Enhanced Biofuel Production from Mixed Marine Microalgae Using UV and UV/H₂O₂ Pretreatment: Optimization of Carbohydrate Release and Fermentation Efficiency

Abstract

The robust structure of algal cell walls presents a major barrier in the recovery of fermentable sugars and intracellular lipids for biofuel production. This study investigates the effectiveness of ultraviolet (UV) radiation and UV-assisted hydrogen peroxide (UV/H₂O₂) pretreatment on a local mixed marine algal culture to enhance biofuel production through cell wall disruption. Local mixed cultures of marine microalgae (LMCMA) were pretreated with UV for various exposure times (5–30 min) and with UV/H₂O₂ using H₂O₂ concentrations ranging from 0.88 to 3.53 mM. The impact of pretreatment was evaluated based on morphological changes (SEM and TEM), elemental composition (C, H, N), sugar release, and downstream fermentation yields of ethanol, methanol, 1-propanol, 1-butanol, and 1-pentanol using Saccharomyces cerevisiae. UV pretreatment at 20–30 min yielded the highest carbohydrate release (up to 0.025 g/gDCW), while UV/H₂O₂ at 1.76 mM achieved maximum sugar liberation (0.0411 g/gDCW). Fermentation performance was enhanced under optimized conditions, with peak ethanol yields of 0.3668 g ethanol/g carbohydrates (UV, 30 min, 48 h) and 0.251 g ethanol/g (UV/H₂O₂, 0.88 mM, 24 h). This study also demonstrated selective production of higher alcohols under varying fermentation temperatures (30–37 °C). These findings highlight the potential of combining oxidative pretreatment and process optimization to enhance biofuel recovery from environmentally relevant algal biomass.

1. Introduction

The rising global demand for renewable energy, coupled with increasing environmental concerns over fossil fuel use, has intensified the search for sustainable and eco-friendly alternatives [1,2]. Microalgae have emerged as a highly promising bioresource for third-generation biofuel production due to their fast growth rates, high photosynthetic efficiency, and ability to accumulate energy-rich compounds such as lipids and carbohydrates [3,4,5]. Unlike terrestrial crops, microalgae do not compete with food production, can thrive in saline or wastewater environments, and offer the advantage of year-round cultivation [6]. However, the commercial viability of microalgae-based biofuels is still hindered by key technical challenges, particularly the robust and recalcitrant nature of algal cell walls, which significantly limits the recovery of intracellular carbohydrates and lipids for biofuel production [7,8,9].

To overcome this bottleneck, several pretreatment strategies have been explored to enhance cell disruption and improve the yield of fermentable substrates. Among them, advanced oxidation processes (AOPs) such as ultraviolet (UV) irradiation and UV-assisted hydrogen peroxide (UV/H₂O₂) pretreatment have shown notable potential [10,11,12]. UV radiation induces oxidative stress in algal cells, damaging cellular membranes and walls, and promoting the leakage of intracellular contents. Studies have shown that controlled exposure to UV radiation can enhance both lipid and carbohydrate yields [10]. For instance, Jongput et al. [13] demonstrated that Botryococcus braunii achieved a biomass yield of 2.45 g/L and a hydrocarbon content of 55.01% after 9 h of UV exposure. Similarly, Kumar et al. [14] reported that UV-C radiation can alter fatty acid profiles, favoring saturated lipid accumulation, which is beneficial for biodiesel quality.

UV/H₂O₂ pretreatment offers an even more effective mechanism for algal cell wall disruption through the generation of highly reactive hydroxyl radicals (•OH). These radicals degrade structural polysaccharides, enhancing cell permeability and promoting the release of fermentable carbohydrates. In a recent review, Liu et al. [15] observed an increase in lipid yield and an upregulation of fatty acid biosynthesis genes in Chlorella sp. following combined UV and H₂O₂ treatment. Di Fraia et al. [10] and Yang et al. [16] have demonstrated that UV-assisted oxidation processes efficiently break down complex polymers and improve substrate availability for downstream processes. However, at higher H₂O₂ concentrations (≥2.65 mM), sugar degradation was observed, likely due to over-oxidation and the formation of inhibitory compounds [17].

Despite promising outcomes from recent studies, several critical gaps in the literature remain. Most current research on UV and UV/H₂O₂ pretreatment focuses on pure algal strains under laboratory conditions, which do not fully reflect the complexity and heterogeneity of natural or industrial algal cultures. There is limited understanding of how these treatments affect mixed marine algal cultures, which are more relevant for large-scale biofuel operations due to their environmental adaptability and reduced cultivation costs. Furthermore, while the impact of UV and UV/H₂O₂ on lipid enhancement has been fairly well explored, fewer studies have examined their role in carbohydrate liberation and subsequent biofuel yields across multiple fermentation products, including ethanol, methanol, 1-propanol, 1-butanol, and 1-pentanol. The link between pretreatment-induced morphological and elemental changes and their effects on fermentation efficiency remains largely unexplored.

This work aims to bridge the aforementioned gaps by systematically investigating the effects of UV and UV/H₂O₂ pretreatment on a local mixed culture of marine microalgae (LMCMA). The research introduces several novel aspects, including the utilization of natural mixed algal cultures to reflect real-world biomass conditions, increasing the scalability and environmental relevance of the findings; evaluation of both lipid and carbohydrate yields following pretreatment, covering biofuel potential from both fermentation and transesterification pathways; quantitative analysis of five bioalcohols (ethanol, methanol, 1-propanol, 1-butanol, and 1-pentanol) under varying fermentation durations and temperatures; and posttreatment morphological characterization of algal cells, using SEM and TEM to directly correlate structural damage with biofuel yield outcomes.

We also conducted elemental composition analysis (C, H, N) of biomass before and after pretreatment, highlighting changes in substrate availability and microbial nutrient profiles. Most previous studies focused on either structural analysis or sugar yields in isolation, whereas this study offers a comprehensive view of how pretreatment conditions impact fermentable sugar release and downstream biofuel production. The novelty of this work lies in its integrative analysis of pretreatment duration, oxidant dosage, fermentation variables, and biofuel product profiles, providing critical insights into optimizing microalgal biomass utilization for bioenergy.

2. Material and Methods

2.1. Microalgae Cultivation and Analysis

Local mixed cultures of marine microalgae (LMCMA) were cultivated under controlled laboratory conditions. The LMCMA used in this study were collected from the Arabian Gulf, specifically near the Doha West Bay area (Qatar). The growth medium was prepared using natural seawater, which was filtered and autoclaved before use to ensure sterility. The medium was supplemented with essential nutrients, including nitrate, phosphate, and trace metals, to support optimal growth conditions. Cultivation was carried out in 500 mL Erlenmeyer flasks containing 250 mL of the prepared medium. The cultures were maintained at 25 ± 2 °C under illumination, with light intensity of 80–100 µmol photons m⁻² s⁻¹, and with a 12:12 light/dark photoperiod. Continuous aeration was provided using filtered air to ensure mixing and adequate CO₂ supply. The LMCMA growth was monitored by measuring the optical density at 680 nm and dry cell weight (DCW). Cells were harvested during the late exponential to early stationary phase for downstream processing and biomass characterization. All cultivation experiments were performed in duplicate, and growth data are presented with a 90% confidence level. Statistical analyses, including means, standard deviations, and confidence intervals, were applied to ensure accurate and reproducible assessment of microalgal growth performance.

The modified phenol–sulfuric acid method was used for carbohydrate extraction from dry biomass, as described by DuBois et al. [18]. The first step was adding 4 mL of acetic acid to a 15 mg dry biomass, and the samples were vortexed at low speed. The samples were kept in a water bath for 1 h at 80 °C. The samples were then cooled. After removing the supernatant, 4 mL of acetic acid was added and kept in the water bath until the samples turned colorless. Next, 5 mL of acetone was added and thoroughly mixed by vortexing at high speed. The samples were centrifuged at 4000 rpm for 10 min. Then, 5 mL of (4 M) HCL was added to the removed supernatants and vortexed at medium speed. The samples were then placed in a boiling water bath for 2 h and were vortexed every 30 min. Then, 900 μL from the sulfuric acid and phenol reagent mixture solution was added to the supernatant, vortexed, and placed in a boiling water bath for 20 min, and 300 μL from the samples was used to measure the carbohydrate concentration from the D-glucose standard calibration at 490 nm. Carbohydrate measurements were performed in triplicate, and the results are statistically represented with a 90% confidence level to ensure data reliability and reproducibility. Standard deviations and confidence intervals were calculated and reported where applicable to reflect the precision of the experimental outcomes.

Bioethanol production was carried out in a 150 mL glass bioreactor containing 100 mL of pretreated LMCMA suspension. The mixture was inoculated with Saccharomyces cerevisiae at a concentration of 10% v/v. The yeast was freshly cultured on an agar plate containing peptone, glucose, and yeast extract to ensure viability. Fermentation was conducted at 30 °C and pH 5.5 under continuous stirring at 135 rpm. The inoculated cultures with pretreated LMCMA were incubated for either 24 or 48 h. Upon completion of the fermentation period, the produced biofuels were collected and analyzed.

Biofuel was quantified and analyzed using a Headspace/Gas Chromatograph/Mass Spectrometer (HS/GC/MS, Agilent, Santa Clara, CA, USA). The gas chromatograph was equipped with a DB-624UI (30 m × 0.25 mm, 1.4 μm) column to determine the following alcohol contents: ethanol, methanol, butanol, 1-pentanol, and 1-propanol. The oven temperature was programmed with an oven temperature of 230 °C, with an initial heating phase at 35 °C for 3 min, and then increased at a rate of 15 °C/min until it reached 170 °C, which was maintained for 2 min. Helium gas was employed as a carrier at a 12 mL/min flow rate and held for 5 min, subsequently transitioning to 1.6 mL/min and holding for 18 min.

The Lowery method [19] was used to determine the percentage total protein content (%). First, 1.5 mL of (0.1 N) NaOH was added to 10 mg of dry biomass, and the total protein extraction was achieved by incubating the samples in an oven at 60 °C for 24 h. Following that, the samples were centrifuged for 5 min at 7000 rpm. Then, the collected supernatants were used to measure the protein content (%) per dry biomass. A protein assay with Folin–Ciocâlteu reagent and bovine serum albumin (BSA) as the standard solution was used to measure the protein content. The protein content measurements were carried out in triplicate, and the results are reported with a 95% confidence level.

The total lipid content was quantified using the modified Folch method [20]. A 15 mg sample of dry biomass was treated with 40 μL of a (1 M) NaCl solution. A solution of chloroform/methanol at a ratio of 2:1 was added to the samples. Initially, 480 μL of methanol was added, and the samples were stored at 4 °C in the dark for 24 h. Following that, chloroform was added to the samples, followed by centrifugation of the samples for 5 min at 7000 rpm. The supernatants were transferred to other tubes to separate the aqueous and organic phases. A second extraction was performed by combining a 2:1 ratio of chloroform and methanol, followed by adding 40 μL of (1 M) NaCl and vigorously swirling the tube. After removing the aqueous top layer, the organic layer was dried, and the tube was weighed to determine the total lipid content per dry biomass (%), as shown in Equation (1) [21]. The total lipid content analysis was conducted in triplicate, and the results are reported with a 95% confidence level.

$$\mathrm{L}\mathrm{i}\mathrm{p}\mathrm{i}\mathrm{d} \mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{e}\mathrm{n}\mathrm{t} \left(\mathrm{\%}\right)={\displaystyle \frac{\mathrm{T}\mathrm{o}\mathrm{t}\mathrm{a}\mathrm{l} \mathrm{l}\mathrm{i}\mathrm{p}\mathrm{i}\mathrm{d}\mathrm{s} \left(\mathrm{g}\right)}{\mathrm{D}\mathrm{r}\mathrm{y} \mathrm{b}\mathrm{i}\mathrm{o}\mathrm{m}\mathrm{a}\mathrm{s}\mathrm{s} \left(\mathrm{g}\right)}}\ast 100$$

(1)

2.2. Pretreatment Methods

The disruption of LMCMA cell walls markedly facilitated the release of intracellular carbohydrates and sugars, thereby enhancing the overall efficiency of the biomass conversion process. As reported in the literature, the sugar yield from microalgal biomass is largely governed by the intrinsic carbohydrate content of the cells [22]. The efficiency of carbohydrate and sugar liberation from LMCMA was further improved through the application of various pretreatment strategies, including ultraviolet (UV) irradiation and UV/H₂O₂ treatment.

The pretreatment of LMCMA biomass was conducted using two approaches: ultraviolet (UV) radiation alone, and UV combined with hydrogen peroxide (UV/H₂O₂). A germicidal UV G20T10 lightbulb (21 watts, 253.7 nm wavelength, two-pin) was employed for both methods, with a constant working volume of 100 mL and continuous stirring at 450 rpm to ensure uniform mixing.

For the UV-only pretreatment, LMCMA biomass with a concentration of 0.23 ± 0.01% (w/v) was exposed to UV light for varying durations of 5, 15, 20, and 30 min to evaluate the effect of exposure time on cell rupture.

For the UV/H₂O₂ pretreatment, 30% (w/w) hydrogen peroxide (Sigma-Aldrich, St. Louis, MO, USA) was added to the microalgae suspension with a biomass concentration of 0.20 ± 0.02% (w/v). The reaction was conducted under UV light exposure for 30 min, using different H₂O₂ concentrations of 0.88, 1.76, 2.65, and 3.53 mM per 100 mL to assess the influence of oxidant dosage on the efficiency of cell rupture.

3. Results and Discussion

3.1. Impact of UV on Algal Cell Destruction

The impact of UV irradiation on LMCMA cell wall disruption was investigated across various exposure durations (5, 15, 20, and 30 min). The physicochemical properties of LMCMA following UV pretreatment and the subsequent fermentation process are summarized in

Table 1. Elemental compositions of microalgae biomass after UV pretreatment and fermentation.

Fermentation Time (h)	UV Duration Time (min)	After Fermentation ppm				After UV Reaction ppm				After UV Pretreatment Dry Microalgae w%
		TOC (Mean ± 95% CI)	TC (Mean ± 95% CI)	IC (Mean ± 95% CI)	TN	TOC	TC	IC	TN	C% (Mean ± 95% CI)	N% (Mean ± 95% CI)	H% (Mean ± 95% CI)
24	5	0.32 ± 0.025	0.36 ± 0.025	0.04 ± 0	0	7.2 ± 0.9	23.4 ± 2.9	16.3 ± 2.0	14.35 ± 0.517	38.3 ± 3.9	6.51 ± 0.48	7.31 ± 0.445
	15	0.0833 ± 0.014	0.0833 ± 0.014	0	0	10.2 ± 1.3	21.1 ± 2.6	10.9 ± 1.4	19.25 ± 0.417	38.5 ± 3.9	6.48 ± 0.48	6.89 ± 0.245
	20	0.153 ± 0.014	0.166 ± 0.014	0.01	0	8.9 ± 1.1	29.1 ± 3.6	20.1 ± 2.5	21.41 ± 0.217	37.9 ± 3.8	6.52 ± 0.48	6.23 ± 0.115
	30	0.163 ± 0.014	0.164 ± 0.014	0	0	16.6 ± 2.1	29.6 ± 3.7	13.0 ± 1.6	10.26 ± 0.22	38.8 ± 3.9	6.52 ± 0.48	6.88 ± 0.22
	45	0.147 ± 0.014	0.153 ± 0.014	0.01 ± 0		15.6 ± 1.9	27.6 ± 3.4	12.0 ± 1.5	9.50 ± 0.35	35.8 ± 3.6	9.51 ± 0.72	7.01 ± 0.2
48	5	0.117 ± 0.014	0.157 ± 0.014	0.02 ± 0	0	7.3 ± 0.92	23.1 ± 2.9	15.7 ± 2.0	19.77 ± 0.57	40.8 ± 4.1	6.70 ± 0.5	7.01 ± 0.15
	15	0.24 ± 0.025	0.123 ± 0.014	0	0	11.7 ± 1.5	23.9 ± 3.0	12.2 ± 1.5	22.87 ± 0.66	40.7 ± 4.1	6.67 ± 0.48	6.89 ± 0.79
	20	0.0640 ± 0.014	0.0662 ± 0.014	0.01 ± 0	0	4.90 ± 0.61	16.6 ± 2.1	11.7 ± 1.5	33.34 ± 0.850	40.4 ± 4.1	6.71 ± 0.51	8.19 ± 0.78
	30	0.154 ± 0.014	0.154 ± 0.014	0	0	8.361 ± 1	26.1 ± 3.3	17.7 ± 2.2	26.6 ± 0.720	41.2 ± 4.1	6.67 ± 0.5	8.14 ± 0.72
	45	0.143 ± 0.014	0.147 ± 0.014	0.01		8.07 ± 1	24.7 ± 3.1	16.4 ± 2.0	24.37 ± 0.71	39.7 ± 4	6.54 ± 0.48	7.9 ± 0.33
Control		-	-	-	-	-	-	-	-	44.3 ± 4.5	7.57 ± 0.57	8.2 ± 0.30

CI: confidence interval.

.

The concentrations of total carbon (TC), total organic carbon (TOC), total nitrogen (TN), and inorganic carbon (IC) (ppm) in the supernatants following UV pretreatment—as well as the elemental composition (C, N, H in wt%) of the dry microalgal biomass—demonstrate the substantial impact of pretreatment conditions on the disintegration of LMCMA cells. The release of intracellular components (sugar, protein, and carbohydrates) into the solution enhances the availability of fermentable substrates, thereby improving the efficiency of S. cerevisiae fermentation. This increased availability is expected to positively influence biofuel production.

The UV exposure facilitated the solubilization of both organic and inorganic carbon, as evident from the measurements after the UV reaction. The highest TOC observed after UV treatment alone was 16.6 ± 2.1 ppm at 30 min of irradiation, followed by 11.7 ± 1.5 ppm at 15 min, clearly demonstrating the time-dependent nature of carbon release. When the UV irradiation time was extended to 45 min, the TOC concentration decreased by approximately 7%. This reduction may be attributed to the further mineralization of the released organic matter under prolonged UV irradiation, leading to its conversion into smaller, non-fermentable compounds such as CO₂.

The total carbon (TC) and inorganic carbon (IC) fractions indicate the speciation of released carbon. For instance, at 20 min, the IC was highest (20.1 ± 2.5 ppm), whereas at 15 min, the IC was much lower (10.9 ± 1.4 ppm), suggesting that UV intensity and duration modulate both the quantity and type of carbon released. These carbon species serve as precursors for fermentation in the subsequent step. The release of total nitrogen (TN) also followed a variable pattern, peaking at 33.34 ± 0.850 ppm at 20 min and 26.6 ± 0.720 ppm at 30 min, confirming that UV effectively disintegrated nitrogen-containing cellular components, such as proteins and amino acids, into soluble forms. This bioavailable nitrogen is critical for yeast growth but must be carefully balanced to optimize alcohol profiles during fermentation. The observed trend suggests that UV treatment should be stopped after 30 min. Extending the irradiation beyond this point not only results in excessive mineralization of the solution but also increases the pretreatment cost and potentially reduces the expected biofuel productivity, due to the reduction in the availability of fermentable material. Limited UV oxidation (e.g., 5–15 min) results in the release of only a small fraction of cellular contents, thus leading to low fermentation yields. On the other hand, controlled UV exposure promotes the disintegration of complex biomolecules such as polysaccharides and proteins into simpler, more bioavailable substrates, enhancing their fermentability. Therefore, it is essential to terminate UV treatment at the optimal point—around 30 min—where simple sugars, proteins, and carbohydrates are most available for microbial conversion to biofuel.

Following UV treatment, the released substrates were fermented for either 24 or 48 h. The values reflect how much of the TOC and TC was retained or transformed. Notably, the highest TOC after fermentation (0.32 ± 0.025 ppm) was observed at 24 h fermentation with 5 min UV, suggesting minimal carbon consumption, likely due to the low carbon release during UV treatment (TOC = 7.2 ± 0.9 ppm in that condition). In contrast, at 48 h/15 min, although the TOC after fermentation was lower (0.24 ± 0.025 ppm), the initial TOC from the UV pretreatment was high (11.7 ± 1.5 ppm), indicating substantial microbial conversion during fermentation.

This trend reflects the fermentability of UV-released substrates and their potential to produce alcohols. Lower post-fermentation TOC values in samples with high pre-fermentation TOC (e.g., 24 h/30 min, 48 h/15 min, and 48 h/30 min) imply effective substrate utilization by S. cerevisiae [23]. Appropriate nitrogen availability drastically enhances Saccharomyces cerevisiae activity, leading to increased ethanol production. Conversely, nitrogen limitation shifts microbial metabolism through the Ehrlich pathway, promoting the synthesis of higher (fusel) alcohols as byproducts. Thus, it illustrates the effectiveness of the released nitrogen in the supernatants after microalgae disintegration. Hence, higher alcohols are synthesized as byproducts under nitrogen limitation due to the shift in microbial pathways via the Ehrlich pathway, resulting in the formation of fusel alcohol. In this study, the maximum TOC yield was obtained after 30 min, followed by 15, 20, and 5 min of UV duration. Nonetheless, the recorded results of the soluble TOC indicate that extending the UV time will significantly increase the TOC, thereby affecting ethanol production.

Elemental analysis of the dry biomass following UV pretreatment further supports carbon and nitrogen mobilization. Compared to the control biomass (C% = 44.3 ± 4.5%, N% = 7.57 ± 0.57%), treated samples showed significant decreases in C% (as low as 37.9 ± 3.8%) and N% (down to 6.48 ± 0.48%), indicating intracellular material loss due to UV-induced disintegration. The hydrogen content (H%) increased in several cases after treatment, particularly at 48 h/20 min (8.19 ± 0.78%) and 48 h/30 min (8.14 ± 0.72%), which may signal the accumulation of hydrogen-rich compounds or residual lipids, and which aligns with conditions favoring higher alcohol synthesis.

The carbon-to-nitrogen (C/N) ratio plays a critical role in guiding yeast metabolism during fermentation. A higher C/N ratio promotes ethanol production by favoring fermentative over anabolic pathways, whereas low or unbalanced ratios may reduce the ethanol yield or lead to fusel alcohol formation. In this study, the optimal C/N ratio of 1.62 was achieved under 30 min of UV pretreatment followed by 24 h of fermentation, with TOC and TN values of 16.64 ppm and 10.26 ppm, respectively. This condition provided sufficient carbon while limiting nitrogen, supporting efficient ethanol biosynthesis. In contrast, lower C/N ratios were observed under other conditions: 0.53 at 48 h/15 min (TOC = 11.7 ± 1.5 ppm, TN = 22.87 ± 0.66 ppm), 0.51 at 24 h/15 min (TOC = 10.2 ± 1.3 ppm, TN = 19.25 ± 0.417 ppm), and the lowest ratio of 0.42 at 24 h/20 min (TOC = 8.9 ± 1.1 ppm, TN = 21.41 ± 0.217 ppm). These lower ratios reflect nitrogen-rich environments, which may favor yeast growth over ethanol production. The findings highlight that a balanced yet carbon-lean nutrient profile—achieved through optimized UV treatment—enhances the utilization of microalgal substrates for bioethanol yield.

The structural and biochemical impact of UV pretreatment on LMCMA was comprehensively assessed through SEM and TEM imaging (Figure 1 and Figure 2) and elemental and biochemical data (Table 1). Together, these results highlight the role of UV exposure in disrupting cellular integrity, enhancing the release of fermentable compounds, and influencing fermentation efficiency.

Figure 1a represents the untreated microalgal biomass, characterized by well-defined, intact cell structures and smooth surfaces, indicating solid cell walls. As UV exposure was applied, progressive morphological deformation was observed in the LMCMA cells across Figure 1b–e, corresponding to increasing UV durations of 5, 15, 20, and 30 min, respectively. After 5 min of UV irradiation (Figure 1b), the cell wall damage was minimal, and some species remained morphologically intact, consistent with a relatively low TOC release of 7.2 ± 0.9 (Table 1). The corresponding dry biomass showed moderate carbon content (C% = 38.3 ± 3.9) and nitrogen content (N% = 6.51 ± 0.48%), indicating limited intracellular leakage. As the UV pretreatment proceeded between 15 and 20 min (Figure 1c,d), the deformation became more pronounced, with increased agglomeration and disrupted surface morphology. This aligns with higher TOC values of 10.2 ± 1.3 ppm and 8.9 ± 1.1 ppm, and TN values of 19.25 ± 0.417 and 21.41 ± 0.217 ppm, respectively, suggesting enhanced release of soluble organics from broken cell walls. At 30 min (Figure 1e), extensive rupture and aggregation were evident, particularly in the surface layers and intracellular contents. This condition corresponds to the highest TOC release (16.6 ± 2.1 ppm) and a significant decrease in TN to 10.26 ± 0.22 ppm, reflecting strong solubilization of cellular contents, most notably carbohydrates and proteins. The obtained results confirmed that UV exposure for 30 min caused the most effective physical disruption, which directly correlates with maximized biochemical release. Liza et al. [24] showed that ozone pretreatment enhanced biohydrogen production from microalgae biomass. It was highlighted that ozone improves cell wall disruption, increases the release of intracellular organics, and optimizes the hydrogen yield.

The TEM images in Figure 2 confirm the internal structural degradation of the LMCMA. Untreated LMCMA samples (Figure 2a) show intact cell membranes and dense internal structures, indicating no disruption. UV-treated cells (Figure 2b–e) reveal progressive membrane thinning, cytoplasmic leakage, and overall collapse of the cellular architecture, with the severity increasing with the exposure duration. Figure 2e (30 min UV) shows fragmented structures and loss of intracellular compartmentalization. Such microstructural damage aligns with the observations of Miao and Tao [25], who documented similar membrane degradation and DOC release after ozonation—a parallel oxidative pretreatment approach. This correlates with the substantial elemental depletion in biomass observed in Table 1: the carbon content dropped from 44.624% (control) to 38.803%, and the nitrogen content declined from 7.569% to 6.516%, reflecting the exodus of organics into the surrounding medium. The morphological degradation observed in Figure 1 and Figure 2 supports the biochemical trends shown in Table 1. The optimal UV exposure (especially 30 min) not only improved the TOC release but also optimized the C/N ratio (1.62 at 24 h/30 min) and enhanced the fermentable substrate availability, particularly carbohydrates and proteins—critical factors for efficient fermentation processes.

3.2. Effects of UV Photolysis Time

Figure 3 illustrates the impacts of UV irradiation time (5, 15, 20, and 30 min) on both the quantity of carbohydrate released from LMCMA and the corresponding release efficiency (%). After 5 min of UV irradiation, the released carbohydrate was approximately 0.014 ± 0.005 g_carb/g_DCW, with a release efficiency of 25%. Increasing the UV exposure to 15 min significantly enhanced the carbohydrate release to 0.018 ± 0.003 g_carb/g_DCW and an efficiency of 32.1%, indicating more effective cell wall disruption and solubilization of polysaccharides. The maximum values were achieved at 20 min, where the released carbohydrate peaked at 0.025 ± 0.004 g_carb/g_DCW and the efficiency reached 44.65%. However, extending the UV treatment to 30 min resulted in a slight reduction, with a carbohydrate release of 0.024 ± 0.003 g_carb/g_DCW and an efficiency of 42.96%. This decline suggests that excessive UV exposure may lead to the degradation of released sugars into non-fermentable compounds or the formation of inhibitory byproducts such as reactive oxygen species or oxidized intermediates. The error bars confirm the reliability of the data, although the variability slightly increases at longer durations. Collectively, the trend highlights that 20 min of UV pretreatment provides the optimal balance between efficient carbohydrate release and minimal degradation, making it the most effective duration for maximizing substrate availability for downstream biofuel fermentation.

The progressive increase in carbohydrate yield from 5 to 20 min reflects enhanced cell wall disruption due to prolonged UV exposure. At shorter UV irradiation periods (5 and 15 min), the liberation of carbohydrates was low, suggesting insufficient energy input to significantly compromise the structural integrity of the cell walls, thereby limiting the release of intracellular polysaccharides. These observations are consistent with the SEM and TEM observations (Figure 1 and Figure 2), where minimal deformation was seen at 5 min and more extensive membrane damage occurred at higher UV irradiation times. However, a slight decline in carbohydrate yield was observed at 30 min compared to 20 min, despite greater morphological damage. This may be attributed to overexposure to UV radiation, which can lead to the degradation or oxidation of released sugars, as well as the possible formation of recalcitrant compounds that increase COD levels while reducing fermentable sugar concentrations. This trend aligns with the observations of Bhushan et al. [26], who reported that extended UV-C exposure can compromise carbohydrate recovery by over-fragmenting cellular contents or initiating photodegradation pathways. It was shown that UV-induced membrane instability is a key factor in this process. As the UV duration increases, membrane permeability improves, facilitating the release of intracellular carbohydrates. However, beyond an optimal time (e.g., 30 min in this study), further exposure may result in structural collapse or oxidative degradation, diminishing the net recoverable sugar yield. The study of Bhushan et al. [26] showed that the maximum sugar content released was 73.61 mg/L after 10 h of exposure. Thoroughly, the cell rupture led to a gradual decrease in sugar liberation and chlorophyll content, as well as weakness in cell stability. Moreover, the study concluded that long-term UV-C exposure results in cell weakness, wherein the cellular contents, including sugars, are discharged into the surrounding liquid, negatively impacting their potential for biofuel production [26]. On the other hand, limited research and a lack of studies describe the effectiveness of the UV-C pretreatment method for microalgae disruption, which provided new insight into biofuel production.

UV radiation is usually combined with other methods, such as H₂O₂, ozone, microwave, and Fenton reaction methods, to enhance hydroxyl radical formation [27]. The results concluded that further combined pretreatment is necessary to improve the cell wall rupture and enhance intracellular liberation, increasing the fermentation process’s efficacy. Liza et al. [24] investigated a combined chemical and UV pretreatment strategy to modify lignin chromophores in wood residues. The chemical pretreatment involved sodium hydroxide (NaOH) and hydrogen peroxide (H₂O₂), followed by UV irradiation for durations ranging from 30 to 120 min. NaOH was used to facilitate the disintegration of the lignin structure, while H₂O₂ generated hydroxyl radicals that contributed to oxidative degradation. This combination resulted in a slight reduction in lignin concentration. Subsequently, the pretreated wood residues underwent mechanical disruption through ball milling and ultrasonic treatment. Additionally, UV radiation was found to further enhance hydroxyl radical formation, intensifying the modification of lignin chromophores and improving the overall efficiency of the pretreatment process [24]. Wen et al. [28] demonstrated an effective combined pretreatment approach to enhance the production of cellulose nanofibers by integrating UV radiation and ozone exposure. In this process, UV irradiation facilitated the decomposition of ozone, leading to the formation of hydrogen peroxide (H₂O₂) as an intermediate, which further decomposed into additional hydroxyl radicals. These radicals played a crucial role in promoting cellulose oxidation and the degradation of polysaccharides. The study specifically found that applying UV radiation for 30 min in the presence of 12 mmol H₂O₂/g fiber significantly enhanced ozone-induced oxidation by increasing hydroxyl radical production. As a result, the structural integrity of the biomass was compromised, leading to improved cellulose accessibility. The authors concluded that UV pretreatment at 20–30 min effectively ruptures cell walls, increases membrane permeability, and enhances the release of intracellular carbohydrates—findings that align closely with the carbohydrate liberation patterns observed in microalgae under similar UV exposure conditions. Overall, these results confirm that UV pretreatment is effective in enhancing carbohydrate release, with 20 min identified as the optimal duration for maximizing the fermentable sugar yield without incurring significant degradation losses. These findings support the integration of UV exposure as a controllable parameter in pretreatment strategies for biofuel feedstock processing [28].

Further investigation is required to study the effects of viscosity on mixed culture cells, their effectiveness in enabling UV light radiation to penetrate microalgae cell walls, and their potential impact on cell disruption by releasing fermentable carbohydrates. In addition, the viscosity of microalgae fluctuates considerably depending on the microalgae species and their composition [22]. Dense microalgae accumulation will increase the microalgae concentration, diminishing the UV light’s ability to penetrate cell walls, and negatively impacting cell disruption.

3.3. Effects of Fermentation Time

Figure 4 presents the effects of fermentation time (24 and 48 h) on carbohydrate consumption following UV pretreatment at various durations (5, 15, 20, and 30 min), using Saccharomyces cerevisiae under optimal fermentation conditions (pH 5.5). The results reveal a progressive decrease in carbohydrate concentration with extended fermentation time, indicating effective microbial utilization of the released proteins and sugars for biofuel production. As fermentation progresses from 24 to 48 h, the carbohydrate content in the fermentation broth declines accordingly. LMCMA treated with UV for 15 min released 0.018 ± 0.003 g_carb/g_DCW. After fermentation, the carbohydrate concentration decreased to 0.010 g_carb/g_DCW at 24 h and further dropped to 0.005 g_carb/g_DCW at 48 h, reflecting carbohydrate consumption of 50.8% and 75.4%, respectively. Similarly, the carbohydrate concentration for LMCMA treated with UV for 5 min declined to 0.009 g/g_DCW at 24 h and to 0.003 g_carb/g_DCW at 48 h, showing 57.0% and 85.1% consumption, respectively. The most pronounced carbohydrate consumption was observed at 20 min of UV pretreatment, where the carbohydrate concentration decreased to 0.0156 g_carb/g_DCW and to 0.0033 g_carb/g_DCW, representing a 25.9% reduction after 24 h of fermentation and a maximum 80% consumption after 48 h. This condition suggests optimal UV-mediated solubilization of fermentable carbohydrates and their efficient microbial uptake.

These results indicate that even low UV durations can partially facilitate cell contents’ release, albeit with less efficiency than longer exposures. Interestingly, at 30 min of UV pretreatment, the carbohydrate concentration after 24 h remained relatively higher, at 0.018 g_carb/g_DCW, but decreased to 0.006 g_carb/g_DCW after 48 h, still marking 70% of the total initial carbohydrate concentration. This confirmed the previous observation that prolonged UV exposure may lead to partial degradation or oxidation of cell contents, thereby reducing their availability for fermentation over time.

The observed trends highlight a balance between UV-induced sugar release and microbial consumption. While UV enhances cell disruption and carbohydrate release, excessive exposure may degrade fermentable sugars or generate inhibitory byproducts, potentially reducing the fermentation efficiency, as seen at 30 min compared to 20 min.

These findings emphasize that while UV pretreatment is effective in enhancing fermentable sugar release, the combination of moderate UV exposure (e.g., 20 min) and sufficient fermentation duration (48 h) offers the best performance, resulting in up to 80.2% carbohydrate conversion. Further improvements in pretreatment strategies, such as combining UV with enzymatic or mild chemical treatments, may help enhance cell wall disintegration and improve the liberation and preservation of fermentable sugars.

Apollo et al. [29] suggested a combined pretreatment of UV-C and photocatalysis at 60 min of UV duration time for improving the anaerobic digestion biodegradation of methylene blue dye, thereby facilitating methane production [29]. The anaerobic digestion pretreatment, which utilizes a 15 min UV duration, enhances the lipid accumulation in microalgae by up to 48.18% [30]. However, microalgae can be resistant to UV radiation via various mechanisms. The integrated pretreatment with UV effectively reduces the cost of the operation by decreasing the UV duration time [30].

Jeon et al. [31] highlighted the effectiveness of ultrasonic pretreatment of the Scenedesmus obliquus YSW15 microalgae strain, facilitating biomass destruction and changing the surface morphology at different sonication duration times, varying from 0 to 60 min, and sonication temperatures of 45 and 65 °C, resulting in releasing 0.12 g/g of carbohydrate at the optimal sonication time of 15 min, which effectively enhanced ethanol and hydrogen generation through fermentation over 23 days. At fermentation temperatures of 35 and 55 °C, the reduction in carbohydrates in the sonicated samples after 23 days of fermentation reduced from 0.37 to 0.08 g_carb/g_biomass; hence, 89% of the total carbohydrates was diminished by 16 days, followed by a gradual decline in the remaining carbohydrates until the end of the fermentation [31].

3.4. Effects of UV Pretreatment on Biofuel Production

Figure 5 presents the biofuel production from microalgae following UV pretreatment and fermentation with Saccharomyces cerevisiae at 30 °C for 24 and 48 h. Five biofuels were quantified: ethanol, methanol, 1-propanol, 1-butanol, and 1-pentanol. Overall, the UV pretreatment facilitated carbohydrate release and subsequent microbial conversion into bioalcohols, with clear differences in product yields depending on the UV duration and fermentation time. The carbohydrate is consumed during fermentation and potentially converted into biofuel with extended fermentation time. The recorded results are described in terms of biofuel produced (g) per gram of carbohydrates for experiments performed at pH 5.5 after 24 and 48 h of fermentation.

Ethanol was the dominant fermentation product (Figure 5a), with the production yield (Y_prod) increasing significantly from 24 to 48 h. The maximum ethanol Y_prod of 0.3668 g_ethanol/g_carb) was recorded at 30 min UV pretreatment and 48 h fermentation. The ethanol Y_prod reached 0.0218 g_ethanol/g_carb at 15 min of UV duration and 24 h of fermentation, increasing to 0.0792 g_ethanol/g_carb after 48 h. A noticeable decrease in ethanol Y_prod to 0.01643 g_ethanol/g_carb was observed for the LMCMA pretreated with UV for only 5 min and fermented for 24 h; increasing the fermentation time to 48 h increased the ethanol Y_prod to 0.15 g_ethanol/g_carb. The observed tend indicates the presence of long-chain carbohydrates in the LMCMA pretreated with UV for only 5 min, which required a long fermentation time to achieve a reasonable ethanol Y_prod. Moreover, a notable decrease in ethanol Y_prod (0.00157 g_ethanol/g_carb) was observed for LMCMA pretreated with UV for 20 min after 24 h of fermentation. Notably, the ethanol Y_prod slightly increased to 0.0325 g_ethanol/g_carb after 48 h of fermentation. An increase in the ethanol Y_prod was observed at 30 min of UV duration, reaching 0.01537 g_ethanol/g_carb after 24 h, and yielding the maximum ethanol yield of 0.3668 g_ethanol/g_carb after 48 h. The ethanol yields were higher than those obtained at 35 and 37 °C. Notably, the 15 min UV pretreatment also resulted in a substantial ethanol Y_prod (0.0792 g/g at 48 h). At 20 min, although UV caused significant sugar solubilization, the ethanol Y_prod was relatively low at 24 h (0.00157 g/g) and improved to 0.0325 g/g at 48 h, possibly due to the initial degradation of fermentable sugars or delayed microbial adaptation. This trend supports previous findings by Jeon et al. [31], where physical pretreatment (ultrasound) increased ethanol yields by improving substrate accessibility. Additionally, longer fermentation times allow yeast metabolism to fully convert available carbohydrates, consistent with the ~84% carbohydrate reduction at 20 min UV duration. It can be inferred that ethanol generation leads to a decrease in hydrogen production due to the shifting in the microbial metabolism pathways of butyrate to ethanol during fermentation. It was observed that ethanol is the main microbial product from the fermentation process, whereas the bacteria degrade the monosaccharides under the anaerobic process [31].

The methanol production yields (Figure 5b) peaked at 5 min UV (0.0564 g/g at 48 h), followed by a slight decline at higher UV durations. The maximum methanol Y_prod of 0.0134 g_methanol/g_carb was recorded at 5 min after 24 h and increased to 0.0564 g_methanol/g_carb after 48 h. Yields of 0.0131 and 0.0203 g_methanol/g_carb were detected at 15 min after 24 and 48 h, respectively. A decline in the methanol Y_prod was observed with increased UV radiation times of 20 and 30 min, yielding 0.0072 and 0.0033 g_methanol/g_carb after 24 h, respectively, which reached slightly similar yields of 0.033 and 0.032 g_methanol/g_carb after 48 h, respectively. This suggests that short-term UV exposure sufficiently disrupted the cell walls to release methanol precursors without extensive sugar degradation. Similarly, the 1-propanol production yield (Figure 5c) peaked at 30 min UV (0.0512 g/g at 48 h), indicating that more intense pretreatment was necessary to release its precursor molecules. However, the lower yields at 15 and 20 min highlight a non-linear relationship, likely due to intermediate compound volatility or microbial pathway competition. Kushwaha et al. [32] produced 8.873 g/L of butanol by applying acidic hydrolysis as a pretreatment of Lyngbya limnetica cyanobacteria using 1.63 M H₂SO₄ at 100 °C for 60 min, mixed with Clostridium beijerinckii ATCC and adding 10 g/L of glucose to the fermentation medium. Another study by Liu et al. [33] produced 62% isobutanol from Microchloropsis salina biomass treated with diluted sulfuric acid using an E. coli consortium. However, it was concluded that the investigated butanol yields were lower than those measured in the literature.

For 1-butanol (Figure 5d), the highest Y_prod (0.0225 g_1-butanol/g_carb) occurred at 15 min UV after 48 h, followed closely by 5 and 20 min. Prolonged UV exposure (30 min) led to a drastic reduction in methanol Y_prod (0.0032 g/g), suggesting possible inhibitory effects of UV byproducts on butanol-producing enzymes, as described by Kushwaha et al. [32] and Liu et al. [33]. 1-Pentanol exhibited a unique pattern: the highest yield occurred at 20 min UV after 24 h (0.0219 g/g), yet this significantly decreased to 0.0016 g/g at 48 h, indicating rapid metabolism or instability over time. The pentanol pathway, linked to the Ehrlich pathway via leucine catabolism [34], may have peaked early and declined due to the depletion of precursor amino acids or the redirection of microbial metabolism toward ethanol. These results affirm that UV pretreatment is effective for enhancing biofuel yields, with each bioalcohol responding differently to the UV duration and fermentation time. Ethanol benefits from prolonged UV and fermentation, while other alcohols, such as methanol and pentanol, show optimal production at moderate durations. However, UV overexposure may generate inhibitory compounds (e.g., HMF, furfural, acetic acid), as noted by Gao et al. [34], affecting S. cerevisiae’s performance. Song et al. [35] also demonstrated UV-induced microbial stress due to hydroxyl radicals, which may explain the declining yields at 30 min UV for butanol and pentanol.

Figure 5e shows that the maximum Y_prod of 1-pentanol was achieved at 30 °C after 24 h of fermentation and 20 min of UV irradiation pretreatment, reaching up to 0.0219 g_1-pentanol/g_carb. A reduction in the 1-pentanol Y_prod was observed with increased fermentation time, resulting in 0.0016 g_1-pentanol/g_carb after 48 h. At 15 min of UV irradiation pretreatment, a decline was recorded, yielding 0.0207 g_1-pentanol/g_carb after 24 h; however, with an increase in fermentation time, the measured yield after 48 h decreased to 0.0016 g_1-pentanol/g_carb. Additionally, the yields decreased with extended UV time, as illustrated at 30 min, which produced 0.0045 and 0.0002 g_1-pentanol/g_carb after 24 and 48 h, respectively.

The effect of the fermentation temperature was also investigated.

Table 2. Summary of biofuel production following UV pretreatment.

Fermentation Time (h)	UV (min)	Methanol (g/gcarb)			Propanol (g/gcarb)			Butanol (g/gcarb)			Pentanol (g/gcarb)
		30 °C	35 °C	37 °C	30 °C	35 °C	37 °C	30 °C	35 °C	37 °C	30 °C	35 °C	37 °C
24	5	0.0134	0.0088	0.0088	0.0023	0.0018	0.0018	0.0158	0.0088	0.0047
	15	0.0131	0.0131	0.012	0.0031	0.002	0.002	0.0073	0.012	0.0047	0.0207	0.016	0.016
	20	0.0072	0.012	0.0056	0.0002	0.0011	0.0011	0.0024	0.0056	0.0047	0.0219	0	0
	30	0.0033	0.0045	0.0045	0.0022	0.0011	0.0011	0.0032	0.0019	0.0047	0.0045	0	0
48	5	0.0564	0.0094	0.0094	0.021	0.0067	0.0067	0.02	0.012	0.006
	15	0.0203	0.012	0.012	0.0111	0.0086	0.0086	0.0225	0.012	0.006	0.0016	0.0003	0.0003
	20	0.033	0.017	0.007	0.0052	0.005	0.005	0.0167	0.009	0.006	0.0016	0	0
	30	0.032	0.007	0.007	0.0512	0.0045	0.0045	0.0032	0.0071	0.006	0.0002	0	0

summarizes the production of different biofuels at 30, 35, and 37 °C. The maximum ethanol Y_prod was at 35 °C, at 5 min, producing 0.02 g_ethanol/g_carb after 24 h, and increasing to 0.085 g_ethanol/g_carb after 48 h. A decline in ethanol Y_prod of 0.017 g_ethanol/g at 15 min was found after 24 h, which reached up to 0.056 g_ethanol/g_carb after 48 h. There was a reduction in yields at 20 min, recording 0.012 and 0.017 g ethanol/g after 24 and 48 h, respectively, along with a slight decline of 0.010 g_ethanol/g_carb after 24 h, and yielding 0.035 g_ethanol/g_carb after 48 h, with the extended UV duration. Compared to the results obtained at 37 °C, as shown in Table 2, the subsequent reduction in the produced yields was investigated, concluding that the yield is potentially affected by the increase in the fermentation temperature. A decline in ethanol yields was recorded.

The maximum produced ethanol was recorded at 5 min—with 0.0130 g_ethanol/g_carb after 24 h, and increasing to 0.05 g_ethanol/g_carb after 48 h—followed by 0.013, 0.012, and 0.01 g_ethanol/g_carb at 15, 20, and 30 min, respectively, after 24 h, which reached up to 0.04, 0.015, and 0.028 g_ethanol/g_carb after 48 h, respectively. Jeon et al. [31] employed ultrasonic pretreatment of the Scenedesmus obliquus YSW15 microalgae strain for 15 min, which effectively destroyed the microalgae biomass and altered its surface morphology. Ethanol and hydrogen were produced through fermentation over 23 days at different temperatures, achieving an 89% reduction in carbohydrates and producing 5.6 g/L ethanol and 2.5 mL/L hydrogen; the microbial fermentation enhanced the organic carbon content, which requires further economic assessment in the utilization of renewable carbon sources to generate biofuel.

It can be inferred that ethanol generation leads to a decrease in hydrogen production due to the shift in the microbial metabolism pathways of butyrate to ethanol during fermentation. It can be observed that ethanol is the main microbial product from the fermentation process, whereas the bacteria degrade the monosaccharides under the anaerobic process [31].

The maximum methanol Y_prod at 35 °C (0.0134 g_methanol/g) was recorded at 5 min after 24 h, and this increased to 0.0564 g_methanol/g_carb after 48 h. Yields of 0.0131 and 0.0203 g_methanol/g_carb were detected at 15 min after 24 and 48 h, respectively. A decline in methanol Y_prod was observed with increased UV radiation times of 20 and 30 min, yielding 0.0072 and 0.0033 g_methanol/g_carb after 24 h, which reached slightly similar yields of 0.033 and 0.032 g methanol/g after 48 h, respectively. A decline in methanol Y_prod was observed by increasing the fermentation temperature from 35 to 37 °C. LMCMA pretreated with UV irradiation for 20 min and fermented at 35 °C showed methanol Y_prod 0.012 g_methanol/g_carb after 24 h and 0.0.017 g_methanol/g_carb after 48 h. At 37 °C, substantial decreases in the yields were measured with increased fermentation temperature.

Similar methanol production yields were recorded at 5 and 15 min, achieving 0.0088 g_methanol/g after 24 h and increasing to 0.0094 and 0.012 g_methanol/g_carb at 5 and 15 min, respectively. A reduction in methanol Y_prod was recorded at 20 min, producing 0.0056 and 0.007 g_methanol/g_carb after 24 and 48 h, respectively. The lowest methanol production yields were at 30 min, with 0.0045 and 0.07 g_methanol/g_carb after 24 and 48 h, respectively. The findings recorded the maximum methanol production yields at a fermentation temperature of 30 °C after 48 h. Nevertheless, few studies have highlighted methanol generation from treated microalgae biomass after fermentation following UV pretreatment using Saccharomyces cerevisiae. The 1-propanol production yields were demonstrated at 30 °C. The obtained production yields decreased with UV durations of 5, 15, 20, and 30 min, achieving values of 0.0023, 0.0031, 0.0002, and 0.0022 g_1-propanol/g_carb after 24 h, respectively. The recorded production yields after 48 h increased to 0.0210, 0.0111, 0.0052, and 0.0512 g_1-propanol/g_carb, respectively. However, there was a decrease in the yields at 35 °C. The maximum yield at 35 °C was obtained at 15 min—reaching 0.002 g_1-propanol/g after 24 h, which subsequently increased to 0.0086 g_1-propanol/g_carb after 48 h—followed by 5 min, resulting in 0.0018 and 0.0067 g_1-propanol/g_carb after 24 and 48 h, respectively.

A reduction in the 1-propanol Y_prod was seen with extended duration time, at 20 and 30 min, achieving similar production yields of 0.0011 and 0.0011 g_1-propanol/g_carb after 24 h, respectively, and attaining 0.005 and 0.0045 g_1-propanol/g_carb after 48 h, respectively. Likewise, similar production yields of 1-propanol were recorded at 37 °C after 24 h, with a decrease in production yields after 48 h. The produced yields of 1-butanol at 30 °C reached a maximum of 0.0158 g_1-propanol/g after 24 h, significantly increasing to 0.02 g_1-propanol/g_carb after 48 h, at 5 min. The next-highest production yield was at 15 min, which achieved 0.0073 g_1-propanol/g_carb after 24 h and increased to 0.0225 g_1-butanol/g_carb after 48 h. Moreover, a reduction in the production yield was found at 20 min after 24 h of fermentation, resulting in 0.0024 g_1-butanol/g_carb, and reaching up to 0.0167 g_1-propanol/g_carb. Notably, an increase in the yield was observed after 30 min, where a 0.0032 g_1-propanol/g_carb yield was obtained.

Low butanol concentrations may impede and inhibit the enzymatic activity associated with butanol synthesis, particularly during anaerobic fermentation [36]. One of the studies generated a new enzyme, EB4.F, by combining a modified E. coli (EB4.0) integrated with the formate dehydrogenase (fdh1) gene from Saccharomyces cerevisiae to enhance the oxidation of formate to CO₂ to produce NADH, which can subsequently be utilized to produce butanol synthesis [37]. The synthesis of butanol depends on the biomass type utilized; for instance, a study employed a batch reactor with Clostridium beijerinckii BA101, achieving a fermentation duration time of 110 h and yielding 18.9 g/L of butanol from an initial starch concentration of 80 g/L of starch derived from peanuts, producing butanol [38], which indicates the impact of prolonged duration on alcoholic fermentation. Compared with the yields shown in Table 2 at 35 °C, a decrease in yields was recorded at 5 and 15 min, with similar 1-butanol yields of 0.0088 and 0.012 g_1-butanol/g_carb after 24 and 48 h, respectively. There was a decline in the yields at 20 min; the obtained yields were 0.0056 and 0.009 g_1-butanol/g_carb after 24 and 48 h, respectively, and yielded a low concentration of 0.0019 and increased to 0.0071 g_1-butanol/g_carb after 24 and 48 h, respectively. However, the yields decreased at 37 °C, as depicted in Table 2 and the highest butanol yields were found at 15 min, yielding 0.0047 and 0.006 g_1-butanol/g_carb after 24 and 48 h, respectively.

As illustrated by Kushwaha et al., 8.873 g/l of butanol was produced by applying acidic hydrolysis as a pretreatment of Lyngbya limnetica cyanobacteria using 1.63 M H₂SO₄ at 100 °C for 60 min, mixed with Clostridium beijerinckii ATCC and adding 10 g/L of glucose to the fermentation medium [32]. Another study by Liu et al. produced 62% isobutanol from Microchloropsis salina biomass treated with diluted sulfuric acid using an E. coli consortium [33]. However, it was concluded that the investigated butanol yields were lower than those measured in the literature.

The maximum production yield of 1-pentanol was achieved at 30 °C after 24 h at 20 min, reaching up to 0.0219 g_1-pentanol/g. A reduction in the production yield was observed with increased fermentation time, resulting in 0.0016 g_1-pentanol/g_carb after 48 h. At 15 min, a decline was recorded, yielding 0.0207 g of g_1-pentanol/g after 24 h; however, with an increase in fermentation time, the measured production yield after 48 h decreased to 0.0016 g_1-pentanol/g_carb. Additionally, the production yields decreased with extended UV time, as illustrated at 30 min, which produced 0.0045 and 0.0002 g_1-pentanol/g_carb after 24 and 48 h, respectively. Following a decrease in production yields at 35 °C, the highest 1-pentanol yield was recorded at 15 min, reaching up to 0.016 g_1-pentanol/g after 24 h, which then decreased to 0.0003 g_1-pentanol/g_carb after 48 h. The production yields gradually reduced with the increased UV radiation time. The low yield of pentanol declined over time, with a further reduction in pentanol yields after 48 hr, ultimately approaching zero at 20 and 30 min. Compared to the results obtained at 37 °C, a reduction in the production yield was observed with the increase in fermentation temperature, highlighting the effect of temperature on microbial metabolic activity to enhance biofuel synthesis. However, the recorded production yield of pentanol after 24 h was higher than the production yields obtained from butanol and propanol, indicating that the enzymatic pathway for pentanol synthesis might dominate during fermentation. The Ehrlich pathway describes the conversion of the amino acid leucine to α-ketoisocaproate, which is then decarboxylated by the enzyme THI3 to form isoamyl aldehyde, which is subsequently reduced to isoamyl alcohol (pentanol).

It was observed that after 48 h at 35 and 37 °C, the recorded results diminished to zero. This observation suggests either a reduction in the α-amino acids isoleucine and leucine during fermentation—where these α-amino acids were dominant in the microbial pathway, leading to increased pentanol yields compared to those found from 1-butanol and 1-propanol—or a shift in microbial activity after 24 h that produced another higher alcohol. UV pretreatment represents a promising process in biofuel production from microalgae. However, UV pretreatment may generate inhibitors that hinder Saccharomyces cerevisiae fermentation. Gao L. et al. emphasized that the formation of inhibitors as byproducts following the pretreatment impacts carbohydrate recovery [34].

Furthermore, the formation of hydroxymethylfurfural, furfural, and acetic acid from glucose hinders the metabolism of bacteria [34]. As a result, it diminishes the Saccharomyces cerevisiae growth activity, decreasing biofuel production. Song, K., M. Mohseni, and F. Taghipour examined the impact of UVA radiation in the generation of hydroxyl radicals, which directly degraded the E. coli cell walls during pretreatment and subsequently influenced the metabolic activity [35].

3.5. UV/H₂O₂ Pretreatment

The combination of UV radiation with hydrogen peroxide (H₂O₂) is a well-established advanced oxidation process (AOP) that significantly enhances the generation of highly reactive hydroxyl radicals (•OH). These radicals effectively degrade microalgal cell walls, thereby improving the release of intracellular compounds and enhancing biofuel production. However, the efficiency of this pretreatment is highly dependent on the dosage of the oxidant; excessive concentrations may adversely affect microbial metabolism during fermentation, ultimately reducing bioenergy yields [27]. Therefore, it is essential to optimize the pretreatment conditions to focus solely on disrupting the microalgal cell walls and facilitating the release of cellular contents suitable for biofuel and valuable bioproduct production [27]. The integration of UV with H₂O₂ also offers a cost-effective alternative to other AOPs. In this combined process, hydroxyl radicals target and oxidize the microalgal cell wall structure, increase the membrane permeability, and promote the breakdown of fermentable carbohydrates. In this study, the UV/H₂O₂ pretreatment was applied at various H₂O₂ concentrations—0.88, 1.76, 2.65, and 3.53 mM—under a fixed UV exposure time of 30 min.

3.6. The Mechanism of UV/H₂O₂ Oxidation

Ultraviolet (UV) irradiation combined with hydrogen peroxide (H₂O₂) is an effective advanced oxidation process (AOP) that generates hydroxyl radicals (•OH) through the photolytic decomposition of H₂O₂. When exposed to UV light, hydrogen peroxide undergoes photolysis, producing two highly reactive hydroxyl radicals, as shown in Equation (2). These radicals readily attack the cell wall components of microalgae, leading to structural disruption and increased membrane permeability. This enhanced permeability facilitates the penetration of H₂O₂ into the cytoplasm, where additional oxidative reactions occur, targeting intracellular biomolecules and promoting the release of fermentable compounds beneficial for biofuel production [39]. The core reactions governing this pretreatment mechanism [39] are illustrated in Equations (2) and (3):

$${\mathrm{H}}_2{\mathrm{O}}_2+\mathrm{h}\mathsf{\upsilon }\to 2\mathrm{H}{\mathrm{O}}^{.}$$

(2)

$$\mathrm{H}{\mathrm{O}}^{.}+\mathrm{H}{\mathrm{O}}^{.}\to {\mathrm{H}}_2{\mathrm{O}}_2$$

(3)

where Equation (2) describes the photolysis of hydrogen peroxide into hydroxyl radicals under UV light, while Equation (3) represents the recombination of these radicals back into hydrogen peroxide under certain conditions, which may reduce the net radical availability. Therefore, optimizing operational parameters such as UV exposure time and H₂O₂ concentration is essential to maximize •OH production and achieve effective microalgal cell disruption. This UV/H₂O₂ pretreatment technique holds significant promise for enhancing the extraction of intracellular compounds and improving downstream processes such as fermentation and biofuel synthesis.

3.7. Elemental Composition

Table 3. Elemental compositions of microalgal biomass after UV/H₂O₂ pretreatment and fermentation.

Fermentation Time (h)	UV/H2O2 (H2O2 mM)		After Fermentation ppm			After UV/H2O2 pretreatment ppm				After UV/H2O2 Pretreatment Dry Microalgae w%
		TOC (Mean ± 95% CI)	TC (Mean ± 95% CI)	IC (Mean ± 95% CI)	TN	TOC (Mean ± 95% CI)	TC (Mean ± 95% CI)	IC (Mean ± 95% CI)	TN (Mean ± 95% CI)	C% (Mean ± 95% CI)	N% (Mean ± 95% CI)	H% (Mean ± 95% CI)
24	0.88	0.5704 ± 0.0217	0.5862 ± 0.0241	0.0126 ± 0.0003	0	21.43 ± 0.79	21.43 ± 0.79	0.00764 ± 0.00016	17.311 ± 0.639	39.344 ± 0.832	6.192 ± 0.229	6.570 ± 0.250
	1.76	0.1312 ± 0.0050	0.2093 ± 0.0086	0.0775 ± 0.0016	0	31.97 ± 1.18	32.29 ± 1.19	0.32301 ± 0.00683	4.300 ± 0.159	35.204 ± 0.745	6.016 ± 0.222	6.350 ± 0.226
	2.65	0.4519 ± 0.0172	0.5023 ± 0.0207	0.0453 ± 0.0010	0	27.13 ± 1.00	27.23 ± 1.01	0.01352 ± 0.00029	14.318 ± 0.529	40.817 ± 0.863	6.800 ± 0.251	7.140 ± 0.236
	3.53	0.0518 ± 0.0020	0.0677 ± 0.0028	0.0157 ± 0.0003	0	28.60 ± 1.06	28.60 ± 1.06	0.00000 ± 0.00000	12.515 ± 0.462	41.889 ± 0.886	6.830 ± 0.252	8.210 ± 0.766
48	0.88	0.0660 ± 0.0025	0.1164 ± 0.0048	0.0501 ± 0.0011	0	39.59 ± 1.46	39.89 ± 1.47	0.29879 ± 0.00632	17.431 ± 0.644	35.908 ± 0.759	5.978 ± 0.221	5.970 ± 0.051
	1.76	0.3554 ± 0.0135	0.4032 ± 0.0166	0.0460 ± 0.0010	0	17.82 ± 0.66	17.83 ± 0.66	0.01226 ± 0.00026	11.694 ± 0.432	36.246 ± 0.767	6.131 ± 0.226	6.510 ± 0.250
	2.65	0.0431 ± 0.0016	0.5862 ± 0.0241	0	0	31.53 ± 1.16	33.82 ± 1.25	2.30274 ± 0.04870	13.397 ± 0.495	39.866 ± 0.843	6.737 ± 0.249	7.010 ± 0.201
	3.53	1.2418 ± 0.0473	1.3251 ± 0.0546	0.0763 ± 0.0016	0	15.31 ± 0.57	24.37 ± 0.90	9.09141 ± 0.19229	16.751 ± 0.619	42.488 ± 0.899	6.426 ± 0.237	6.890 ± 0.296
Control		-	-	-	-	-	-	-	-	45.829 ± 0.969	7.831 ± 0.289	8.450 ± 0.388

shows the concentrations of TC, TOC, TN, and IC (ppm) from the supernatants obtained following UV/H₂O₂ pretreatment and subsequent fermentation. Additionally, the elemental composition (C, N, H in wt%) of dry microalgal biomass before and after pretreatment is shown, revealing the extent of cellular disruption and internal content release caused by this oxidative strategy.

The application of UV/H₂O₂ significantly enhanced the solubilization of cellular carbon and nitrogen, as reflected by the increased TOC and TN values in the supernatant compared to the untreated control.

For instance, at a concentration of 0.88 mM H₂O₂, the TOC and TC values reached 21.43 ± 0.79 and ppm, with negligible IC (0.0076 ppm) and a relatively high TN of 17.311 ± 0.639 ppm. The dry biomass still retained 39.344 ± 0.832% carbon, 6.192 ± 0.229% nitrogen, and 6.570 ± 0.250% hydrogen, indicating moderate cell rupture and content release. When the same H₂O₂ dose was applied under different conditions (presumably longer exposure or different UV intensity), TOC and TC increased significantly to 39.59 ± 1.46 ppm and 39.89 ± 1.47 ppm, respectively, while TN rose slightly to 17.431 ± 0.644 ppm. However, the carbon content in the biomass dropped to 35.908 ± 0.759%, confirming more efficient content leaching. At 1.76 mM H₂O₂, an optimal release of organic contents was observed under one condition, with TOC and TC peaking at 31.97 ± 1.18 ppm and 32.29 ± 1.19 ppm, respectively, while IC rose to 0.32301 ± 0.00683 ppm, and TN decreased sharply to 4.300 ± 0.159 ppm. This indicates effective oxidation and solubilization of carbon-rich compounds, with lower nitrogen retention in the medium. The biomass carbon content also dropped to 35.204 ± 0.745%, with nitrogen at 6.016 ± 0.222%, suggesting substantial cell disruption. Interestingly, under another condition at the same concentration, the TOC and TC values were considerably lower (17.82 ± 0.66 ppm and 17.83 ± 0.66 ppm, respectively), although biomass carbon (36.246 ± 0.767%) and nitrogen (6.131 ± 0.226%) remained similar—possibly due to experimental variation or oxidative degradation of organics. For 2.65 mM H₂O₂, the TOC and TC levels were 27.13 ± 1.00 ppm and 27.23 ± 1.01 ppm in one trial, respectively, with the biomass carbon content increasing to 40.817 ± 0.863%, which may indicate less extensive biomass oxidation despite effective cell rupture. In a separate condition, TC rose to 33.82 ± 1.25 ppm, accompanied by a significant increase in IC (2.30274 ± 0.04870 ppm) and a moderate TN of 13.397 ± 0.495 ppm, pointing to enhanced oxidation of organics into inorganic forms. The biomass retained 39.866 ± 0.843% carbon and 6.737 ± 0.249% nitrogen, suggesting partial but effective treatment. At the highest concentration, 3.53 mM H₂O₂, TOC and TC reached 28.60 ± 1.06 ppm, with zero IC, and TN was 12.515 ± 0.462 ppm in one case, while another condition showed reduced TC (24.37 ± 0.90 ppm) but a substantial IC of 9.09141 ± 0.19229 ppm and TN of 16.751 ± 0.619 ppm. These results imply that at high oxidant concentrations, further oxidation may lead to increased formation of inorganic byproducts (e.g., CO₂), diminishing the availability of fermentable organics. Interestingly, despite higher carbon release, the residual carbon in the biomass remained high (up to 42.488 ± 0.899%), showing that complete breakdown was not achieved. Compared to the untreated control biomass (C = 45.829 ± 0.969%, N = 7.831 ± 0.289%, H = 8.450 ± 0.388%), all pretreated samples showed a decrease in carbon and nitrogen contents, validating the effectiveness of UV/H₂O₂ in rupturing cells and leaching intracellular materials. Overall, moderate H₂O₂ concentrations (1.76–2.65 mM) were the most effective for maximizing organics’ release into the supernatant while minimizing excessive oxidation into non-fermentable forms. This balance is crucial for optimizing substrate availability in the subsequent fermentation stage.

The supernatants obtained from UV/H₂O₂-pretreated LMCMA were subjected to fermentation using Saccharomyces cerevisiae (S. cerevisiae), with the objective of utilizing the released organic carbon for biofuel production. The efficiency of fermentation was assessed by measuring the remaining total carbon (TC), inorganic carbon (IC), and total nitrogen (TN) in the medium after 24 and 48 h of microbial activity. The data clearly demonstrate that S. cerevisiae effectively consumes the available carbon sources released during pretreatment. After 24 h of fermentation, the TC levels decreased dramatically across all conditions: From an initial TC of 21.4–32.25 ppm (as seen in Stage 1), the post-fermentation TC values dropped from 21.43 ± 0.79 to 0.5704 ± 0.0217 ppm for tests carried out with 0.88 mM H₂O₂. Increasing the concentration of H₂O₂ exhibited lower TC values after fermentation. The TC values dropped from 32.29 ± 1.19 to 0.2093 ± 0.0086 ppm, from 27.23 ± 1.01 ppm to 0.5023 ± 0.0207 ppm, and from 28.60 ± 1.06 ppm to 0.0677 ± 0.0028 ppm for H₂O₂ concentrations of 1.76 mM, 2.65 mM, and 3.53 mM, respectively. These reductions correspond to carbon removal efficiencies exceeding 98%, indicating a high degree of substrate uptake and metabolic activity. The IC levels remained low (ranging from 0.0126 ± 0.0003 to 0.0775 ± 0.0016 ppm), suggesting minimal conversion of organics to inorganic carbon or CO₂ within the liquid phase. Importantly, TN dropped to 0 ppm in all conditions, implying complete nitrogen assimilation by the yeast or conversion into undetectable forms. Extending the fermentation to 48 h improved the uptake of TC, and lower concentrations were reported at all initial concentrations of H₂O₂: from 39.89 ± 1.47 to 0.1164 ± 0.0048 ppm at 0.88 mM, from 17.83 ± 0.66 ppm to 0.4032 ± 0.0166 ppm at 1.76 mM, from 33.82 ± 1.25 to 0.5862 ± 0.0241 ppm at 2.65 mM, and from 24.37 ± 0.90 ppm to 1.3251 ± 0.0546 ppm at 3.53 mM. These values still represent a >95–99% reduction in TC, with the lowest carbon residue observed for the 0.88 mM pretreatment, suggesting that this condition supports the most complete fermentation after 48 h. The IC values remained low (between 0 and 0.0775 ± 0.0016 ppm), reinforcing that only a small fraction of carbon ends up in inorganic form. TN remained consistently at 0 ppm in all conditions, confirming that nitrogen compounds released during pretreatment were fully utilized by the microbial cells.

The variation in total nitrogen (TN) across the two stages—UV/H₂O₂ pretreatment (Stage 1) and fermentation using Saccharomyces cerevisiae (Stage 2)—provides important insights into the efficiency of nutrient release and utilization, both of which are directly linked to biofuel production outcomes.

In Stage 1, the UV/H₂O₂ pretreatment is designed to rupture microalgae cells and release their intracellular components into the surrounding medium. The TN concentrations in the supernatant following this treatment varied widely depending on the H₂O₂ concentration and processing conditions. For example, at 0.88 mM H₂O₂, the TN values were relatively high, reaching 17.311 ± 0.639 ppm and 17.431 ± 0.644 ppm, indicating that milder oxidative conditions are effective at releasing nitrogenous compounds like amino acids and peptides without degrading them. In contrast, at 1.76 mM H₂O₂, TN decreased sharply to 4.300 ± 0.159 ppm, suggesting that more aggressive oxidative conditions may either degrade nitrogen-containing compounds into undetectable forms or volatilize them. Intermediate values were observed at 2.65 mM (e.g., 13.38–14.3 ppm) and at 3.53 mM (e.g., 12.5–16.73 ppm), indicating a balance between release and oxidative degradation. These variations highlight that TN release does not increase linearly with H₂O₂ concentration, and that excessive oxidant levels may compromise the bioavailability of nitrogen for microbial processes.

In Stage 2, after inoculating the pretreated supernatants with S. cerevisiae, the TN levels dropped to 0 ppm across all treatments and fermentation durations (both 24 and 48 h). This complete nitrogen depletion indicates that the nitrogen released during Stage 1 was readily bioavailable and fully assimilated by the yeast. Nitrogen is essential for microbial growth, particularly for protein synthesis, enzyme activity, and biomass formation. The ability of S. cerevisiae to consume all available TN confirms active metabolic processes and robust cell proliferation—both prerequisites for efficient fermentation and bioethanol production.

The observed TN dynamics have important implications for biofuel generation. In fermentation systems, the carbon-to-nitrogen (C/N) ratio plays a critical role in determining the balance between microbial growth and biofuel synthesis. Excess nitrogen can shift metabolism toward biomass accumulation at the expense of ethanol production, while insufficient nitrogen can hinder microbial activity altogether. The data suggest that the TN levels released during pretreatment were within a favorable range, supporting both microbial activity and carbon utilization. The complete TN removal, coupled with a >95–99% reduction in total carbon (TC), indicates that the system achieved an optimal nutrient balance that facilitated the efficient conversion of microalgal biomass into biofuel precursors [40].

3.8. Effects of UV/H₂O₂ Pretreatment on Microalgae Cell Wall Morphology

The SEM images of untreated and treated LMCMA (Figure S1, in the Supplementary Materials) show the effects of UV/H₂O₂ treatment on surface morphology under different treatment conditions. Figure S1a depicts untreated biomass, characterized by uniform, intact cell structures and a well-defined morphology. The image confirms the presence of mixed algal species with smooth, coherent cell walls, indicating the absence of any physical or chemical disruption. Upon UV/H₂O₂ treatment, significant morphological changes were observed. Using 0.88 mM H₂O₂ (Figure S1b), the LMCMA began to exhibit early signs of surface disruption; however, many cells still retained their original shapes, suggesting limited damage. With increasing H₂O₂ concentrations, the severity of the morphological deformation became more pronounced. At 1.76 mM and 2.65 mM (Figure S1c,d), cells exhibited wrinkling, fragmentation, and partial collapse, indicating progressive oxidative degradation of the cell walls. The most dramatic changes occurred at 3.53 mM H₂O₂ (Figure S1e), where the cells appeared aggregated and severely ruptured, with distorted surfaces and irregular shapes. These deformations confirm the efficient breakdown of the microalgae cell walls and the subsequent release of intracellular compounds, especially carbohydrates, which can then be utilized in fermentation processes.

Similarly, the TEM images (Figure S2, in the Supplementary Materials) further elucidate the internal structural degradation of the microalgae. Figure S2a shows the untreated control, where the cells maintain an intact and well-defined membrane and internal structure. In contrast, Figure S2b through Figure S2e reveal progressive damage corresponding to increasing H₂O₂ concentrations. At 0.88 mM H₂O₂ (Figure S2b), the cell wall begins to fracture, and the intracellular density decreases. At higher concentrations, particularly 2.65 and 3.53 mM (Figure S2d,e), there is clear evidence of extensive cytoplasmic leakage, total loss of membrane integrity, and collapse of the cellular architecture. The granular and dispersed appearance in these images signifies oxidative degradation and confirms the destruction of the cell envelope. Thus, this suggests that UV/H₂O₂ pretreatment can destroy and affect the entire microalgal structure, oxidizing the microalgae’s walls and increasing the release of fermentable carbohydrates [41].

These SEM and TEM results collectively demonstrate the effectiveness of UV/H₂O₂ pretreatment in disrupting microalgae’s structure. The treatment facilitates the breakdown of complex macromolecules such as polysaccharides into simpler, fermentable sugars. The progressive increase in cell damage with higher oxidant concentrations confirms that this pretreatment method enhances the bioavailability of intracellular materials, which directly supports increased yields in subsequent biofuel production. Furthermore, the correlation between morphological damage and increasing H₂O₂ concentration underscores the importance of optimizing the oxidant dosage to balance cell lysis efficiency with the preservation of fermentable substrates.

The morphological alterations observed via SEM and TEM validate the role of UV/H₂O₂ in cell wall degradation and content release. These structural changes are essential for improving enzymatic accessibility and maximizing carbohydrate recovery, which, in turn, enhances the efficiency of bioethanol or biohydrogen generation from microalgal biomass.

3.9. Effect of UV/H₂O₂ on Carbohydrate Release from LMCMA

Figure 6 illustrates the impact of UV/H₂O₂ pretreatment at varying hydrogen peroxide (H₂O₂) concentrations (0.88, 1.76, 2.65, and 3.53 mM) on carbohydrate liberation from LMCMA. The results demonstrate that pretreatment significantly enhanced the carbohydrate release, with the yield reaching a maximum at 1.76 mM H₂O₂ before declining at higher concentrations.

The specific carbohydrate release observed at each H₂O₂ concentration was 0.0215 ± 0.0003 g/g_DCW (0.88 mM), 0.0411 ± 0.0004 g/g_DCW (1.76 mM), 0.0360 ±0.0004 g/g_DCW (2.65 mM), and 0.0348 ± 0.0003 g/g_DCW (3.53 mM). The efficiency of carbohydrate release was found to be 43.0, 82.2, 72.0, and 69.6%, respectively. These values clearly show that the 1.76 mM treatment resulted in the highest carbohydrate release, nearly doubling the yield compared to the lowest concentration (0.88 mM). The increase in carbohydrate liberation up to this point suggests that UV activation of H₂O₂ effectively generated hydroxyl radicals (•OH), which are known for their strong oxidative potential and their ability to disrupt cell walls, solubilize polysaccharides, and convert complex carbohydrates into fermentable sugars. However, at concentrations beyond 1.76 mM, the carbohydrate yield began to decline, indicating the onset of over-oxidation. At 2.65 mM and 3.53 mM, although the cell walls were visibly ruptured (as confirmed by the SEM and TEM micrographs), the excess oxidant likely degraded some of the liberated carbohydrates into smaller, non-fermentable fragments or volatile carbon compounds, thereby reducing the effective sugar yield. This tradeoff between cell disruption and carbohydrate preservation underscores the importance of optimizing the H₂O₂ dosage.

The pretreatment mechanism involves UV-induced photolysis of H₂O₂, which produces highly reactive hydroxyl radicals. These radicals attack the microalgae cell wall components—such as cellulose, hemicellulose, and glycoproteins—leading to structural collapse and increased membrane permeability. This facilitates the leakage of intracellular contents, especially polysaccharides. At controlled concentrations (notably 1.76 mM), this process enhances carbohydrate availability without significant degradation. In contrast, higher concentrations (≥2.65 mM) introduce excessive oxidative stress, damaging both the cellular structure and the liberated sugars.

The supporting literature reinforces these findings. For instance, Yang et al. [16] demonstrated that a UV-assisted alkaline H₂O₂ pretreatment of sisal waste (pH 10, UV for 6 h) improved enzymatic hydrolysis efficiency, converting 76% of total sugars into 91.6% glucose. Similarly, combining UV exposure for 30 min with 12 mmol/g fiber of H₂O₂ significantly increased hydroxyl radical generation, enhancing cellulose oxidation and leading to polysaccharide degradation [28]. These studies confirm that UV/H₂O₂ treatment is most effective under controlled conditions where radical formation is maximized for cell wall rupture but limited enough to avoid excessive sugar loss. Wen et al. [28] demonstrated that combining UV radiation for 30 min with 12 mmol H₂O₂/g fiber significantly enhanced the oxidative efficiency by generating additional hydroxyl radicals. This synergistic effect amplified ozone-induced oxidation, accelerating cellulose breakdown and leading to effective polysaccharide degradation. Their findings support the conclusion that UV/H₂O₂ pretreatment is a powerful method for disrupting microalgae’s cell walls, increasing membrane permeability, and facilitating the release of intracellular carbohydrates. In alignment with these observations, the current study identified 1.76 mM H₂O₂ as the optimal concentration for maximizing carbohydrate liberation. Beyond this point, higher oxidant levels led to a decline in sugar yield, likely due to over-oxidation and degradation of released carbohydrates.

3.10. Effects of Fermentation Time on Carbohydrate Yield

Figure 7 illustrates the effects of fermentation time (24 and 48 h) on the carbohydrate release (gcarb/g_DCW) following UV/H₂O₂ pretreatment of microalgal biomass at various H₂O₂ concentrations (0.88, 1.76, 2.65, and 3.53 mM). The figure provides a comparative assessment of how effectively fermentable carbohydrates released during pretreatment were consumed over time by Saccharomyces cerevisiae, highlighting the interplay between oxidant concentration and fermentation duration.

At 24 h of fermentation, the carbohydrate concentration dropped drastically, especially at higher H₂O₂ concentrations. The most notable reductions were observed at 2.65 mM and 3.53 mM, where the carbohydrate concentrations decreased to approximately 0.0023 ± 0.0003 g_carb/g_DCW and 0.0018 ± 0.0002 g_carb/g_DCW, respectively. These values reflect a 93.6% and 93.1% reduction, respectively, relative to the released carbohydrate, indicating rapid and efficient microbial utilization of sugars. This performance even exceeded the 91.6% glucose conversion reported by Yang et al. [16], who utilized UV/H₂O₂ treatment on sisal waste with 6 h of exposure.

In contrast, at lower H₂O₂ concentrations (0.88 and 1.76 mM), the carbohydrate reduction was less pronounced at 24 h, reaching 57.2% and 63%, respectively. However, after extending the fermentation to 48 h, the yields at these lower concentrations further declined—indicating continued microbial activity—with final reductions of 63.5% (0.88 mM) and 71% (1.76 mM). Interestingly, while the yield at 2.65 mM slightly increased to a final reduction of 72.8%, the yield at 3.53 mM continued to drop to 0.0006 g_carb/g_DCW, reflecting a 98.4% total reduction—the highest observed across all conditions. This suggests that while higher H₂O₂ concentrations may initially generate inhibitory compounds that slow microbial metabolism, prolonged fermentation enables S. cerevisiae to adapt and further metabolize the available sugars.

These trends demonstrate the dual influence of oxidative pretreatment and fermentation time. At moderate oxidant levels (e.g., 1.76 mM), UV/H₂O₂ pretreatment effectively ruptures the cell wall and enhances sugar availability without severely degrading carbohydrates, facilitating sustained fermentation. However, at higher oxidant concentrations (≥2.65 mM), over-oxidation may initially impair fermentability but does not entirely prevent microbial utilization over extended periods.

Several studies support these findings. For example, Scenedesmus obliquus, after sonication and 16 days of fermentation, achieved an 89% reduction in carbohydrates [31]. In another study, UV/H₂O₂ pretreatment followed by ZSM-5-catalyzed pyrolysis yielded 6.4% sugar after 2 h of UV radiation using 4 mL/g H₂O₂ [42]. Similarly, a UV–alkaline H₂O₂ pretreatment followed by enzymatic hydrolysis of corn stover led to 90.2% lignin removal, with enhanced hemicellulose recovery (+48.9%) [43]. These examples reaffirm that H₂O₂ concentration, treatment time, and posttreatment processes significantly affect saccharification efficiency.

The current study reveals that excessive H₂O₂ concentration can suppress carbohydrate release and subsequent fermentation due to potential sugar degradation and the formation of inhibitory byproducts such as furfural, hydroxymethylfurfural (HMF), and phenolic compounds. These inhibitors are known to hinder microbial growth and ethanol yield [44]. Additionally, the observed trends emphasize that microalgae species, pretreatment conditions, and microbial strains all contribute to sugar yield variability.

3.11. Effects of UV/H₂O₂ Pretreatment on Biofuel Production

Figure 8 presents the yields of various biofuels—ethanol, methanol, 1-propanol, 1-butanol, and 1-pentanol—produced through fermentation of UV/H₂O₂-pretreated microalgae biomass. The treatments involved four concentrations of hydrogen peroxide (0.88, 1.76, 2.65, and 3.53 mM) and fermentation durations of 24 and 48 h at 30 °C, showcasing the influence of oxidant dose and fermentation time on biofuel production efficiency. The results depict that carbohydrates are consumed during fermentation and potentially converted into biofuel.

Figure 8a shows that highest ethanol Y_prod was recorded at 1.76 mM H₂O₂ after 48 h, reaching 0.268 g_ethanol/g_carb, which was a 6.8% increase from 0.251 g_ethanol/g_carb at 24 h. At 0.88 mM, the ethanol Y_prod improved slightly, from 0.211 to 0.219 g_ethanol/g_carb (3.8% increase). At 2.65 mM, the ethanol Y_prod increased from 0.202 to 0.232 g_ethanol/g_carb (14.9% increase), while at 3.53 mM, the ethanol Y_prod increased from 0.148 to 0.179 g_ethanol/g_carb (21.0% increase). These trends suggest continued carbohydrate conversion over time, especially at moderate concentrations. The ethanol Y_prod at 1.76 mM was 27% higher than at 3.53 mM at 24 h, confirming 1.76 mM as optimal for ethanol production. Zhang et al. [42] revealed that a combined pretreatment pyrolysis of treated water hyacinth using UV/H₂O₂ resulted in a 32% increase in hydrocarbons, facilitating bio-oil production.

The methanol Y_prod (Figure 8b) showed a distinct peak at 2.65 mM after 24 h, yielding 0.1311 g_methanol/g_carb, followed by a dramatic drop to 0.00012 g_methanol/g_carb after 48 h, reflecting a 99.9% reduction. At 0.88 mM, the methanol Y_prod decreased from 0.0495 g_methanol/g_carb to 0.00055 g_methanol/g_carb (98.9% reduction). At 1.76 mM, the methanol Y_prod declined from 0.0228 g_methanol/g_carb to 0.00021 g/g (99.1% decrease). No significant methanol was detected at 3.53 mM. These results confirm that methanol production peaks early and rapidly depletes, possibly due to volatilization or microbial assimilation. This behavior underscores the need for short fermentation times to capture methanol efficiently.

Figure 8c shows that the 1-propanol Y_prod increased with higher H₂O₂ concentrations. At 3.53 mM, the maximum 1-propanol Y_prod of 0.139 g_1-propanol/g_carb was achieved after 24 h, decreasing to 0.028 g_1-propanol/g_carb at 48 h (79.9% reduction). At 2.65 mM, the 1-propanol Y_prod dropped from 0.086 to 0.0096 g_1-propanol/g_carb (88.8% decrease). At 0.88 mM, 1-propanol Y_prod declined from 0.036 to 0.021 g_1-propanol/g_carb (41.7% decrease), while at 1.76 mM, the 1-propanol Y_prod remained constant at 0.022 g_1-propanol/g_carb over both time points. Thus, 1-propanol is rapidly generated and then depleted at high oxidant levels. The 1-propanol Y_prod at 3.53 mM of H₂O₂ and fermentation for 24 h was 4.36× higher than the Y_prod at 1.76 mM, making this treatment the most favorable for fast, high-yield 1-propanol production.

The 1-butanol Y_prod (Figure 8d) peaked at 2.65 mM after 24 h, with a Y_prod of 0.1058 g_1-butanol/g_carb, which dropped to 0.00725 g_1-butanol/g_carb at 48 h (93.2% reduction). At 0.88 mM, the 1-butanol Y_prod decreased from 0.0466 to 0.0199 g_1-butanol/g_carb (57.3% decrease). At 1.76 mM, the 1-butanol Y_prod remained relatively stable (0.0133 to 0.0125 g_1-butanol/g_carb; a 6.0% decrease). At 3.53 mM, the 1-butanol Y_prod fell from 0.026 to 0.0122 g_1-butanol/g_carb (53.1% decrease). The 1-butanol Y_prod yield at 2.65 mM was 127% higher than the yield at 0.88 mM after 24 h, highlighting that moderate oxidative pretreatment significantly boosts the initial butanol yield, although this is not sustained.

Unlike the other alcohols, 1-pentanol’s Y_prod (Figure 8e) increased over time at lower concentrations. At 1.76 mM, the 1-pentanol Y_prod rose from 0.0174 g_1-pentanol/g_carb (24 h) to 0.046 g_1-pentanol/g_carb (48 h), representing a 164% increase. At 0.88 mM, the Y_prod increased from 0.010 to 0.034 g_1-pentanol/g_carb (240% increase). At 2.65 mM, the 1-pentanol Y_prod rose from 0.0093 to 0.018 g_1-pentanol/g_carb (93.5% increase). At 3.53 mM, however, the Y_prod decreased from 0.0235 to 0.0051 g_1-pentanol/g_carb (78.3% decrease). The highest 1-pentanol Y_prod was 0.046 g_1-pentanol/g_carb at 1.76 mM after 48 h, which was 360% higher than the yield at 0.88 mM after 24 h. This behavior suggests that 1-pentanol may be a secondary fermentation product favored under prolonged, moderate oxidation.

These findings align with those of studies such as the work of Yang et al. [16], who achieved 91.6% glucose conversion using UV/H₂O₂ pretreatment, as well as additional research on ZSM-5 catalysis and enzymatic hydrolysis [42,43]. However, unlike lignocellulosic systems, where the biofuel yield increases with time, the current study reveals that, for microalgae, the biofuel type, H₂O₂ concentration, and fermentation time must be tailored specifically to the target product.

The impact of fermentation temperature on the conversion efficiency of UV/H₂O₂-pretreated microalgal biomass into biofuels was systematically evaluated at 30 °C, 35 °C, and 37 °C over fermentation durations of 24 and 48 h, using fixed hydrogen peroxide concentrations (0.88, 1.76, 2.65, and 3.53 mM). This study focused on the production of ethanol, methanol, 1-propanol, 1-butanol, and 1-pentanol, with a particular emphasis on the metabolic and thermodynamic principles influencing yield outcomes.

Ethanol biosynthesis via Saccharomyces cerevisiae relies on the glycolytic breakdown of glucose into pyruvate, followed by decarboxylation and reduction to ethanol under anaerobic conditions. The maximum ethanol yield was observed at 30 °C, peaking at 0.251 g/g at 1.76 mM after 24 h, and further increasing by 6.7% to 0.268 g/g after 48 h.

In contrast, at 35 °C, the yield at 0.88 mM was 0.18 g/g (24 h) and 0.21 g/g (48 h)—28% lower than the corresponding 30 °C value. At 37 °C, the yields were even more suppressed, rising from 0.09 g/g to 0.15 g/g, marking a 40–50% reduction compared to optimal conditions. The decline in ethanol yield with increasing temperature can be attributed to the thermal sensitivity of S. cerevisiae’s fermentative enzymes and the shift toward stress-response pathways that divert carbon flux away from ethanol synthesis.

Methanol was detected only at 30 °C, with the highest yield of 0.1311 g/g recorded at 2.65 mM after 24 h. However, this yield plummeted by >99.9% to 0.00012 g/g after 48 h. At lower H₂O₂ concentrations, methanol similarly decreased with time. No methanol was detected at 35 °C or 37 °C, indicating rapid volatilization or metabolic assimilation at elevated temperatures. This aligns with the known low boiling point and volatility of methanol, and suggests that methanol accumulation is thermodynamically unstable above 30 °C, reinforcing the need for early harvest strategies in methanol-focused processes.

1-Propanol is produced through amino acid catabolism (notably threonine) and may proceed via the Ehrlich pathway. At 30 °C, the yield reached a maximum of 0.139 g/g (3.53 mM, 24 h) but fell to 0.028 g/g (48 h)—a 79.9% reduction. At 2.65 mM, the yield decreased from 0.086 g/g to 0.0096 g/g (88.8% decrease). At 35 °C, the peak yield dropped to 0.05 g/g (24 h) and 0.01 g/g (48 h). Interestingly, a reverse trend was observed at 2.65 mM, with the yields increasing from 0.035 g/g to 0.067 g/g, suggesting temperature-induced activation of secondary fermentative enzymes. At 37 °C, the highest recorded yield was 0.009 g/g at 0.88 mM (24 h), decreasing to 0.006 g/g, marking an over 90% reduction compared to the 30 °C peak. These results highlight the thermal fragility of 1-propanol biosynthesis, which is optimal under moderate temperatures and is highly suppressed by heat stress.

1-Butanol formation is typically governed by the CoA-dependent ABE (acetone–butanol–ethanol) pathway. At 30 °C, the highest yield was 0.1058 g/g (2.65 mM, 24 h), decreasing sharply to 0.00725 g/g (48 h)—a 93.2% drop. At 0.88 mM, the yield declined from 0.0466 to 0.0199 g/g, while at 1.76 mM it remained relatively low (0.00796 to 0.00985 g/g). At 35 °C, the maximum yield was 0.05 g/g (24 h), declining to 0.012 g/g (48 h), while at 37 °C, the maximum yield did not exceed 0.01 g/g, with most values <0.009 g/g. This represents a 5–10× reduction compared to the optimal yield at 30 °C. This temperature sensitivity is consistent with the well-documented thermolability of the key enzymes involved in acetyl-CoA conversion and butanol elongation, as well as the inhibitory effect of heat on NADH regeneration necessary for butanol synthesis.

In contrast to the other biofuels, 1-pentanol exhibited an increasing trend over time and appeared to benefit from elevated fermentation temperatures. At 30 °C, the yield at 3.53 mM decreased from 0.0235 to 0.0051 g/g (78% reduction), but at lower concentrations, the yields increased: from 0.010 to 0.034 g/g (0.88 mM) and from 0.0174 to 0.046 g/g (1.76 mM) over 48 h. At 35 °C, the yields continued this trend: 0.011 to 0.021 g/g (1.76 mM) and 0.009 to 0.015 g/g (3.53 mM), reflecting a 91% increase in the former case. At 37 °C, the most notable rise was at 1.76 mM, from 0.00174 to 0.022 g/g—a 1164% increase, the highest relative growth among all biofuels in the study. This pattern suggests that 1-pentanol may be regulated through heat-responsive pathways, possibly involving amino acid catabolism (e.g., leucine-derived keto acid elongation) or the Ehrlich pathway, which is known to be activated under oxidative or thermal stress. The delayed accumulation implies that 1-pentanol may function as a secondary or overflow metabolite, emerging when primary fermentative pathways are saturated or inhibited. These results indicate that 1-pentanol production benefits from higher temperatures and longer fermentation times, suggesting that it may follow an alternative secondary metabolic pathway (e.g., Ehrlich pathway) that is activated under stress or extended fermentation.

The previous results reveal that fermentation temperature plays a pivotal role in determining the profile and yield of biofuels from pretreated microalgal biomass. It was observed that 30 °C is optimal for ethanol, methanol, 1-propanol, and 1-butanol, supporting high enzymatic activity and maintaining redox balance for efficient fermentation Methanol formation is exclusive to 30 °C, likely due to its volatility and instability at higher temperatures. 1-Propanol and 1-butanol yields are suppressed by 80–95% at 37 °C compared to 30 °C, reflecting thermally sensitive metabolic bottlenecks. 1-Pentanol production uniquely increases at higher temperatures and extended durations, suggesting temperature-induced redirection of carbon flow into alternative alcohol pathways. These findings align with those of prior studies, including the works of Yang et al. [16], Zhang et al. [42], and Qureshi et al. [44], which highlight the importance of fine-tuning the fermentation parameters—particularly temperature and retention time—to maximize the yield of targeted biofuels. Understanding the enzymatic and regulatory basis of these temperature-dependent shifts is crucial for developing optimized microalgae-based biorefineries.

Table 4. Summary of biofuel yields from microalgae following UV/H₂O₂ pretreatment.

Fermentation Time (h)	H2O2 mM	Methanol (g/gcarb)			Propanol (g/gcarb)			Butanol (g/gcarb)			Pentanol (g/gcarb)
		30 °C	35 °C	37 °C	30 °C	35 °C	37 °C	30 °C	35 °C	37 °C	30 °C	35 °C	37 °C
24	0.88	0.0495	ND	ND	0.036	0.03	0.009	0.0466	0.03	0.0093	0.01	0.01	0.009
	1.76	0.0228	ND	ND	0.022	0.01	0.006	0.0133	0.01	0.00796	0.0174	0.011	0.00174
	2.65	0.1311	ND	ND	0.086	0.035	0.0072	0.1058	0.05	0.008	0.0093	0.009	0.0067
	3.53	ND	ND	ND	0.139	0.05	0.0067	0.026	0.018	0.01	0.0235	0.009	0.006
48	0.88	0.00055	ND	ND	0.021	0.026	0.006	0.0199	0.026	0.0076	0.034	0.017	0.014
	1.76	0.00021	ND	ND	0.022	0.067	0.005	0.0125	0.012	0.00985	0.046	0.021	0.022
	2.65	0.00012	ND	ND	0.0096	0.045	0.0055	0.00725	0.012	0.0075	0.018	0.015	0.0094
	3.53	ND	ND	ND	0.028	0.01	0.0053	0.0122	0.006	0.01	0.0051	0.015	0.009

highlights the impacts of H₂O₂ concentration, fermentation time, and temperature on alcohol production from UV/H₂O₂-pretreated microalgae. Methanol was only detected at 30 °C, peaking at 0.1311 g/g (2.65 mM, 24 h), but dropped by >99% after 48 h. No methanol was detected at 35 °C or 37 °C, likely due to volatilization. The 1-Propanol yield was highest at 30 °C, reaching 0.139 g/g (3.53 mM, 24 h), but declined by 85–95% at 37 °C. At 35 °C, some delayed increases were observed, suggesting moderate stress-induced metabolism. 1-Butanol peaked at 0.1058 g/g (30 °C, 2.65 mM, 24 h), but the yields dropped by over 90% at higher temperatures and longer durations. Unlike the other alcohols, 1-pentanol increased over time and with temperature. At 37 °C and 1.76 mM, the yield rose from 0.00174 to 0.022 g/g (a 1164% increase), indicating a stress-activated pathway, possibly the Ehrlich pathway. Overall, 30 °C favored methanol, propanol, and butanol production, while higher temperatures enhanced 1-pentanol yields, demonstrating the need to tailor the fermentation conditions based on the target biofuel.

The current study achieved a maximum ethanol Y_prod of 0.3668 g/g, along with notable production yields of other alcohols, including methanol (0.0564 g/g), 1-propanol (0.0512 g/g), 1-butanol (0.0225 g/g), and 1-pentanol (0.0219 g/g). These results were obtained using UV/H₂O₂ pretreatment under optimized fermentation conditions (temperatures of 30–35 °C and durations of 24–48 h). Table 5 provides a comparative overview of the biofuel production results from this study in relation to previous works employing different microalgal pretreatment and fermentation strategies.

Compared to prior studies using UV pretreatment alone, the current findings demonstrate enhanced biofuel production. For example, Bhushan et al. [26] investigated UV-C irradiation (254 nm) as a standalone pretreatment for wastewater-grown algal biomass, with exposure durations ranging from 15 to 60 min. Their highest reported ethanol yield was 0.084 g/g volatile solids (VS), with a maximum hydrogen yield of 30.1 mL/g VS following 60 min of UV-C exposure. These values are notably lower than the ethanol yield obtained in the present study, highlighting the efficacy of incorporating H₂O₂ to generate additional hydroxyl radicals for improved cell wall disruption.

The work of Ma’mun et al. [45] further supports the importance of effective pretreatment. In their study, a two-step pretreatment process—comprising thermal acid hydrolysis (1% H₂SO₄ at 121 °C for 15 min) followed by enzymatic hydrolysis with cellulase—was applied to Arthrospira platensis biomass. This combination significantly enhanced sugar release and fermentability, resulting in an ethanol yield of 5.24 g/L, corresponding to 62.7% of the theoretical maximum. While the yield was substantial, it reflected the concentration per liter rather than per gram of biomass, and it required a longer fermentation duration (72 h) compared to the current study.

Bhushan et al. [46] offered a comprehensive review of advancements in microalgae-based bioenergy. They emphasized that chemical and oxidative pretreatments, including UV/H₂O₂, are particularly effective for breaking down microalgal cell walls to enhance fermentable sugar release. This aligns closely with the outcomes of the present study, where UV/H₂O₂ pretreatment facilitated substantial production of ethanol and other alcohols from microalgal biomass. Common operational conditions across these studies include fermentation temperatures ranging from 30–60 °C and pH values between 4 and 6. Saccharomyces cerevisiae remains the preferred fermenting organism for ethanol production, while Clostridium spp. are more effective for butanol and hydrogen. As reported in the review, yields vary with species and conditions—such as up to 5.6 g/L ethanol from Scenedesmus obliquus after ultrasonic pretreatment and lipid contents reaching 57% of dry cell weight (DCW) for biodiesel, with biohydrogen yields of up to 112 mL H₂/g VS via dark fermentation.

Table 5. Comparison of biofuel production with other studies.

Feedstock	Pretreatment Method	Pretreatment/Fermentation Conditions	Ethanol Yield	Other Biofuels	Reference
Mixed culture	UV and UV/H2O2	30–35 °C; 24–48 h; S. cerevisiae	0.3668 g/gcarb	Methanol (0.0564), 1-propanol (0.0512), 1-butanol (0.0225), 1-pentanol (0.0219) g/g	Current study
Mixed culture	UV-C	60 min UV; fermenter not specified	0.084 g/gVS	Hydrogen: 30.1 mL/g VS	[26]
Arthrospira platensis	Microwave-assisted acid hydrolysis (H2SO4)	48 h hydrolysis, 72 h fermentation at 30 °C; S. cerevisiae	5.24 g/L (~0.35 g/g carbohydrate)	Not reported	[45]
Various microalgae (e.g., Chlorella, Scenedesmus, Arthrospira)	Various (UV/H2O2, acid, alkaline, ultrasonic, enzymatic)	Varies widely across studies	Up to 5.6 g/L (Scenedesmus)	Butanol (18.9 g/L), H2 (112 mL/g VS), lipids (up to 57% DCW)	[26]
Sisal waste (lignocellulosic)	UV photocatalysis + alkaline H2O2	Enzymatic hydrolysis post pretreatment	Not directly reported; 71.2% sugar conversion	None reported	[16]
Ulva reticulata (seaweed)	CEM microwave + H2SO4	5 days at 30 °C; 12% S. cerevisiae; pH 4.5	42.53% ethanol content (~5.02 g/L est.)	Not reported	[47]

Table 5. Comparison of biofuel production with other studies.

Feedstock	Pretreatment Method	Pretreatment/Fermentation Conditions	Ethanol Yield	Other Biofuels	Reference
Mixed culture	UV and UV/H2O2	30–35 °C; 24–48 h; S. cerevisiae	0.3668 g/gcarb	Methanol (0.0564), 1-propanol (0.0512), 1-butanol (0.0225), 1-pentanol (0.0219) g/g	Current study
Mixed culture	UV-C	60 min UV; fermenter not specified	0.084 g/gVS	Hydrogen: 30.1 mL/g VS	[26]
Arthrospira platensis	Microwave-assisted acid hydrolysis (H2SO4)	48 h hydrolysis, 72 h fermentation at 30 °C; S. cerevisiae	5.24 g/L (~0.35 g/g carbohydrate)	Not reported	[45]
Various microalgae (e.g., Chlorella, Scenedesmus, Arthrospira)	Various (UV/H2O2, acid, alkaline, ultrasonic, enzymatic)	Varies widely across studies	Up to 5.6 g/L (Scenedesmus)	Butanol (18.9 g/L), H2 (112 mL/g VS), lipids (up to 57% DCW)	[26]
Sisal waste (lignocellulosic)	UV photocatalysis + alkaline H2O2	Enzymatic hydrolysis post pretreatment	Not directly reported; 71.2% sugar conversion	None reported	[16]
Ulva reticulata (seaweed)	CEM microwave + H2SO4	5 days at 30 °C; 12% S. cerevisiae; pH 4.5	42.53% ethanol content (~5.02 g/L est.)	Not reported	[47]

The findings of Yang et al. [16] further corroborate the potential of UV/H₂O₂ pretreatment. They demonstrated that combining UV photocatalysis with alkaline H₂O₂ significantly enhanced the enzymatic hydrolysis of sisal waste, achieving a 62.4% increase in reducing sugar yield. Although their substrate was lignocellulosic rather than algal, the underlying mechanism—generation of hydroxyl radicals and disruption of biomass structure—supports the strategy used in the current study to improve carbohydrate accessibility.

Additional studies provide further context. For example, Kushwaha et al. [32] reported high butanol production (18.9 g/L) using Clostridium spp., although their process required 110 h of fermentation under acidic conditions, making it less time-efficient. Kolo et al. [47] found that UV pretreatment of Chlorella vulgaris for 30 min improved cell disruption, leading to a 16% increase in biogas yield and an increase in methane content from 48% to 54%, underscoring the broad applicability of UV-based pretreatment in various bioenergy contexts.

In summary, the UV/H₂O₂ pretreatment applied in this study effectively enhanced bioethanol and higher alcohol production from microalgae. The results not only demonstrate comparable or superior yields to those reported in other studies but also underscore the efficiency of oxidative pretreatment in improving fermentation outcomes through improved cell disruption and sugar release.

While this study successfully demonstrated the effectiveness of UV and UV/H₂O₂ pretreatments in enhancing carbohydrate release and biofuel production from mixed marine microalgal cultures, several future directions should be acknowledged. First, the current work was conducted using batch fermentation and did not incorporate kinetic analysis or real-time monitoring of biotransformation rates. Future research should include kinetic studies and real-time biotransformation measurements to better understand the substrate utilization dynamics and process efficiency. Furthermore, pigment and chlorophyll degradation analyses were not conducted but could serve as important indicators of microalgal digestion and pretreatment effectiveness. Incorporating such spectrophotometric measurements in future studies would enhance our understanding of cell wall disruption and biomass utilization efficiency.

The obtained results show that the efficiency of carbohydrate release from LMCMA was significantly influenced by the type of pretreatment applied. UV irradiation alone achieved low carbohydrate release efficiencies in the range 25.0% to 44.6%. In contrast, UV/H₂O₂ pretreatment demonstrated markedly superior performance, with efficiencies ranging from 43.0% to 82.2%. This enhanced performance can be attributed to the generation of more hydroxyl radicals (•OH), which effectively degraded structural polysaccharides and disrupted cell membranes, resulting in a more complete solubilization of intracellular contents.

The significant increase in carbohydrate release directly translated into improved biofuel production. UV/H₂O₂ pretreatment not only increased the availability of fermentable sugars but also facilitated the liberation of additional biomolecules such as proteins and lipids—critical for supporting microbial growth and the synthesis of diverse biofuels, including ethanol, methanol, and higher alcohols (1-propanol, 1-butanol, and 1-pentanol). This comprehensive cell disruption enabled more complete biomass utilization and improved the overall carbon conversion efficiency during fermentation. Compared to UV-only pretreatment, which resulted in limited substrate availability due to partial cell disintegration, the UV/H₂O₂ strategy led to higher ethanol yields and broader bioalcohol production profiles. Moreover, the UV/H₂O₂ approach provided a balanced nutrient environment for Saccharomyces cerevisiae, enhancing the fermentation performance and product selectivity. By maximizing the fermentable substrate availability while minimizing over-oxidation and inhibitory byproduct formation, UV/H₂O₂ pretreatment offers a robust and scalable strategy for sustainable biofuel production. These findings highlight its potential as a key enabling technology for the full valorization of algal biomass within an integrated biorefinery framework, supporting the transition toward a circular and low-carbon bioeconomy.

4. Mechanistic Pathways for Biofuel Production from Microalgae Biomass

The utilization of microalgae as a renewable and sustainable biofuel feedstock has gained significant attention due to its high photosynthetic efficiency, rapid biomass accumulation, and biochemical flexibility. During the stationary growth phase, microalgae accumulate considerable quantities of carbohydrates and proteins, which serve as vital substrates for fermentation-based biofuel production processes [48]. However, the rigid and polysaccharide-rich cell wall structure poses a substantial barrier to intracellular substrate accessibility. To overcome this limitation, UV and UV/H₂O₂ pretreatment strategies have been widely adopted to enhance cell wall disruption and facilitate the release of fermentable intracellular contents [45,49].

Figure 9 presents an overview of the UV/H₂O₂-assisted pretreatment mechanism within the biofuel production pathway. The photolytic decomposition of hydrogen peroxide under UV light generates reactive hydroxyl radicals (•OH), which induce oxidative cleavage of cell wall polymers. This enhances membrane permeability and promotes the release of intracellular metabolites, particularly carbohydrates and amino acids. These liberated compounds serve as critical precursors for the microbial biosynthesis of alcohols such as ethanol, methanol, 1-propanol, 1-butanol, and 1-pentanol during anaerobic fermentation.

The core route for ethanol biosynthesis involves the glycolytic pathway, wherein glucose derived from the disrupted algal cells is metabolized to pyruvate. Under anaerobic conditions, Saccharomyces cerevisiae catalyzes the decarboxylation of pyruvate to acetaldehyde, followed by its reduction to ethanol via NADH-mediated pathways. This process regenerates NAD⁺, enabling the continuation of glycolysis. The conversion reaction is illustrated as follows [50]:

$${{\mathrm{C}}_6\mathrm{H}}_{12}{\mathrm{O}}_6\to {2{\mathrm{C}}_2\mathrm{H}}_5\mathrm{H}\mathrm{O}+\mathrm{C}{\mathrm{O}}_2$$

(4)

Fusel alcohols, which contain more than two carbon atoms, include aliphatic alcohols like 1-propanol, isobutanol, and isoamyl alcohol, and they are produced via the Ehrlich pathway. This catabolic route involves the transamination of amino acids into corresponding α-keto acids, decarboxylation into aldehydes, and reduction to higher alcohols. The enzymes BAT1/BAT2, PDC1/PDC5/PDC6, and ARO10 are key components of this pathway, whose activity is regulated by cellular nitrogen status and oxidative stress, both of which are influenced by UV/H₂O₂ treatment [23,51].

Figure 10 illustrates the Ehrlich pathway, detailing the transformation of amino acids into fusel alcohols. For example,

• Valine is converted into isobutanol via α-ketoisovalerate and isovaleraldehyde.
• Leucine is converted into isoamyl alcohol via α-ketoisocaproate and isoamylaldehyde.
• Isoleucine and methionine serve as precursors for amyl alcohol and methanethiol, respectively.
• Threonine and methionine are precursors to 1-propanol, while α-ketovalerate serves as a key intermediate for 1-butanol synthesis.

Methanol production exhibited a distinct profile, with a peak yield of 0.1311 g/g at 30 °C and 2.65 mM H₂O₂ after 24 h. This decreased dramatically—by over 99.9%—after 48 h. No methanol was detected at 35 °C and 37 °C, which can be attributed to its volatility, rapid microbial uptake, or thermal degradation under elevated conditions. Among the higher alcohols, 1-propanol peaked at 0.139 g/g (30 °C, 3.53 mM H₂O₂, 24 h) but declined by 79.9% to 0.028 g/g at 48 h. At 37 °C, the yields dropped by >90%, highlighting the sensitivity of the associated enzymes to heat. For 1-butanol, the maximum yield of 0.1058 g/g was recorded at 30 °C (2.65 mM H₂O₂, 24 h), followed by significant reductions at higher temperatures and longer fermentation periods. This pattern aligns with the ABE fermentation pathway, which involves NADH-dependent enzymes that are sensitive to thermal stress. Conversely, 1-pentanol biosynthesis followed a delayed but amplified trend. At 30 °C and 1.76 mM, the yield increased from 0.0174 g/g to 0.046 g/g between 24 and 48 h. At 37 °C, a 1164% increase was recorded, reaching 0.022 g/g from 0.00174 g/g over the same period. This suggests that 1-pentanol production is governed by secondary metabolic routes, possibly linked to overflow metabolism or stress-induced expression of the Ehrlich pathway enzymes. Collectively, these results underscore the importance of temperature, oxidative stress level, and fermentation duration in modulating biofuel yields. While ethanol and methanol require mild conditions and timely harvesting, 1-propanol and 1-butanol necessitate optimized oxidative stress and moderate temperatures to maintain enzymatic activity. 1-Pentanol, on the other hand, benefits from prolonged fermentation under thermal stress, potentially offering an alternative avenue for enhanced biofuel output.

5. Conclusions

This study demonstrates that UV and UV/H₂O₂ pretreatment methods are effective in disrupting the cell walls of mixed marine microalgae cultures, leading to enhanced release of fermentable carbohydrates and subsequent biofuel production. UV pretreatment at 20–30 min significantly increased the total organic carbon (TOC) and reducing sugars, with SEM and TEM analyses confirming structural degradation and cytoplasmic leakage. Similarly, UV/H₂O₂ pretreatment at 1.76 mM achieved the highest sugar release, while higher concentrations led to reduced efficiency due to over-oxidation. Elemental analysis revealed changes in carbon and nitrogen contents, suggesting enhanced substrate availability for fermentation. Ethanol was the dominant product across conditions, with maximum yields obtained at 30 °C and 48 h of fermentation. Other alcohols, including methanol, 1-propanol, 1-butanol, and 1-pentanol, were also produced at variable levels depending on the temperature, fermentation time, and pretreatment method. This study provides a comprehensive understanding of how oxidative pretreatment enhances microalgal biomass conversion, and it identifies optimal conditions for maximizing biofuel yields. The findings offer scalable insights for the development of sustainable microalgae biorefineries and contribute to bridging the gap between laboratory research and industrial biofuel production using mixed cultures of marine microalgae.