Low-Concentration Chemical Pretreatment of Lignocellulose Biomass Derived from Hemp Hurds, Agricultural Waste: Comparative Characterisation of Various Pretreatment Methods

Article type: Research Article

Keywords: agricultural waste, HH, pretreatment, crystallinity, delignification

Authors: Ziningi Rosebud Myeni ⚬, Farai Dziike ⚬, Tshwafo Elias Motaung ⚬, Nirmala Deenadayalu ⚬

Affiliations: Technology Transfer and Innovation Directorate, Durban University of Technology, Durban 4000, South Africa; Department of Chemistry & Chemical Technology, Sefako Makgatho Health Sciences University, Ga-Rankuwa 0204, South Africa

License: © 2026 by the authors. CC BY 4.0 Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license.

Article links: DOI: 10.3390/ma19050843  |  PubMed: 41828111  |  PMC: PMC12986429

Relevance: Relevant: mentioned in keywords or abstract

Full text: PDF (2.2 MB)

1. Introduction

Lignocellulosic biomass (LB) has been a significant area of interest in the hunt for sustainable energy sources due to rising energy demands and the need to reduce greenhouse gas emissions (GHGs) [ref. 1]. Bio-based products, including biochemicals and biofuels derived from industrial and agricultural waste, present a compelling alternative to fossil fuels and provide new avenues for mitigating environmental damage [ref. 2]. LB is a viable renewable resource for producing valuable bio-based products due to its abundance and sustainability. The primary sources of LB are agro-industrial residues, energy crops, woody biomass, and municipal solid wastes [ref. 3,ref. 4].

Lignin, cellulose, and hemicellulose are essential constituents of LB, crucial for plant cell wall structure. Cellulose (C₆H₁₀O₅)_n is a linear polysaccharide consisting of about 10,000 D-glucose units bonded together by β-(l → 4) glycosidic bonds [ref. 5,ref. 6]. Lignin[C₉H₁₀O₃(OCH₃)_0.9–1.7]_n is a complex hydrophobic polymer belonging to polyphenols and offers structural support. It forms a protective barrier through covalent bonding with cellulose and hemicellulose, rendering them highly resistant to chemical and biological degradation needed for the bioconversion process [ref. 7]. Hemicellulose(C₅H₈O₄)_m, a polymeric amorphous carbohydrate, is made up of 100 to 200 sugar units, such as glucose, xylose, mannose, and galactose, linked together by β-(1,4)- and/or β-(1,3)-glycosidic bonds [ref. 4].

The presence of the three polymers in LB reduces enzymes’ access to cellulose and hemicellulose, which leads to bioproducts manufacturing [ref. 5]. Cellulose is hydrolysed into fermentable sugars which serve as primary carbon and energy source for fermenting microorganisms [ref. 8]. These microorganisms, such as bacteria and yeast, metabolise sugars through glycolysis and convert them into pyruvate. This pyruvate is then converted into final product depending on the biological pathway in the particular microorganism [ref. 9]. Figure 1 demonstrates the pathway followed by LB for bioconversion into various bioproducts.

Within this process, pretreatment is the most energy-intensive step, accounting for approximately 40% of the total manufacturing cost. Pretreatment technologies must be simple, green, and both economically and technically viable. The cost associated with LB pretreatment and hydrolysis processes pose a significant barrier to industrial LB bioconversion into bioproduct. The selection of the pretreatment technology depends on the type of biomass, the intended bio-product, and existing technological capabilities. Therefore, there is a need for effective pretreatment technologies to remove lignin and hemicellulose and give access to cellulose for bioconversion [ref. 10].

Various pretreatment technologies have been investigated, differing in their effectiveness and cost implications [ref. 11]. Mechanical pretreatment reduces the LB through grinding, milling, or chipping. The final choice of size reduction depends on the specifications for the biomass, which is often reduced by 10–30 mm during chipping and 0.2–2 mm during grinding and milling operations [ref. 12,ref. 13]. Mechanical pretreatment is beneficial for altering the physical condition of LB, but it is insufficient alone, particularly in bioproduct manufacturing, as it does not entirely remove lignin [ref. 14]. Therefore, mechanical pretreatment alone is insufficient and requires the addition of chemical or biological methods to enhance the process; however, the inclusion of another method increases operating costs. Additionally, excessive grinding can generate undesirable waste and inconsistent particle sizes, which negatively affect operations [ref. 15].

The chemical pretreatment method uses acids, alkalis, organic solvents, and ionic liquids to degrade plant cell walls. The method solubilises hemicellulose and removes lignin [ref. 16]. This increases accessibility to cellulose for conversion into bioproducts. The choice of chemical pretreatment method depends on the type of biomass, application, cost, and environmental impact [ref. 17]. The benefits of using chemical pretreatment methods include high efficiency and increased reaction rates compared to biological approaches [ref. 13]. Nonetheless, the method has disadvantages, including the production of toxic chemicals that negatively affect subsequent processes [ref. 18]. Additionally, the techniques increase operational costs associated with the disposal of toxic chemical waste.

Biological pretreatment is an environmentally friendly method that involves the action of microbes, enzymes, bacteria, fungi, insects, and worms, which can eliminate the hemicellulose and lignin content without affecting the cellulosic content [ref. 19]. The microorganisms decompose LB into its constituents, which is crucial for enzymatic hydrolysis. This method requires no chemicals and yields high results; however, it has drawbacks, including time consumption and the need for continuous monitoring of growth conditions [ref. 20]. These considerations influence the economic feasibility and efficacy of the biological pretreatment method.

Physicochemical pretreatment combines mechanical and chemical methods to break down LB matrix. The method reduces energy consumption compared to physical pretreatments, making it a cost-effective option. Effective physicochemical methods for LB include steam explosion, ammonia fibre explosion, carbon dioxide (CO₂) explosion, and liquid hot water (hydrothermal) pretreatment [ref. 21]. A steam explosion utilises increased pressure and temperature, causing water molecules to penetrate the LB structure and disrupt the bond between hemicellulose and lignin. The steam explosion pretreatment is environmentally friendly and uses less energy; however, it produces toxic waste [ref. 22].

The ammonia fibre explosion uses pressurised liquid ammonia at temperatures ranging from 60 to 100 °C for a duration of 90 min, facilitating the cleavage of C–O–C bonds between lignin and cellulose. Ammonia is corrosive and harms the environment, making it necessary to recover it. The cost associated with recovering ammonia is high for industrial applications [ref. 23]. CO₂ explosion operates like steam and ammonia fibre explosions, but it offers more benefits than these two approaches. Under high pressure, CO₂ breaks through the cell wall structure of LB, penetrating and disrupting the cellulose and non-cellulose structure. The CO₂ explosion method is less expensive than the ammonia method and safer for the environment, as it reduces the amount of greenhouse gases in the air.

Traditional methods, although successful, typically use harsh chemicals that produce toxic waste. The methods lack environmental sustainability and require significant energy consumption. In recent years, the focus has shifted towards using sustainable procedures due to their environmental benefits and minimal waste production.

This study compares and assesses various environmentally friendly pretreatment methods for HH, a type of agricultural waste. The methods include organic solvent pretreatment with 60% ethanol (EtOH), a combined oxidative-alkaline method utilising 3% hydrogen peroxide and ammonia (H₂O₂/NH₃), and ultrasound-assisted sodium hydroxide (NaOH) pretreatment, under fixed experimental conditions. Differences in the recovered HH had pronounced changes that were observed across characteristics techniques used. This study examines the biomass change in structure and surface morphology following each treatment, illustrated in Figure 2. The observations allow for a comparative assessment of pretreatment methods under the fixed experimental conditions, without isolating individual chemical or physical contribution. The findings provide a preliminary guidance for selecting pretreatment routes for HH. The pretreatment methods emphasised low reagent concentrations, aqueous systems, moderate temperatures, short reaction times, and avoidance of strong mineral acids. Energy input was minimised using mild heating and ultrasound-assisted treatment.

2. Materials and Methods

2.1. Materials

The HHs were sourced from hemp stalks during the hemp trial studies at Cedara (Hilton, South Africa), in collaboration with the Agricultural Research Council and the Department of Agriculture, Pietermaritzburg, South Africa. The HHs were obtained from a mixed cultivation of ARC-CAN-01 and ARC-CAN-03 strains in an approximately 50:50 ratio. D-(+)-glucose standard of >99.5% was used as a standard reference material for creating the standard curve as well as a control. Chemicals and reagents used were of analytical grade (AR) and sourced from Sigma-Aldrich a life science business of Merk kGaA, Darmstadt, Germany. The following chemicals, Sulphuric acid concentration of 98.0%, Absolute ethanol 99.9%, Sodium acetate AR, Sodium hydroxide pellets (AR), 30% Hydrogen peroxide, and D-(+)-glucose standard of >99.5% were used.

2.2. Methods

2.2.1. Sample Preparation

The HHs were separated from the hemp stalk via manual decortication, ground and sieved through a 1 mm mesh, and particles passing through the sieve (<1 mm) were collected for all experiments. This process increased the surface area and ensured sample homogeneity. The final product of HH was stored in an airtight container to prevent moisture uptake before use.

2.2.2. Polysaccharide Quantification

In this study, total polysaccharides were quantified using the phenol–sulphuric acid colorimetric method, differing from the National Renewable Energy Laboratory (NREL) method, which uses HPLC after acid hydrolysis to quantify individual carbohydrates. The phenol–sulphuric acid method is recognised for screening total carbohydrates in LB, emphasising overall carbohydrate availability rather than specific sugar types. For polysaccharide content analysis, the phenol–sulphuric acid method was used. A total of 5% of phenol crystals was prepared by weighing 5.0 g of phenol crystals and slowly dissolving them in around 80 mL of deionised water at room temperature with gentle stirring. After that, the solution was put into a 100 mL volumetric flask and filled to the top with deionised water to make a 5% (w/v) phenol solution. Sodium acetate was prepared by dissolving 0.82 g of sodium acetate in about 80 mL of deionised water and it was put in a 100 mL volumetric flask. Deionised water was used to bring volume up to the mark. For 60% ethanol, from 99.9% absolute ethanol, 90 mL of absolute ethanol was mixed with 60 mL of deionised water to make 150 mL. To make 2% NaOH, 1.0 g of NaOH pellets were slowly added to around 40 mL of deionised water while stirring and cooling. When everything was dissolved, the solution was put into a volumetric flask and filled with deionised water to make 50 mL. A commercial hydrogen peroxide stock solution of 30% (w/v) s used to generate 3% hydrogen peroxide. To make 50 mL of 3% hydrogen peroxide, 5 mL of the stock solution was added to 45 mL of deionised water. To make 3% aqueous ammonia, 6 mL of commercial ammonia with 25% (w/v) was added to 44 mL of deionised water. This made 3% (w/v) of ammonia in water. D-(+)-glucose standard of >99.5% was used as a standard reference material for creating the standard curve.

A total of 50 mg of HH was mixed with 2 mL concentrated 72% sulphuric acid and this was conducted in triplicates and glucose control. The mixture was capped, vortexed, and placed in an ice water bath for 1 h. A total of 4 mL of deionised water was added, and the mixture was kept for 10 s. An additional 6 mL of deionised water was added, followed by loosening the caps and placing the tubes in a 100 °C water bath for 2 h. After 2 h, the mixture was transferred into a 50 mL volumetric flask and made up to the mark with sodium acetate. Glucose standards for a calibration curve were prepared concurrently with the treated sample, where 2 mL from the sample mixture and different standards were pipetted and 1 mL of 5% phenol was added to each tube for condensation, producing a yellow-orange chromogen. This was followed by 5 mL concentrated sulphuric acid, vortexed, and allowed to stand for 10 min in a room-temperature water bath for 20 min. The reactive intermediates underwent condensation with phenol, producing a yellow-orange chromogen. The intensity of the produced colour, spectrophotometrically determined at approximately 490 nm, is directly proportional to the total carbohydrate content. A calibration curve generated with glucose solutions enabled accurate quantification.

2.2.3. Organic Solvent Pretreatment

To conduct a comparative evaluation of the changes in structure of hemp hurds after exposure to various pretreatment methods, an organic solvent pretreatment was performed, involving 5 g of HH submerged in 50 mL of 60% (v/v) ethanol and heated at 80 °C for 2 h. This treatment was used to partially dissolve lignin and other amorphous components, which changed the structure of the biomass, giving access to cellulose. It is important to look at these physical changes to compare how each pretreatment method affects the structure of hemp hurds and what that means for future biomass use and conversion processes.

2.2.4. Combined Hydrogen Peroxide and Ammonia Pretreatment

A total of 25 mL of 3% hydrogen peroxide and 25 mL of 3% aqueous ammonia were mixed in a 1:1 ratio and added to 5 g of HH for three hours at room temperature. This approach aimed to promote oxidative reaction whilst enhancing the solubilisation of hemicellulose in the HH. The combination of these two agents underscores the method’s focus on modifying the chemical structure of HH to improve their processing and application in preceding treatments.

2.2.5. Ultrasound-Assisted Sodium Hydroxide Pretreatment

A total of 5 g of HH were submerged in 50 mL of 2% sodium hydroxide solution, subjected to ultrasound treatment using a Qsonica Q700 probe Sonicator (maximum rated power 700 W, Qsonica, LLC, Newtown, CT, USA) equipped with a 12.7 mm titanium probe, and operated at 60% amplitude for 30 min. The temperature was maintained at 50 °C throughout the sonication to ensure consistent treatment conditions. This ultrasound NaOH-assisted treatment method was used to enhance lignocellulose structure modification through chemical processing and mechanical induced shear. The method was included to conduct comparative assessment of its effectiveness compared to other pretreatment methods.

No individual controls were included in the study to isolate the chemical and mechanical (sonication) effects. The results are therefore solely comparative observations under fixed conditions.

2.3. Characterisation Method

2.3.1. Crystalline Structure and Crystallinity Index Determination Using XRD

XRD analyses were conducted using Rigaku MiniFlex 600 X-ray generator (Rigaku Corporation, Tokyo, Japan) set at 40 kV and 15 mA, operating in one-dimensional scanning mode. Data acquisition was performed at a scan speed of 40.00°/min using a MiniFlex 600 goniometer (Rigaku Corporation, Tokyo, Japan), resulting in a step width of 0.01° 2θ. A scan rate of 40° min⁻¹ was chosen to find a good balance between acquisition time and data quality for comparing crystallinity. This rate gives enough resolution for determining the Segal crystallinity index, as a guide on the crystalline and amorphous structure.

The system used an ASC-8 attachment for a θ/2θ scanning configuration, encompassing a 2θ scan range of 3° to 90°. A diffraction angle of 1.25° was used, and the diffracted beam was adjusted using an IHS aperture (Hikvision, Hangzhou, China) of 10 mm. The D/teX Ultra2 detector operated in an open SS mode without the implementation of filters, hence preserving the RS and other optical characteristics. These attributes facilitated the acquisition of a precise representation of the diffracted intensities. This enabled an accurate characterisation of the crystalline structure of the HH. The crystallinity index (CrI) was assessed using the Segal technique (Segal, New York, NY, USA), which calculates the ratio of crystalline cellulose by comparing the intensity of the crystalline peak (I₀₀₂) to the amorphous region (I_am) and the following equation:

2.3.2. Functional Group Analysis Using FTIR

Shimadzu Fourier-transform infrared spectroscopy (FTIR) (Shimadzu, Kyoto, Japan) was used to determine the functional groups present in HH. The sample was ground into a fine powder and analysed using an FTIR spectrometer equipped with an attenuated total reflectance (ATR) attachment. The spectra were obtained in the 4000–400 cm⁻¹ region at a resolution of 4 cm⁻¹ over 32 scans to enhance signal quality. Critical peaks were studied to determine the chemical composition and to understand the composition, structural alterations, and degradation patterns of HH powder.

2.3.3. Thermal Stability and Decomposition Behaviour Using TGA

PerkinElmer Thermogravimetric Analyser (TGA) STA 6000 (PerkinElmer, Inc., Waltham, MA, USA) was used to determine the thermal decomposition properties of HH in an inert nitrogen (N₂) atmosphere, preventing undesirable oxidation. The HHs were subjected to a temperature range of 30–105 °C for 5 min to remove moisture. The temperature was increased at a rate of 10 °C/min from 105 °C to 200 °C to facilitate hemicellulose dehydration. Heating at a rate of 10 °C/min from 200 to 400 °C caused cellulose decomposition, followed by an increase from 400 to 800 °C at the same rate, during which lignin degradation and ash formation occurred. Nitrogen gas was used in place of oxygen to prevent oxidative combustion, hence facilitating the thermal breakdown of HH without the impact of oxidation reactions. This method provides various identification of moisture loss, hemicellulose, cellulose, and lignin degradation, offering insights into the thermal stability and composition of HH.

2.3.4. Surface Morphology Using SEM

Scanning electron microscopy (SEM) (JEOL Ltd., Akishima, Japan) was used to assess the morphology and surface structure of the HH powder. The sample was desiccated and coated with a thin layer of gold/palladium via a sputter coater to improve conductivity. The analysis was conducted using SEM equipment at an accelerating voltage of 5–15 kV. High-resolution images were taken at various magnifications, supporting the examination of fibre arrangement, porosity, and surface roughness. SEM provided insights on structural modifications, degradation effects, and the accessibility of HH powder to hydrolysis and enzymatic processes.

3. Results and Discussion

3.1. Quantification of Polysaccharide Content

The quantification of total carbohydrates using the phenol–sulphuric colorimetric method was discovered in 1956 by Dubois and his colleagues [ref. 24]. The results demonstrate measurable changes in total carbohydrate content compared to HH and other pretreatment methods. A glucose calibration curve was generated with glucose standards of 99.5% purity and values ranging from 0 to 0.995 mg/mL, as indicated in Table 1 below. The standards demonstrated a linear correlation between glucose concentration and absorbance at 490 nm, as seen in Figure 3. The blank showed no absorbance; nevertheless, when glucose concentrations increased, the absorbance values correspondingly increased. The calibration curve was subsequently used to quantify carbohydrates in HH expressed as glucose equivalents.

Table 2 shows that untreated hemp hurds (HHs) had the highest mean absorbance (0.513), which meant that the hydrolysate had a glucose-equivalent concentration of 0.56 mg/mL and a total carbohydrate content of 28 mg. The absorbance and glucose-equivalent concentrations of pretreated samples were lower. NaOH-treated HH had 24.3 mg, H₂O₂/NH₃-treated HH had 22.8 mg, and EtOH-treated HH had 14.5 mg of total carbohydrates. The glucose control had an average absorbance of 0.912, which was close to its purity-corrected concentration of 0.995 mg/mL. This showed that the calibration and analytical method were correct.

When compared to the original sample mass of 50 mg (Table 3), the untreated HH had 56.0% polysaccharide content, which went down after pretreatment. The NaOH treatment kept the most polysaccharides (48.5%), followed by H₂O₂/NH₃ (45.5%). The EtOH treatment, on the other hand, kept the least polysaccharides (28.30%). These results show that alkaline pretreatment had the highest effect in modifying the LB matrix and keeping a lot of cellulose, while the organosolvent treatment had the least effect on how easy it was to get to carbohydrates.

The reduction in polysaccharide content is attributed to the combination of solubilisation of hemicellulose, degradation of amorphous carbohydrates, and loss of sample during treatment wash. The NaOH-treated HH had the highest remaining carbohydrate content of 48.5%. This outcome demonstrates successful lignin removal while maintaining a high polysaccharide content (cellulose). The second most effective method was chemical oxidation using H₂O₂/NH₃, resulting in a carbohydrate content of 45.5%, which demonstrated the ability to remove the lignin barrier, albeit less effectively than the NaOH. In contrast, the EtOH treatment had a lower impact, accounting for only 28.30% of total carbohydrates, indicating that this organosolvent method was the least effective in removing lignin, resulting in a substantial fraction of cellulose and hemicellulose remaining protected.

In the absence of HPLC analysis, the origin of carbohydrate loss cannot be further resolved. Solid recovery yields were not quantified; compositional data refer only to the purity of the remaining solid fraction, not the efficiency of the recovery.

3.2. Effect of Pretreatment Crystallinity Structure and Index Using XRD

XRD is used to determine the degree of crystallinity in semi-crystalline substances, such as HH, which have both crystalline (cellulose) and amorphous (hemicellulose) structures [ref. 25,ref. 26]. The crystalline index (CrI) was calculated according to the Segal method. The Segal method was adapted to compute a confidence interval for cellulose II, using I_am = 16° 2θ, as a representative of amorphous structure and I₀₀₂ = 22° 2θ, representing crystalline structure. The Segal CrI is a popular tool for figuring out how CrI changes after physicochemical or biological treatments of cellulose materials because it is user-friendly. It is important to note that the Segal method was designed as a “time-saving empirical measure of relative crystallinity values” and therefore does not provide any additional information about the cellulose’s properties, such as paracrystallinity, which is a distorted or loose layered structure found on the surface of crystallites [ref. 27]. The graph in Figure 4 displays crystalline regions indicated by sharp peaks and broad “humps” indicate amorphous areas, which gives information to calculating Crl [ref. 28].

The diffraction patterns show characteristic cellulose I reflections at approximately 15–16° 2θ, associated with the ($\overline{11}0$)/(110) planes, and a noticeable peak at around 22° 2θ, linked to the (200) plane. The broad halo centred at around 18° 2θ is associated with amorphous cellulose, hemicellulose, and lignin. The increase in Crl following pretreatment is attributed to the removal of amorphous parts.

The graph illustrates that each sample exhibits a unique central crystalline peak, highlighting the impact of different treatments on the crystallinity of HH. The baseline sample, represented by untreated HH, shows a broad, low-intensity peak at approximately 22°, indicating a significant presence of amorphous hemicellulose and lignin, which results in a crystallinity index of 39.26% (Figure 5). EtOH-treated sample (light blue) shows a marked increase in the intensity of the 22° peak compared to the untreated sample, with a crystallinity index of 55.00% indicating the removal of some amorphous components and a relative increase in crystalline cellulose. The sample treated with H₂O₂/NH₃ exhibits a strengthened and sharper peak at 22°, with a crystallinity index of 58.29%, indicating superior removal efficacy of lignin and hemicellulose compared to EtOH treatment. In contrast, the NaOH-treated sample exhibits the tallest and sharpest peak at 22°, with a crystallinity index of 65.80%, indicating the highest index. This treatment yields a higher quantity of stable crystalline cellulose. The increase in CrI indicates the removal of amorphous structure, leading to a more resistant crystalline residue that may necessitate more severe conditions for subsequent treatment.

3.3. Analysis of Chemical Structure Post-Treatment Using FTIR

The FTIR spectra in Figure 6 are a spectral representation of untreated HH and treated HH from various pretreatment methods.

Key infrared absorption peaks in cellulose are identified as follows: 3337 cm⁻¹ indicates –OH group stretching, common in polysaccharides as well as residual moisture [ref. 29]. Hemicellulose is a branching polysaccharide consisting of sugars such as galactose, glucose, mannose, and xylose. While it shares characteristics with cellulose, there are distinct spectral differences. The peak at 2583 cm⁻¹ is linked to the stretching vibrations of C–H bonds in alkyl groups, commonly seen in lignin within LB. It reflects the asymmetric stretching of –CH₃ and –CH₂– groups, indicating lipids and aliphatic structures [ref. 30]. The 2945 cm⁻¹ peak is associated with lignin [ref. 31], while peaks in the 2920–2932 cm⁻¹ range relate to C–H stretching in both lignin and cellulose [ref. 32,ref. 33]. The peak’s intensity and presence vary with biomass treatment or degradation [ref. 34].

A peak at approximately 1734 cm⁻¹ indicates the stretching of the C=O bond in acetyl and ester groups, distinguishing hemicellulose from cellulose [ref. 35]. The C–O stretching of the acetyl group is observed at approximately 1245 cm⁻¹ [ref. 36]. Lignin, an aromatic polymer, differs from polysaccharides, exhibiting distinct peaks due to its aromatic rings. Key peaks are 1510 cm⁻¹ and 1245 cm⁻¹, with the 1510 cm⁻¹ peak being a unique indicator of lignin.

For HH, the peak 3337 cm⁻¹ is broad, whereas for NaOH, H₂O₂/NH_3, and EtOH, the peaks are less broad and sharper, indicating cellulose hydroxyl group exposure. The aliphatic C-H stretching at 2850 cm⁻¹ was observed in all samples, associated with polysaccharides. There was not much difference among all treatment methods. The C=O peak at 1734 cm⁻¹, representing hemicellulose, is visible in HH and slightly reduced in EtOH. On the contrary, with NaOH and H₂O₂/NH₃ treatments, the peak is below FTIR detection limits, and the total polysaccharides are likely to include residual hemicellulose not detected by surface-sensitive FTIR. The aromatic skeletal vibration at 1510 cm⁻¹ is visible in HH, slightly reduced with EtOH, and quite reduced with NaOH and H₂O₂/NH₃ treated samples.

Peaks at 1245 cm⁻¹ weaken in treated samples, indicating effective lignin removal. The NaOH treatment is the most effective, showing decreased peaks representing both hemicellulose and lignin, as defined in Table 4.

3.4. Influence of Pretreatment on Thermal Behaviour Using TGA

Thermogravimetric analysis (TGA) and derivative thermogravimetric (DTG) analyses, presented in Figure 7a,b, respectively, were used to determine the thermal stability and decomposition behaviour of both untreated and treated HH. Figure 7a shows the thermogravimetric analysis (TGA) profiles of untreated HH and those that have been treated using various methods. Figure 7b shows the corresponding derivative thermogravimetric (DTG) curves. In Figure 7a, all samples show a slight loss of mass at temperatures below about 120 °C. This is because moisture and volatile components with low molecular weight were removed. This dehydration stage is only shown in Figure 7b as a small, broad DTG feature, which shows that there is not much chemical degradation at low temperatures.

The major mass-loss region illustrated in Figure 7a occurs between approximately 250 and 400 °C, corresponding with the thermal decomposition of hemicellulose and cellulose, with simultaneous contributions from lignin. This is due to hemicellulose having a lower degree of polymerisation with its amorphous structure, as compared to cellulose, making it less thermally stable and prone to degradation [ref. 37].

Figure 7b shows this area separated down into clear DTG peaks that show the highest rates of mass loss. HHs show a broader DTG peak at a reduced decomposition temperature, indicating a heterogeneous lignocellulosic matrix wherein hemicellulose and amorphous cellulose degrade simultaneously. This behaviour matches the TGA curve’s more gradual mass-loss profile. Lignin degradation is slower over a broad temperature range due its thermal stability. It is responsible for the trail of the curve above 400 °C and the formation of residual char [ref. 28,ref. 38].

Pretreated samples show sharper DTG peaks in Figure 7b, demonstrating changes in structural organisation following pretreatment. The NaOH-treated sample shows a pronounced DTG peak. The noticeable DTG peak in NaOH-treated samples indicates enhanced compositional uniformity due to the removal of hemicellulose and lignin. The temperature at maximum decomposition rate (T_max) showed a modest shift to higher temperatures for pretreated samples, particularly for NaOH-treated hemp hurds, indicating improved thermal stability following lignin and hemicellulose removal.

The thermal behaviour is related to the crystallinity indices observed in XRD. NaOH and H₂O₂-/NH₃-treated samples show enhanced thermal stability, evidenced by delayed major weight loss. This is attributed to the increased proportion of cellulose with its crystallinity. The narrowing and temperature shift in DTG peaks are related to the increased surface accessibility seen in SEM images and the higher crystallinity indices found in XRD analysis. These results show that ultrasound-assisted NaOH pretreatment modifies the surface the most, followed by H₂O₂/NH₃, and that EtOH pretreatment is the least effective. Consequently, TGA and DTG analyses not only confirm but also reinforce the comparative conclusion regarding the effectiveness of each pretreatment method in modifying the surface properties of hemp hurds, as seen in Table 5.

3.5. Morphological Changes Caused by Pretreatment Using SEM

The SEM analyses demonstrate the surface morphology of HH after various treatments. The images show a noticeable difference in surface morphology and fibre exposure at 2000× magnification. The untreated HH illustrated in Figure 8a have a compact and smooth surface resulting from a lignin and hemicellulose matrix coverage. Figure 8b shows HHs treated with EtOH, which have minimal surface disruption and a moderately rougher surface that maintains the integrity of the fibre bundles. In contrast, the HHs shown in Figure 8c, post-treatment with H₂O₂/NH₃, have increased surface roughness and disruption of the compact fibre structure, indicating physical biomass change post-treatment. The HH image shown in Figure 8d, post-treatment with NaOH, demonstrates a similar pattern with H₂O₂/NH_3, but with more increased surface roughness and disruption. This is another observable indication of changes in the surface of HH post-treatment. Morphological changes serve as a reference for comparison research aimed at evaluating the patterns or behaviours of HH after different pretreatment methods. In accordance with XRD and FTIR analyses, NaOH-treated HH exhibited the most disrupted surface, indicating a substantial modification in the physical structure of HH.

This confirms that HHs subjected to NaOH treatment represent the optimal pretreatment method and this can be confirmed by the comparison data in Table 6. This provides access to cellulose, which is required for the bioconversion of HH.

4. Conclusions

The study investigated the following three pretreatment methods: organo-solvent, oxidative-alkaline, and sonication. The three methods included 60% ethanol, 3% hydrogen peroxide 3% ammonia, and 2% sodium hydroxide. Post-treatment, the hemp hurds were subjected to various techniques and analyses to determine the structural modifications and understand their thermal behaviour under different temperatures.

Under the specific experimental conditions investigated, NaOH-based pretreatment resulted in more identified changes in polysaccharide content of 48.5%, crystallinity index of 65.80%, thermal behaviour, and most disrupted surface morphology, compared to the other pretreatment routes evaluated. These observations show that there was modification on the HH matrix rather than selective removal of individual biomass components. The observed changes are a result of the combined physicochemical environment; the specific contribution of the NaOH versus the ultrasonic cavitation cannot be isolated.

To determine the practical relevance of the observed structural and compositional modifications, further research will concentrate on the assessment of downstream performance measures, particularly enzymatic hydrolysis efficiency, under controlled conditions. The purpose of these investigations is to find links between changes in biomass caused by pretreatment and changes in functional bioconversion results.

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