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BY-NC-ND 3.0 license Open Access Published by De Gruyter January 29, 2016

Lignin reinforcement of urea-crosslinked starch films for reduction of starch biodegradability to improve slow nitrogen release properties under natural aerobic soil condition

  • Zahid Majeed EMAIL logo , Nurlidia Mansor EMAIL logo , Zakaria Man and Samsuri Abd Wahid
From the journal e-Polymers


The urea-crosslinked starch (UcS) film has a major drawback of very rapid biodegradability when applied as slow release fertilizer in soil. Lignin reinforcement of the UcS was used to prepare composite films, aimed to reduce the starch biodegradability and slow the release of nitrogen in aerobic soil condition. Study results revealed that mineralization of the composite films was delayed from 6.40 to 13.58% more than UcS film. Inhibition of composite films mixing with soil, the Michaelis-Menten reaction rates for α-amylase were inhibited ~1.72–2.03 times whereas the Michaelis-Menten reaction rates for manganese peroxidase were increased ~1.07–1.41 times compared to UcS film. Saccharides–glucose, maltose and maltotriose demonstrated that their rates of formation (zero-order reaction) and depletion (first-order reaction); both were slowed more in aerobic soil which received the composite films. Increasing of lignin in composite films, the acid to aldehyde ratios of vanillyl and syringyl phenols of the lignin declined from 1.18 to 1.17 (~0.76%) and 1.59–1.56 (~1.78%), respectively. The diffusivity of nitrogen was effectively slowed 0.66–0.94 times by the lignin in composite films and showed a “Fickian diffusion” mechanism (release exponent n=0.095–0.143).

1 Introduction

Thus far, urea is well-known for nitrogen recovery lower than 50% by most cropping systems due to its major part being lost on farmland for various reasons like runoff, leaching and volatilization (1). These losses restrict the continuous supply of nitrogen to growing plants for building new biomass, which results in stunted plant growth. These factors not only lead to financial losses but also pose risks to environment through contamination of ground and surface water and emission of nitrous gases into the air (2). Currently, commercial production of slow or control release fertilizer (S/CRFs) depends on non-renewable petroleum derived non-biodegradable polymers, e.g. polyolefin (Meister™ Chisso-Asahi Fertilizer Corporation, Japan), polyalkenes (Nutricote®, Chisso-Asahi Fertilizer Corporation, Japan) and polyurethane-like resins (Haifa Chemicals Co. Ltd, Israel) (3). These synthetic polymers are the main environmental pollution concerns which have justified finding other alternate green solutions for the fertilizer industry. Therefore, the key motives of exploring biodegradable polymers from renewable resources have been envisaged for effective urea encapsulation and to prolong the supply of nitrogen to plants through delaying its fast release in soil (4). This way, plant nitrogen demand could possibly be catered for at optimum rates required by the crops at different growth stages to increase the yield under specific soil conditions.

Starch is a semicrystalline polymer which is derived from plants. It consist of glucose units joined together by glycosidic bonds. These glycosidic bonds are highly susceptible to the enzymatic hydrolysis under the natural environment and makes the starch a highly biodegradable polymer (5). Starch is the most widely utitlized biodegradable biopolymer for the coating of fertilizer, e.g. starch-g-poly (vinyl acetate) (6), starch-urea matrix (7). Green natural rubber-g-modified starch (8), starch-g-acrylic acid (9), starch-g-poly (l-lactide) (10) and starch films plasticized with urea (11). However, starch has a very high biodegradability which comes from the free hydroxyl groups. There were some studies reported on reducing the starch composite films biodegradability out of the context of slow release fertilizer by addition of other natural polymers. For example, composite films comprised of starch reinforced with cellulose, improved the tensile strength about 47%, and decreased the biodegradability as a result of its increased resistance to microbial attack and improvement in its half-life (5). The Michaelis-Menten reaction rates of amylase enzyme were reduced to 40%. In another work, Guohua et al. (12) reported that polyvinyl alcohol hindered the starch biodegradability in starch/PVA film. The replacement of starch in starch/PVA film with methylated-cornstarch has increased the water resistance and amplified the mechanical properties. This would have caused the low enzymatic, microbiological biodegradation in these films under soil burial conditions. Thus, starch requires more research particularly to achieve better biodegradability resistance properties to find its effective applicability towards S/CRFs.

Lignin has a filler role in starch films due to its highly complex aromatic heteropolymer network of phenylpropane units (13). The complex spatial network structure of lignin is formed from three phenylpropane units: guaiacyl, syringyl, and p-hydroxyphenyl (14). The β-O-4 are the most common chemical linkages in lignin which upon cleavage could increase the proportion of carboxylic acid relative to aldehyde. This leads to an increase in acid-to-aldehyde ratios of vanillyl and syringyl phenols upon biodegradation of lignin in soils (15). Syringyl and cinnamyl phenols of the lignin are more susceptible to biodegradation compared to vanillyl phenols, resulting in a decrease of syringyl/vanillyl (S/V) and cinnamyl/vanillyl (C/V) ratios (15). In this connection, an earlier work of Sarwono (16) reported urea-crosslinked starch (UcS) film reinforcement with 5–20% (wt/wt) lignin as a blending component enhanced the tensile properties and reduced the hydrophilicity. The soil burial degradation test found 75% of weight loss in UcS reinforced with 10% lignin film during 2 weeks of the burial soil test.

In this research, starch was functionally cross-linked with urea and transformed into composites with different lignin percentages in order to test reduction of its biodegradability and minimization of nitrogen release under a natural aerobic soil microcosm.

2 Experimental

2.1 Experimental design

Figure 1 shows the experimental scheme followed to conduct the experiment. Approximately, 10–15 composite samples of soil were collected at 0–20 cm depth from the top of the soil from a paddy field at Titi Gantung (4.36°N, 100.84°E), Perak, Malaysia. Soil samples were tested for their chemical and physical characteristics as had already been cited in our earlier research work (17). A soil microcosm was set-up at a laboratory scale according to Massardier-Nageotte et al. (18).

Figure 1: Experimental scheme.
Figure 1:

Experimental scheme.

The experiments were set-up in a randomized complete block design with two factors as the blocking criteria, namely percentage of lignin (5, 10, 15 and 20%) and incubation time (0, 1, 2, 4, 6, 10, 14, 21, 28, 42, 49, 62 days). Black colored wide mouth pots (width 8.5 cm×length 14 cm) were used to conduct soil incubation tests for film biodegradation. Such pots were filled with approximately 75–200 g soil (60% moisture). Each composite film was cut into small pieces, i.e. 1 cm×1 cm and mixed with soil at the rate of approximately 84.66 mg g-1 dry weight soil (8.46%).

2.2 Materials

Tapioca starch (Kapal ABC, Malaysia) was purchased from the local market, urea (46% total nitrogen) was received from Petronas Fertilizer [(Kedah) Sdn. Bhd., Malaysia]. Loamy sand soil was collected from Titi Gantung Bota paddy field, Perak, Malaysia. Alkaline kraft lignin (Indulin AT) was purchased from Sigma Aldrich (USA). All other chemicals of analytical grade (>99%) were used in this research.

2.3 Composite film synthesis

Composite films were synthesized under optimized conditions according to the conditions detailed in the work of Sarwono (16). Briefly, tapioca starch (5 g) was mixed with 100 ml of deionized water and heated at 80°C for 30 min. After this, urea (2 g) and disodium tetra-borate (0.45 g) was added. Then lignin (L) was added at 5, 10, 15 and 20% of initial tapioca starch weight. The reaction mixture was stirred for a further 3 h at 80°C. The solution of each composition was poured into square polypropylene plastic bowls (12 cm×12 cm) and dried at 40°C overnight in a vacuum oven to obtain composite films. Composite films were dried further after setting for 2 h at a post curing temperature of 120°C in an air drying oven to complete cross-linking reactions of urea with starch (Figure S1). Composite films of urea-crosslinked starch (UcS) was named as UcS5%L, UcS10%L, UcS15%L and UcS20%L. The UcS film which received no lignin served as an experimental control.

2.4 Analytical methods

2.4.1 Mineralization

Aerobic mineralization of composite films was assessed following the ASTM D 5988-03 standard method (19) according to the experimental set-up as shown in Figure S2. Briefly, the CO2 evolved was captured in a 1 m NaOH (Merck, Germany) solution, under closed bottle conditions maintained at 27–28°C and ~72% relative humidity. At a particular incubation interval, the amount of CO2 produced was determined by titration of 50 ml of 0.1 m NaOH with 0.5 m HCl. The CO2 absorbed in the alkaline solution was titrated to two end points at pH 8.3 and pH 3.7 detected by phenolphthalein (Merck, Germany) and methyl orange (Bendosen, Norway) indicators respectively. Each endpoint was cross validated precisely with a pH probe while titration was in progress. To the sample, one to two drops of the phenolphthalein indicator was added first and then titrated until the pink color disappeared at pH 8.3 according to Equations [1] and [2]. Then one to two drops of the methyl orange were added and the titration continued until the color disappeared due to the reaction as shown in the equation [3].

[1]CO2+NaOHNa2CO3+H2O [1]
[2]Na2CO3+HClNaHCO3+NaCl(pH8.3) [2]
[3]NaHCO3+HClCO2+NaCl+H2O(pH3.7) [3]

The mass of CO2 (g) was obtained after multiplying the volume of the titrant (L)×molarity of the standard acid×molecular mass of CO2. The CO2 concentration was fitted into the Equation [4] to calculate the percentage mineralization (20).

[4]%Mineralization=gCO2-gCO2bmf×%Cf1004412×100 [4]

The % mineralization is the percent of carbon molecules converted to CO2, gCO2 is the mass of CO2 evolved from soil amended with composite films, gCO2b is the mass of total evolved CO2 in blank soil, mf is the mass of composite film (g), %Cf is the percent carbon content of the composite film. The molecular weight (g/mol) of carbon dioxide is depicted as 44, the molecular weight of carbon atom is taken as 12 (g/mol).

In order to assess the mineralization kinetics in aerobic soil microcosm, the experimental data were fitted into a Hill model (21) according to Equation [5] using OriginPro software, version 9.0.0 (OriginLab Corporation, Northampton, MA, USA).

[5]y=ymax×tnkminn+tn [5]

Here, y is the percentage of mineralization at time t (days), ymax is the percentage of mineralization at infinite time, kmin is the half-life time (days) and n′ is the curve radius of the sigmoidal function.

2.4.2 Michaelis-Menten reaction rates

The effect of composite films on the reaction reates of α-amylase and manganese peroxidase (MnP) was tested in soil after 21 days of incubation. The Michaelis-Menten equation [6] was applied to measure maximum soil enzyme reaction velocity (Vmax) and the substrate affinity constant (Km).

[6]v=Vmax[S]Km+[S] [6]

where v is the enzyme reaction velocity (a function of enzyme concentration), [S] is the substrate concentration. The catalytic efficiency (kcat) of the enzyme was calculated by Vmax/Km.

Using the starch as a substrate, the activity of the α-amylase was measured by the release of glucose (Sigma Aldrich, USA) in a phosphate buffer [pH 6.9, 6.70 mm NaCl (Merck, Germany) as activator] at 30°C. Glucose was quantified by its reaction with dinitrosalicylic acid at 534 nm wavelength on a UV-visible spectrophotometer (Shimadzu, Japan) (22–24).

MnP assay is based on the oxidative coupling of 3-methyl-2-benzothiazolinone hydrazone (MBTH, Fluka, Switzerland) and 3,3-(dimethylamino) benzoic acid (Sigma Aldrich, USA) and the extinction coefficient value was taken 53,000 M-1 cm-1 according to Castillo et al. (25).

2.4.3 Activation energies

The activation energies were calculated by determining the enzyme reaction rates of soil α-amylase and MnP enzyme at three different temperatures, 20, 30 and 40°C. Enzyme reaction rates were fitted into the Arrhenius equation (26).

[7]k=A.e-EaRT [7]

where k, the reaction rate as a function of temperature; A, the collision frequency factor; Ea, the activation energy; R, the universal gas constant (8.314 J mol-1 K-1/mol.K); T, the temperature in Kelvin (K). After taking the natural log of Equation [7], transformed into linear expression as shown in Equation [8].

[8]lnk=(-EaR)(1T)+lnA [8]

Plotting lnk against 1/T (K), Ea was calculated directly from the slope (-Ea/R) of the linear regression.

2.4.4 Starch biodegradability

Total starch biodegradation within composite films was measured by the method of Fernandes et al. (27). Approximately, 1 g freeze dried soil was washed with 80% of 5 ml hot ethanol (Merck, Germany) for 30 s to remove soluble sugars. The 4 ml of distiled water and 5 ml of 52% cold perchloric acid (Merck, Germany) were added to the soil prior to extraction in ice cooled water bath for 1 h. Starch absorbance was monitored at 625 nm on a UV-visible spectrophotometer (Shimadzu, Japan) and the concentration was determined using a calibration curve of starch (0–200 μg/ml). Equation [9] was used to calculate the starch concentration in soil treated with different composite films at different incubation times.

[9]Starch(mgg-1soil dw)=[(Cs-Cb)×ai]×Dfsoil(g)dw×11000 [9]

Cs is starch concentration (μg) of soil added with films; Cb is starch concentration (μg) of blank soil; ai is the relative theoretical starch fraction in each film composition; Df is the dilution factor.

First-order reaction was applied to calculate the rate constant of starch biodegradability after fitting total starch concentration into Equation [10].

[10]Cs,t=Cs,0.e-kst [10]

Cs,0(mg) is the starch initial concentration; Cs,t(mg) is the starch concentration at a particular time t. ks (day-1) is the kinetic constant of starch biodegradability. The value of ks from Equation [10] was inserted into Equation [11] in order to calculate half-life (t1/2).

[11]t1/2=ln2ks0.693ks [11]

Release of saccharides, i.e. glucose, maltose (Merck, Germany) and maltotriose (Acros Organics, Belgium) as the products of starch biodegradation were also determined through reverse phase-high pressure liquid chromatography (RP-HPLC). The RP-HPLC (Agilent 1100) system assembled with ZORBAX Eclipse XDB-C18 column (250 mm length, 4.6 mm internal diameter and 5 μm particle size, Agilent, USA) was set at 30°C to resolve and quantify the concentration of saccharides. Saccharides were derivatized with 1-phenyl-3-methyl-5-pyrazolone (Sigma Aldrich, USA) and eluted with 0.1 m phosphate buffer (pH 6.7) and acetonitrile (Merck, Germany) (83:17% v/v; pH 7.1) at an isocratic flow rate of 1 ml/min and detected on a UV detector at a wavelength of 245 nm. Rates of formation (kf) and depletion ((kd) of each saccharide were determined for the composite films under aerobic soil through applying zero- and first-order reaction kinetics.

2.4.5 Lignin biodegradability

Low molecular weight phenolic compounds were removed before processing the soil for total lignin measurement using the method of Lepifre et al. (28). Homogenized freeze dried soil (2 g) was suspended in 30 ml of 50/50 (% v/v) of diethyl ether (R & M chemicals, Malaysia) /ethyl acetate (Merck, Germany) mixture. The mixture was vortexed for 2 min and then allowed to settle for 10 min in order to allow the soil fine particles to settle down under gravitational force. Then the organic phase was removed carefully and samples were allowed to dry under a fume hood.

The sum of lignin oxidation products, vanillyl-phenols [vanillin (Fluka, Switzerland), acetovanillone (Merck, Germany) and vanillic acid (Fluka, Switzerland)], syringyl-phenols [syringaldehyde (Santa Cruz Biotechnology, USA), acetosyringone (Sigma-Aldrich, USA) and syringic acid (Merck, Germany)] and cinnamyl-phenols [ferulic (Merck, Germany) and p-coumaric acids (Sigma-Aldrich, USA)] was used to determine total lignin concentration. Thus, lignin was oxidized by the CuO (Merck, Germany) oxidation technique (29). Trimethylsilylether and esters of CuO oxidation products of lignin were analyzed by GC-FID using a 60 m×0.320 mm (internal diameter) J&W DB-1 column (0.25 μm film thickness). Chromatographic separation was achieved using a temperature program: initial temperature, 100°C; ramp temperature, 4°C min-1; final temperature, 320°C and a final holding time, 10 min.

The decrease in the total lignin was calculated as sum of vanillyl-, syringyl- and cinnamyl-phenols (30). In addition, the increase in ratios of acidic to aldehydic forms of the vanillyl-(Ad/Al)V and syringyl-(Ad/Al)S phenols and decrease in the ratio of syringyl-to-vanillyl (S/V) phenol and cinnamyl-to-vanillyl (C/V) phenols were calculated as indices of lignin’s degree of biodegradation (30, 31) within composite films.

2.4.6 Slow release of urea-nitrogen

The Liang and Liu (32) method was followed to study the slow release of urea-nitrogen (urea-N) from composite films. Approximately 10 g soil was extracted with 30 ml of 0.5 m K2SO4 for 1 h at 25°C. Release of nitrogen in the soil extract was determined using a total nitrogen analyzer (Shimadzu, Kyoto, Japan) according to the Equation [12].

[12]NitrogenreleaseμgNg-1urea-Ng-1soildw=Ns-Nb×E×Df(mf×%N100)×soildw(g) [12]

Ns and Nb is the concentration of urea-N release in soil added with film and blank soil, respectively. E is the exractant volume (ml), Df is a dilution factor, mf is the mass (g) of the film, N is the theoretical total nitrogen in the film determined by the CHNS analyzer. The slow urea-N release properties of each composite film was investigated using a semi-empirical Korsmeyer-Peppas model (33) Equation [13].

[13]MtM=kϕtn [13]

where Mt/M, is the fraction of urea-nitrogen-released; is a release factor which is a characteristic of each composite film; t, release time (days); n is a release exponent determines the release mechanism. Diffusion coefficient (D) was calculated from Equation [14].

[14]MtM=4(Dtπl2)0.5 [14]

l is the thickness of each composite film which is taken 0.54 mm.

2.5 Statistical analysis

Mean data of each parameter was tested with a significance threshold level set at p<0.05 for all statistical analysis. One-way analysis of variance (ANOVA) was used to test the significant difference of lignin reinforced composite film on Michaelis-Menten reaction rates and Ea of each soil α-amylase and MnP enzymes. One-way repeated measures ANOVA was tested for significance difference of lignin on reduction of total starch biodegradability reduction in composite films and release of saccharides. ANOVA test was followed by a post-hoc Dunnet’s test. The goodness of experimental data fit to the empirical models was reported as error mean square (MSE) and coefficient of determination (R2). All statistical tests were run using the statistics package of OriginPro version 9.0.0 (OriginLab Corporation, Northampton, MA, USA).

3 Results and discussion

3.1 Mineralization

As shown in Figure 2, mineralization of composite films showed a reducing trend with increasing of lignin under aerobic soil conditions. Maximum mineralization of all films was reached at 21 days after which it was leveled-off until 28 days. Percentage mineralization, 66.84%, in control film was higher in comparison with composite films, where experienced mineralization in the ranged of 62.56 to 57.76%. This difference showed reduction of mineralization from 6.40 to 13.58% in composite films in response to an increase of 5–20% lignin as compared to control film. One-way repeated measures ANOVA showed mineralization was reduced for composite films more significantly (F1,12=50.73, p<0.001) by addition of lignin. Post-hoc Dunnett’s test of mean comparison further had showed all percentages of lignin could reduce mineralization very significantly (p<0.001) as compared to control film.

Figure 2: Effect of lignin on mineralization of composite films under aerobic soil.
Figure 2:

Effect of lignin on mineralization of composite films under aerobic soil.

Table 1 shows the prediction of mineralization by the Hill model. The MSE was not more than 6.90% and R2 had values>0.990 which jointly indicates a reliable predictability of the Hill model for mineralization kinetics of each composite film in aerobic soil. A very fast change in mineralization was seen from 2 to 4 days in all composite films. The maximum rate change of mineralization, rmax; required about 4 days when lignin was set ≥15% in composite films whereas it was only 2 days when lignin was ≤10% in composite films. It was further observed that rmax showed a pronounced delay from 19.79 to 40.76% with increasing lignin in composite films in contrast to control film. This was due to high mineralization of starch by soil microorganisms.

Table 1

Mineralization parameters of composite films in aerobic soil.

FilmsTime (days)ymax (%)rmax (%/day)kmin (days)nMSER2

The reduction in the mineralization of composite films occurred due to the lignin stabilizing effect on the starch matrix possibly through increasing the strength and resistance against α-amylase activity in aerobic soil. The maximum mineralization, ymax, reduced from 63.87 to 59.04% with increase of lignin, which explained the reduction of 6.18–12.94% as compared with control film. The mineralization half-life, kmin, escalated from 2.98 to 3.40 days after increase of lignin in composite films and this change was 2.68–14.70% higher than in control film. The curvature of the mineralization curve, n′, was found exponentially increase as lignin’s percentage increased in composite films. Increase in kmin, and decrease in rmax caused a more exponential shift of n′ at the initial time in response to a delay in mineralization of composite films compared to control film. Domenek et al. (34) reported the Hill model for mineralization kinetic studies in wheat gluten (protein) based bioplastic materials under aerobic farmland soil conditions. Compared with the current study, they predicted a higher ymax, 104.5 days; higher kmin, 4.91 days; lower n′, 1.28 over 50 days. The difference could be due to the nature of polymers and environmental conditions of the soil under which mineralization studies were conducted.

3.2 Michaelis-Menten reaction rates

3.2.1 α-amylase

Figure 3 shows Michaelis-Menten reaction rates of α-amylase in response to mixing composite films in aerobic soil. The Michaelis-Menten reaction rates of soil α-amylase were reduced more in soil after adding the composite films.

Figure 3: Michaelis-Menten reaction rates of α-amylase in aerobic soil mixed with different composite films.
Figure 3:

Michaelis-Menten reaction rates of α-amylase in aerobic soil mixed with different composite films.

Michaelis-Menten reaction rate constants of α-amylase enzyme in soil were determined for each composite film (Table S1). The maximum reaction rate, Vmax of α-amylase, responded negatively to the increase in percentages of lignin in composite films. Further, Vmax of α-amylase for the composite films was reduced ~1.72–2.03 times than its values found for control film. This confirmed that starch biodegradability in the composite films is protected by lignin through its reduction in Michaelis-Menten reaction rates of α-amylase in soil.

The substrate half-saturation constant, Km of α-amylase generally followed a decreasing trend after mixing composite films with aerobic soil. Low values of Km is an indication that α-amylase has more affinity against the composite films. The Km of α-amylase for the composite film was 0.93–1.87 times lower than the control film.

The catalytic efficiency, kcat of α-amylase showed a tendency of slow starch biodegradation in the composite films than the control film in aerobic soil. For composite film, the kcat of α-amylase was 1.16–1.83 times slower than control film. However, higher loadings of lignin percentages in the composite films, was observed with a general tendency of increase in kcat of α-amylase.

The lignin inhibition of α-amylase in composite films showed a competitive mode of enzyme inhibition. The decrease in Km (increase in substrate affinity) of α-amylase and the reasonably minor difference in Vmax confirmed that lignin favorably competed to bind α-amylase active site in composite films. This is in agreement with the competitive lignin inhibition of cellulase which has been proved through the increase in the value of Km with increasing concentration of lignin (35).

From the above prediction of Michaelis-Menten reaction rate constants, it was confirmed that lignin reduced significantly the α-amylase action on the starch in the composite films. Similar findings have been reported by Spiridon et al. (5) who found a 40% reduction in the Michaelis-Menten reaction rate of pure α-amylase while testing the biodegradation of the starch in its cellulosic fibers reinforced composite film than in the pure starch film.

As shown in Figure 4, Michaelis-Menten reaction rates of the soil MnP enzyme was enhanced by the lignin addition into the composite films under aerobic soil conditions. Percentages of lignin in the composite films showed a non-significant difference (one-way ANOVA, F5,54=2.31, p>0.05) on the MnP reaction rates.

Figure 4: Michaelis-Menten reaction rates of MnP in aerobic soil mixed with different composite films.
Figure 4:

Michaelis-Menten reaction rates of MnP in aerobic soil mixed with different composite films.

3.2.2 Manganese peroxidase

Michaelis-Menten reaction rate constants of MnP were determined in aerobic soil for different composite films (Table S1). It was found that 5–15% of lignin in the composite films increased the Vmax of MnP particularly higher than the control film. The lignin addition into the composite films correlated positively for increase in Vmax of MnP. The Vmax of MnP was 1.07–1.41 times faster in the soil, which was added with composite films as compared with control film. Thus, Vmax of MnP provided affirmative evidence of lignin influence on speed of reaction rates of MnP possibly due to more enzyme and substrate collision at higher percentages of lignin. Increase in Km of MnP was correlated negatively to the lignin percentages of the composite films under aerobic soil conditions. Compared to control film, the Km of MnP was 1.05–1.20 times higher for 5–15% of lignin based composite films while 1.05 times lower for 20% lignin added composite film. In fact, a gradually lower estimate of Km of MnP, explained that the affinity of enzyme increased for increasing percentages of lignin in composite films.

The inhibition of MnP has not been observed over the range of lignin percentages tested in composite films in aerobic soil. The increase of Vmax and decrease of Km (increase in substrate affinity) has confirmed the lignin-preferable binding with MnP and this is in agreement with the increase of Kcat.

The Kcat of MnP revealed a positive response to the increase in lignin percentages of composite films. Kcat of MnP was noticed 1.19–1.33 times faster and 1.12 times slower respectively for 10–20%L and 5%L reinforced composite films in comparisons with control film under aerobic soil environment. The reason for this increase in Kcat was due to lignin which could serve as a natural substrate of MnP.

In the current study, neither the starch nor the lignin increase (up to 20%) has any inhibition effect on MnP in aerobic soil. This supported the findings of higher Vmax, Kcat and lower Km (higher substrate affinity) of MnP in response to mounting lignin percentages in composite films. This MnP response could be explained by the oxidizabiltiy of Mn2+ into Mn3+ in MnP and thus it could form a complex with organic acid products of biodegradation and/or generate a diffusible oxidant which could attack the lignin further (36). It has been reported that lignin could reduce the release of nitrogen in the soil from starch-urea-borate film (16). Low nitrogen concentration in the soil might have led to the increase in reaction rates of MnP (37).

3.3 Activation energies

3.3.1 α-Amylase

The Ea of soil α-amylase was determined in a soil mixed with composite films (Table S3). The Ea of α-amylase increased in the range 1.10–1.41 times for the composite films as compared to control film as the biodegradability proceeds under the aerobic soil. However, those soils incubated with 5–15% lignin composite films, showed not much difference in their Ea of α-amylase.

The composite films showed a higher Ea of α-amylase in comparison with control film in aerobic soil. The reason for higher Ea, of α-amylase is its competitive inhibition by the lignin. In addition, a slightly descending trend in Ea was noticed for the shift in lignin from 5 to 20% in composite films. It could be due to the phase-separation between lignin and starch in composite films, which leads to adsorption of soil α-amylase enzyme, further ease the binding of the enzyme to starch, and hence possibly slightly declined the Ea. In general, analysis of Ea showed high sensitivity of soil α-amylase to temperature increase for composite films as compared to control film. The effect of different composite films on the Ea of α-amylase showed a significant difference (one-way ANOVA F5.12=7.56, p<0.05). The post-hoc Dunnet’s test of mean comparison showed a significant difference of Ea of α-amylase only at 5% lignin in composite film as compared to control film. Similar findings have been reported for increases in Ea of α-amylase during hydrolysis of cellulosic fibers reinforced starch (composite) film than pure starch film (5).

3.3.2 Manganese peroxidase

The Ea of soil MnP for the composite films are shown in Table S4. The Ea of MnP was reduced more in 10–15% lignin composite films than in the control film while its response in 20% lignin composite film was more similar to the control film. Low Ea of MnP was related to its being less temperature sensitive for initiating lignin biodegradability in composite films.

The Ea of MnP for composite films was found to be 1.02–1.11 times higher than control film. The effect of different composite films mixing into aerobic soil on Ea of MnP showed a significant difference (one-way ANOVA F5.12=5.27, p<0.05). The post-hoc Dunnet’s test of pairwise means comparison established a non-significant (p>0.05) difference of lignin on Ea of composite films. The low Ea of MnP observed in this study could be supported with poor dependence of MnP reaction on the external energy and less-driven by the increase of lignin concentration due to its similar chemical composition (38) within composite films.

3.4 Total starch decay and half-life

Figure 5 shows the biodegradability of starch in composite films as compared to control film under aerobic soil environment. It was found that starch decreased exponentially in all films. It was observed that starch required ~21–28 days to reach its maximum biodegradability. It was estimated that lignin minimized the starch biodegradability within the composite films from 86.83 to 95.65% compared to 97.04% in control film. Further, lignin addition into the composite films had showed a non-significant difference (one-way repeated measures ANOVA, F1,9=8.00, p>0.05) on the reduction of starch biodegradability.

Figure 5: Effect of lignin on starch biodegradability in composite films under aerobic soil.
Figure 5:

Effect of lignin on starch biodegradability in composite films under aerobic soil.

In the current study, alkaline kraft lignin would have been a reason for lower water uptake into the composite film with increase of lignin percentages (13, 39). This behavior of the composite films could be anticipated to understand the starch biodegradability in real soil conditions. Soil water naturally contained rich inoculums of various microbial species, which enters into the composite films through the advection process. Therefore, higher percentages of lignin could resist more water uptake which results in less microbial species per unit volume of the composite films. It thus restricted the microbial access to the hydrophilic starch surrounded by hydrophobic lignin.

Estimation of starch biodegradability rate (ks) and its half-life (t1/2) was calculated from Figure 5 (Table S5). It was shown that starch biodegradability rates reduced and t1/2 improved in composite films as compared to control film. Starch biodegradability rates were 0.122–0.168 day-1 in composite films as compared to 0.254 day-1 in control film. Starch t1/2 was enhanced up to 4.12–5.68 days in composite films than 2.72 days in control film. Thus, starch t1/2 was improved 33.86–51.98% than control film in aerobic soil.

3.4.1 Release of saccharides

Saccharides like glucose (Figure S3: a), maltose (Figure S3: b) and maltotriose (Figure S3: c) were determined as products of starch biodegradation from each composite film in soil. It was observed that the yield of each saccharide reached linearly to their maximum peak values within 4 days irrespective of lignin percentages added into composite films. Afterwards, the yield of each saccharide exponentially reached to lowest values until 14 days with no further remarkable change till 28 days. A negative correlation was confirmed between the lignin and each saccharide as findings showed increase in lignin loadings in composite films, yield of each saccharide decreased more. One-way repeated measures ANOVA explained a significant difference in the yield of glucose (F1,8=7.16, p<0.05) and maltose (F1,8=8.51, p<0.05) while a non-significant difference in the yield of maltotriose (F1,8=4.78, p>0.05). Further, a post-hoc Dunnett’s test of pairwise means comparison highlighted a highly significant (p<0.001) difference in the yields of glucose in 15 and 20% lignin composite films whereas maltose and maltotriose in 10–20% lignin composite films.

It was noticed that the maximum yield of saccharides was reduced by lignin in composite films. It was found that the maximum yield of glucose decreased from 694.12 to 490.24 μg g-1 soil dw (7.55–34.71% lower than control film) (Figure S3: a); maltose from 790.56 to 642.54 μg g-1 soil dw (4.14–22.11% lower than control film) (Figure S3: b) and maltotriose from 1515.46 to 1131.51 μg g-1 soil dw (3.22–25.33% lower than control film) (Figure S3: c). Therefore, it can be anticipated that large fragments of starch formed more frequently due to breaking of glycosidic linkages randomly at multiple sites within very long starch polymer chains. It resulted in the maximum yield of saccharides in the sequence maltotriose>maltose>glucose among all films. Based on the results, it could be anticipated that lignin protected the starch through conservation of large fragments and their slow conversion into saccharides.

Thus, the initial maximum yield of each saccharides; glucose, maltose and maltotriose over 0–4 days was considered as a “saccharide formation” step whereas the maximum to the minimum yield over 4–28 days was considered as a “saccharides depletion” step (Figure S3: a–c and Table S6). It was found that saccharides formation proceeds according to zero-order reaction kinetics whereas saccharides depletion involved the first-order reaction kinetics under aerobic soil conditions.

Increase of lignin in composite films, the zero-order kinetic rates for saccharides formation decreased for glucose from 197.87 to 150.99 μg day-1 (8.21–29.96% lower than control film); maltose from 208.42 to 187.50 μg day-1 (2.49–12.28% lower than control film); maltotriose from 422.01 to 359.04 μg day-1 (0.96–15.74% lower than control film) (Table S6). The zero-order kinetic rates of maltotriose were higher than maltose and glucose. The percent difference in the zero-order kinetic rates of maltotriose and maltose was less different than the glucose in the composite films as compared with the control film.

Table 6 shows the first-order kinetic rates which demonstrated that saccharides depletion supersedes their formation in the events of starch biodegradation in all films. Upon increase of lignin’s loadings into composite films, the rates of saccharides depletion become slower than that of control film. This was confirmed from the decrease in first order kinetic rates of glucose from 0.351 to 0.252 day-1 (~6.312–16.27% lower than control film); for maltose from 0.318 to 0.298 day-1 (~1.88–6.28% lower than control film); maltotriose from 0.480 to 0.454 day-1 (~9.53–14.46% higher than control film. The more percentage change in the kinetic rates of maltotriose and glucose depletion over the maltose predicted that possibly large fragments of starch deplete faster into low molecular weight saccharides.

Based on reduction of yield and kinetic rates of saccharides formation and depletion, it is confirmed that rates of starch saccharification were reduced by lignin strengthening of starch network through its hydrophobic filler characteristics (13, 40) and inhibition of α-amylases. In addition, lignin through the condensation reactions with starch could produce more dimers and oligomeric structures under radiation (28). Therefore, under the current study, thermal radiation during post-curing process of composite films might have promoted the reactivity of the lignin within the starch matrix. Consequently, such process could reduce the maximum yield and kinetic rates of saccharides, favored through condensation reaction of large fragments of starch with lignin in composite films. It ultimately might have reduced the number of bonds breakdown along the starch polymer in composite films.

3.5 Lignin biodegradation

Figure 6 shows that lignin biodegradability in composite films under soil conditions. It was found that biodegradability of lignin was slowed in composite films with a higher percentage of lignin. It was in agreement with conservation of lignin biodegradability about 6.20% more with increase of 5–20% lignin in composite films. The higher starch to lignin ratio at low percentages of lignin in composite film is assumed to increase the lignin hydrophilic character upon starch biodegradation, which thus also increased the lignin susceptibility to further biodegrade easily.

Figure 6: Lignin biodegradability in composite films under aerobic soil.
Figure 6:

Lignin biodegradability in composite films under aerobic soil.

First-order reactions were confirmed that increase of 5–20% lignin in composite films gradually reduced the lignin biodegradability rates from 0.0063 to 0.0042 day-1 (33.33%). The t1/2 of lignin was higher in those composite films reinforced with higher percentages of lignin. The t1/2 was calculated 109.75 days, 120.16 days, 139.06 days and 161.93 days, respectively, for 5, 10, 15 and 20% lignin in composite films (Linear eq. 8.77x+97.64; R2, 0.974). In addition, biodegradability of lignin in different composite films showed a highly significant difference (one-way repeated measures ANOVA, F1,9=978.58, p<0.001).

Results explained that increasing lignin from 5 to 20% in composite films slowed the loss of vanillyl phenols, 5.42% (Figure S4a); cinnamyl-phenols, 5.72% (Figure S4b); syringyl-phenols, 6.86% (Figure S4c). From Table 2, it was observed that indices of lignin’s biodegradation like (Ad/Al)v and (Ad/Al)s ratios decreased with increase of lignin loadings of composite films. The (Ad/Al)v ratio was decreased from 1.18 to 1.17 (~0.76%). The (Ad/Al)s ratio decreased from 1.59 to 1.56 (~1.78%) which showed more significant and higher change in values than found for (Ad/Al)v ratio. This is in agreement with the findings of Thevenot et al. (15) where they have reported lower values of (Ad/Al)v ratio as compared to (Ad/Al)s ratio in different origins of soil. This analysis also confirmed that the vanillyl-phenols were more resistant than syringyl phenols during the biodegradability of lignin in composite films. The ratio of cinnamyl/vanillyl (C/V) increased from 1.198 to 1.199 (~1.00%) and ratios of syringyl/vanillyl (S/V) increased from 1.33 to 1.53 (~1.15%). These indices provide not only the evidence of lignin biodegradability but also confirmed a higher percentage of lignin in composite films reduced its biodegradability. Thus, it is possible that starch increased the lignin’s hydrophilicity and increased the release of more acidic phenols from aldehyde and syringyl units in lignin. This is supported with earlier observations of fast Michaelis-Menten kinetic reactions of soil MnP (see Section 3.2.2) against those composite films comprised of low lignin percentages (i.e. higher starch to lignin ratio). It also supports the possibility that higher starch to lignin ratio in composite films could increase the cleavage of Cα-Cβ bonds of phenylpropane units within the lignin structure (15).

Table 2

Indices of lignin biodegradability in composite films under aerobic soil.

Films(Ad/Al)v Mean±SD(Ad/Al)s Mean±SDS/V Mean±SDC/V Mean±SD

3.6 Nitrogen release

Nitrogen release as log(Mt/M) versus its log time was plotted for different composite films to determine the values of the nitrogen release exponent, n; release factor, kϕ; and diffusion coefficient, D; as shown in Table 3. The kinetic mechanism of urea-nitrogen release showed a “Fickian diffusion” process in all biodegraded films as it was confirmed from the value of n which was <0.5. However, increase of lignin had showed a relative increase in the value of n and kϕ which together confirmed a strong control of lignin exerted on the overall geometry of the composite films and hence release of cross-linked nitrogen was reduced in soil. It was found that the value of D for nitrogen reduced with increasing of lignin in biodegraded composite films. The composite films showed lignin reduced the D values from 0.66 to 0.94 times its value for control film.

Table 3

The effect of composite films on released exponent (n), release factor (kϕ) and diffusion coefficient (D) of nitrogen release in aerobic soil.

FilmsRelease exponent, nRelease factor, kϕCorrelation coefficient, R2Diffusion Coefficient, D (cm2/s)

In chitosan/polyacrylic acid-co-acrylamide coated NPK fertilizer, Liang et al. (41) noticed higher values of n, 0.54; much lower values of kϕ, 0.051; higher values of D, 2.96×10-4 for nitrogen release through coated fertilizer granules into soil. In the current study, the difference could be due to the process of urea-crosslinking with starch in composite films instead of coating urea as reported by Liang et al. (41).

However, nutrient release from biodegradable composite films very much depends on environmental conditions like soil pH, ionic strength and clay-polymer coating interaction, which determines the release of trapped nitrogen through osmotic processes and enzymatic hydrolysis of amide linkages in urea cross-linked starch. Slow release of chemically trapped nitrogen in composite films relates to lignin resistance to soil microorganisms activity and its contribution to reduction of starch fast biodegradability under aerobic soil.

4 Conclusion

This research work concludes that addition of lignin into composite films could effectively reduce the biodegradability as compared to control film. Thus, slow biodegradability of the composite films reduced the fast release of crosslinked urea-nitrogen with starch. Slow mineralization of composite films explained the evidence for lignin controlling the biodegradability of starch in composite films. Higher starch to lignin ratio decreased the hydrophobicity of the lignin, which caused fast lignin’s acid to aldehyde turnover. Decrease of Kcat and increase of Ea provided insight into the idea that lignin inhibition of soil α-amylase shifts higher the external energy barrier involved in starch biodegradability in composite films. For future study we recommend to study the surface modifications of composite films with enzyme inhibitors to reduce their further biodegradability in soil.

Corresponding authors: Zahid Majeed and Nurlidia Mansor, Department of Chemical Engineering, Universiti Teknologi PETRONAS, Bandar Seri Iskandar, 32610 Tronoh, Perak, Malaysia, e-mail: (Z. Majeed); (N. Mansor)


Authors gratefully acknowledge the Ministry of Higher Education (MOHE), Malaysia, for the research funding through OneBaja Project (Grant No. 0153AB-C71-4) under the Long Term Research Grant Scheme (LRGS). The first author is also grateful to Universiti Teknologi PETRONAS, Malaysia for providing research facilities and financial support through Graduate Assistantship.


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Supplemental Material

The online version of this article (DOI: 10.1515/epoly-2015-0231) offers supplementary material, available to authorized users.

Received: 2015-10-14
Accepted: 2015-11-28
Published Online: 2016-1-29
Published in Print: 2016-3-1

©2016 by De Gruyter

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