INTRODUCTION
Soil salinization-alkalization has emerged as a critical global environmental crisis, threatening agricultural sustainability and food security. Currently, over 1.08 billion hectares of salinized soil are distributed across >100 countries, with China facing severe challenges, particularly in regions like the Yellow River Delta where 620,000 hectares of saline-alkali land remain underutilized (Li et al. 2012, Zheng et al. 2020). By 2050, nearly 50% of the world’s arable land is projected to be degraded by salinity, causing crop yield losses of 18–43% (Ronga et al. 2019, Shah et al. 2022). Saline-alkali soils impose multifaceted stresses on plants, including osmotic imbalance, ionic toxicity (Na+ accumulation), hydroxyl ion (OH−) damage to root membranes, nutrient deficiencies (e.g., P, Fe, Zn), and oxidative stress, collectively inhibiting seed germination, seedling development, and physiological functions (Carillo et al. 2019, Wang and Zhang 2022).
Conventional remediation strategies—such as chemical leaching, synthetic fertilizers, and engineered soil treatments—are limited by high costs, environmental pollution, and unsustainable outcomes (Cui et al. 2020, Alvarez et al. 2021). For instance, excessive fertilizer use exacerbates soil structure deterioration and microbiota depletion (Liu et al. 2021). Consequently, there is an urgent need for eco-friendly and sustainable approaches to rehabilitate saline-alkali lands while enhancing crop resilience.
The limitations of conventional saline-alkali land remediation methods necessitate the exploration of sustainable biological alternatives. Microalgae emerge as a compelling solution due to their unique physiological adaptability and ecological functionality. Native halotolerant strains, such as Chlorella spp. isolated from the Yellow River Delta, exhibit exceptional resilience under high salinity stress (200–600 mM NaCl), accumulating biomass up to 1.2 g L−1 while actively reducing soil sodium content through extracellular polymeric substance secretion (El Arroussi et al. 2018, Yu et al. 2025). These indigenous microorganisms not only bind Na+ ions but also enhance soil organic carbon and aggregate stability, creating a synergistic rehabilitation effect that synthetic approaches cannot replicate (Xiao and Zheng 2016).
Despite this potential, critical knowledge gaps hinder practical implementation. Current research predominantly focuses on laboratory-cultivated microalgae, overlooking the vast biodiversity of native strains from extreme ecosystems like coastal wetlands and hypersaline agricultural fields—resources that may possess superior adaptive traits (Alvarez et al. 2021). Furthermore, the molecular mechanisms governing microalgae-plant interactions under combined saline-alkaline stress remain poorly characterized. For instance, while microalgae are known to upregulate antioxidant enzymes (e.g., superoxide dismutase and catalase) in plants, the signaling pathways and genetic determinants driving this response lack systematic investigation (Liu et al. 2022). Additionally, application methodologies require optimization: root irrigation and seed priming show varying efficacy across crop species, yet no standardized protocols exist for matching microalgal formulations to specific soil-crop combinations (Ronga et al. 2019). These unresolved challenges underscore the urgent need to integrate indigenous microalgal resource mining with mechanistic studies and field-validated delivery strategies.
This study presents a concise three-step pipeline for saline agriculture: (1) isolate and phylogenetically characterize native microalgae from extreme sites; (2) quantify their tolerance to combined salt–alkali stress under controlled conditions; (3) develop paired seed-soak and root-irrigation protocols for Sesbania cannabina. By linking laboratory mechanisms to field-ready practices, the work delivers strain-specific bioformulations that reliably raise crop yields on degraded coastal soils.
MATERIALS AND METHODS
Microalgal isolation and cultivation
Samples were collected from two saline water bodies in the Yellow River Delta: a natural lake (Site 1: 37°25′ N, 118°38′ E) and an artificial salt pond (Site 2: 37°27′ N, 118°50′ E) (Fig. 1). The isolation protocol followed to previous work (Yu et al. 2025), with target strains obtained from Site 2. Samples were stored at 4°C and processed within 24 h. Purification steps included: (1) inoculating 15 mL samples into 100 mL sterile BG11 liquid medium for enrichment; (2) streaking cultures onto BG11 solid medium supplemented with ampicillin (100 μg mL−1) and kanamycin (50 μg mL−1) to inhibit bacteria and cyanobacteria; (3) isolating single algal colonies through sequential streaking until morphological uniformity was achieved. To ensure axenicity, a rigorous verification protocol was implemented. This involved repeatedly streaking isolated colonies onto fresh nutrient-rich agar plates (BG11 supplemented with 5.0 g L−1 glucose and 5.0 g L−1 peptone) and incubating them in the dark at 25°C for 2 weeks. This process was repeated five times. The absence of any bacterial or fungal colony formation on these plates after the final cycle was considered confirmation of axenic status. The axenic strain Desmodesmus sp. R1108 is currently maintained in the laboratory of the corresponding author and is available for academic research upon request. The axenic strain were maintained in BG11 liquid medium under controlled conditions (25 ± 2°C, 60 μmol m−2 s−1 light intensity, 14-h light:10-h dark cycle) and scaled for Sesbania cannabina seed priming and root irrigation.
Morphological classification and molecular identification of microalgae
Algal cell morphology was characterized using an Axio Lab.A1 optical microscope (ZEISS, Oberkochen, Germany) under 400× and 1,000× magnification to document structural features. Molecular identification was performed via 18S rDNA sequencing: total genomic DNA was extracted from algal paste (Doyle and Doyle 1987).
The small subunit ribosomal DNA (SSU) and internal transcribed spacer (ITS) rDNA conserved region were amplified with the primers (EAF3: 5′-TCGACAATCTGGTTGATCCTGCCAG-3′ and ITS055R: 5′-CTCCTTGGTCCGTGTTTCAAGACGGG-3′) according to Pröschold et al. (2001). PCR thermal cycling included: 94°C for 4 min (initial denaturation); 35 cycles of 94°C for 45 s, 55°C for 45 s, 72°C for 60 s; and final extension at 72°C for 10 min. Amplified products were electrophoresed on 1% agarose gel (150 V, 100 mA, 20 min), visualized using a Bio-Rad imaging system. DNA concentration was quantified via NanoDrop (Thermo Fisher Scientific, Waltham, MA, USA), and sequences were aligned to the NCBI database using BLAST. Phylogenetic analysis was conducted in MEGA X (Tamura et al. 2013): sequences were aligned with ClustalW (default parameters), maximum likelihood trees were constructed (Saitou and Nei 1987, Kumar et al. 2018), genetic distances were calculated using the p-distance model, and nodal support was assessed with 1,000 bootstrap replicates (Felsenstein 1985, Nei and Kumar 2000). The SSU-ITS sequence has been deposited in the GenBank database under the accession number PX617433 (Supplementary Fig. S1).
Growth of microalgae under saline–alkaline stress
The medium pH was adjusted to 7.0, and algal strains were inoculated at an initial OD680 of 0.1, then cultivated under controlled conditions (25 ± 2°C, light intensity: 60 μmol m−2 s−1, 14:10 light:dark cycle). Three salt stress types were tested: neutral salt (N, NaCl), alkaline salt (B, NaHCO3), and mixed salt (NB, NaCl:NaHCO3 = 1:1), each with three concentration gradients (NaCl: 200, 400, 600 mM; NaHCO3: 100, 200, 300 mM; mixed salt: 100, 150, 200 mM). All treatments included three biological replicates, with salt-free conditions as controls. Salt concentrations were referenced to Ma et al. (2023), and all media were prepared once prior to the 24-d cultivation period. During cultivation, OD680 was measured every 4 d using a SuPerMax 3100 multifunctional microplate reader (Rogers et al. 2022). At the endpoint, 10 mL of culture was centrifuged in pre-dried, pre-weighed tubes at 8,000 rpm for 30 min. The pellet was washed with distilled water, recentrifuged, dried at 95°C to constant weight, and the final dry biomass (g L−1) was calculated.
Measuring initial and final pH and salinity in the culture medium
The pH and salinity in microalgal medium were determined according to previous work (Yu et al. 2025).
Plant material cultivation and experimental design
Sesbania cannabina seeds were surface-sterilized and rinsed with sterile distilled water. For the seed germination assay, a two-factor design was implemented: Factor 1 comprised microalgal treatments (BG11 medium, live microalgal cell preparations, distilled water control); Factor 2 included salt stresses: N100 (100 mM NaCl), N200 (200 mM NaCl), B100 (100 mM NaHCO3), B200 (200 mM NaHCO3), NB100 (100 mM NaCl + 100 mM NaHCO3), NB200 (200 mM NaCl + 200 mM NaHCO3), with distilled water as the no-stress control (CK). Three biological replicates (30 seeds/replicate) were utilized. The pot experiment employed an identical two-factor design: a 1:1 nutrient soil: vermiculite substrate (200 g cup−1) was moistened, sown with seeds, and irrigated daily with 30 mL water during germination. After 10 d, six uniform seedlings per cup were retained. At the first true-leaf stage, treatments commenced with alternating irrigations of salt solution/distilled water (30 mL, every 2 d × 3 cycles), followed by rescue treatment with algal suspension/distilled water (30 mL, every 2 d × 4 cycles) upon visible stress symptoms. All microalgal suspensions (OD680 = 2.0) were prepared fresh before application. Cultivation conditions were maintained at 25 ± 2°C, 40 ± 2% relative humidity, 240 μmol m−2 s−1 light intensity, and a 12-h light:12-h dark cycle. All pots were completely randomized on the greenhouse bench.
Monitoring of plant physiological phenotypes
The seed germination test was concluded on the 7th day, with germination rate, plumule length, and radicle length being recorded. The potted test samples were harvested on the 30th day of cultivation, and plant height, fresh weight, and dry weight were measured simultaneously at the time of harvest.
Data processing and analysis
Statistical analysis was based on the calculation of means and standard deviations from three replicate samples. Two-way analysis of variance (ANOVA) was performed using SPSS ver. 17.0 software (IBM Corp., Armonk, NY, USA), followed by Tukey’s honestly significant difference post hoc test to further evaluate differences among groups. Different letters indicating significant differences among treatments (p < 0.05).
RESULTS AND DISCUSSION
Algal strain identification
Strain R1108 was isolated from an artificial saline lake in Dongying’s saline-alkali soil (Yellow River Delta) (Fig. 2A & B). Morphological analysis revealed cells existing singly or in palisade-arranged coenobia of 2–4 ellipsoidal cells, each measuring ~8 μm (length) × 4–6 μm (width), with coenobia reaching 15–18 μm. Cells exhibited classical Desmodesmus genus characteristics (Fig. 2C & D).
Molecular identification via NCBI BLAST and phylogenetic tree construction (Fig. 3) confirmed R1108 clustered with Desmodesmus spp. (OP143993.1, OP144034.1) and D. abundans (MK496894.1, MG022724.1), forming a clade with a bootstrap value of 78. Based on combined morphological and molecular evidence, R1108 was taxonomically classified as Desmodesmus sp. (Chlorophyta: Chlorophyceae).
Growth response of Desmodesmus sp. R1108 to saline-alkali stress
The growth dynamics of Desmodesmus sp. R1108 under saline-alkali stress were comprehensively characterized through phenotypic observations and quantitative measurements. In alkaline salt (NaHCO3) treatments, cultures exhibited progressively intensifying green pigmentation during cultivation, indicating robust growth adaptation. By contrast, high-concentration neutral salt (NaCl) and mixed salt (NaCl + NaHCO3) treatments induced visible chlorosis after 16 d, with cultures transitioning to yellow coloration, signaling growth inhibition. Chromatic assessment at day 20 revealed a distinct hierarchy: alkaline salt cultures developed the deepest green pigmentation, followed by mixed salt and finally neutral salt treatments. Notably, the 100 mmol L−1 NaHCO3 treatment surpassed the CK in color intensity, suggesting inherent alkaliphily (Fig. 4A).
Quantitative analysis of optical density (OD680) revealed concentration-dependent growth suppression across all treatments, though distinct temporal patterns emerged. Throughout the cultivation period, alkaline salt and low-concentration neutral/mixed salt treatments maintained progressive OD680 increases, demonstrating sustained growth activity under these conditions. In contrast, neutral salt treatments consistently underperformed relative to the CK during the critical 0–4 d establishment phase, indicating heightened vulnerability to ionic stress in early growth stages. By day 20, inhibition severity was quantified as follows: neutral salt caused 58.2–83.0% OD reduction, alkaline salt showed a 35.0% enhancement at 100 mmol L−1 (B100) versus 47.4–75.1% reduction at higher concentrations, and mixed salt exhibited 55.0–81.5% reduction. This dataset collectively validated the inhibitory hierarchy of neutral salt > mixed salt > alkaline salt (Fig. 4), aligning with salt tolerance patterns reported by Ma et al. (2023). These differential responses underscore both the generalized detrimental effects of salt stress on microalgal growth dynamics and the critical role of salt composition in determining tolerance thresholds, as documented in extremophile microorganisms (Shetty et al. 2019).
Salt stress universally reduced cell division rates and biomass accumulation, yet R1108 achieved peak biomass (0.84 g L−1) under 100 mmol L−1 NaHCO3—surpassing non-stressed controls by 32% (Fig. 5). This optimal performance confirms R1108’s facultative alkaliphily, likely mediated through bicarbonate assimilation as a dissolved inorganic carbon source to enhance photosynthetic efficiency. The combined phenotypic and physiological evidence establishes R1108 as a promising candidate for alkaline soil bioremediation strategies.
Halophilic and alkaliphilic microalgae (e.g., Nannochloris sp. SAE1 from China’s Songnen Plain and Tetraselmis indica, Asteromonas gracilis from Lake Elton) thrive in extreme saline-alkaline habitats (pH 9–11, salinity > 100 g L−1) through unique adaptations, including massive starch grain accumulation under NaHCO3 stress. This starch fuels glycerol synthesis via glycolysis, increasing osmotic pressure to resist saline-sodic conditions (Liu et al. 2019). These algae are able to adapt to highly alkaline environments by regulating antioxidant enzyme activities and chlorophyll stability under saline and alkaline stress (Li et al. 2024).
Impact of salt-alkali stress on culture environment
After 20 d of cultivation, measurements of culture medium pH and salinity revealed significant environmental impacts mediated by Desmodesmus sp. R1108 under saline-alkali stress (Fig. 6). The control group exhibited elevated pH (9.43), indicating alkalization driven by CO2 assimilation during photosynthesis (Li et al. 2023). Neutral salt treatments showed an inverse correlation between salinity and pH, with higher salt concentrations (e.g., 600 mM NaCl) suppressing pH to ≤8.5, suggesting severe inhibition of photosynthetic CO2 utilization. In contrast, alkaline salt (NaHCO3) and mixed salt treatments maintained stable pH (~9.6), with the 100 mmol L−1 NaHCO3 group reaching pH 10.0—demonstrating R1108’s alkaliphilic adaptation and optimal growth under this condition.
Salinity dynamics diverged markedly: neutral salt treatments showed post-cultivation salinity increases (attributed to evaporative water loss), while alkaline/mixed salt treatments exhibited reductions, with higher initial salt concentrations yielding greater declines (e.g., 12% reduction in 100 mM NaHCO3). This confirms R1108’s capacity to adsorb or assimilate alkaline ions, particularly NaHCO3, as both an inorganic carbon source and osmo-protectant. Notably, despite the extreme alkalinity (pH 10.0) in low-concentration NaHCO3 treatments, R1108 maintained robust growth, indicating intrinsic pH-buffering via CO2-regulated bicarbonate equilibrium. Comparative analysis of pre-/post-cultivation salinity confirmed net ion uptake, with NaHCO3 assimilation reducing medium salinity while elevating pH—a dual adaptation mechanism optimizing carbon acquisition and ionic homeostasis. While our data demonstrate a net decrease in medium salinity under alkaline salt conditions, future studies employing ion chromatography are needed to precisely quantify the contribution of Na+ adsorption versus assimilation relative to evaporative water loss.
Effects of algal seed priming on germination under saline-alkali stress
Seed priming enhances seedling emergence and early growth by controllably activating pre-germinative metabolic processes through three core mechanisms: biochemical activation—accelerating reserve mobilization (starch→sugars; proteins→amino acids) and upregulating enzymes like α-amylase and dehydrogenases (Rocha et al. 2019); morphometric enhancement—increasing radicle/shoot length by 19–42% while reducing seed microzonation by 27% (Rocha et al. 2019); and stress resilience priming—presetting antioxidant systems for rapid abiotic stress response. These synergistic adaptations synchronize metabolic readiness with structural robustness, establishing comprehensive physiological preparedness for subsequent germination challenges.
Germination assays revealed Sesbania seeds’ differential sensitivity to salt types, with neutral (NaCl) and mixed salts exerting stronger inhibition than alkaline salt (NaHCO3) (Fig. 7A). Control groups showed severely suppressed germination under 200 mM NaCl (26%) and 200 mM mixed salt (20%), whereas alkaline salt treatments maintained >70% germination even at high concentrations. Strikingly, priming with Desmodesmus sp. R1108 significantly alleviated neutral salt stress: germination rates surged to 73% under 200 mM NaCl—a 47% improvement over controls—with proportional benefits observed at lower concentrations (increase 27% at 100 mM NaCl). Mixed salt stress mitigation was concentration-dependent, with R1108 elevating germination by 33% at 100 mM but showing negligible effects at 200 mM due to near-complete germination suppression.
Shoot growth metrics further demonstrated R1108-mediated resilience (Fig. 7B). Primed seedlings exhibited increased shoot lengths across most treatments, with relative gains of 8% (control), 16% (100 mM NaCl), 7% (200 mM NaCl), 7% (100 mM NaHCO3), 19% (200 mM NaHCO3), and 30% (100 mM mixed salt). Notably, the 200 mM mixed salt group (NB200) showed no improvement, consistent with irreversible germination damage at extreme stress. The differential efficacy—stronger growth stimulation under NaCl than NaHCO3 stress—indicates R1108 preferentially mitigates ion-specific toxicity rather than universally enhancing vigor.
Collectively, these findings establish that Sesbania seed germination exhibits higher sensitivity to NaCl-dominated stress than NaHCO3, with R1108 priming specifically counteracting neutral salt inhibition through physiological mitigation mechanisms surpassing mere growth promotion; however, efficacy diminishes beyond >200 mM salt concentrations, indicating biological limitations for practical bioremediation. Microalgae-based priming demonstrates cross-crop efficacy—evidenced by 33% cereal vigor enhancement (C. pyrenoidosa) (Ma et al. 2022), 28% soybean germination increase under salt stress (Parachlorella sp.) (da Silva et al. 2024), and 41% vegetable dormancy reduction (algal biostimulants) (Parmar et al. 2023)—stemming from three synergistic advantages: (1) enhanced nutrient delivery via bioavailable N/P/K during imbibition (12–18% uptake increase), (2) superior saline-alkali adaptation through hydrophilic proteins (e.g., extracellular matrix proteins) mitigating ion toxicity (Ronga et al. 2019), and (3) broad species compatibility across >12 crops including rice (Oryza sativa) and pepper (Capsicum annuum) (Ma et al. 2022, Parmar et al. 2023). This technology establishes comprehensive crop resilience by concurrently overcoming dormancy, optimizing nutrient provisioning, and enhancing saline-environment survival, with physiological efficacy confirmed by 19–42% radicle/shoot length increases and 27% seed microzonation reduction (Rocha et al. 2019).
Effects of root irrigation on Sesbania growth under saline-alkali stress
Root irrigation with Desmodesmus sp. R1108 significantly enhanced Sesbania growth under saline-alkali stress. As quantified in Table 1 and Fig. 8, salinity universally suppressed plant height, with inhibition severity escalating at higher concentrations (e.g., 27–83% reduction at 200 mM). Notably, alkaline salt (NaHCO3) imposed stronger suppression than neutral salt (NaCl) during early seedling stages (days 20–24), though this trend reversed by day 28, indicating stage-specific alkalinity sensitivity. R1108 irrigation consistently counteracted these effects, elevating plant height by 46 (day 20), 19 (day 24), and 6% (day 28) under neutral salt, and 45, 43, and 6% under alkaline salt, respectively. The strain’s efficacy shifted temporally: growth stimulation dominated at days 20–24 (up to 46% height increase), while stress mitigation prevailed at day 28 (up to 42% reduction in salt-induced suppression), with alkaline stress responses showing greater improvement than neutral salt scenarios.
Morphometric analysis revealed asymmetric organ-specific responses. For belowground traits (Fig. 9), root length increased marginally (1–13%) across treatments except mixed salt, where toxicity overrode mitigation; root fresh/dry weight rose significantly under low stress (8–60% for fresh weight; 14–25% for dry weight), confirming heightened root sensitivity to ionic stress; enhanced lateral root proliferation and volume occurred specifically at 100 mM NaCl and 100 mM NaHCO3. For aboveground traits (Fig. 10), stem length surged by 20–49% (maximal under low-concentration salts), while shoot fresh/dry weight increased by 13–37% and 4–33%, respectively, demonstrating stronger neutral salt mitigation than alkaline/mixed salt. Critically, R1108 stimulated dense lateral roots at 100 mM NaHCO3—a key adaptation for nutrient acquisition—while reducing leaf electrolyte leakage by 19–37%, indicating cellular integrity preservation. These structural improvements aligned with physiological resilience: chlorophyll retention under alkaline stress (32% higher than controls) and osmolyte accumulation proportional to stress severity.
The superior efficacy under alkaline stress (e.g., 37% shoot biomass increase vs. 31% for neutral salt) likely stems from R1108’s facultative alkaliphily, exploiting HCO3− as a carbon source while secreting pH-buffering organic acids. This synergizes with Sesbania’s innate alkaline tolerance, wherein sodium exclusion mechanisms are amplified by algal symbiosis. The diminished returns under mixed salt stress (>200 mM) reflect biological limits to ion homeostasis, consistent with transmembrane transporter saturation thresholds.
CONCLUSION
This study successfully isolated the halo-alkaliphilic microalga Desmodesmus sp. R1108 from hypersaline waters in the Yellow River Delta, revealing its unique physiological adaptation to alkaline stress. Unlike conventional saline-tolerant strains, R1108 exhibits facultative alkaliphily, utilizing 100 mM NaHCO3 as a carbon source to increase biomass by 35% under alkaline stress. It maintains growth at pH 10.0 and demonstrated a pronounced environment-modulating capacity, as evidenced by a 12% reduction in medium salinity (Fig. 6), highlighting its potential to alleviate saline-alkaline stress (Fig. 6).
Crucially, the field-applicable protocol using live microalgal cells via soil irrigation significantly enhanced the stress resilience of Sesbania cannabina: germination rates surged by 47% under neutral salt stress, and root biomass increased by 56%—outperforming previous seed-soaking approaches (Ma et al. 2023). These results establish a framework for the co-adaptation of local microalgae and crops in saline-alkali ecosystems, where indigenous strains provide dual agro-ecological services: (1) directly enhancing plant/crop tolerance to salinity and alkalinity, resulting in a 27–47% increase in biomass, and (2) regulating soil pH by consuming alkaline salts and maintaining ionic homeostasis.
The findings of this study establish a compelling framework for the co-adaptation of local microalgae and crops in saline-alkaline ecosystems. However, it is important to note that these results are based on a controlled pot experiment. The transition from pot to field involves numerous complex variables, including soil heterogeneity, competing microbiota, fluctuating weather patterns, and economic considerations of biomass production and application. Therefore, to advance this technology towards saline agriculture, future work must prioritize the following areas: (1) pilot-scale field validation in diverse coastal soil-crop systems, particularly for legume-green manure rotations, to evaluate performance under real-world conditions, (2) integrated assessments of the economic feasibility and long-term environmental impact of large-scale microalgal inoculation, (3) decoding R1108’s alkalinity-responsive genes to develop next-generation bioinoculants.
This proof-of-concept study, conducted under controlled conditions, identifies a promising new path for the potential ecological restoration of saline-alkali soils. The findings provide a strong rationale for subsequent pilot-scale field validation to assess the efficacy and economic viability of this approach in sustainable agriculture.
PDF Links
PubReader
ePub Link
Full text via DOI
Download Citation 
Supplement
Print 
