1 Introduction

Hemicelluloses are heteropolysaccharides with very complex molecular structures consisting of acetylated or methylated heteropolymers of mannose, glucose arabinose, xylose, and galactose [1]. It is the second largest component of lignocellulose—the most abundant biomass in nature. They are mostly classified based on the primary monosaccharide in their overall backbone. Hemicelluloses are found at the interface between cellulose and lignin [1]. Certain properties of hemicelluloses are closely linked to their structure and chemical composition. Hemicellulose extraction methods that alter the natural structure of the polymer will decrease its industrial relevance. The structure of hemicellulose influences its mechanical characteristics, rate of water absorption, swelling ratio and ability to retain and slowly release drugs in a dose-dependent manner [1, 2]. For example, acetylation of hemicelluloses improves their physical strength and thermal stability, while deacetylation makes the polymer more prone to hydrolysis and degradation [3].

Hemicelluloses are becoming increasingly popular in the production of xylooligosaccharides which serves as prebiotics and supports the growth of healthy gut bacteria [4], production of biopolymers such as hydrogels [1] and bio-composites [2], for use as carriers in drug delivery and tissue engineering, in the production of enzymes for bio-bleaching of pulp [5] deinking of waste paper [6], enhancing the digestibility of animal feed [7] and dough quality [8], and in biofuel production [9].

Biodegradation is an important characteristic of biopolymers and one of the major advantages biomaterials have over conventional polymers, although several attempts are being made to produce biodegradable synthetic polymers [10]. It refers to the chemical decomposition that occurs via enzymatic cleavage of bonds within the polymer by microbial action, yielding low molecular weight compounds and non-toxic environmentally friendly byproducts [11, 12]. The structural and chemical properties of a biopolymer have a direct impact on its biodegradability. And this informs its use in various biotechnological processes such as in food and packaging, production of medical implants and more efficient tailored drug delivery systems in biomedicine and tissue engineering processes [13].

Biodegradation of hemicelluloses requires the synergic and concerted action of a group of enzymes classified as glycoside hydrolases which act cooperatively due to the heterogeneity of the polymer [14, 15]. This group of enzymes include endo-1,4-β-D- glycoside hydrolases which randomly cleave the hemicellulose backbone, β-D-glycosidases which cleaves monomers from the non-reducing end of the oligosaccharides [14, 15]. The removal of the side groups is catalyzed by α-L-arabinofuranosidases (EC 3.2.1.55), α-D-glucuronidases (EC 3.2.1.139), acetyl esterases (EC 3.1.1.72) and ferulic acid esterases (EC 3.1.1.73). These enzymes are widespread among fungi, actinomycetes and bacteria. Some of the most important xylanolytic enzyme producers include the Aspergilli, Trichodermi, Streptomycetes, Phanerochaetes, Chytridiomycetes, Ruminococci, Fibrobacteres, Clostridia and Bacilli [14, 16].

Biodegradable polymers can be divided into two major groups: Biodegradable polymers of native/natural origin and biodegradable polymers of synthetic origin [17, 18]. Native biodegradable polymers such as polysaccharides, lipids and proteins are synthesized naturally over several years of evolution to meet specific needs in nature [12]. Synthetic biopolymers on the other hand refer to man-made copies of these polymers synthesized via chemical routes. Examples include polylactic acid (PLA) and poly(ϵ-caprolactone) (PCL). However, not all native polymers are biodegradable, and not all synthetic polymers are non-biodegradable for example, PCL and PLA are synthetic but biodegradable, whereas fossil-based polyethylene (PE) although bio-based, is not biodegradable [17].

The number of hydrolysable linkages present on the polymer that are susceptible to microbial or enzymatic cleavage and how tightly packed their arrangement is within the biopolymer affects the biodegradability of the polymer [18]. Tightly packed polymers may conceal catalytic groups, which makes it resistant to biodegradation. Water is necessary for the initiation of biodegradation and stabilization of the transition state as the H+ and OH− of water are essential for bond breakage and product formation during enzymatic hydrolysis [19]. The ability of the biopolymer to absorb and interact more freely with water molecules will enhance its biodegradability [20].

Resistance to biodegradation depends on the crystallinity, hydrophobicity, and the spatial arrangements on the atoms of the side groups that create a steric effect in the polymer backbone [21, 22]. Linear polysaccharides with high regular intermolecular interactions form largely insoluble crystalline structures. One major mechanism by which the presence of branch points on the polysaccharide chain enhances biodegradability is by weakening steric-induced intramolecular interactions. Hydrophobic environments can also be created by the presence of hydrophobic groups like O-Me and O-Ac on the biopolymer [20]. Polysaccharides containing the same type of monosaccharides held together by different linkages will respond differently to enzymatic degradation. For example, β-glucan will respond differently to enzymatic degradation than cellulose because one or more β-1,3-linkages are inserted into the β-1,4- chain. [17, 23].

Other current applications of these biopolymers that involve temporary biomedical systems cannot directly utilize microbes. Microorganisms employ the use of the extracellular enzymes secreted to efficiently degrade biopolymers; such enzymes can be exploited for use in the biodegradation of biopolymers [24]. These extracellular enzymes can be incorporated into the synthesis/assembly of biomaterials in concentrations sufficient for a time-bound biodegradation which must be based on established in vitro experiments [24]. So far, many of the polymers used are made via synthetic crosslinking and the native bonds which these hydrolases attack is either unavailable or tightly embedded within the synthesized polymer [25].

Certain parameters can be used to monitor the rate of degradation of biopolymers. Gradual diffusion of water molecules into the biopolymer marks the first stage of degradation which is followed by chemical hydrolysis of the molecule. These chemical changes can be monitored using spectroscopic analysis (IR, X-ray). However, to analyze later stages of degradation, the most suitable method depends on the unique properties of the biomaterial being analyzed. Certain characteristics when monitored can give an idea on the degradation properties of the biopolymer, including the degree of water absorption, weight loss measurements, molecular weight analysis, crystallinity, contact angle measurements before and after degradation and monitoring the degradation products released in the solution [26].

In this study, the enzymatic degradation of extracted hemicellulose (EH) from ARW and AW by hemicellulase from Bacillus trypoxylicola was monitored. The enzymatic biodegradation of the extracted hemicellulose was monitored by measuring the concentration of reducing sugars released into the incubation medium over time.

2 Methodology

2.1 Chemicals

All chemicals used in the present study were of analytical grade from Sigma-Aldrich, Qualikems and BDK.

2.2 Alkaline-extraction and characterization of hemicelluloses

Previously extracted and characterized hemicellulose from Beech, African rose and Agba woods were used for this study [27]. Milled lignocellulosic biomass was soaked in 10 % NaOH (1:10), incubated overnight in a shaker (120 rpm at 60 °C) and boiled for 3 h. The supernatant was recovered by filtration. Solubilized hemicellulose was precipitated in 1:2 volume of ethanol (99.8 %), decantated and oven-dried at 60 °C. the extract was weighed and stored in air-tight containers at room temperature.

Percentage hemicellulose yield was calculated as:

$$Percentage\, yield \left(\% w/w\right)=\frac{dry\, weight\, of \,extracted\, xylan (g)}{dry \,weight\, of\, sample (g)} X 100$$

2.2.1 FTIR analysis

The isolated hemicelluloses were subjected to FTIR analysis at room temperature over a spectra wavelength of 4000 to 500 cm-1 at 4 cm-1 resolution and an interval of 1 cm−1 [27].

2.2.2 HPLC analysis

EH samples were hydrolyzed under acidic conditions. 0.5 g of EH was dissolved in 1 mL of water. H2SO4 (1 mL of 98 % v/v) was added to an aliquot of sample (500 μL) with mixing at 80 °C for 1 h. To 100 μL aliquots 1 mL of distilled water was added. The pH was adjusted to 5–6 with calcium carbonate and the hydrolysate filtered and analyzed (10 μL) using HPLC (Shimadzu) with distilled water as mobile phase at a flow rate of 0.4 mL/min and a refractive index detector at 50 °C. Quantification was referenced to glucose, galactose, mannose, xylose, and arabinose standards [27].

2.3 Isolation of hemicellulase-producing organism

One gram each of soil samples were obtained from three points at the local timber-processing site with observed wood decay was suspended in 9 mL of sterile distilled water and then diluted tenfold. From each dilution, an aliquot (0.1 mL) of the diluted mixture was inoculated onto plates containing sterilized basal salt medium (pH 9.0) charged with 0.2 % w/v NaNO3, 0.1 % w/v KH2PO4, 0.05 % w/v MgSO4.7H2O, 0.05 % w/v KCl, 0.2 % w/v protease peptone, 2 % w/v agar-agar and 1 % w/v extracted hemicellulose (EH). The plates were incubated at 37 °C for 48 hr. All morphological contrasting colonies were purified by repeated streaking and sub-culturing on separate plates until pure isolates were obtained [28].

2.4 Screening for enzyme production

Plates containing isolates were flooded with 1 % Congo red dye for 1 h, destained with 1 M NaCl and the zone of hydrolysis around the colonies recorded. The selected Isolates based on their zone of clearance were further cultured onto a sterile liquid basal saline medium charged with 0.2 % w/v NaNO3, 0.1 % w/v KH2PO4, 0.05 % w/v MgSO4.7H2O, 0.05 % w/v KCl, 0.2 % w/v protease peptone and 1 % w/w EH in several 100 mL Erlenmeyer flasks and incubated at 37 °C with constant shaking (120 rpm) for 7 days. The fermentation broth was harvested each day and centrifuged at 8,000 rpm for 10 minutes to remove the cells and assayed for enzyme activity.

2.5 Standardization of cell suspensions

Bacterial colonies were transferred aseptically into a test tube containing 5 mL of sterile inoculum broth. The mixture was vortexed to obtain a uniform consistency. The density of the bacterial cell suspension was compared to 1.0 McFarland turbidity standard (containing 0.1 mL of 1% w/v BaCl2 and 9.9 mL of 1% v/v H2SO4) by holding the bacterial suspension and the standard in a well lightened area against a Wickerham card (Hardy Diagnostics). The turbidity of the bacterial cell suspension was adjusted to 1.0 Mcfarland turbidity standard containing a microbial suspension of approximately \(3.0\times {10}^{8}\text{ CFU}/\text{mL}\) by the addition of either bacterial colonies or sterile inoculum broth under aseptic conditions [29]. The standard was tightly sealed and stored in the dark.

2.6 Molecular identification of selected isolate

Extraction of genomic DNA was carried out by strictly adhering to the AccuPrep® Genomic DNA extraction kit (K-3032, Bioneer, Republic of Korea) user’s guide. Microbial cells were gently collected from 3-day old plates and placed in a microtube. The PCR amplification of the 16S r-RNA gene was carried out using the primers forward (5'-AGAGTTTGATCCTGGCTCAG-3') and reverse (5'-GCGCTTTTT GAGATTCGCTC-3'). The PCR containing 15 µL of nuclease free water, corresponding PCR primers (1 µL each of forward and reverse), and 3 µL of DNA was carried out under the following conditions: initial denaturation at 95 °C for 5 min, 25 cycles of denaturation at 94 °C for 1 min, annealing at 52 °C for 1 min, extension at 72 °C for 7 min, and final extension at 72 °C for 7 min [30]. Homology search for the sequence was carried out using the Basic Local Alignment Search Tool (BLAST) program of the National Centre of Biotechnology Information (NCBI) as described by [31].

2.7 Enzymatic degradation of hemicellulose

2.7.1 Preparation of incubation medium

A total of twenty 250 mL Erlenmeyer flasks containing 100 mL each of sterile cultivation medium charged with 0.2 % w/v NaNO3, 0.1 % w/v KH2PO4, 0.05 % w/v MgSO4.7H2O, 0.05 % w/v KCl, 0.2 % w/v protease peptone and 1 % w/v extracted hemicellulose (EH) were stoppered and autoclaved at 121 °C (15 psi) for 20 min. The flasks were inoculated with 1 % standardized cell suspensions and incubated at room temperature under shaking at 150 rpm for 7 days [32].

2.7.2 Assay for enzyme activity

Enzyme activity was assayed by measuring the amount of reducing sugar liberated from the extracted hemicellulose (EH) using the dinitrosalicyclic acid (DNS) method [33], with the following modifications. An aliquot, 0.1 mL of 1 % w/v xylan was mixed with 0.4 mL of 0.1 M sodium acetate buffer, pH 5.0, and 0.5 mL of enzyme. The reaction mixture was incubated at 50 °C for 50 min, after which the reaction was stopped by adding 2 mL of DNS reagent and heated in boiling water for 10 min for color development [34]. The mixtures were cooled at room temperature and the optical density was measured at 550 nm using 721-vis spectrophotometer. One unit (U) of enzyme activity was defined as the amount of enzyme that released 1μmole of xylose per minute under assay conditions. Protein concentration was determined by Lowry method [35] using (2mg/mL) bovine serum albumin (BSA) as a standard for the calibration curve.

Enzyme activity was calculated as:

$${\text{Units}}\,{ = }\,\frac{{\mu \,{\text{mol}}\,{\text{of}}\,{\text{xylose}}\, \times \,{\text{reaction}}\,{\text{volume}}}}{{{\text{Sample}}\,{\text{volume}}\, \times \,{\text{reaction}}\,{\text{time}}}}$$

2.7.3 Release of reducing sugars

The concentration of reducing sugars released upon hydrolysis was monitored by harvesting the fermentation broth each day over the 7-day period. The optical density of the solution was obtained for each day and the concentration of the reducing sugar released was measured using the dinitrosalicyclic acid (DNS) method as described above (section 2.7.2). The concentration of reducing sugars released xylose equivalent under assay conditions via extrapolations from xylose standard curve.

$$\text{Concentration of reducing sugar }\left(\text{mM}\right)=\frac{\text{Absorbance of test}}{\text{equivalent concentration}}$$

2.8 Statistical analysis

Data obtained were analyzed using SPSS version 25. Independent student t-test was used for analysis and means were separated using Leven’s test for equality of variance. Significance was accepted at p-value ≤ 0.05

3 Results

3.1 Quantitative screening of microbial isolates for enzyme production

Two isolates, one collected from point one, S14, and another from point three, S38, at the timber-processing site, may have evolved the ability for more efficient enzyme production. Screening of these isolates, S14 and S38 yielded enzyme activities of 269.04 U and 222.66 U, respectively (Fig. 1). This guided our choice of the isolate, S14 for the present study.

Fig. 1
figure 1

Quantitative screening of microbial isolates for enzyme production

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3.2 Molecular identification

The microbial isolate appeared white, round, and granular. Agarose gel electrophoresis of the 16S r-DNA amplicon showed a band at approximately 800 bp. A phylogenic tree constructed using the neighbor-joining method showed the location of the isolate among its phylogenic neighbors. The phylogenic tree showed one major lineage clustered among other Bacillus trypoxylicola stains in one clade. The morphology, physiology, and molecular data of the isolate showed 99% similarity to Bacillus trypoxylicola strains with only 1% genetic variation (Fig. 2).

Fig. 2
figure 2

Molecular Identification of Bacillus trypoxylicola, containing the pure culture, agarose gel electrophoresis and Phylogenetic tree (observed genetic variation 0.01 = 1%)

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3.3 Alkaline extraction and characterization of hemicelluloses

The percentage yield of hemicellulose extraction was 45.5% for African rose wood and 36.7% for Agba wood. Absorption band between 4000 cm−1 to 1500 cm−1 (the functional group region) indicated the presence of non-hydrogen bonded O–H stretching, normal polymeric OH-stretching, C–H2 stretching, C–H stretching, C–O stretching, C=O stretching, and C–H deformation in the EH. The FTIR analysis of EH from ARW indicated absorption bands at 1161.1, 930.8 and 923.7 cm−1 at the fingerprinting region which indicates an asymmetric β-1,3-glycosidic stretch, β-1,4-glycosidic stretch and an asymmetric β-1,6-glycosidic stretch, respectively. Agba wood EH showed absorption bands of 1265.39 and 923.75 cm−1 at the fingerprinting region corresponding to the absorption of β-1,3-glycosidic and β-1,4-glycosidic C–O–C stretch, respectively (Table 1). The HPLC analysis indicated that the hemicellulose extracted from ARW was a mannan with 96% mannose, and AW was a galactoglucomannan consisting of 20.82% glucose, 14% mannose, and 9.22% w/w galactose (Table 2).

Table 1 FTIR analysis of ARW and AW
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Table 2 Identified monosaccharides present in EH from ARW and AW
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3.4 Enzyme activity on EH from African rose wood

There was a rapid increase in the specific activity of hemicellulase from B. trypoxylicola on EH from ARW between D0 to D2 with mean values of 66.81 ± 2.27 and 694.44 ± 1.61 U/mg, respectively, which plateaued out and slightly decreased on D7 (668.95 ± 14.12) (Fig. 3, Table 3). Generally, the means of the specific activity of the enzyme on EH from ARW was significantly higher than that of AW (Fig. 4), from D0 to D7 with P-values ≤ 0.05 (Table 3).

Fig. 3
figure 3

Hemicellulase activity on ARW and AW biopolymers

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Table 3 Statistical means for enzyme activity analyzed using student t-test (2 significant digits) for both EH from ARW and AW
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Fig. 4
figure 4

Concentration of reducing sugar (RS) released upon enzymatic hydrolysis of ARW and AW biopolymers

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3.5 Enzyme activity on EH from Agba wood

A rapid rise in enzyme activity was also observed between D0 and D2 with mean values of 22.80 ± 0.76 and 355.07 ± 3.76 U/mg, respectively. This increase slowed down but was maintained on D3 (434.09 ± 8.59 U/mg), attained its peak on D4 (493.03 ± 6.92 U/mg), and began to gradually decline from D5 (443.72 ± 10.94 U/mg) to D7 (387.36 ± 25.65 U/mg) (Fig. 3, Table 3).

3.6 Release of reducing sugar from ARW hemicellulose

Initially, the concentration of RS released from ARW increased rapidly from 0.21 ± 0.01 mM to 2.53 ± 0.01 mM from D0 to D2 and obtained its peak of 2.92 ± 0.08 mM on D4. A gradual decrease in the concentration of the reducing sugar released into the broth was observed from D3 (2.65 ± 0.06 mM) to D7 (2.18 ± 0.03 mM) (Fig. 4, Table 4).

Table 4 Statistical means for reducing sugar release analyzed using student t-test (2 significant digits) for both EH from ARW and AW
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3.7 Release of reducing sugar from AW hemicellulose

A rapid increase in the release of reducing sugars from AW EH into the fermentation broth from D0 (0.16 ± 0.01 mM) to D2 (3.46 ± 0.04 mM) which obtained its peak on D3 (3.56 ± 0.07 mM). This was followed by a decline in reducing sugar release from 3.54 ± 0.05 mM (D4) to 2.51 ± 1.15 mM (D7 (Fig. 4, Table 4). The reducing sugar release from AW hemicellulose was generally significantly higher than that of ARW hemicellulose except for D7 which had a p-value of 0.185.

4 Discussion

The ability of an organism to utilize certain substrates influences its production of the corresponding hydrolytic enzymes [15, 36]. Since the isolates were collected from soil at wood-decomposing sites, it was inferred that such organisms should have acquired the capacity to utilize wood components to survive in such an environment [36, 37]. Microorganisms in their natural habitat develop specific adaptations to survive. Such organisms evolve mechanisms through which they can degrade the carbon sources and nutrients available in that environment for their metabolism, survival, and growth. Soil microbes from timber-processing sites may have evolved mechanisms needed for wood degradation such as secretion of extracellular hemicellulases for the complete hydrolysis of the hemicellulose component of wood. Hemicellulase production is dependent on two main factors: the organism as well as the agricultural waste [36]. The variations in enzyme production with different carbon sources may be attributed to the individual compositions of the heterogeneous polysaccharide. Since hemicelluloses are heteropolymers of five major sugars; xylose, arabinose, glucose, mannose and galactose, the presence and concentrations of these sugars may to a large extent influence enzyme production by the organism which may be inhibited by the presence of simple sugars in the fermentation medium because simple sugars are readily available energy sources [37].

To perform phylogenetic analysis of prokaryotes, short nucleotide sequences, usually the 16s rRNA gene sequences are used. The amplification and analysis of this gene provides evolutionary relationships in bacteria due to their ubiquitous presence in bacteria and the slow evolution rates of these gene sequences, their genetic information is highly conserved [38]. The challenge with using conserved genes like trpA, trpB, and pabB for the identification of Shigella, Salmonella and E. coli is that they are non-ubiquitous among bacterial species. This means that they could have undergone certain evolutionary changes, hence, unreliable. The 16s rRNA was ideal for identifying the isolated soil bacteria in the present study as it is highly conserved across all bacterial species. The phylogenic tree indicates that isolate was a Bacillus strain with 99 % sequence similarity to Bacillus trypoxylicola and only 1 % genetic variation. Phylogenic trees are most used to show the relationship between bacterial genomes. The phylogenic tree in the present study diverged into one major lineage clustered among several closely related Bacillus trypoxylicola strains all sharing a common ancestor. This indicates that the 16s sequence of the isolate was highly conserved with minimal variation in the gene.

The FTIR functional group analysis of EH from both ARW and AW which indicated the presence of several hydroxyl groups of polysaccharides also accounted for the solubility of EH water. Soluble biopolymers indicate hydrophilicity [23]. Biopolymers that easily absorb water/moisture are more prone to enzymatic hydrolysis because the H+ and OH- of water molecules interact with the bond on the biopolymer encouraging transition state formation and stabilization (Figure 3). Aromatic rings absorb IR. This is indicated by the absorption bands of 1592 and 1525 cm-1 in EH from ARW. This suggests the presence of residual lignin on EH from ARW. The presence of the absorption bands at 1734 and 1644.93 cm-1 in the IR spectra of ARW and AW, respectively, which indicates the presence of a C=O stretching. This suggests that the acetylation. Acetylation of hemicellulose increases hydrophobicity of the molecule and creates a resistance to water [39]. The C–H stretch of aldehydes suggests the presence of free aldehyde groups at the end of the heteropolymer. Hydrolysis of glycosidic bonds, and hydroxyl groups show characteristic IR absorption bands at 1720 cm-1, the absence of this band indicates that there was minimal attack on the glycosidic linkages during extraction.

FTIR fingerprinting region shows specific IR absorption patterns for glycosidic bonds mainly for glucans and mannans. They include IR absorption bands at 825, 855, 930, 923, 1161 cm-1 located between 1200 cm-1 to 800 cm-1 [40, 41]. The EH from ARW showed IR absorption bands at 1161.1, 930.8 and 923.7 cm-1 indicating an asymmetric β-1,3-, β-1,4-glycosidic stretch and an asymmetric β-1,6-glycosidic stretch, respectively. A major IR band was observed for EH from AW at 923.75 cm-1 that corresponds to a β-1,4-glycosidic C–O–C stretch [42]. The asymmetric β-1,6-glycosidic stretch indicates the presence of branched points in EH from ARW. In polysaccharides, β-1,4- and β-1,3- glycosidic linkages primarily make up the linear backbones, although β-1,3- glycosidic linkages may sometimes appear at branched points [43].

From the HPLC analysis, EH from ARW comprised mainly of mannose (96 %). These mannan units linked via β-1,3-, β-1,4-glycosidic and β-1,6-glycosidic linkages may present as a straight-chain molecule with some branched points. The EH from AW is a primarily a straight chain galactoglucomannan consisting of 20.82 % glucose, 14 % mannose and 9.22 % galactose. The unidentified sugar (56.47 %) could either be a disaccharide or an oligosaccharide that may have occurred due to incomplete hydrolysis.

At the initial stage of the reaction, the hemicellulase cleaves the biopolymers randomly, more oligomers are available to bind the enzyme. This caused a rapid rise in enzyme activity between D0 and D2 for both EH from ARW and AW. Overtime, there was a saturation of the enzyme molecules by the substrate, hindering further increase in rate of the reaction [44]. Generally, the enzyme’s specific activity was higher on ARW than AW. Branched polysaccharides could be more resistant to enzymatic hydrolysis and may require more specific branched-points enzyme. The β-1,4-glycosidic linkages of mannans in the EH from both ARW and AW may be the primary target of the enzyme. Although it has been observed that mannans can be hydrolyzed by both mannoside and glucoside hydrolases [45]. Since the experimentation was carried out under the same environmental conditions of temperature, pH and substrate/enzyme concentrations, the difference in enzyme activity may be a result of the chemical and structural disparities of the biopolymer from ARW and AW. The rate of degradation of a biopolymer relies both on the enzyme as well as the chemical composition and structural organization of the biopolymer [46].

The concentration of reducing sugars released upon degradation initially increased until it got to its peak for both EH from ARW and AW, after which a decline was observed. The reducing sugar concentrations in the medium between D0 and D1 was originally higher for ARW than AW. The presence of β-1,6- branch points in EH from ARW initially made the polymer appear more susceptible to biodegradation. However, as the reaction progressed, the biodegradation of EH from AW seemed to be significantly higher than that of ARW. Generally, EH from ARW presented to be more crystalline in nature than that from AW because linear mannans are preserved by steric effects. Hence, the reducing sugar concentrations for EH from ARW were generally lower. The concentration of reducing sugar in the broth began to drop steadily after D3 for EH from AW and D4 for EH from ARW. The hemicellulose in the medium depleted because no additional substrate was introduced into the medium and the sugar released could have been incorporated into the organism’s metabolism, hence the steady decline in reducing sugar concentration. Although EH from AW consist of linear β-1,3-, β-1,4-glycosidic, the heterogeneity of the galactoglucomannan may have lowered the crystallinity of molecule by reducing the intramolecular interactions. This reduces the resistance that could have occurred due to steric hinderance. Thus, EH from AW demonstrated higher susceptibility to enzymatic biodegradation indicated by the generally higher values of reducing sugar concentrations in the broth than EH from ARW. Structural studies of galactomannans showed that the hydrogen molecule on the galactosyl side chain interacts with the mannan backbone in such a manner that encourages intramolecular associations that ensures stability of the structure [47, 48].

5 Conclusion

The growing applications of biopolymers require adequate understanding of their properties and behavioral responses to enzymatic/microbial biodegradation. From the present experiment we note that biodegradation is dependent on several factors such as the biomaterial; monomeric component and structural arrangement, enzyme activity and environmental factors which play important roles in the dynamics of biodegradation. Although ARW and AW are both hemicelluloses, each responded differently to enzymatic degradation by hemicellulase from B. trypoxylicola. The unique properties and characteristics of polymer must be studied along with its biodegradation parameters for each unique biotechnological application.

6 Future prospect

Enzymatic hydrolysis of biopolymers is a field that should be given attention to properly address biodegradation of biopolymers. Highly efficient microbial hydrolases are needed for controlled biodegradation of biopolymers. The specific interactions of these enzymes on a well characterized biopolymer can provide insights into polymer-specific enzyme interactions unique to each biopolymer. This precision is required for the construction of temporary biomedical systems that could self-degrade after the time required for tissue healing or drug delivery.