Exploring the Potential of Desmodesmus sp. KNUA231 for Bioenergy and Biofertilizer Applications and Its Adaptability to Environmental Stress

Abstract

As global energy demand continues to rise, microalgae have gained attention as a promising feedstock for biofuel production due to their environmental adaptability and renewable nature. This study investigated the growth performance and stress tolerance of Desmodesmus sp. KNUA231 under varying pH and salinity conditions to evaluate its potential as a biofuel candidate. The strain was cultivated under controlled laboratory conditions and exhibited stable growth across a broad pH range (4–10) and moderate salinity levels (up to 5 g L⁻¹ NaCl), indicating its resilience to diverse environmental conditions. Fatty acid methyl ester (FAME) analysis revealed that the biodiesel properties of Desmodesmus sp. KNUA231 comply with ASTM and EN standards in specific parameters, reinforcing its feasibility as a renewable biofuel feedstock. Additionally, its high calorific value (CV) suggests its potential as an energy-dense biomass source. The results of inductively coupled plasma mass spectrometry (ICP) analysis show that the soil is supplied with essential nutrients while minimizing heavy metal contamination, suggesting the possibility of biofertilizers. Although Desmodesmus sp. KNUA231 demonstrated promising characteristics for biofuel applications, further research is required to optimize large-scale cultivation and improve productivity for industrial applications. These findings highlight the potential of Desmodesmus sp. KNUA231 as a biofuel resource, particularly in non-optimal environmental conditions where pH and salinity fluctuations are common, contributing to the ongoing search for sustainable bioenergy solutions.

1. Introduction

As annual global energy consumption continues to rise, there is increasing interest in renewable energy sources as alternatives to fossil fuels. Non-renewable energy consumption, such as fossil fuels, increased by 84% from 2000 to 2023, while the capacity of renewable energy expanded by 414% [1]. Non-renewable energy encompasses fossil fuels, including coal and oil, as well as nuclear energy. The reliance on non-renewable energy has heightened interest in renewable energy due to its significant contribution to greenhouse gas emissions and global warming [2]. Global warming can also have a negative impact on major crops in many agricultural regions [3,4]. As a result, interest in biofertilizers using microalgae is also increasing [5,6]. Despite the rapid growth in renewable energy capacity, climate-related challenges associated with non-renewable energy persist [7]. Addressing these issues requires sustained efforts to enhance the adoption and utilization of renewable energy resources. The first generation of renewable energy sources, such as corn, wheat, and soybeans, faced limitations, including competition with food production and extensive farmland requirements. To address these drawbacks, second-generation renewable energy resources, primarily derived from woody plants, were introduced. However, their reliance on land required for food production posed additional challenges [8,9]. As a solution, microalgae have been identified as third-generation renewable energy resources due to their ability to overcome the limitations of the previous generations [8,10].

Microalgae have garnered significant interest due to their ability to accumulate substantial lipid levels and convert them into biodiesel via the transesterification process, highlighting their potential for biofuel production [11,12,13]. Microalgae are microorganisms that grow by photosynthesizing in various aquatic environments. They can grow in a variety of culture conditions, including salt water, fresh water, and wastewater, so they offer great production flexibility [14,15]. Microalgae, which can also be cultivated in wastewater, utilize nutrients such as nitrogen and phosphorus from wastewater for growth, thereby contributing to wastewater purification [16]. Furthermore, microalgae are recognized as a sustainable resource for various biotechnological applications, including the production of high-value compounds such as omega-3 fatty acids and lutein [17]. For instance, microalgae species like Nannochloropsis and Schizochytrium are rich in eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) and are widely utilized as raw materials in health supplements and pharmaceuticals [18]. Unlike omega-3 fatty acids derived from fish, microalgae-based omega-3 production offers a sustainable alternative that does not compromise marine ecosystems [19,20,21]. Microalgae have the advantages of fast growth rate, efficient carbon dioxide fixation, and the inclusion of various high-value compounds, including pigments that can be utilized as bioactive ingredients in food, pharmaceutical, and cosmetic applications [22,23]. In addition, microalgae can accumulate lipids, carbohydrates, proteins, various bioactive compounds, and essential inorganic nutrients, enabling their utilization in the production of bio-based products such as biofuels and biofertilizers [24,25]. Furthermore, as a biofertilizer, microalgae contribute to soil fertility enhancement and reduce dependence on synthetic fertilizers, positioning them as a sustainable alternative to conventional fertilization practices [22].

Despite these advantages, scaling up microalgae cultivation and processing to meet industrial demands poses significant challenges. Large-scale production encounters numerous technical and economic barriers, such as optimizing growth conditions, developing cost-effective harvesting techniques, and streamlining downstream processing. Furthermore, the physiological and biochemical characteristics of microalgae vary considerably among species, requiring targeted research to identify optimal strains for biofuel production and other biotechnological applications. A thorough understanding of each strain’s growth dynamics, metabolic pathways, and responses to environmental stresses is crucial for enhancing cultivation efficiency and maximizing biomass yield in commercial settings.

Desmodesmus sp. KNUA231, the focus of this study, was collected from Ulleungdo, located east of the Korean Peninsula. This region consists of a central island and several smaller islands, characterized by steep slopes that prevent the formation of lakes and ponds, instead giving rise to rivers. These rivers, while originating from the same freshwater source, display varied characteristics based on the surrounding environmental conditions. The unique geographical features of Ulleungdo provide distinct environmental niches, facilitating research into various microalgal species [26,27].

In this study, the growth characteristics of the Desmodesmus sp. KNUA231 strain were monitored through cultivation to elucidate its fundamental properties. Experiments were also conducted to assess the strain’s tolerance to pH and salinity stress. Biomass harvested from these experiments underwent physicochemical and biochemical analysis to further investigate the strain’s traits. Proximate and ultimate analyses were performed to determine the calorific value, evaluating its potential as a bioenergy source. Additionally, fatty acids were extracted to assess its potential as biodiesel, and inductively coupled plasma (ICP) analysis was utilized to explore its capabilities as a biofertilizer. Consequently, these varied analyses tested the potential applications of the Desmodesmus sp. KNUA231 microalgal strain.

2. Materials and Methods

2.1. Isolation and Identification

2.1.1. Isolation and Molecular Identification

Desmodesmus sp. KNUA231 was isolated from fresh water, Ulleungdo, East Sea, Republic of Korea. It was inoculated into 100 mL of BG-11 medium and incubated at 25 °C, 160 rpm, with a 16:8 light/dark cycle for one week to allow sufficient growth. After harvesting the culture medium at the stationary growth phase and centrifuging it at 4032× g for 5 min, the supernatant was removed, and the penicillin–streptomycin–neomycin (PSN) antibiotic mixture (Sigma Aldrich, St. Louis, MO, USA) was added at a final concentration of 100 μg mL⁻¹. The resulting pellet was plated onto the BG-11 agar medium, and single colonies were subsequently isolated until distinct colonies were obtained. Genomic DNA was extracted using a DNeasy Plant Mini kit (Qiagen, Hilden, Germany), and PCR was performed with ITS, 18S rRNA, and tufA primers (Table S1). The resulting sequences were compared to genetically related species using the National Center for Biotechnology Information (NCBI) database.

2.1.2. Morphological Identification

During the growth experiment, the morphology of the microalgae was observed on day 9 using a light microscope with a camera (Axiocam 208; Zeiss, Jena, Germany) at a magnification of 1000×.

2.2. Growth Measurements

2.2.1. Growth Rate and Biomass Productivity

Growth experiments were conducted using 500 mL of the BG-11 medium in a 1 L flask. Cultures were incubated at 25 °C, 200 μmol m⁻² s⁻¹ light intensity, with a 16:8 light/dark cycle and agitation at 120 rpm (VS-202D shaker; Vision Scientific, Bucheon, Republic of Korea) for 9 days. Optical density (OD) at 680 nm was measured daily using a spectrophotometer (UV 1900i; Shimadzu, Kyoto, Japan). Cell numbers were determined using a Neubauer Improved Hemocytometer and an optical microscope (Axio Vert.A1; Zeiss, Jena, Germany). pH was measured using a SevenDirect SD20 pH meter (Mettler Toledo, Shanghai, China). Microalgae growth was supplemented with 1–1.5% carbon dioxide and injected through a filter for pH control. On day 9, dry weight, chlorophyll, and total carotenoid content were measured.

Dry weight was determined by filtering 5 mL of culture medium through GF/C Whatman filter paper (0.7 µm pore size; Cytiva, Shanghai, China), followed by drying at 105 °C for 24 h (WOF-W155; Daihan Scientific, Wonju, Republic of Korea). Chlorophyll and total carotenoid content were analyzed after centrifuging 2 mL of medium at 12,000 rpm at 4 °C. The resulting pellet was resuspended in methanol and subjected to ultrasonication (VC-505; Sonics & Materials, Newtown, PA, USA) with 5 s on/off intervals for 1 h. Samples were stored overnight at 4 °C in darkness, centrifuged, and the supernatant was diluted with methanol. OD values at 470, 653, and 666 nm were measured, and chlorophyll-a (Chl-a), chlorophyll-b (Chl-b), and carotenoid (Car) concentrations were calculated using Wellburn’s equations (1994):

Chl-a = 15.65A666 − 7.34A653

Chl-b = 27.05A653 − 11.21A666

Car = (1000A470 − 2.86 Chl-a − 129.2 Chl-b)/221

2.2.2. Carotenoid Analysis

After incubation, the sample is harvested using a centrifuge (5810 R; Eppendorf, Hamburg, Germany) at 4000 rpm for 5 min and a freeze dryer is used (PVTFD20R; Ilshin Lab, Suwon, Republic of Korea) to obtain lyophilized biomass. Then, 10 mg of lyophilized biomass is used for carotenoid analysis. First, 1 mL of 60% KOH (w/v) was added, followed by bead beating for a total of 16 min and heating in a 40 °C water bath for 40 min, and saponification was performed by shaking at 4 °C overnight in the dark. After that, the sample is washed with a solvent of Dichloromethane, Methanol 1:1 and filtered using a PVDF 0.22 μm syringe filter (PV1322; Chromdisc, Hwaseong, Republic of Korea) and the liquid is evaporated with nitrogen gas. The resulting product was then dissolved in high-performance liquid chromatography (HPLC)-grade MeOH for analysis using an Agilent 1200 series gradient HPLC system (Agilent Technologies, Santa Clara, CA, USA) with a photodiode array detector (200–800 nm). This study employed lutein, zeaxanthin, β-carotene, fucoxanthin astaxanthin, and canthaxanthin, obtained from Sigma–Aldrich (St. Louis, MO, USA), as the standard materials.

2.3. pH and Salinity Stress Tolerance

The following experimental conditions were employed to evaluate pH and salt tolerance. Both pH and salt tolerance were assessed using 100 mL of BG-11 medium in 250 mL flasks. To evaluate the effects of pH, the BG-11 medium was first prepared and adjusted to target pH values of 4, 6, 8, 10, and 12 using either 1 N hydrochloric acid (HCl) or 1 N sodium hydroxide (NaOH). After pH adjustment, the medium was autoclaved to maintain sterile conditions throughout the cultivation. For salinity stress experiments, BG-11 medium was prepared by directly adding NaCl to reach final concentrations of 5, 10, 20, and 30 g L⁻¹ in 100 mL volumes. The salt-containing media were also autoclaved prior to use to maintain sterility. Cultures were maintained under the following conditions: 25 °C, 200 μmol m⁻² s⁻¹ light intensity, a 16:8 light/dark cycle, and 160 rpm. The pH and OD at 680 nm were monitored over 12 days period.

2.4. Biochemical Composition

2.4.1. Carbohydrate, Protein and Lipid

To determine carbohydrates, 50 mg of lyophilized biomass is dissolved in 2.5 mL of 2 N H₂SO₄ and boiled in a 100 °C water bath for 3 h. Then, the sample is neutralized by adding CaCO₃ until there is no foam, made up with 50 mL of dH₂O, and centrifuged at 4000 rpm for 5 min, using only the supernatant. Next, 0.1–0.5 mL of the sample is placed in each of five glass tubes and the remaining 1 mL is filled with dH₂O. Also, 0.05 mL of phenol solution and 5 mL of sulfuric acid are added to each tube, inverted, and placed in an incubator set at 25 °C for 20 min. After 20 min, the OD is measured at 490 nm [28].

To measure protein, transfer 20 mg of lyophilized biomass to a 1.7 mL tube, add 0.5 mL of RIPA lysis buffer solution (GenDEPOT, Katy, TX, USA), and sonicate for 1 h (at 5 s internally). Then, incubate at 100 °C for 30 min and centrifuge at 4 °C, 11,000× g for 15 min. Transfer 600 µL of the supernatant to a 5 mL tube, add four times the volume of the supernatant to the tube with a solution containing 10% (w/v) trichloroacetic acid (Sigma-Aldrich, Bengaluru, India), and incubate at −20 °C overnight. Divide into equal volumes in a tube and centrifuge at 18,400× g for 15 min. Discard the supernatant, loosen the pellet with 500 µL of ice-cold acetone, and centrifuge again at 18,400× g for 15 min. Discard the supernatant, add 200 µL of ice-cold acetone, collect in one tube, spin the centrifuge, and discard the resulting supernatant. Air dry and dissolve the sample in 2 mL of 8 M urea (Sigma-Aldrich, St. Louis, MO, USA) (pH 8.8) in 60 mM Tris-HCl (Biosesang, Yongin, Republic of Korea). Place 0, 20, 40, 60, 80, and 100 µL of sample in each of six glass tubes and fill the rest with dH₂O. Fill with 2 mL of BCA working reagent (WR) and allow it to react for 30 min in an incubator at 37 °C; then, measure the OD at 562 nm [29]. All of the protein samples were quantified using Pierce BCA protein assay kits (Thermo Scientific, Waltham, MA, USA), with bovine serum albumin (BSA) (Thermo Scientific, Vilnius, Lithuania) used as the standard.

Based on the sulfo-phospho-vanillin (SPV) method [30], 10 mg of biomass and 1 mL of Dh₂O were added and sonicated. Five glass tubes were filled with 10, 20, 30, 40, and 50 µL of the sample, respectively, and the rest was filled with dH₂O to make a total of 100 µL. Then, 2 mL of concentrated sulfuric acid (H₂SO₄) was added to each tube, and the samples were incubated at 100 °C for 10 min, followed by cooling on ice for 5 min. A phospho-vanillin (PV) reagent solution of 5 mL was prepared and added to the samples. The samples were shaken at 200 rpm for 15 min at 37 °C. Finally, the optical density (OD) was measured at 530 nm using a spectrophotometer. A calibration curve was generated using canola oil as the lipid standard at concentrations of 0, 10, 20, 30, 40, 50, 60, and 70 µL mL⁻¹, each measured in triplicate. The resulting regression equation was y = 0.005x − 0.0031 (R² = 0.9938), where y represents the absorbance at 530 nm and x is the lipid concentration in µL mL⁻¹.

2.4.2. Monosaccharide Analysis

The HPLC (Prominence; Shimadzu, Kyoto, Japan) analysis was outsourced to determine monosaccharide composition. Prior to the analysis, the carbohydrate samples were pre-processed. The analysis utilized sucrose, lactose, glucose, galactose, fructose, arabinose, mannitol, fucose, mannose, rhamnose, and ribose as standards. The analysis was conducted at a flow rate of 0.5 mL min⁻¹ and at 90 °C using an injection volume of 20 µL.

2.4.3. Free Amino Acid Analysis

Free amino acid analysis was conducted by mixing 0.1 g of biomass with 0.1 N HCl and 5% TCA, each at 1 mL, followed by separation of various amino acids using an amino acid autoanalyzer (LA8080; Hitachi, Chiyoda, Japan). The separated amino acids were reacted with ninhydrin to form chromogenic compounds, which were analyzed by measuring absorbance at wavelengths of 570 nm and 440 nm.

2.4.4. Fatty Acid Methyl Ester Analysis

Lipids were extracted from 10 mg of freeze-dried microalgal biomass using a biphasic solvent system composed of chloroform/methanol (4:5, v/v) and an aqueous phase containing 50 mM Tris and 1 M NaCl. After vortexing and ultrasonication for 15 min, the mixture was centrifuged at 4032× g for 10 min to facilitate phase separation. The lower chloroform-rich organic layer containing the lipids was carefully collected. The extraction was repeated three times, and all organic layers were pooled and washed with distilled water to remove residual salts. The lipid-containing solution was then concentrated using a rotary evaporator to obtain the crude lipid extract, which was stored at −20 °C until further use.

The extracted lipids were converted into fatty acid methyl esters (FAMEs) via acid-catalyzed transesterification. Specifically, 3 mL of methanol containing 5% (v/v) sulfuric acid (H₂SO₄) was added to the dried lipid extract, and the reaction mixture was incubated at 70 °C for 3 h in a water bath, with vortexing carried out every 30 min. After cooling to room temperature, 3 mL of distilled water and 3 mL of hexane were added, followed by vortexing and orbital shaking for 15 min. The mixture was centrifuged at 1200× g for 5 min, and the upper hexane layer containing FAMEs was carefully collected. To remove residual impurities, the hexane layer was washed with 2 mL of distilled water, vortexed, and centrifuged again under the same conditions. The resulting hexane phase was filtered through a 0.22 µm PVDF syringe filter, concentrated using a rotary evaporator, and stored at −20 °C until GC/MS (7890B-5977B; Agilent Technologies, Santa Clara, CA, USA) analysis [31].

2.5. Biodiesel Quality Assessment

The quality of biodiesel was evaluated by determining various fuel characteristics, such as saponification value (SV), iodine value (IV), cetane number (CN), degree of unsaturation (DU), long-chain saturated factor (LCSF), cold filter plugging point (CFPP), oxidative stability (OS), kinematic viscosity (υ), and density (ρ). These parameters were calculated based on the FAME profile of the biodiesel using the equations outlined below, in accordance with the fuel quality criteria defined by ASTM D6751 and EN 14214 standards [32].

SV = ∑ (560 × P)/MW

IV = ∑ (254 × P × D)/MW

CN = (46.3 + 5458/SV) − (0.225 × IV)

DU = ∑ MUFA + (2 × PUFA)

LCSF = (0.1 × C16) + (0.5 × C18) + (1.0 × C20) + (1.5 × C22) + (2.0 × C24)

CFPP = (3.1417 × LCSF) − 16.477

OS = 117.9295/X + 2.5905 (0 < 100)

ln(υ) = ∑ −12.503 + 2.493 × ln(MW) − 0.178 × D

ρ = ∑ 0.8463 + 4.9/MW + 0.0118 × D

2.6. Proximate Analysis and Ultimate Analysis

For thermogravimetric analysis, 5 mg of biomass was analyzed using a thermal analyzer (DTG-60A; Shimadzu, Kyoto, Japan). α-Alumina (α-Al₂O₃) powder (30 mg; Shimadzu, Kyoto, Japan) was employed as a reference material. Nitrogen gas was supplied at a flow rate of 25 mL min⁻¹, and the sample temperature was increased at a rate of 10 °C min⁻¹ from 50 °C to 900 °C.

Carbon (C), hydrogen (H), oxygen (O), nitrogen (N), and sulfur (S) contents were analyzed using an elemental analyzer (FlashSMART; Thermo Fisher Scientific, Milan, Italy). The obtained data were subsequently used to calculate the calorific value (CV) based on the equation provided below.

CV (MJ kg⁻¹) = 0.3278C + 1.419H + 0.09257S − 0.1379O + 0.637

2.7. Nutrient and Metal Composition Analysis in Biofertilizer

For nutrient analysis, approximately 0.1 g of freeze-dried microalgal biomass was digested with concentrated nitric acid using a microwave digestion system (UltraWAVE; Milestone, Italy). The digested samples were diluted with 2% nitric acid and subsequently analyzed using inductively coupled plasma mass spectrometry (ICP-MS) (NexION 2000; PerkinElmer, Waltham, MA, USA) to quantify macronutrients (P, K, Ca, Mg, S, Si, Na) and micronutrients (Fe, Zn, Cu, Mn, Mo, B) [33].

As heavy metals (Cd, As, Pb) were not detected at mg L⁻¹ levels by ICP-MS, these elements were reanalyzed using inductively coupled plasma optical emission spectrometry (ICP-OES) (Avio 500 and 55; PerkinElmer, Waltham, MA, USA) under standard conditions to confirm their presence or absence.

3. Results and Discussion

3.1. Identification of KNUA231

3.1.1. Molecular Identification

NCBI-based ITS, 18S rRNA, and tufA sequence analysis of the gene showed that KNUA231 is close to the Desmodesmus sp. group in molecular identification (

Table 1. Identification of microalgae KNUA231 using molecular markers.

Marker Gene	Length (bp)	Closest Match (Accession No.)	Query Cover	Identification
ITS	653	Desmodesmus sp. KNUA024 (MT603589)	100%	100%
18S rRNA	407	Desmodesmus sp. (OM893304)	100%	99.51%
tufA	920	Desmodesmus spinosus (MN218421)	94%	96.08%

). The primer ITS phylogeny was also plotted with Coelastrum as an outgroup (Figure S1), revealing that it was closest to Desmodesmus sp. KNUA024 (MT603589), followed by Desmodesmus multivariabilis strain TAU-MAC 2517 (OK642366).

3.1.2. Morphological Identification

Desmodesmus sp. KNUA231 was cultivated in a liquid medium for 9 days and observed under an optical microscope at 1000 × magnification on the final day. A microscopic examination revealed that Desmodesmus sp. KNUA231 primarily exhibited a linear arrangement of four cells, with some two-celled structures also observed. The length of individual cells, both horizontally and vertically, did not exceed 5 μm (Figure S1).

A distinguishing feature of this species is the presence of short terminal spines, which varied among specimens within the sample [34,35]. These morphological characteristics were only discernible under optical microscopy.

The measurements of Desmodesmus sp. KNUA231 in this study were consistent with previously reported size ranges for the genus Desmosomes [36], supporting its taxonomic similarity to the species documented in the literature.

3.2. Growth Measurements

3.2.1. Growth Rate and Biomass Productivity

The 9-day incubation period was selected to capture the most active growth phase of the microalgae, during which optical density (OD) and pH were monitored daily (Figure S2). The OD of the initial microalgae culture was 0.103 ± 0.003 and increased to 4.487 ± 0.315 after 9 days of cultivation (Figure 1). The dry weight of the biomass at the end of the cultivation period was measured at 1.54 g L⁻¹.

On day 9, chlorophyll a, chlorophyll b, and total carotenoids were quantified, yielding concentrations of 36.29 ± 1.63 μg mg⁻¹, 6.55 ± 0.84 μg mg⁻¹, and 12.98 ± 0.13 μg mg⁻¹, respectively.

3.2.2. Carotenoid Analysis

A HPLC analysis of carotenoids identified only lutein and β-carotene, with lutein measured at 0.272 mg L⁻¹ and β-carotene at 0.085 mg L⁻¹. Among these pigments, lutein is a key component of the macula in the human eye, whereas oxidative stress is considered a major factor contributing to age-related macular degeneration [37,38]. Additionally, lutein has been reported to play a role in preventing atherosclerosis in cardiovascular disease due to its antioxidant properties [39,40].

3.3. pH and Salinity Stress Tolerance

This experiment was conducted over a 12-day period under various pH and salinity conditions (Figure 2) to assess stress tolerance based on growth rates. The rationale behind testing these stress conditions was to evaluate the strain’s physiological adaptability to environments with fluctuating pH and salt concentrations, which are commonly encountered in large-scale cultivation systems, including outdoor and industrial settings.

Obtaining such baseline data is essential for identifying the environmental limits of the strain and assessing its feasibility for broader biotechnological applications. Among the tested conditions, the most acidic condition, pH 4, resulted in a rapid shift in the culture medium toward alkalinity (Table S2). Meanwhile, pH 12 led to the lowest growth rate, whereas pH 10 showed a similar growth pattern to the control. By day 12, the growth rate at pH 10 surpassed that of the control, making it the highest among all pH treatments—highlighting the strain’s enhanced performance under slightly alkaline conditions. These results suggest that Desmodesmus sp. KNUA231 has the ability to utilize bicarbonate (HCO₃⁻) as an alternative carbon source under high-pH conditions, where free CO₂ is limited due to carbonate equilibrium. In alkaline environments, the concentration of hydrogen ions (H⁺) is low, and CO₂ in the medium tends to decrease as it shifts toward bicarbonate and carbonate forms. However, if KNUA231 efficiently converts HCO₃⁻ into intracellular CO₂, it can maintain photosynthetic activity and sustain growth even as external CO₂ diminishes. As more inorganic carbon is assimilated for photosynthesis, CO₂ levels in the medium drop further, reducing H⁺ generation and causing a continued rise in pH rather than a decrease. This internal carbon acquisition mechanism likely provides a physiological advantage for growth in high-pH environments.

Furthermore, previous studies have reported that Desmodesmus species tend to exhibit relatively stable growth across a wide range of environmental conditions rather than displaying exceptionally high growth rates under specific conditions [41,42]. Given these findings, Desmodesmus sp. KNUA231 is more likely to maintain stable growth across diverse environmental fluctuations rather than exhibiting rapid growth under optimal conditions. This physiological adaptability enhances its potential for large-scale cultivation in various cultural environments, suggesting that it could be utilized in a broad range of industrial applications without being restricted to narrowly defined optimal conditions.

To assess salinity tolerance, the growth rate at 5 g L⁻¹ NaCl exhibited a continuous increase over 12 days, although it remained lower than that of the control. Under 10 g L⁻¹ NaCl conditions, growth increased slightly for the first three days but stagnated thereafter (Table S3). At higher salinity levels, growth was entirely inhibited. These findings suggest that Desmodesmus sp. KNUA231 can sustain growth under moderate salinity conditions but experiences significant growth inhibition at higher salt concentrations.

As shown, Desmodesmus sp. KNUA231 exhibits the broadest pH tolerance range (4–10), while maintaining moderate salinity tolerance (up to 10 g L⁻¹ NaCl). This suggests that Desmodesmus sp. KNUA231 is particularly well-suited for environments with highly variable pH and moderate salinity, in contrast to species with narrower pH tolerance limits (Table S4) [43,44,45,46,47,48,49,50,51]. These findings underscore the ecological robustness of the strain and support its potential use in outdoor photobioreactors, wastewater-based cultivation systems, and other industrial platforms where environmental conditions are not strictly controlled.

3.4. Biochemical Composition

3.4.1. Carbohydrate, Protein and Lipid Contents

After 9 days of cultivation under CO₂-enriched conditions, the carbohydrate, protein, and lipid contents of the harvested biomass were analyzed. The results indicated that the macromolecular composition followed the following order: carbohydrates (24.36 ± 1.41 wt%), proteins (26.18 ± 1.14 wt%), and lipids (17.21 ± 0.57 wt%) (Figure 3). The microalgal carbohydrate and protein content suggests potential applications in the food and aquaculture industries [52,53].

3.4.2. Monosaccharide Analysis

To investigate the composition of carbohydrates, a monosaccharide analysis was performed. The results showed that glucose was 561 μg mL⁻¹ and mannose was 595 μg mL⁻¹. Glucose is a product of photosynthesis in microalgae and serves as an energy source and metabolic intermediate. It also acts as a precursor to storage carbohydrates such as starch, playing an important role in cell growth and environmental adaptation [37]. Mannose is mainly known as a component of the cell wall of microalgae. The presence of mannan, a polysaccharide containing mannose, in the cell walls of green algae has been reported, suggesting that it may play a structural role in the microalgae in this experiment. Therefore, the detection of glucose and mannan may reflect their functions as energy sources and structural components, respectively [54].

3.4.3. Free Amino Acid Analysis

Amino acid analysis was conducted to characterize the protein composition of the microalgal strain. Amino acids play a critical role in physiological metabolism, particularly in protein synthesis and energy production. Essential amino acids, which cannot be synthesized by the human body and must be obtained through dietary intake, were identified in this strain. Specifically, valine (1.607 µg mL⁻¹), leucine (1.138 µg mL⁻¹), lysine (4.219 µg mL⁻¹), isoleucine (0.819 µg mL⁻¹), histidine (0.309 µg mL⁻¹), phenylalanine (0.925 µg mL⁻¹), and tyrosine (0.665 µg mL⁻¹) were detected, highlighting the potential application of this strain as a valuable feedstock in the food and pharmaceutical industries (

Table 2. The constituent amino acid composition of the Desmodesmus sp. KNUA231 biomass.

Amino Acids		Content (µg mL−1)
Essential	Threonine	1.802
	Valine	1.607
	Methionine	0.230
	Isoleucine	0.819
	Leucine	1.138
	Phenylalanine	0.925
	Lysine	4.219
	Histidine	0.309
Conditionally essential	Arginine	2.371
	Glycine	2.272
	Tyrosine	0.665
	Proline	2.576
Non-essential	Aspartic Acid	0.480
	Serine	1.490
	Glutamic acid	41.567
	Alanine	20.280
	Gamma-Aminobutyric Acid (GABA)	2.344

) [55].

In addition to essential amino acids, non-essential amino acids contribute significantly to cellular functions. Among them, glutamic acid (41.567 µg mL⁻¹) was the most abundant, widely recognized for its role as a natural flavor enhancer and a neurotransmitter in brain function, making it relevant for applications in the food and pharmaceutical sectors [56,57]. Moreover, proline (2.576 µg mL⁻¹) was identified as one of the major amino acids in this strain. Proline is classified as a conditionally essential amino acid that plays a key role in osmotic regulation and oxidative stress mitigation, particularly under environmental stress conditions [58].

The presence of gamma-aminobutyric acid (GABA, 2.344 µg mL⁻¹), a key inhibitory neurotransmitter, suggests additional functional applications of this strain in neurological health-related products [59]. Additionally, alanine (20.280 µg mL⁻¹) and glycine (2.272 µg mL⁻¹) were detected at relatively high concentrations compared to other amino acids, highlighting this strain’s distinctive amino acid profile and its potential implications for metabolic functions and industrial applications [60].

The diverse presence of essential, non-essential, and conditionally essential amino acids underscores the potential industrial applications of this strain, particularly in the development of functional food ingredients, feed supplements, and pharmaceutical compounds.

3.4.4. Fatty Acid Methyl Ester Analysis

Fatty acid methyl ester (FAME) analysis provides information on the carbon chain length and degree of unsaturation, as well as the ratio of saturated fatty acids (SFA) to unsaturated fatty acids (UFA). The results indicate that the highest proportion of ω-3 fatty acids was 53.59%, comprising 1.88% C16:3 (ω-3), 16.38% C16:4 (ω-3), 29.56% C18:3 (ω-3), and 5.78% C18:4 (ω-3) (

Table 3. The composition of fatty acid methyl esters (FAMEs) in Desmodesmus sp. KNUA231.

(%)	Desmodesmus sp. KNUA231
C16:0	22.54
C16:1	2.40
C16:2 (ω6)	2.40
C16:3 (ω3)	1.88
C16:4 (ω3)	16.38
C18:0	0.78
C18:1	9.12
C18:2	9.17
C18:3 (ω3)	29.56
C18:4 (ω3)	5.78
Saturated fatty acid	19.99
Monounsaturated Fatty Acids	11.81
Polyunsaturated Fatty Acids	68.31

).

Omega-3 fatty acids have been investigated for their potential therapeutic applications, particularly in mitigating brain damage, reducing oxidative stress, and alleviating neuroinflammation [61].

3.5. Biodiesel Quality

The quality of biodiesel is determined by the type and concentration of FAME, and Desmodesmus sp. KNUA231 exhibits a kinematic viscosity (υ) of 3.28 mm⁻² s⁻¹ (

Table 4. Biodiesel quality properties of Desmodesmus sp. KNUA231 compared with EN14214 and ASTM D6751 standards.

	Desmodesmus sp. KNUA231	EN14214	ASTM D6751
Saponification value (mg KOH g−1)	167.29
Iodine value (g I2 100 g−1 fat)	169.04	≤120
Cetane number	38.19	≥51	≥45
Degree of unsaturation	148.4
Cold filter plugging point (°C)	−9.4	−20~0
Oxidation stability (110 °C, h)	5.0	≥6	≥3
Kinematic viscosity (mm2 s−1)	3.28	3.5	1.9~6.0
Density (15 °C) (g cm−3)	0.89	0.872~0.878

). Kinematic viscosity is a physical property that quantifies a fluid’s resistance to flow, influencing fuel injection and combustion in the engine. Excessive viscosity can lead to issues in the fuel injection system, while insufficient viscosity may compromise the lubricating properties of the fuel. The results indicate that the measured values fall within the acceptable range of 1.9 to 6.0 mm⁻² s⁻¹, as specified by the ASTM D6751 standard [32]. IV (Iodine Value) is determined by the reaction of iodine with double bonds in unsaturated fatty acids, allowing for the quantification of unsaturation levels. A higher number of double bonds increases the susceptibility to oxidation, which can negatively impact fuel stability. To ensure long-term storage and maintain fuel quality, EN 14214 sets a limit of IV ≤ 120. However, a high IV also indicates lower viscosity and better fluidity, which are desirable properties for bio-lubricant applications. The oxidation stability of the biodiesel was measured at 110 °C and recorded at a value of 5.0 h, which represents the time the fuel remains stable before oxidation begins. This result meets the ASTM D6751 requirement of at least 3 h, demonstrating sufficient oxidative resistance. Additionally, the Cold Filter Plugging Point (CFPP), which indicates the lowest temperature at which fuel can pass through a filter without clogging, was −9.4 °C. This value complies with the EN 14214 standard range of −20 to 0 °C, confirming its suitability for use in cold climates.

The high iodine value (IV) observed in this study led to a relatively low cetane number (CN), which may pose a risk of not meeting the ignition quality standards. However, a high IV also indicates low viscosity and high fluidity, suggesting that Desmodesmus sp. KNUA231 holds considerable potential for application in the bio-lubricant sector. In addition, the biodiesel analyzed in this study met the ASTM D6751 standards for both kinematic viscosity and oxidative stability. This implies that further optimization, such as process control or the addition of antioxidants, could enhance the fuel’s overall stability and broaden its application potential.

3.6. Proximate Analysis and Ultimate Analysis

Thermogravimetric analysis (TGA) provides critical insights into the composition of biomass and serves as a key parameter in evaluating its suitability as a biofuel feedstock by quantifying moisture, volatile matter (VM), and ash content. Among these, the moisture content should remain below 10% to avoid the need for additional drying, which would increase processing costs and reduce conversion efficiency [62]. Desmodesmus sp. KNUA231 exhibited a moisture content of 4.04 ± 0.51% (wt) (

Table 5. Proximate and ultimate analyses of Desmodesmus sp. KNUA231 biomass.

(%)	Desmodesmus sp. KNUA231
Proximate analysis (wt%)
Moisture	4.04 ± 0.51
Volatile matter	89.56 ± 1.11
Ash	6.40 ± 1.40
Ultimate analysis (wt%)
Carbon (C)	53.14 ± 0.16
Hydrogen (H)	7.67 ± 0.16
Oxygen (O)	25.16 ± 0.01
Nitrogen (N)	8.80 ± 0.04
Sulfur (S)	0.30 ± 0.05
CV * (MJ kg−1)	25.49 ± 0.28

* CV: Calorific value.

), which falls within the acceptable range, ensuring its efficient utilization in biofuel applications without supplementary drying. A high volatile matter (VM) content enhances fuel reactivity, lowers the ignition temperature, and improves overall combustion efficiency. Many conventional lignocellulosic biomass materials, such as agricultural residues (65.15%) [24] and fuel briquettes (65.99%) [63] derived from biomass residues, exhibit lower VM levels compared to microalgae-derived biomass. In contrast, Desmodesmus sp. KNUA231 demonstrated a VM content of 89.56 ± 1.11%, indicating superior combustion efficiency and enhanced thermal energy output.

A detailed analysis also allows for the determination of CV, which represents the energy released during combustion. Desmodesmus sp. KNUA231 exhibited a CV of 25.49 ± 0.28 MJ kg⁻¹, which is substantially higher than that of conventional biomass feedstocks such as bamboo (15.66 MJ kg⁻¹) and corn cobs (19.23 MJ kg⁻¹) [64]. A higher CV indicates greater energy density, positioning Desmodesmus sp. KNUA231 as a promising candidate for biofuel production. The CV of biofuels is predominantly influenced by carbon and hydrogen content, with higher values correlating with increased energy density. The exceptional CV observed in Desmodesmus sp. KNUA231 underscores its potential as a high-energy-density biofuel that is comparable to or exceeds existing biomass feedstocks (Table S4).

3.7. Nutrient and Metal Composition Analysis

Trace elements and heavy metals in biological samples can be analyzed using ICP-MS and ICP-OES, which are widely applied in assessing the potential of biofertilizers. In this study, the macronutrient, micronutrient, and heavy metal concentrations in the microalgal-derived biofertilizer were evaluated against EU fertilizer standards [65]. Macronutrients were converted into their oxidized forms for direct comparison with EU regulations. The results confirmed that key nutrients such as phosphorus, potassium, calcium, magnesium, sodium, and sulfur were present at sufficient levels (

Table 6. Nutrient and metal composition analysis of Desmodesmus sp. KNUA231.

	Element	Concentration (mg kg−1)
Macronutrient	K	8495.826
	Ca	1558.945
	Mg	2148.444
	Na	1519.323
	S	5595.413
	P	5877.109
	Si	74.623
Micronutrient	Zn	55.919
	Cu	20.072
	Mn	407.678
	Fe	1972.932
	B	N.D. *
	Mo	N.D.
Metal	As	0.482
	Cd	0.02
	Pb	0.366

* N.D.(Not Detected): below 10 mg L⁻¹.

). For comparison, the EU requires a minimum of 10,000 mg kg⁻¹ P₂O₅ for phosphorus, 30,000 mg kg⁻¹ K₂O for potassium, 12% CaO for calcium, 5% MgO for magnesium, and 10% SO₃ for sulfur. Sodium remained within the acceptable limits, ensuring no risk of excessive salt accumulation.

Micronutrient analysis showed that zinc, copper, manganese, and iron concentrations were above the EU threshold for inorganic micronutrient fertilizers, which requires at least 2% for liquid fertilizers and 5% for solid fertilizers. However, boron and molybdenum were not detected, which does not meet the minimum EU requirement for micronutrient fertilizers (boron ≥ 10 mg kg⁻¹, molybdenum ≥ 5 mg kg⁻¹). While the biofertilizer provides a rich source of trace elements required for plant metabolism, supplementation with boron and molybdenum may be necessary to achieve a more balanced nutrient profile.

Heavy metal concentrations were within safe limits according to EU organic fertilizer regulations. Arsenic was well below the EU limit of 40 mg kg⁻¹, cadmium remained significantly lower than the EU limit of 1.5 mg kg⁻¹, and lead was well below the threshold of 120 mg kg⁻¹. These results suggest that the microalgal biofertilizer offers a high nutrient profile while maintaining environmental safety, highlighting its potential for sustainable agricultural use.

The overall findings indicate that Desmodesmus sp. KNUA231 could serve as a promising alternative to conventional fertilizers. The high macronutrient and micronutrient content ensures an adequate nutrient supply for plant growth, while the low heavy metal levels minimize environmental contamination risks. Moreover, microalgae-based biofertilizers can be produced sustainably, reducing dependency on synthetic fertilizers and mitigating the ecological impact of agricultural practices. However, the absence of certain essential micronutrients, such as boron and molybdenum, suggests the need for further optimization in nutrient formulation. Future research should explore the supplementation of these missing elements and assess long-term soil and crop responses to microalgal biofertilizer application.

4. Conclusions

This study examined the growth performance and adaptability of Desmodesmus sp. KNUA231 under pH and salinity stress conditions to assess its potential as a biofuel resource. The strain exhibited stable growth across a broad pH range (4–10) and moderate salinity levels (up to 5 g L⁻¹ NaCl), demonstrating its ability to thrive under diverse environmental conditions. This environmental adaptability suggests that Desmodesmus sp. KNUA231 may contribute to cost-effective large-scale cultivation by reducing the sensitivity to fluctuating culture conditions. FAME analysis indicated that the biodiesel properties of Desmodesmus sp. KNUA231 comply with ASTM and EN standards in specific parameters, supporting its feasibility as a renewable bioenergy feedstock. The strain exhibited a favorable fatty acid profile, contributing to desirable biodiesel characteristics such as kinematic viscosity, oxidative stability, and cold filter plugging point (CFPP). Furthermore, its high calorific value (CV) confirms its potential as an energy-dense biomass source. In addition, the strain accumulated essential nutrients while maintaining low heavy metal levels, supporting its potential as a biofertilizer. Overall, the findings highlight Desmodesmus sp. KNUA231 as a valuable candidate for third-generation bioenergy production, particularly in non-optimal environmental conditions where pH and salinity fluctuations are common.