utilized to treat the samples for 20 min. The samples were embedded with a mixture of embedding medium and acetone, heated at 70 °C overnight, and afterward sliced up to obtain 70–90 nm sections, which were stained for 10 min each with lead citrate solution and 50% ethanol-saturated uranyl acetate solution. The dried samples were finally observed using a 120 kV transmission electron microscope (Talos L120c, Thermo Fisher, Waltham, MA, USA).

2.3.3. Measurement of Photosynthetic Pigment Content

Chlorophyll a and carotenoid content: The algal samples were poured into a vacuum pump filter equipped with a 0.45 μm glass fiber filter membrane for filtration. The filter membrane was added to an 80% acetone solution, thoroughly shaken, and extracted at 4 °C in the dark for 24 h. The supernatant was then collected by centrifugation (5000 r/min, 15 min), and the absorbance of the extract was measured at wavelengths of 470, 630, 645, 663, and 750 nm. The mass concentrations (mg·L⁻¹) of chlorophyll a (Chl a) and carotenoids (Car) in the samples were calculated using Formulas (3) and (4), respectively.

$$\mathrm{C}\mathrm{h}\mathrm{l} \mathrm{a}=11.64 \times \left({\mathrm{OD}}_{663}-{\mathrm{OD}}_{750}\right)-2.16 \times \left({\mathrm{OD}}_{645}-{\mathrm{OD}}_{750}\right)+0.1 \times \left({\mathrm{OD}}_{630}-{\mathrm{OD}}_{750}\right)$$

(3)

$$\mathrm{Car}={\displaystyle \frac{(1000 \times {\mathrm{OD}}_{470}-2.05 \times \mathrm{Chl} \mathrm{a})}{245}}$$

(4)

Phycobiliprotein content: The algal samples were centrifuged to collect algal cell pellets (5000 r/min, 15 min) and washed twice with 0.1 mol/L PBS buffer; the algal cells were resuspended in PBS buffer and pre-cooled at −20 °C for 12 h; under ice-water bath conditions, an ultrasonic cell disruptor (JY92-2D, Ningbo Scientz Biotechnology Co., Ltd., Ningbo, China) was used to disrupt the cells for 10 min (power 300 W, ultrasonication 5 s, interval 5 s). The resulting homogenate was diluted to a constant volume with PBS buffer, and the crude enzyme solution was collected by centrifugation. The absorbance of the crude enzyme solution was measured at wavelengths of 565, 620, and 650 nm. The mass concentrations (mg·L⁻¹) of phycocyanin (PC), allophycocyanin (APC), phycoerythrin (PE), and phycobilin (PB) in the samples were calculated using the following formulas:

$$\mathrm{PC} = {\displaystyle \frac{{\mathrm{OD}}_{620}-0.7 \times { \mathrm{OD}}_{650}}{7.38}}$$

(5)

$$\mathrm{APC}={\displaystyle \frac{{\mathrm{OD}}_{650}-0.19 \times { \mathrm{OD}}_{620}}{5.65}}$$

(6)

$$\mathrm{PE}={\displaystyle \frac{{\mathrm{OD}}_{565}-2.8 \times \mathrm{PC}-1.34 \times \mathrm{APC}}{1.27}}$$

(7)

$$\mathrm{PB}=\mathrm{PC}+\mathrm{APC}+\mathrm{PE}$$

(8)

2.3.4. Measurement of Chlorophyll Fluorescence

Chlorophyll fluorescence measurements were conducted using a pulse-amplitude modulated fluorometer (Water-PAM, Walz, Effeltrich, Germany) to assess the photosynthetic activity of C. raciborskii. Fresh samples were immediately transferred to the instrument’s temperature-controlled cuvette with the detector gain standardized to 400. For light response curves (Light Curve), nine actinic light intensities (24, 79, 117, 179, 263, 401, 598, 849, and 1178 μmol photons m⁻² s⁻¹, 30 s each) were applied sequentially while recording the relative electron transport rate (rETR). Using the Eilers and Peeters model [39], we performed non-linear curve fitting (Origin 2018) to derive the maximum electron transport rate (rETRmax) and light utilization efficiency (α). Another sample was taken and treated in the dark for 15 min, and then the light-induced curve (Induction Curve) was measured under a photochemical light intensity of 849 μmol photons m⁻² s⁻¹, during which minimum fluorescence yield (Fo) and maximum fluorescence yield (Fm) were determined. These values were used to calculate the maximum quantum yield (Fv/Fm) and potential activity (Fv/Fo). All measurements were performed in triplicate with WinControl V3.25 supporting software.

$$\mathrm{Fv}/\mathrm{Fm}=(\mathrm{Fm}-\mathrm{Fo}) / \mathrm{Fm}$$

(9)

$$\mathrm{Fv}/\mathrm{Fo}=(\mathrm{Fm}-\mathrm{Fo}) / \mathrm{Fo}$$

(10)

2.3.5. Metabolomics Analysis

Non-targeted metabolomics was used to detect changes in the composition and content of metabolites during the algal suppression process. The algal samples were centrifuged to collect algal cell pellets (5000 r/min, 15 min), washed three times with 0.1 mol/L PBS buffer, rapidly cooled in liquid nitrogen for 10 min, and stored at −80 °C for 24 h. The samples were ground and sonicated with an appropriate amount of extraction solution and magnetic beads, followed by centrifugation to collect the supernatant, which was then vacuum-dried. The supernatant was resuspended with an appropriate amount of extraction solution and analyzed using the detection platform. The detection platform was an ultra-high-performance liquid chromatography–tandem high-resolution mass spectrometer. Data were collected using MassLynx V4.2, and data processing, such as peak extraction and peak alignment, was performed using Progenesis QI software (v2.4, Waters Corporation, Milford, MA, USA). Metabolite identification was conducted using the online METLIN database and public databases via Progenesis QI software (v2.4).

2.4. Statistical Analysis

Data processing and graphing were performed using Microsoft Excel 2019 and Origin 2018, respectively. The Probit program in SPSS Statistics 25 software was used to calculate EC₅₀, and statistical significance was accepted when p < 0.05 using SPSS Statistics 25 for correlation analysis and one-way ANOVA.

3. Results

3.1. The Inhibitory Effect of Oleic Acid on the Growth of C. raciborskii

Oleic acid demonstrated a concentration-dependent inhibitory effect on C. raciborskii growth (initial density: 2.39 × 10⁹ cells/L). Figure 1a,b show that, while algal density in the control group increased steadily with culture time, oleic acid-treated groups exhibited dose-dependent growth inhibition. The 72 h half-maximal inhibitory concentration (EC₅₀) was 0.903 mg·L⁻¹, with the 1.6 mg·L⁻¹ treatment achieving 99.5% inhibition within 48 h, demonstrating rapid and potent algicidal activity at higher concentrations. At subthreshold concentrations, C. raciborskii maintained normal growth potentially through cellular repair mechanisms. However, when concentrations exceeded the threshold, inhibition increased sharply, showing a characteristic dose–response relationship. After 7 days, control cultures remained green while treated cultures showed increased transparency with yellow precipitates (Figure 1c), suggesting that oleic acid not only inhibits proliferation but also induces structural damage, leading to cell lysis and sedimentation.

3.2. The Effect of Oleic Acid on the Morphological Structure of C. raciborskii

3.2.1. Observation Under an Optical Microscope

C. raciborskii is a multicellular filamentous cyanobacterium characterized by non-constricted or slightly constricted cell walls [40]. Quantitative analysis indicates that oleic acid treatment disrupts the filamentous structure of C. raciborskii, leading to significant fragmentation of the filaments into short segments. As shown in Figure 2a, after 24 h of exposure, the average length of algal filaments in each treatment group decreased in a dose-dependent manner. The average lengths of algal filaments in the control group and the oleic acid-treated groups at 0.4, 0.6, 0.8, and 1.0 mg·L⁻¹ were 85.54, 81.43, 77.89, 52.30, and 46.62 μm, respectively, and the proportion of short filaments (<50 μm) significantly increased from 50.0% to 52.7%, 63.8%, 67.6%, and 66.7%, respectively. Microscopic examination demonstrated characteristic pathological alterations, including filament dissociation, pigment degradation, and cellular atrophy (Figure 2b). The observed morphological damage pattern closely resembled that induced by Bacillus cereus L7 (ATCC 14579) [41], suggesting that oleic acid may exert its algal inhibitory effect by interfering with algal filament intercellular connections or cell wall synthesis pathways.

3.2.2. Scanning Electron Microscope (SEM) Observation

SEM observation demonstrated progressive surface alterations in C. raciborskii filaments following oleic acid treatment. As shown in Figure 3a–c, normally growing C. raciborskii exhibited smooth surfaces with uniform diameter and clearly visible cellular septa. After 4 h of exposure, initial morphological changes included filament twisting and localized breakage (Figure 3d–f). By 8 h, extensive surface roughening was observed, accompanied by the appearance of irregular flocculent deposits and partial cellular atrophy (Figure 3g–i). The damage progressed to complete filament fragmentation after 12 h, with the formation of short segments displaying pronounced surface wrinkling and collapse of cellular architecture (Figure 3j–l). These structural modifications closely resembled the damage patterns induced by Bacillus cereus L7 [41] and N-TiO₂ photocatalysts [42], suggesting that oleic acid similarly compromises cellular integrity through surface structure disruption. The time-dependent progression of surface damage correlates with the observed inhibition of algal growth, supporting the algicidal effect of oleic acid through the disruption of filament integrity.

3.2.3. Transmission Electron Microscopy (TEM) Observation

TEM observation revealed severe ultrastructural alterations in C. raciborskii following oleic acid treatment (1.6 mg·L⁻¹) (Figure 4). Under normal growth conditions, C. raciborskii possesses a complete cellular structure, including an ordered three-layer membrane structure (outer membrane–cell wall–cytomembrane), uniformly distributed cytoplasmic organelles, clearly visible thylakoid membrane structures and photosynthetic lamellae, as well as intact vacuoles and phycobilisomes [43,44] (Figure 4a–e). After treatment with 1.6 mg·L⁻¹ oleic acid, the algal cells exhibited progressive degradation. Initial changes included cell shrinkage, vacuole proliferation, and membrane disruption. Advanced damage featured complete thylakoid disorganization, cell wall rupture, cytoplasmic leakage, and eventual cellular disintegration (Figure 4f–o). As shown in Figure 4j, after 24 h of treatment, a large amount of cell matrix leaked out, leaving only incomplete membrane structures, and almost no structurally intact algal cells could be observed. These results confirm that oleic acid can disrupt the integrity of the cell membrane system and interfere with the normal functions of algal cells, ultimately leading to the complete disintegration of algal cell structure.

3.3. The Effect of Oleic Acid on the Photosynthetic System of C. raciborskii

3.3.1. Photosynthetic Pigment Content and Proportion

Chlorophyll a and carotenoids are the primary lipophilic photosynthetic pigments in C. raciborskii, participating in the processes of light energy capture, absorption, and transfer. Experimental data showed that oleic acid stress significantly reduced chlorophyll a and carotenoid content, exhibiting a clear dose–response relationship (Figure 5a). After 144 h of treatment, chlorophyll a levels decreased from 1.236 mg·L⁻¹ (control group) to 0.024 mg·L⁻¹ (1.2 mg·L⁻¹ oleic acid), while carotenoids declined from 0.833 mg·L⁻¹ (control group) to 0.043 mg·L⁻¹ (1.2 mg·L⁻¹ oleic acid). Notably, high concentrations (≥1.0 mg·L⁻¹) induced a characteristic shift in pigment ratios, with carotenoids becoming proportionally more abundant than chlorophyll a, which may be related to the protective accumulation of carotenoids as antioxidants under stress conditions [45,46]. This dynamic change suggests that oleic acid may influence algal cell photosynthetic efficiency through two pathways: on the one hand, it directly inhibits pigment synthesis, and on the other hand, it accelerates pigment degradation by inducing ROS bursts.

Oleic acid exposure induced significant alterations in the phycobiliprotein composition of C. raciborskii, causing both quantitative and qualitative changes (Figure 5b). After 144 h treatment, total phycobiliprotein content showed a dose-dependent decline from 10.989 μg·L⁻¹ (control group) to 0.459 μg·L⁻¹ (1.2 mg·L⁻¹ oleic acid). Notably, phycocyanin (PC) demonstrated greater susceptibility to oleic acid stress compared to allophycocyanin (APC) and phycoerythrin (PE), with a reduction pattern consistent with previous reports on fatty acid interactions [47]. Phycobiliproteins are located on the outer side of PSII. Light energy is captured by PE and then sequentially transferred to PC, APC, and chlorophyll a and, finally, to the D1 protein, where photochemical reactions convert light energy into energy used for cell growth and reproduction. This selective PC reduction impairs both light-harvesting capacity and energy conversion efficiency, directly contributing to the observed photosynthetic inhibition and growth suppression through compromised phycobilisome function and reaction center activity.

3.3.2. Photosynthetic Activity

Chlorophyll fluorescence parameters are sensitive indicators of algal cell photochemical activity. As shown in Figure 6, oleic acid significantly impaired PSII photochemical activity in C. raciborskii, as evidenced by rapid declines in key chlorophyll fluorescence parameters (rETRmax, α, Fv/Fm, and Fv/Fo) at high concentrations (0.8–1.2 mg·L⁻¹). The initial depression of these parameters indicates substantial damage to reaction centers and electron transport chains, similar to effects observed with other stressors like 4-nonylphenol [48], bisphenol A [49,50], and cadmium [51]. Notably, partial parameter recovery after 72 h occurred despite ongoing pigment degradation, suggesting activation of photoprotective mechanisms that optimize energy allocation and electron transport efficiency. This adaptive response highlights the photosynthetic apparatus’s plasticity under fatty acid stress, though ultimately insufficient to prevent growth inhibition. The temporal dynamics of fluorescence parameters provide critical insights into both the immediate impacts and acclimation responses to oleic acid exposure.

3.4. Metabolomics Analysis of C. raciborskii Exposed to Oleic Acid

Metabolomic analysis revealed profound metabolic reprogramming in C. raciborskii following 24 h of oleic acid exposure, with 1351 differentially expressed metabolites mapped to 124 metabolic pathways (Figure 7). The observed metabolic shifts demonstrate a multi-target inhibition mechanism: (1) the upregulation of membrane-protective metabolites (Zeaxanthin diglucoside, Galactosylsphingosine, and 5-Hydroxyectoine) reflects cellular responses to membrane destabilization [52,53,54], while (2) the downregulation of proliferation-related metabolites (Cyclic GMP, Dihydrofolic acid, and Guanosine diphosphate mannose) indicates impaired signal transduction [55,56], nucleic acid synthesis [57], and energy metabolism [58]. These systemic alterations collectively disrupt cellular homeostasis through four synergistic pathways, namely, membrane integrity loss, energy metabolism interference, nucleic acid synthesis inhibition, and signaling pathway blockade, ultimately leading to growth arrest. The metabolomic profile provides molecular-level evidence for oleic acid’s algicidal efficacy, revealing both the stress response mechanisms and the metabolic vulnerabilities exploited by fatty acid exposure.

The complex metabolic reactions and their regulatory mechanisms in organisms are mediated through an intricate network of genes, proteins, and metabolites. Our study demonstrates that oleic acid stress induces systemic alterations in the metabolic network of C. raciborskii, with significant dysregulation of the ABC transporter pathway identified as a primary target. ABC transporters, serving as central mediators of transmembrane transport, not only facilitate the translocation of metabolites (including carbohydrates, lipids, amino acids, and nucleotides) [59] but also participate in xenobiotic detoxification [60], thereby playing a pivotal role in cellular homeostasis maintenance. KEGG enrichment analysis (Figure 8) revealed that oleic acid disrupts ABC transporter function through a tripartite cascade: (1) increased membrane permeability and toxicant accumulation; (2) dysregulated transport of essential metabolites (e.g., amino acids and nucleotides); and (3) consequent disturbances in energy metabolism, signal transduction, and cellular integrity. These multi-level disruptions ultimately lead to intracellular environment imbalance, causing structural damage and functional impairment that manifests as growth inhibition or even cell death.

4. Discussion

The present study demonstrates that oleic acid exhibits rapid and potent inhibitory effects on C. raciborskii, with a 72 h EC₅₀ of 0.903 mg·L⁻¹—superior to that of many reported algicides, including linoleic acid (1.288 mg·L⁻¹) [21] and N-phenyl-2-naphthylamine (1.02 mg·L⁻¹) [19,20]. High-concentration treatment (1.6 mg·L⁻¹) achieved 99.5% growth inhibition within 48 h, accompanied by cell lysis and sedimentation (Figure 1c). These findings align with Nakai et al.’s [25] observations in Microcystis aeruginosa, confirming the broad-spectrum algicidal activity of long-chain unsaturated fatty acids (C18:1) against cyanobacteria.

Microscopic and ultrastructural observations revealed that oleic acid primarily targets cellular membranes in C. raciborskii, causing dose-dependent structural damage that progresses from initial filament fragmentation (light microscopy) to surface roughening and twisting (SEM), ultimately culminating in complete cellular disintegration (TEM) through rupture of the trilaminar membrane system (Figure 2, Figure 3 and Figure 4). This outside-in damage progression mirrors the membrane-targeting effects of ferulic acid in Microcystis aeruginosa [61], suggesting oleic acid similarly disrupts cellular integrity via (1) altered membrane potential and fluidity [28,61], (2) ABC transporter-mediated osmoregulatory dysfunction (Figure 8) [59,60], evidenced by abnormal bubble enlargement (Figure 4), and (3) specific inhibition of cell wall synthesis or intercellular junctions, as indicated by fracture patterns resembling Bacillus cereus L7 [41] and N-TiO₂ [42] treatments.

Functional assessment of the photosynthetic apparatus revealed significant impairment in C. raciborskii under oleic acid exposure, characterized by a biphasic ‘acute damage–partial repair’ response in PSII activity (Figure 6). The initial (<72 h) sharp decrease in Fv/Fm and rETRmax reflected the inactivation of the reaction center or the blockage of electron transfer, which was consistent with the Q_A-accumulation mechanism reported by Xu et al. [21], while the recovery of the parameters at the later stage might be due to the turnover of D1 protein or the activation of non-photochemical quenching (NPQ) [48,49,50,51]. Notably, the carotenoid/chlorophyll a ratio was elevated in the high-concentration group (Figure 5a), which may mitigate oxidative damage by activating the carotenoid cycle, consistent with its photoprotective function under oxidative stress [45,46]. The preferential degradation of PC in phycobilisomes (Figure 5b) supports the ‘phycocyanin–photosynthetic chain’ targeting hypothesis [47], which may be related to oleic acid-specific interference with the energy transfer efficiency of phycobilisomes.

Metabolomic analysis of C. raciborskii under oleic acid stress revealed comprehensive metabolic reprogramming (Figure 7 and Figure 8), demonstrating three key adaptive responses: (1) the upregulation of membrane-protective metabolites (e.g., Zeaxanthin diglucoside), indicating activated antioxidant defenses [52,53]; (2) the downregulation of nucleic acid precursors (Dihydrofolic acid) and energy carriers (GDP-mannose), suggesting cell cycle arrest [57,58]; and (3) significant enrichment of the ABC transporter pathway, whose dysfunction may simultaneously impair xenobiotic efflux and nutrient uptake [59,60]. These multi-target effects elucidate oleic acid’s superior algicidal efficiency compared to single-mechanism inhibitors and provide molecular validation for the fatty acid synergistic algal control hypothesis.

Our study systematically elucidates the multi-target algicidal mechanism of oleic acid against C. raciborskii through structural, functional, and metabolic analyses. The results reveal a complete mechanistic chain from initial structural damage to ultimate cell death: membrane disruption (SEM/TEM observations) triggers ion imbalance, subsequently followed by photosynthetic system impairment (PSII inactivation evidenced by fluorescence parameters), leading to energy metabolism obstruction (GDP-mannose downregulation) and signal transduction disruption (cGMP reduction), ultimately culminating in cell death.

Our findings demonstrate oleic acid’s potential for mitigating cyanobacterial blooms, especially in emergency scenarios requiring rapid response, while further formulation improvements (e.g., microencapsulation) could enhance its environmental stability for practical applications. However, despite its eco-friendly profile, the ecotoxicological risks of oleic acid to non-target aquatic organisms remain unclear. Future studies should evaluate these effects to ensure their safe and sustainable application.

5. Conclusions

This study confirms oleic acid as a highly efficient algicide against C. raciborskii, demonstrating exceptional efficacy (72 h EC₅₀ = 0.903 mg·L⁻¹) and rapid action (99.5% inhibition at 1.6 mg·L⁻¹ within 48 h). Its potency stems from a distinctive multi-target mechanism that simultaneously disrupts cellular membranes, impairs photosynthesis, and interferes with critical metabolic pathways. Compared to similar allelochemicals, oleic acid demonstrates superior efficacy due to this comprehensive mode of action. These findings highlight its strong potential for applications in ecological prevention and emergency bloom control, though further research should focus on formulation optimization and environmental safety assessments to facilitate practical use.