Investigating the Potential of Grass Biomass (Thysanolaena latifolia) as an Alternative Feedstock for Sugar Platforms and Bioethanol Production

Abstract

Bioethanol, a lignocellulosic biofuel, has increased energy sustainability and lessened the environmental effects associated with energy production. Thysanolaena latifolia is a common weed found in the northern part of Thailand that is considered non-food biomass, with a high biomass productivity of approximately 10.2 kg/year. Here, we evaluated the potential of T. latifolia biomass as an environmentally friendly material source for producing alternative bioethanol. To this end, we treated the feedstock under mild conditions using various concentrations of phosphoric acid to create ideal conditions for enzymatic hydrolysis. Pretreatment with 75% phosphoric acid yielded the highest solid recovery (55.8 ± 0.6%) and glucans (93.0 ± 0.3%). Additionally, the hydrolysis efficiency and glucose yield of treated biomass were significantly improved. As a result, the liquid hydrolysate from T. latifolia used for ethanol fermentation by Saccharomyces cerevisiae TISTR 5339 generated 8.9 ± 0.0 g/L ethanol. These findings demonstrate that glucose derived from liquid hydrolysate is a promising sustainable carbon source for producing ethanol from T. latifolia feedstock. Thus, using T. latifolia as a feedstock for generating ethanol can improve the efficiency of bioenergy production.

1. Introduction

In response to the growing demand for energy, the need to mitigate environmentally negative effects, and national energy security concerns associated with conventional energy sources, alternative renewable energy sources such as bioethanol have become increasingly attractive [1,2,3,4]. Specifically, bioethanol is a sustainable, ethanol-based biofuel that could result in significantly reduced detrimental pollution emissions due to its highly efficient combustion efficiency [5,6]. Consequently, bioethanol manufactured through lignocellulose-based processing represents a viable solution that may reduce pollution while ameliorating associated environmental and regional security concerns [2,7,8]. Lignocellulosic biomass is considered an abundant and renewable energy substitute capable of providing green, clean energy [9,10,11]. Lignocellulosic materials are derived from plants and include agricultural waste, hardwood and softwood, non-food crop plants, and weed biomass. Plant cell wall material comprises cellulose, hemicellulose, and lignin [5,7]. Therefore, on this basis, lignocellulosic biomass, in particular weeds, has been proposed as a feedstock for bioethanol production. Weedy biomass has the advantages of ubiquity and relatively low cost, as well as being a non-food crop. A significant number of different weed species with a high biomass yield consisting of low water content and nutrient composition, growing under poor conditions, can be found globally [12]. Thus, weedy lignocellulosic biomass has excellent potential as feedstock for biofuel production [13,14,15].

Thysanolaena latifolia (formerly Thysanolaena maxima) [16] is an inedible weed commonly found in the northern region of Thailand and is native to Southeast Asia. T. latifolia is a member of the Poaceae family and is commonly known as tiger grass or broom grass. This weed grows in large clumps, has a cycle of up to 10 years, thrives in both temperate and subtropical regions, and can endure a wide range of conditions, from hillsides and valleys to rocks, degraded land, and wasteland, or can also be cultivated on agricultural land [17,18]. T. latifolia has an average aboveground biomass productivity of approximately 10.2 kg/year [19]. The uses of this plant are diverse; its leaves and tips are used as fodder for animals, while its bushy panicles are used in broom manufacturing, owing to their appearance, which resembles a fox tail. However, the stem is usually discarded by burning after removal of the panicles, releasing large amounts of carbon monoxide and nitrogen dioxide as air pollutants [15,20]. Therefore, utilizing T. latifolia biomass in sugar platforms to produce biofuels, such as bioethanol, will contribute to significantly reducing air contamination.

Nevertheless, the complex structure of lignocellulosic biomass is a major bottleneck in saccharification, which prevents biomass from being converted into biofuels. Thus, lignocellulose pretreatment is necessary to circumvent the inadequate accessibility of the lignocellulosic fibrous cell wall architecture for fermentation. Additionally, pretreatment helps improve the enzymatic digestion of treated substrates and produces monomeric sugars appropriate for further fermentation to produce bioethanol [4,21].

Several pretreatment processes involving physical, chemical, physicochemical, and biological reactions are applied to lignocellulosic materials [7,9]. One such method, acid-based pretreatment, is most often used because of its efficacy in disrupting the lignocellulosic structure, thereby improving cellulose conversion [9]. In 2006, Zhang et al. [22] developed a promising method based on phosphoric acid. Cellulose, a widely distributed natural polymer, demonstrates a distinctive property of being soluble in phosphoric acid. According to Zhang et al.’s study findings, cellulose, particularly Avicel, can only be dissolved in phosphoric acid when the concentration of the acid is higher than 80.5%. This concentration level is considered a key limit for the dissolving process. The process of disintegration takes place in two separate stages. The process begins with esterification, where phosphoric acid interacts with the hydroxyl groups (OH) in cellulose molecules to form cellulose phosphate (cellulose-O-PO₃H₂). The acid becomes attached to the cellulose molecular chain. In the next phase, hydrogen bonds are formed as cellulose hydroxyl groups and solvent (phosphoric acid) molecules compete for bonding opportunities. This competition causes the phosphoric acid and cellulose to separate, with the cellulose remaining largely unchanged chemically. This process requires maintaining low temperatures, typically between 30 and 70 degrees Celsius, to prevent hydrolysis, a byproduct that can hinder its effectiveness. Hydrolysis can degrade cellulose molecules, which is not the desired outcome. By carefully controlling the temperature and acid concentration, researchers can improve cellulose dissolution in phosphoric acid, generating valuable insights for various scientific and industrial applications [22,23]. It was reported that phosphoric acid is an excellent agent for pretreating various biomass species because it is capable of breaking the linkages between lignocellulose components, and it partially breaks the hydrogen bonds in cellulose fibers under mild conditions. Furthermore, compared to dilute acid pretreatment, phosphoric acid pretreatment effectively removed a greater amount of lignin [23,24]. Compared to the more popular acid pretreatment procedures, phosphoric acid pretreatment is a more practical option that results in lower levels of activity and hazard. Furthermore, phosphoric acid has been progressively utilized in the method of mild acid pretreatment, due to its utilization of pretreatment wastes as a source of nutrients and neutralizers [25]. Investigations of various agricultural wastes, grasses, eucalyptus, and poplar wood types indicate that various amounts of phosphoric acid used in treatments are efficient [26,27,28]. Various optimization approaches have been implemented to enhance the conditions during pretreatment and hydrolysis via enzymes [29,30]. Premjet et al. [31] used phosphoric acid solutions ranging from 70% to 80% to treat weed biomass, breaking down cell walls before enzymatic hydrolysis. They found that S. acuta and A. aspera provided the highest glucose when treated with 75% and 80% phosphoric acid, respectively. This result demonstrates that different weed species have varying acid tolerances. Additionally, we demonstrated that 75% phosphoric acid is effective in treating bark and cores fibers of Thai kenaf, resulting in significantly improved enzymatic hydrolysis yields [32]. Applying 85% phosphoric acid to wheat straw resulted in a significant increase in enzymatic glucan conversion. This conversion rate reached 83% within a 12 h hydrolysis period using 3.5 FPU of cellulase/gram of solid [33]. Chen et al. [34] employed an orthogonal experimental design and found that enzymatic hydrolysis yield of poplar powder was maximally improved (73.44%) when it was treated with phosphoric acid 1.5% (v/v) at 190 °C for 150 min. Tong et al. [27] found that incorporating a surfactant called JFC with dilute phosphoric acid and using a steam explosion method for pretreatment of poplar biomass resulted in the highest enzymatic saccharification yields of 84.62%. At present, there is limited comprehensive data available on the impacts of phosphoric acid pretreatment on various types of weed biomass. This encompasses variations in weed biomass composition, structural alterations, and the consequent enzymatic digestibility. Moreover, there is insufficient information regarding the scale-up of this pretreatment method for industrial uses of weed biomass, as well as efficient techniques for the recovery and recycling of phosphoric acid.

The purpose of the present investigation was to determine the optimal phosphoric acid pretreatment conditions and evaluate the potential of T. latifolia as an alternative feedstock for ethanol production.

2. Materials and Methods

2.1. Sample Collection

T. latifolia sample collection was conducted in the Bo Kluea District of Nan Province, Thailand, in August 2022. T. latifolia specimen accuracy was confirmed by the herbarium of Naresuan University’s Biology Department, in which the specimen was deposited under record number 05935. The fresh biomass was cleaned with water, after which it was kept in the shade for 7 d. Subsequently, the material was optionally cut into small segments of approximately 5 cm and milled into a fine powder using a milling machine (Retsch, Haan, Germany). Thereafter, all samples were sieved using a 150–300-µm laboratory test sieve and deposited in vessels at 25 °C for further testing and analysis [32].

2.2. Analytical Methods

Sugar and ethanol quantity were determined using a high-performance liquid chromatography system (HPLC, Agilent 110; Agilent Technologies, Santa Clara, CA, USA) connected to a refractive index detector (Agilent Technologies). After heating the system to 55 °C, analytical separation was conducted using a 300 mm × 7.8 mm Aminex HPX-87P column (Bio-Rad Laboratories, Hercules, CA, USA), heated to 80 °C, and a 20-µL sample. The mobile phase comprised 0.005 M sulfuric acid at a flow rate of 0.6 mL/min [32]. Sugars, acid-soluble and -insoluble lignin, ash, and ethanol extracts in the raw and treated feedstocks were determined using methods developed by the National Renewable Energy Laboratory (NREL) [35,36,37].

2.3. Pretreatment Procedure

A 300 mg dry sample was mixed with 24 mL of 70%, 75%, and 80% phosphoric acids (v/v) in a 50-mL centrifuge tube. Thereafter, the mixture was mixed by stirring vigorously, after which the tubes were heated to 60 °C in a water bath (Grant W28; Grant Instrument Ltd., Cambridge, UK) for 60 min. Next, approximately 25 mL of acetone was added to the tubes and thoroughly mixed using a stirring rod. The liquid mixture was precipitated using a fixed-angle centrifuge (ScanSpeed 1248; LaboGene, Allerød, Denmark) set at 7000× g for 10 min, and the supernatant was discarded. This procedure was repeated three times. Finally, the solid fraction was washed with deionized water until the pH of the solid fraction was approximately 6.5–7 [32]. The lignin removal percentage was computed using Equation (1):

$$\mathrm{L}\mathrm{i}\mathrm{g}\mathrm{n}\mathrm{i}\mathrm{n} \mathrm{r}\mathrm{e}\mathrm{d}\mathrm{u}\mathrm{c}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}\left(\%\right)=100-\mathrm{l}\mathrm{i}\mathrm{g}\mathrm{n}\mathrm{i}\mathrm{n} \mathrm{r}\mathrm{e}\mathrm{c}\mathrm{o}\mathrm{v}\mathrm{e}\mathrm{r}\mathrm{y}$$

(1)

The recovery yield percentage was determined using Equation (2):

$$\mathrm{R}\mathrm{e}\mathrm{c}\mathrm{o}\mathrm{v}\mathrm{e}\mathrm{r}\mathrm{y} \mathrm{y}\mathrm{i}\mathrm{e}\mathrm{l}\mathrm{d}\left(\%{\mathrm{D}}_{\mathrm{w}}\right)=\mathrm{S}\mathrm{R}\left(\%\right)\times \mathrm{t}\mathrm{r}\mathrm{e}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{d} \mathrm{c}\mathrm{o}\mathrm{m}\mathrm{p}\mathrm{o}\mathrm{n}\mathrm{e}\mathrm{n}\mathrm{t} (\%{\mathrm{D}}_{\mathrm{W}})/\mathrm{u}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{e}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{d} \mathrm{c}\mathrm{o}\mathrm{m}\mathrm{p}\mathrm{o}\mathrm{n}\mathrm{e}\mathrm{n}\mathrm{t} (\%{\mathrm{D}}_{\mathrm{W}})$$

(2)

where SR and %D_w are solid recovery and percentage dry weight, respectively [32].

2.4. Crystallinity and Morphology Analysis

The treated and untreated feedstock were rinsed three times with acetone and allowed to dry in the air overnight at 25 °C. Next, the dried sample was pulverized and screened by passing through a 150-μm sieve. Thereafter, the crystalline structure of the sample was scanned using an X-ray diffractometer (PANalytical X’pert Pro, PW 3040/60; PANalytical, Almelo, The Netherlands) at a rate of 0.02° s⁻¹/min from 10° to 40°. The crystallinity index (CrI) was determined using the Segal et al. [38] formula (Equation (3)) [32]:

$$\mathrm{C}\mathrm{r}\mathrm{I}\left(\%\right)={\displaystyle \frac{{\mathrm{I}}_{002}-{\mathrm{I}}_{\mathrm{a}\mathrm{m}}}{{\mathrm{I}}_{002}}}\times 100$$

(3)

The crystalline region is most intense at 2θ = 22.0°, denoted as I₀₀₂, whereas the amorphous zone is least intense at 2θ = 18.2°, denoted as I_am.

The treated and untreated samples were freeze-dried and attached to stubs using dual-sided glue. Thereafter, a thin layer of gold was applied to all samples before analysis, and imaging was conducted using a field-emission scanning electron microscope (SEM, Apero S; Thermo Fisher Scientific, Waltham, MA, USA) [39].

2.5. Biomass Enzyme Saccharification

Treated and untreated samples were subjected to an enzymatic hydrolysis process. Briefly, 0.1 g of biomass (dry weight) was added to a 50-mL Erlenmeyer flask. The reaction mixture comprised 0.1 mL of 0.2% (w/v) of sodium azide solution and 0.05 M sodium citrate buffer with a pH of 4.8. The total reaction volume was 10 mm. For each gram of dry biomass, the enzyme mixture comprised 30 filter paper units of Trichoderma reesei C2730 cellulase (Sigma-Aldrich, St. Louis, MO, USA) and 60 units of β-glucosidase (Oriental Yeast Co., Ltd., Tokyo, Japan). The reaction mixture was agitated for 72 h at 150 rpm and 50 °C using a rotary shaker (Innova 4340; New Brunswick Scientific Co., Inc., Edison, NJ, USA). Subsequently, 20 µL of hydrolysates were collected for HPLC sugar analysis at 12, 24, 48, and 72 h [32]. Hydrolysis efficiency (HE) and glucose recovery (GR) were calculated using Equations (4) and (5), respectively:

$$\mathrm{H}\mathrm{E}={\displaystyle \frac{\mathrm{g}\mathrm{l}\mathrm{u}\mathrm{c}\mathrm{o}\mathrm{s}\mathrm{e} \mathrm{r}\mathrm{e}\mathrm{l}\mathrm{e}\mathrm{a}\mathrm{s}\mathrm{e} \left(\mathrm{g}\right)\times 0.9\times 100}{\mathrm{i}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l} \mathrm{g}\mathrm{l}\mathrm{u}\mathrm{c}\mathrm{a}\mathrm{n}\mathrm{b}\mathrm{i}\mathrm{o} \mathrm{m}\mathrm{s}\mathrm{s} \left(\mathrm{g}\right)}}$$

(4)

$$\mathrm{G}\mathrm{R}\left(\%\right)=\mathrm{S}\mathrm{R} (\%)\times \mathrm{g}\mathrm{l}\mathrm{u}\mathrm{c}\mathrm{a}\mathrm{n} (\%)\times 1.11\times \mathrm{H}\mathrm{E} (\%)\times 100$$

(5)

where SR is solid recovery and 0.9 and 1.11 represent the converting coefficients of cellulose to glucose [32].

2.6. Preparation of Biomass Hydrolysate

The biomass hydrolysate of T. latifolia (BHT) was obtained after saccharification and further treated at 100 °C for 20 min in a water bath (Grant W28; Grant Instruments, Ltd.). Afterward, the BHT was centrifuged at 12,000× g, 10 °C for 2 h and filtered through a glass microfiber filter. Next, the solution was evaporated to attain approximately 20 g/L glucose, and 1 M NaOH was added to increase the pH to 6. Finally, the BHT was stored at 4 °C for further experiments [40].

2.7. Ethanol Fermentation

BHT and control media (CM) were used to produce ethanol. Commercial glucose was used as the carbon source in CM medium. Biomass-produced glucose was present in BHT media. Both media comprised 20 g/L glucose, 10 g/L yeast extract, 10 g/L peptone, 2 g/L K₂HPO₄, and 2 g/L MgSO₄, and pH was set to 6. A 0.02-µm Millipore filter (Millipore Sigma, Burlington, MA, USA) was used to sterilize both media. Fifty milliliters of BHT medium, CM, and 2% yeast inoculum were combined, and the mixture was subsequently placed in a shaker incubator (Innova 4340; New Brunswick Scientific) calibrated at 150 rpm and 30 °C to produce ethanol. Following incubation for 3, 6, 9, 12, 15, 18, 21, and 24 h, the liquid fraction was collected for HPLC to determine the amount of sugar consumed and ethanol produced. The ethanol yield percentage was determined using Equation (6) [40,41]:

$$\mathrm{E}\mathrm{t}\mathrm{h}\mathrm{a}\mathrm{n}\mathrm{o}\mathrm{l} \mathrm{y}\mathrm{i}\mathrm{e}\mathrm{l}\mathrm{d}\left(\%\right)={\displaystyle \frac{\mathrm{a}\mathrm{m}\mathrm{o}\mathrm{u}\mathrm{n}\mathrm{t} \mathrm{o}\mathrm{f} \mathrm{e}\mathrm{t}\mathrm{h}\mathrm{a}\mathrm{n}\mathrm{o}\mathrm{l} \mathrm{g}\mathrm{e}\mathrm{n}\mathrm{e}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{d} \left(\mathrm{g}\right)\times 100}{0.511\times \mathrm{a}\mathrm{m}\mathrm{o}\mathrm{u}\mathrm{n}\mathrm{t} \mathrm{o}\mathrm{f} \mathrm{i}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l} \mathrm{g}\mathrm{l}\mathrm{u}\mathrm{c}\mathrm{o}\mathrm{s}\mathrm{e} \left(\mathrm{g}\right)}}$$

(6)

2.8. Microbial Strain

The yeast strain Saccharomyces cerevisiae TISTR 5339 used for ethanol fermentation was obtained from the Thailand Institute of Scientific and Technological Research (TISTR), Khlong Luang, Thailand. To obtain the inoculum, yeast on an agar slant was transferred to 10 mL of liquid yeast malt medium, after which it was incubated at 30 °C for 18 h in a rotary shaker incubator (Innova 4340; New Brunswick Scientific) at 180 rpm. UV spectrophotometry (SP-830 Plus; Metertech, Taipei, Taiwan) was used to measure yeast cell growth at an optical density (OD) of 600 nm. Approximately 1.5 × 10⁷ cells/mL are present when the OD at 600 nm equals one [40].

2.9. Quantitative Analysis

Data are presented as means ± standard deviation, and tests were performed in triplicate. Data were analyzed using ANOVA followed by Tukey’s test to compare treatment means at a 5% significance level in SPSS version 26 (SPSS Inc., Chicago, IL, USA).

3. Results

3.1. Characterization of T. latifolia Biomass

The chemical composition (% Dw) of the raw experimental biomass used in the present investigation was 32.9 ± 0.5% glucan, 23.8 ± 0.2% xylan, 5.7 ± 0.0% arabinan, 22.2 ± 0.5% acid-soluble lignin (AIL), 5.9 ± 0.1 acid-insoluble lignin (ASL), 9.4 ± 0.1% ash, and 4.3 ± 0.2% ethanol extractives.

3.2. Effect of Phosphoric Acid Concentration on Chemical Composition

To determine the optimal pretreatment conditions, we treated the T. latifolia feedstock with various phosphoric acid concentrations. When the phosphoric acid concentration increased, both the relative xylan and arabinan contents decreased substantially. The lowest xylan (9.0 ± 0.0%) and arabinan (3.2 ± 0.0%) yields were obtained at 80% phosphoric acid. However, a gradual increase in relative glucan percentage was observed upon treatment with 70%, 75%, and 80% phosphoric acid. The maximum relative glucan content (95.7 ± 0.5%) was produced with 80% phosphoric acid (

Table 1. Chemical composition of feedstock before and after phosphoric acid pretreatment.

Composition (% dw)	Raw Material	H3PO4 Concentration (% v/v)
		70	75	80
Glucan	32.9 ± 0.5 d	50.4 ± 0.3 c	54.9 ± 0.2 b	59.7 ± 0.5 a
Xylan	23.8 ± 0.2 a	12.4 ± 0.1 b	10.2 ± 0.2 c	9.0 ± 0.0 d
Arabinan	5.7 ± 0.0 a	3.4 ± 0.0 b	3.3 ± 0.0 c	3.2 ± 0.0 d
AIL	22.2 ± 0.5 a	18.0 ± 0.6 b	13.6 ± 0.2 c	12.0 ± 0.6 d
ASL	5.9 ± 0.1 a	3.7 ± 0.0 b	3.6 ± 0.1 b	3.3 ± 0.1 c
Total lignin	28.2 ± 0.4 a	21.7 ± 0.6 b	17.1 ± 0.1 c	15.3 ± 0.7 d
Solid recovery	100.0 ± 0.0 a	62.2 ± 0.7 b	55.8 ± 0.6 c	50.7 ± 0.9 d
Glucan recovery	100.0 ± 0.0 a	95.3 ± 0.5 b	93.0 ± 0.3 c	91.9 ± 0.8 c
Xylan recovery	100.0 ± 0.0 a	32.4 ± 0.2 b	23.8 ± 0.4 c	19.2 ± 0.1 d
Arabinan recovery	100.0 ± 0.0 a	37.1 ± 0.2 b	32.4 ± 0.1 c	28.5 ± 0.3 d
AIL recovery	100.0 ± 0.0 a	50.4 ± 1.6 b	34.0 ± 0.5 c	27.4 ± 1.3 d
ASL recovery	100.0 ± 0.0 a	39.1 ± 0.5 b	33.4 ± 0.8 c	27.9 ± 0.7 d
Total lignin recovery	100.0 ± 0.0 a	47.9 ± 1.3 b	33.9 ± 0.2 c	27.5 ± 1.2 d
Total lignin removal	n.d.	52.0 ± 1.3 d	66.1 ± 0.2 c	72.5 ± 1.2 b

The superscript characters (a, b, c, and d) indicate statistically significant differences in the variation of each parameter at a 5% significance level. n.d. = not determined. AIL and ASL are acid-insoluble lignin and acid-soluble lignin, respectively.

). In addition, increased phosphoric acid concentration led to significant declines in the relative amounts of AIL, ASL, and total lignin (p < 0.05). Although the relative ASL content was reduced by treatment with 70% (3.7 ± 0.0%) and 75% (3.6 ± 0.1%) phosphoric acid, this outcome did not demonstrate statistical significance (p < 0.05). However, relative ASL decreased remarkably upon treatment with 80% (3.3 ± 0.1%) phosphoric acid. Meanwhile, total lignin removal (72.5 ± 1.2%) was most effective at a concentration of 80% phosphoric acid (Table 1).

Additionally, total lignin, glucan, xylan, arabinan, AIL, and ASL recovery yields decreased significantly with increasing phosphoric acid concentrations. Feedstock treatment with 70% phosphoric acid resulted in the highest recovery of solids (62.2 ± 0.7%), glucan (95.3 ± 0.5%), xylan (32.4 ± 0.2%), arabinan (37.1 ± 0.2%), AIL (50.4 ± 1.6%), ASL (39.1 ± 0.5%), and total lignin (48.0 ± 1.3%). Nevertheless, the decline in glucan recovery yield was not significantly different (p < 0.05) at phosphoric acid concentrations of 75% (93.0 ± 0.3%) and 80% (91.9 ± 0.8%). However, treating the sample with 80% phosphoric acid resulted in minimal recovery yields for solids (50.7 ± 0.9%), glucan (91.9 ± 0.8%), xylan (19.2 ± 0.1%), arabinan (28.5 ± 0.3%), AIL (27.4 ± 1.3%), ASL (27.9 ± 0.7%), and total lignin (27.5 ± 1.2%) (Table 1).

3.3. Impact of Phosphoric Acid Concentration on Cellulose Crystallinity

The relative CrIs and X-ray diffraction (XRD) patterns of both untreated and treated feedstocks are presented in Figure 1 and

Table 2. Crystallinity index of raw and phosphoric acid-pretreated feedstock.

	Untreated	Concentration of Phosphoric Acid
		70%	75%	80%
CrI (%)	54.1	57.7	61.2	59.1

. When the raw material was treated with phosphoric acid at different concentrations, changes in cellulose crystallinity were shown by the XRD patterns and CrI values. The XRD pattern of raw materials, which included three prominent peaks at 2θ = 15.9°, 22.0°, and 34.5° (Figure 1), and the relative CrI value of 54.1%, led to a classification of cellulose I. After treating the sample with 70% and 75% phosphoric acid, the CrIs rose to 57.7% and 61.1%, respectively. However, after treatment with 80% phosphoric acid, the CrI dropped to 59.1%. Furthermore, the XRD patterns of the cellulose I crystallinity sample treated with 70%, 75%, and 80% phosphoric acid stayed the same.

3.4. Impact of Phosphoric Acid on Biomass Morphology

The results shown in Figure 2A reveal that the morphological surface structure of the raw material is uniform, integrative, and manifests close bundles. The alteration in the actual feedstock’s surface structure became more distinct for samples subjected to treatment with varying concentrations of phosphoric acid. For example, exposure to 70% phosphoric acid resulted in an initial roughing, splinting, expanding, and loosening of the surface (Figure 2B). As indicated, the surfaces of all samples treated with 75% phosphoric acid concentration behaved similarly, but the peeling, delamination, loosening, and surface disintegration increasingly intensified (Figure 2C). Moreover, increasing phosphoric acid concentrations led to cellulose fiber disintegration and impaired morphological structures (Figure 2D).

3.5. Enzymatic Saccharification Yields

Our results indicated that the HE and GR yields of both untreated and pretreated samples dramatically increased within 12 h of incubation and further slightly improved in 24, 48, and 72 h (Figure 3). The untreated and pretreated samples exhibited maximum HE and GR yields at 72 h incubation. The untreated sample achieved the lowest HE and GR yields of 29.5 ± 0.7% and 12.0 ± 0.3%, respectively. However, all pretreated samples displayed higher HE and GR yields compared with those of raw material. Treatment with 70%, 75%, and 80% phosphoric acid resulted in HE yields of 65.3 ± 0.6, 86.9 ± 0.7, and 87.2 ± 0.2%, and GR yields of 25.3 ± 0.2, 32.8 ± 0.3, and 32.5 ± 0.1%, respectively. However, the HE and GR yields of 75% and 80% phosphoric acid treatment did not significantly differ.

3.6. Bioethanol Fermentation

The BHT medium produced by enzymatic hydrolysis without a detoxification process was subjected to fermentation using S. cerevisiae TISTR 5339 to assess its potential for bioethanol production. The results indicated that glucose consumption, ethanol synthesis, yeast cell proliferation, and pH levels exhibited similar trends in both the CM and BHT media (Figure 4 and Figure 5). During fermentation, yeast cell growth profiles and the pH observed in both the CM and BHT media differed slightly. CM demonstrated rapid growth, reaching the stationary phase at 15 h; the growth rate in BHT medium was slower than that in CM (Figure 4). The pH of both CM and BHT media decreased from 6.0 to 5.2. In addition, S. cerevisiae TISTR 5339 rapidly utilized glucose while concurrently producing ethanol. In both CM and BHT media, glucose was entirely consumed in 15 h. Ethanol production observations revealed that CM produced ethanol (1.0 ± 0.1 g/L or 26.7 ± 1.7% of the ethanol yield) within 6 h, whereas BHT medium exhibited delayed ethanol production (1.2 ± 0.0 g/L or 29.0 ± 0.9% of the ethanol yield) to 9 h. Nevertheless, after 15 h of fermentation, maximum ethanol yields were produced from both CM (9.4 ± 0.0 g/L or 91.5 ± 0.3% of the ethanol yield) and BHT medium (8.9 ± 0.0 g/L or 86.6 ± 0.1% of the ethanol yield). Thereafter, ethanol yields declined steadily in both media (Figure 5).

4. Discussion

4.1. Characterization of T. latifolia Biomass

The chemical composition of biomass significantly influences the bioenergy production performance [42]. Therefore, analyzing chemical composition reveals whether lignocellulose can be a suitable alternative. As shown in our results, the chemical composition of T. latifolia biomass comprised three major components: lignin, hemicellulose, and cellulose. The amount of total lignin significantly differed from that reported in previous studies, while amounts of glucan and xylan were similar to previous reports [43]. Differences in the composition of lignocellulosic biomass are due to multiple factors, including growth conditions, sources, period of harvest/stage of development, and other influencing factors [8,44,45]. However, the lignin content of T. latifolia was lower than that of switchgrass (31.2%) or eucalyptus (29.4%) [46], and several other agricultural wastes that have excellent lignocellulosic properties, including sorghum straw, (30.4%) [44], cotton stalks (30.0%), and walnut shells (37.5%) [1]. In addition, the total carbohydrate content of T. latifolia biomass (62.4 ± 0.7%) was higher than that of lignocellulosic materials such as banana pseudo-stems (51.5%), bean straw (55.0%), switchgrass (56.8%), olive tree (57.8%), and sugar cane (60.7%) [4], which are considered potential feedstocks for biofuels owing to a total carbohydrate percentage appropriate for producing cellulosic ethanol [47]

4.2. Effect of Phosphoric Acid Concentrations on Chemical Composition

The resistance of lignocellulosic material to enzymatic degradation necessitates pretreatment to modify biomass properties [5]. This study demonstrated that an increase in phosphoric acid concentration led to a rise in the relative glucan content while simultaneously causing a substantial decrease in the content of relative xylan, arabinan, AIL, ASL, and total lignin. During the pretreatment process, phosphoric acid reacted to break down the glycosidic linkages and partially break the hydrogen bound within the lignin–carbohydrate complex structure of the biomass, resulting in the partial solubilization of hemicellulose, lignin, and cellulose components [4,48,49]. In addition, the removal of hemicellulose and lignin also increases the relative glucan content in the treated sample [33,50].

Additionally, as the concentration of phosphoric acid increased, the carbohydrate and lignin constituents in complex structure dissolved more readily [4,48,49,51], resulting in a substantial decrease in the recovery yields of solids, glucan, xylan, arabinan, and total lignin. Consequently, the amount of xylan, arabinan, and entire lignin recovery remained below 50%. In contrast, the total recoveries of solids and glucans exceeded 50% and 90%, respectively. This study indicated that treating the feedstock with 70–80% phosphoric acid only partially reduced hemicellulose (xylan and arabinan). However, hemicellulose of white jute [52], roselle [32], and both chaff flower, and common wireweed [31] was entirely removed when these feedstocks were treated with 70%, 75%, and 80%, phosphoric acid, respectively. In addition, phosphoric acid pretreatment of poplar revealed that acid concentrations had less impact on decreased hemicellulose than pretreatment temperature [34].

4.3. Impact of Phosphoric Acid Concentration on Cellulose Crystallinity

Due to the presence of both intra- and intermolecular hydrogen bonds, cellulose exhibits exceptional crystalline structure stability, significantly influencing the efficiency of enzymatic hydrolysis [35,53]. Cellulose degradation in phosphoric acid typically corresponds to esterification, which prevents the formation of hydrogen bonds throughout the cellulose chain [23,54]. Consequently, phosphoric acid induces the swelling and disintegration of the crystalline region inside the cellulose structure [54,55]. In the present study, treating samples with 70% and 75% phosphoric acid led to increased relative CrI values, attributed to the partial elimination of amorphous components such as hemicellulose and lignin during the pretreatment stage [50,56,57,58]. However, when treated with 80% phosphoric acid, CrI values dropped slightly to 59.1%. Concentrations of phosphoric acid higher than 80% are capable of completely dissolving cellulose and disrupting its highly ordered hydrogen bonding network [22,23]. Therefore, we can attribute the decrease in relative CrI values to the disintegration of some crystalline sections of cellulose. However, several studies have observed that raising the crystallinity index (CrI) of lignocellulosic materials does not always impact enzymatic hydrolysis yield [31,59,60]. Consequently, these findings suggest that morphological changes caused by removing hemicellulose and lignin have a greater impact on enzymatic hydrolysis efficiency than simply increasing CrI [31].

4.4. Impact of Phosphoric Acid Concentration on Biomass Morphology

The recalcitrant nature of untreated T. latifolia feedstock’s SEM images (Figure 2A) indicates that the material originated from a complex of cellulose and hemicellulose biopolymers, as well as lignin heteropolymers [8,61,62]. The SEM images showed that the feedstock’s morphological structure progressively loosened, delaminated, and fibrillated after phosphoric acid treatment, leading to a quite disorganized arrangement. The severity of these modifications was dependent on phosphoric acid concentrations, suggesting that alterations in the resistance structure of T. latifolia biomass were due to the influence of phosphoric acid, which disrupted the intermolecular connections among cellulose, hemicellulose, and lignin. Consequently, hemicellulose and lignin were separated from the cellulose microfibers [33,49,53,63]. During pretreatment, esterification caused cellulose swelling, which extends cellulose chains, also leading to fiber structure disruption [22,53,55]. In addition to modifying the physical structure of the feedstock, phosphoric acid enhances its surface area and porosity, thereby facilitating enzymatic hydrolysis [27,31,64,65,66,67]. The implications of phosphoric acid concentration on modifying the morphological structure and cellulose crystalline pattern in biomass correspond to alterations in the chemical composition, as discussed in Section 4.2. Furthermore, these effects have been observed in various types of lignocellulosic biomes, including durian peel [68], java jute [69], chaff flower and common wireweed [31], elephant grass [70], and cotton fibers [65].

4.5. Enzymatic Saccharification Yields

We treated the feedstock with 70%, 75%, and 80% phosphoric acid, and the HE and GR yields substantially improved compared with those of the untreated sample. However, the level of improvement varied depending on phosphoric acid concentration in the pretreatment process. These results suggested that the chemical linkages present in the lignin–carbohydrate complex structure were disrupted when the feedstock was treated with 70–80% phosphoric acid. Owing to this disintegration, hemicellulose and lignin were partially eliminated during the pretreatment process, indicating that the physical barriers (hemicellulose and lignin) surrounding cellulose were removed [23,33,49]. Additionally, these effects increased the pore size and cellulose accessibility surface, leading to improved HE and GR yields for each treated sample [41,53,57,68]. Consequently, the greatest amounts of HE and GR were achieved by treatment with 75% and 80% phosphoric acid. The HE yields of samples treated with 75% and 80% phosphoric acid did not significantly differ.

The best pretreatment conditions are characterized by a very high GR value approaching the initial glucan content of the biomass [41,71]. As such, 75% phosphoric acid was the most appropriate pretreatment condition for T. latifolia feedstock because it maximized both the HE (86.9 ± 0.7%) and GR (32.8 ± 0.3%) yields, which were enhanced by approximately 2.9- and 2.7-fold, respectively, compared with the raw material. Moreover, the solid (55.8 ± 0.6%) and glucan (93.0 ± 0.3%) recovery levels were high. Under the aforementioned conditions, the lignin (33.9 ± 0.2%) retained in the treated biomass demonstrated that partially removing lignin from the biomass is necessary. In addition, increasing the percentage of CrI values (61.2%) in the treated samples did not impact the hydrolysis yield. Modifications in the chemical components and morphological and crystalline structures of the treated feedstock correlated with enhanced HE and GR yields, and these observed events are similar to those described for other feedstocks, including durian peel [68], Vietnamosasa pusilla [41], java jute [68], chaff flower and common wireweed [31], and Glycyrrhiza residue [51].

4.6. Bioethanol Fermentation

The non-detoxified BHT medium and CM considerably affected yeast cell proliferation. In addition, CM produced a marginally higher yield compared with that of BHT medium. These results indicated that the non-detoxified BHT medium might contain furfural, 5-hydroxymethyl furfural, levulinic acid, and other aromatic compounds. Some inhibitors are produced by biomass disintegration during the acid pretreatment of raw feedstock [4,23,72]. These inhibitors adversely affect microbial growth, leading to a reduction in the rate of sugar uptake, which might subsequently decrease ethanol production [7,23]. Hydrolysate detoxification is considered imperative to prevent inhibitory effects against fermenting organisms, before use as a substrate in bioethanol fermentation [73,74].

We assessed the material balance of T. latifolia feedstock by employing the most effective phosphoric acid pretreatment strategy for producing cellulosic ethanol, as shown in Figure 6. The use of 1000 g of T. latifolia feedstock pretreated with 75% phosphoric acid resulted in the recovery of approximately 558 g of the treated material. Subsequently, both the untreated and treated samples underwent enzymatic hydrolysis to investigate the influence of pretreatment on enzymatic saccharification, which substantially improved both the HE and GR yields. The treated samples exhibited the highest HE (86.9%) and GR (328 g) yields after 72 h of saccharification. In contrast, untreated samples exhibited the lowest production of HE (29.5%) and GR (120 g). Compared with the untreated material, HE and GR yields increased by approximately 2.9- and 2.7-fold, respectively. Subsequently, a non-detoxified BHT medium containing 20 g/L glucose from the treated biomass was fermented using S. cerevisiae TISTR 5339 for ethanol production. Glucose from both untreated and treated feedstocks resulted in an estimated 53 and 145 g of cellulosic ethanol, respectively. The ethanol yield from the treated feedstock demonstrated an enhancement of approximately 2.7-fold compared with that of the untreated samples. The outcomes of our study emphasize the potential of utilizing a non-detoxified BHT medium as a practical source of carbon for S. cerevisiae TISTR 5339.

5. Conclusions

Based on these findings, treating raw materials with phosphoric acid alters feedstock morphology, crystalline makeup, and chemical composition, increasing HE and GR yields. These changes could increase the surface area of treated cellulose, which is favorable for the use of this renewable biomass. Pretreating T. latifolia biomass with 75% phosphoric acid for 60 min at 60 °C achieved the maximum theoretical yield of HE (86.9%) and GR (328 g) based on 1000 g (dry matter) of T. latifolia feedstock. Under these conditions, lignin removal was approximately 66%. Raw samples exhibited the lowest HE (29.5%) and GR (120 g) yields. Subsequently, the hydrolysate from T. latifolia feedstock was used for ethanol production by S. cerevisiae TISTR5339. The maximum ethanol yield (86.6%) was obtained at 15 h of fermentation. Moreover, raw and treated feedstock-based cellulosic ethanol production can be estimated at approximately 53.1 and 145.1 g, respectively. Therefore, the results of this study establish that T. latifolia represents a low-cost, renewable material from lignocellulosic feedstock that can produce green biofuels and bio-based chemicals without disrupting the world’s food supply. In future studies, analyzing the effects of pretreatment parameters using response surface methodology or similar approaches will be crucial. Furthermore, bioethanol yields can be increased by enhancing fermentation efficiency by modifying microbial strains to improve their efficiency and tolerance to inhibitors. Furthermore, investigating the scalability and economic feasibility of producing cellulosic ethanol from T. latifolia on a larger scale could provide valuable insights for commercial applications. Exploring the potential use of by-products for creating value-added chemicals or materials can improve the process’s economic viability. Assessing the environmental impact and carbon footprint of the entire bioconversion process is essential for evaluating its overall sustainability.