Degradation of disperse blue 2BLN by oleaginous C. sorokiniana XJK

Abstract

In this study, an oil-producing freshwater microalgae Chlorella sorokiniana XJK was identified and used for the degradation of disperse blue 2BLN. According to the decolorization rate, the optimal conditions were found to be as follows: initial dye conc. = 60 mg L⁻¹, initial biomass density = 8 × 10⁶ cell per mL, initial pH = 4, light intensity 470 μmol (m² s)⁻¹ and CO₂/air (v/v) ratio = 1%. The decolorization rate reached up to 83% in 6 days cultivation with Chlorella sorokiniana XJK, which also obtained 570 mg L⁻¹ of biomass and 43% of lipid content under the optimum conditions. Moreover, the enzyme activities (laccase and manganese peroxidase) in XJK were determined to be 3.89 U mL⁻¹ and 4.86 U mL⁻¹, respectively, which played a significant role in dye decolorization. The degradation products of the dye were analyzed by UV-vis, FTIR and GC-MS methods. This revealed that the major chromophore group was broken down and disperse blue 2BLN was converted into several low-molecular compounds including dibutyl phthalate, ethylbenzene and ethyl acetate during the microalgae-based treatment. This indicated that XJK shows a potential for dye degradation and providing materials for biofuel.

---

1. Introduction

Nowadays, with the rapid development of economy and industry, synthetic dyes are commonly and widely applied in energy sources, agriculture and all types of industries. According to their chemical structure,¹ anthraquinone dyes are the critical groups after azo dyes to have attracted increasing attention to their toxicological and environmental effects with their increasing applications. It has been reported that some losses of the dyes are incurred during the dyeing process and a large quantity of partial dyes are emitted from the textile industry each year.² These effluents are usually discharged in high concentrations and the toxicity of multiple pollutants, such as the complex aromatic structure of dyes, pigments, heavy metals, biocides, surfactants, multifarious organic and inorganic matter can give rise to complexation phenomena.^3,4 Moreover, the contaminants may cause carcinogenic and teratogenic effects on human health, such as dermatitis, ulceration of skin and cancer and pollute water bodies too.⁵ Hence, there is a significant need to treat dye wastewater prior to discharge. Thus far, physical, chemical and biological processes are available for dyestuff treatment. Nevertheless, the development of physicochemical methods⁶ has been blocked due to the high-cost, time and material consumption, lack of management and disturbance by other wastewater components on the settlement of dye wastewater. There is a considerable need to put forward biological processes to dispose wastewater, particularly in light of their economic efficiency, eco-friendliness and extensive adaptability.⁷

Various microorganisms in all biological processes such as fungi, bacteria and algae display efficiency in the degradation of synthetic dyestuff.⁸ Different types of bacteria such as Escherichia coli⁹ and Enterobacter sp. F NCIM 5545 (ref. 10) are employed for different dyes and dye wastewater. Next to bacteria, the role of fungi, such as Tinctoporellus sp. CBMAI 1061, Ganoderma sp. En3 and Irpex lacteus F17,^11–13 in dye removal have been comprehensively studied. Certain microalgae including Gonium sp.⁸ and Chlorella pyrenoidosa¹⁴ have also been effectively applied in the disposal of dye-containing wastewater and water purification. Among microorganisms, microalga have potential applications in treating wastewater because of their adaptability in hostile conditions.⁷ According to Kim,¹⁵ algae-based piggery effluent treatment efficiently removed inorganic nutrients and accumulated algal biomass using an ozonation pretreatment with a dose of 1.1 mg O₃ per mg C or light transmittance magnification in liquor. In view of the recycling utilization of the resource, algae-based treatment has been widely used on wastewater, which could make full use of the wastewater as a growth medium and promote carbon dioxide fixation. Besides the treatment of wastewater with significantly low cost, a great amount of microalgae can serve as crude materials of renewable fuels such as bio-oil.¹⁶

At present, some synthetic dyes, involving anthraquinone groups, have been processed by fungi and bacteria. Liu et al.¹ found that alizarin red (anthraquinone) was destroyed and transformed into small molecules using chloroperoxidase obtained from Caldariomyces fumago. The biodecolorization and biodegradation of the dye simultaneously occurred during the treatment process with mixed fungi.¹⁷

In this study, the decolorization efficiency of one oleaginous Chlorella sorokiniana XJK was investigated with dye (disperse blue 2BLN) wastewater in an air-lift column photobioreactor. Various parameters including the dye concentration, biomass density, initial pH value, light intensity, and CO₂ concentration were assayed, and the decolorization rate served as an index to achieve the optimal conditions. Considering the role of the enzyme on dye removal, we attempted to assay the ligninolity enzyme activities of Chlorella sorokiniana XJK during the algal growth. Furthermore, another important aspect of this work was to confirm the degradation mechanism of anthraquinone dye using UV-vis, FTIR and GC-MS analysis.

2. Materials and methods

2.1 Dye

Disperse blue 2BLN (1,5-diamino-2-chloro-4,8-dihydroxy-9,10-anthracenedione) was purchased from Zhejiang Runtu Co. Ltd in a chemically pure form. It is one of the anthraquinone dyes and belongs to the group of the most durable dyes. It is widely utilized in dyeing polyester fibre and its blended fabrics. The chemical structure of disperse blue 2BLN used for decolorization research is shown in Fig. 1.

Fig. 1 The chemical structure of disperse blue 2BLN.

2.2 Identification of microalga

One freshwater oleaginous microalga XJK was isolated from Keketuohai, XinJiang Province, China. The microalga was preliminary identified through the observation of its morphological characteristics, including cell size, shape, and chloroplast core protein number. Nucleic acid extraction of the algal cells was performed using the NuClean PlantGen DNA kit (Beijing Com Win Biotech Co., Ltd., China) according to the manufacturer's instructions. The internal transcribed spacer (ITS) was amplified by polymerase chain reaction (PCR) using the universal primer ITS-F (5′-GGAAGTAA AAGTCGTAACAAGG-3′) and ITS-R (5′-TCCTCCGCTTATTGATATGC-3′) according to the procedure of Katia et al.¹⁸ The amplified sequence was sent for analysis. The sequence obtained for the ITS gene was aligned to the published sequences obtained from GenBank using Clustal X 1.83. A neighbor-joining tree was constructed from the data by the bootstrap method (1000 replicates) using Mega 5.0 software.

XJK was then conserved in Key Laboratory for Green Process of Chemical Engineering of Xinjiang Bingtuan, Shihezi University.

2.3 Culture medium and conditions

The basal culture medium BG-11 was modified as follows: NaNO₃ – 0.83 g L⁻¹, K₂HPO₄·3H₂O – 0.09 g L⁻¹, MgSO₄·7H₂O – 0.12 g L⁻¹, CaCl₂ – 27.2 mg L⁻¹, citric-acid – 6.6 mg L⁻¹, Fe(NH₄)₃(C₆H₅O₇)₂ – 6.0 mg L⁻¹, NaCO₃ – 0.02 g L⁻¹, Na₂EDTA·2H₂O – 1.1 mg L⁻¹, 0.2 mol L⁻¹ and trace metal mix (H₃BO₃ – 2.86 g L⁻¹, MnCl₂·4H₂O – 1.86 g L⁻¹, ZnSO₄·7H₂O – 0.22 g L⁻¹, NaMoO₄·2H₂O – 0.39 g L⁻¹, CuSO₄·5H₂O – 0.08 g L⁻¹ and Co(NO₃)₂·6H₂O – 0.05 g L⁻¹).

The dye medium was prepared by adding disperse blue 2BLN at different concentrations into the modified medium BG11.

Seed cultured photoautotrophically in a 2 L Erlenmeyer flask containing 1.3 L medium was placed in an illumination chamber under 830 μmol (m² s)⁻¹ with a 12/0 h light/dark photoperiod in a constant temperature (25 °C) for 5 days. Aeration and mixing were achieved by sparging air through a 0.2 μm gas filter (Millipore) at a flow rate of 0.4 m³ h⁻¹.

Photobioreactor and photoautotrophic cultivation conditions: A 1 L air-lift column photobioreactor (diameter: 5.00 cm; height: 55.00 cm) containing 700 mL working volume was used for the photoautotrophic cultivation. The inoculum biomass density was 9.9 × 10⁶ cell mL⁻¹. The culture conditions were maintained at room temperature (25 °C) under a white fluorescent light with 12/0 day/dark photoperiod for 6 days. The light intensity was approximately 830 μmol (m² s)⁻¹ at the surface of the photobioreactor for two sides illumination. Aeration and mixing were achieved by sparging air with CO₂ through a 0.2 μm gas filter (Millipore) at a flow rate of 0.4 m³ h⁻¹.

2.4 Optimization on decolorizing ability

According to biomass, lipid production, and the best dye decolorization rate of microalga XJK, the initial concentration of disperse blue 2BLN, incubating biomass density, initial pH, light intensity and CO₂/air (v/v) ratio were optimized in sequence. All the experiments were conducted in triplicate.

The optimum initial concentration of disperse blue 2BLN was firstly selected from 60–110 mg L⁻¹ at intervals of 10 mg L⁻¹. Next, an inoculum biomass density ranging from 3 × 10⁶ to 1.5 × 10⁷ cell per mL on the decolorization efficiency was examined, based on the selected dye concentration.

The dye medium with the dye was then prepared at pH 2, 4, 6, 8, 10 and 11, adjusted using 1 mol L⁻¹ NaOH and HCl solution and measured with a pH meter (PHS-3C, INESA). The effect of the initial pH value on the decolorization efficiency was examined using the optimized initial dye concentration and biomass density.

Subsequently, the effect of light intensity on the decolorization efficiency was investigated, where the dye concentration (60 mg L⁻¹), biomass density (8 × 10⁶ cell mL⁻¹) and initial pH value at 4 were kept constant. Different light densities were chosen: 830 μmol (m² s)⁻¹ (designated as high light HL), 470 μmol (m² s)⁻¹ (designated as low light LL), natural light (designated as NL), 0 μmol (m² s)⁻¹ (designated as photophobism PP), 830 μmol (m² s)⁻¹ for the first three days and 470 μmol (m² s)⁻¹ for the last three days (designated as decreasing photoperiod DP), 470 μmol (m² s)⁻¹ for the first three days and 830 μmol (m² s)⁻¹ for the last three days (designated as increasing photoperiod IP).

Finally, six levels of CO₂/air (v/v) ratios (0%, 1%, 3%, 5%, 7%, and 9%) were used to determine the best concentration for maximum decolorization, where the dye concentration (60 mg L⁻¹), biomass density (8 × 10⁶ cell mL⁻¹), initial pH (4) and 470 μmol (m² s)⁻¹ of light intensity were kept constant.

2.5 Determination of ligninolytic enzyme

2.5.1 Preparation of the crude enzyme. Broths at regular cultural times were sampled and centrifugated at 10 000 rpm for 5 min. The supernatant fractions were used to determine the extracellular enzyme. Cells were frozen and ground with liquid nitrogen and reconstituted in buffer. The intracellular enzyme was then obtained by centrifugation at 10 000 rpm for 5 min, after removing the accumulated sedimentation. The enzymatic activities were determined spectrophotometrically. In all cases, the reaction mixture, in a final volume of 4 mL, contained 400 μL of sample.

2.5.2 Determination of laccase. Laccase activity was determined by the oxidation of 2,2′-azinobis(3-ethylbenzthiazoline)-6-sulfonate (ABTS).¹⁹ The reaction mixture was incubated in Britton–Robinson (pH 2) containing 0.5 mM ABTS at 45 °C. In addition, the oxidation was followed via the increase in absorbance at 420 nm (ε₄₂₀ = 36 000 M cm⁻¹).

2.5.3 Determination of lignin peroxidase. The reaction system comprised of lignin peroxidase (LiP) was established according to Iqbal et al.,²⁰ detecting the absorbance at 310 nm (ε₃₁₀ = 9300 M cm⁻¹). The assay mixture was incubated in tartaric acid buffer (pH 3) containing 15 mM veratryl alcohol at 25 °C. The reaction was started upon the addition of 0.1 mL H₂O₂ until a final concentration of 15 mM H₂O₂.

2.5.4 Determination of manganese peroxidase. MnP activity was determined using the method of Bermek et al.²¹ The reaction mixture included 0.11 M malonate–sodium malonate (pH 4.5) and 40 mM MnSO₄, and the reaction was started by the addition of 0.1 mL H₂O₂ until a final concentration of 1.6 mM H₂O₂. The formation of Mn(III)–malonate complexes was followed at 270 nm (ε₂₇₀ = 11 590 M cm⁻¹) at 25 °C.

2.6 Analysis

2.6.1 Determination of the decolorization rate. Dye decolourization was monitored using a spectrophotometer (UV-vis spectrophotometer, spectrumlab S22_PC). Aliquots from different test tubes were taken at regular intervals and centrifuged at 10 000 rpm for 5 min to remove the cells. Absorbance of supernatant was first scanned by UV-vis from 400 to 800 nm using the culture medium as the control. The decolorization rate at the maximum absorption wavelength of 560 nm was defined as the changes of initial absorbance (A₀) and absorbance of t moment (A_t) calculated as follows:

2.6.2 Determination of biomass and lipid. The total weight of the collected wet biomass was dried for 12 hours using a lyophilizer at −55 °C and re-weighed to obtain the dried weight. The biomass (mg L⁻¹) was defined as the dried weight (m₀) in mg and training volume (V₀) in L calculated as follows:

The total lipids were extracted following the protocol of Bligh and Dyer.²² Briefly, 10 mg of the lyophilized algal powder and distilled water of 1 mL served as layering and were added to 6 mL of a chloroform : methanol (2 : 1, v/v) solution. The mixture was shaken at 40 °C and 200 rpm for 4 hours. The solution was centrifuged at 10 000 rpm for 5 min. The chloroform layer was evaporated using a rotary evaporator (model RE-201D, Shanghai Dongxi) at 40 °C for 10 min. Herein, m₁ was defined as the weight of extracted liquid and the evaporation flask, and m₂ was defined as the weight of the evaporation flask dried in an oven at 105 °C for 2 hours.

The lipids content in % and lipids yield in mg (L d)⁻¹ were determined according to the difference value of m₁ and m₂. The calculations were as follows:

2.6.3 Experimental evidence for dye decolorization. Sample pretreatment for FTIR and GC-MS analysis was as follows: supernatants of the broth were extracted with an equal volume of ethyl acetate and the ethyl acetate layer concentrated using a rotary evaporator (model RE-201D, Shanghai Dongxi) at room temperature. The extract was filtered with 0.22 μm Millex syringe filters and stood for 12 hours after removing the water upon the addition of Na₂SO_4.

Infrared spectra of the preprocessing liquid samples were analyzed using a Fourier transform infrared spectrometer (FTIR) (Magna-IR 750, Thermo Nicolet). KBr was used as the background; liquid samples were pointed at potassium bromide tableting to obtain their infrared spectra. The resolution and scan number were set at 8.0 cm⁻¹ and 16, respectively. The scan range of the wavenumber was set at the mid IR region (4000–400 cm⁻¹).

The degradation products of disperse blue 2BLN were also analyzed by GC-MS, which comprised of a GC system (model 7890A, Agilent) equipped with a HP-5MS capillary chromatographic column (30 × 0.32 × 0.25) and coupled to a MS system (model 5975C, Agilent). Helium served as the carrier gas at a flow rate of 0.8 mL min⁻¹ and the injector temperature was 250 °C in splitless mode. The extracted samples were measured under the following temperature gradient:²³ the initial column temperature was kept constant at 40 °C for 10 min, increased to 100 °C at a rate of 12 °C min⁻¹, then ramped at 5 °C min⁻¹ to 200 °C, further raised at 20 °C min⁻¹ to 270 °C and held constant for 5 min, then with increase to 300 °C at 10 °C min⁻¹ remained constant for 5 min. The degradation products in different periods were analyzed by comparing the GC-MS spectra patterns with those of standard mass spectra in the National Institute of Standards and Technology (NIST) library.

3. Results and discussion

3.1 Microalgae identification

According to its morphological characteristics, including cell size, shape, and chloroplast core protein number, the screened oleaginous microalga was named as Chlorella sp.-XJK. A micrograph of the microalga XJK is shown in Fig. 2a. Besides the characterization of lipid production (20%), it was found that XJK could also grow in the dye medium with a lipid production of 24% (Fig. 2b). When compared to the basal modified medium BG11, the accumulation of biomass evidently decreased in the dye medium because the presence of chlorides, complex aromatic structures, heavy metals, and synthetic dyes limit the light penetration and inhibit the growth of the microalgae.⁵ However, the production of lipids was slightly increased in XJK with the existence of the dye. Gopalakrishnan et al. found that lipids act as a secondary metabolite, maintaining specific cell signaling pathways and membrane functions while responding to changes in the environment.⁷ That is, the screened microalga XJK can be used to remove the synthetic dye from water and accumulate lipids for biofuels at the same time.

Fig. 2 Identification of the microalga XJK. (a) Micrograph (400×) of microalgae XJK; (b) the accumulation of biomass and lipid in the basal and dye mediums; (c) the gel electrophoresis map of products of ITS from microalgae XJK and (d) the phylogenetic tree of microalgae XJK based on its ITS sequence.

Further identification of XJK was carried out by PCR amplification and subsequent DNA sequencing. The length of the ITS sequence was obtained as 751 bp (Fig. 2c). The ITS sequence of XJK showed 99% sequence homology with Chlorella sorokiniana RP JX014220.1 (Fig. 2d). It was thus named Chlorella sorokiniana XJK (C. sorokiniana XJK). Related sequences were downloaded and used to build a phylogenetic tree using Mega 5.0 software.

3.2 Optimization on dye decolorization

Environmental variables can remarkably affect the dye removal process by microalga. A series of experiments were carried out and the optimum conditions of disperse blue 2BLN removal were acquired according to the decolorization rate.

3.2.1 The effect of dye concentration. It was shown that the decolorization rate and accumulation of biomass decreased with increasing dye concentration (Fig. 3A and a). After 6 days of cultivation, a maximum decolorization rate of 72% was found with an initial dye concentration of 60 mg L⁻¹ (Fig. 3A). Moreover, a maximum of 1243 mg L⁻¹ biomass was also obtained at 60 mg L⁻¹(Fig. 3a). However, the highest lipid content of 44% was obtained when XJK was subjected to an initial dye concentration of 90 mg L⁻¹. The decreasing trend of decolorization rate was similar to that reported by M. Cerboneschi⁹ and R. Waqas.²⁴ Exceedingly high concentration of dye caused less light penetration for photosynthesis and inhibited the impacted growth, metabolite production and photosynthetic activity in the XJK cells.²⁴ Hence, a 60 mg L⁻¹ dye concentration was selected for further studies.

Fig. 3 Optimization on the decolorization and accumulation of biomass of disperse blue 2BLN using Chlorella sorokiniana XJK. (A) Dye concentration (biomass density = 9.9 × 10⁶ cell per mL was constant); (B) biomass density (dye conc. = 60 mg L⁻¹ was constant); (C) initial pH (dye conc. = 60 mg L⁻¹ and biomass density = 8 × 10⁶ cell per mL were constant); (D) light density (dye conc. = 60 mg L⁻¹, biomass density = 8 × 10⁶ cell per mL and pH = 4 were constant); (E) CO₂/air (v/v) ratio (dye conc. = 60 mg L⁻¹, biomass density = 8 × 10⁶ cell per mL, pH = 4 and light intensity = 470 μmol (m² s)⁻¹ were constant).

3.2.2 The effect of the inoculums biomass density. The effects of the inoculums biomass density on the decolorization rate and growth of XJK are shown in Fig. 3B and b, respectively. A maximum decolorization rate of 80% was obtained by C. sorokiniana XJK when the inoculums biomass density was found to be 8 × 10⁶ cell per mL on the 4th day of the culture (Fig. 3B). The decolorization rate was decreased with further increases in the inoculum concentration. The decreasing decolorization rate was attributed to the algae cells receiving less light to support their quick growth and metabolism when the cells were inoculated at higher initial cell density.²⁵ Furthermore, the competition for nutrients with microalgae caused slower growth, thus affecting the decolorization and accumulation of biomass production. After 6 days of cultivation, a maximum of 1184 mg L⁻¹ biomass was obtained with 1.3 × 10⁷ cell per mL biomass density (Fig. 3b). Moreover, it only obtained a lipid yield of 71 mg (L d)⁻¹ and lipid content of 41%. In comparison, XJK had accumulated the highest lipid content (44%) and lipid yield of 81 mg (L d)⁻¹ with a low inoculum biomass density (8 × 10⁶ cell per mL). The results demonstrate what Ale et al. proposed to define an optimal balance between inoculums biomass density and algal density,²⁶ rather than the concept of increasing the inoculum density to increase the biomass. This corresponds with the previous study reported by Qiaoning and Feng who observed the negative relationship between inoculum size and lipid accumulation.^25,27 In view of decolorization rate and lipid content, 8 × 10⁶ cell per mL inoculum biomass density was used for further studies.

3.2.3 The effect of the initial pH on decolorization. C. sorokiniana XJK was found to be sensitive to pH ranging from 2 to 11 during the dye removal process. The decolorization rate of disperse blue 2BLN was determined under different pH conditions (Fig. 3C). Acidic conditions were more favorable to dye removal when compared with alkaline condition and a maximum decolorization rate of 78% was found at pH 4. The quick increase in decolorization rate in the first 4 days was caused by the increasing growth of microalgae, which was beneficial for the attractive forces between the dye molecules and algae, leading to fast diffusion into the algae cells to decolorize, and then attain a rapid equilibrium.²⁸ The decolorization rate of disperse blue 2BLN was kept stable after 4 days, which could be attributed to the pH affecting the enzymatic activity participating in the decolorization rather than the growth of the algae.²⁴ The acidic conditions were favorable for algal growth and accumulation. It can be speculated that the cell surface and dye molecules rarely interact at a pH beyond 4.²⁹ When the pH was less than 3, which is the isoelectric point for algae sp.,³⁰ it caused a decrease in the decolorizing efficiency due to the competition of H⁺ ions and dye cations. The decolorizing efficiency was less at pH 2, where only 19% efficiency was observed over 6 days. An increase in the pH results in a decrease of accumulation of both biomass and lipids (Fig. 3c). Moreover, a maximum biomass (838 mg L⁻¹) was obtained at pH 4 and further acquired 37% of lipid content. In addition, the maximum lipid content (44%) was observed at pH 6. Hence, an optimal pH of 4 was used in our further experiments.

3.2.4 The effect of the light intensity on decolorization. Given that the light intensity was limited in the C. sorokiniana XJK culture grown in the tubular reactor with dye, different lighting strategies were determined for 8 days (Fig. 3D). With the low light strategy (470 μmol (m² s)⁻¹), the maximum decolorization rate reached 86%. The natural light strategy could also obtain a high decolorization rate of 78%. However, too much light may induce photoinhibition and thus could be detrimental to the decolorization process. The decolorizing process was initially increased and then decreased, affecting the growth of the microalgae, which experienced three phases as follows: light limitation, light saturation, and light inhibition.³¹ It was found that an appropriately increased light intensity can enhance metabolism to boost the decolorization efficiency. Moreover, a light–dark regime for efficient photosynthesis can contribute to a good decolorizing efficiency.²⁴ The highest value of biomass and lipids (Fig. 3d) were 1259 mg L⁻¹ and 40%, respectively under 830 μmol (m² s)⁻¹. This might be due to CO₂ fixation being enhanced under the high light intensity and positively achieved biomass and neutral lipids.³² Therefore, a light intensity of 470 μmol (m² s)⁻¹ was applied to the subsequent runs of the optimization experiments.

3.2.5 The effect of CO₂ concentration on decolorization. The effect of different CO₂ concentrations on the dye removal using C. sorokiniana XJK was investigated under different CO₂/air (v/v) ratios, as shown in Fig. 3E. The decolorization rate curve had a lag phase over the first 3 days and a maximum of 83% for the decolorization rate was observed at a 1% CO₂/air (v/v) ratio over 6 days of cultivation. The result showed that³³ increasing the CO₂/air (v/v) ratio benefited the photosynthesis, growth and nutrient uptake, and therefore supplying an appropriate amount of CO₂ would enhance the decolorizing efficiency. It is necessary to notice that a lag phase was observed by the variation of pH under various CO₂ concentrations.³⁴ Biomass and lipid of C. sorokiniana XJK was obtained under supplemental of CO₂ (Fig. 3E), where 570 mg L⁻¹ of algal biomass and 43% of lipids were harvested for 6 days at 1% CO₂/air (v/v) ratio. The cultures of C. sorokiniana XJK supplied with a higher CO₂ concentration meant that an increased biomass concentration was obtained. However, cells in the presence of a 5–9% CO₂/air (v/v) ratio did not accumulate a better biomass than that under a lower CO₂/air (v/v) ratio due to the dead cells that are gradually produced when the nutrition was insufficient for growth, which was also supported by the work of Fan.³⁵ Hence, the optimal CO₂/air (v/v) ratio was 1%.

According to the degradation rate of dye using C. sorokiniana XJK, the optimum conditions were described as follows: dye conc. = 60 mg L⁻¹, biomass density = 8 × 10⁶ cell per mL, pH = 4, light intensity 470 μmol (m² s)⁻¹ and CO₂/air (v/v) ratio = 1%, which gave efficient decolorization and the desired decolorization rate was 83%.

3.3 Analysis

3.3.1 UV-vis analysis. The UV-vis spectrum was used for detecting the degradation of the dye in an algae-based treatment. Fig. 4a shows that the change of the wavelengths from 400 to 800 nm before and after C. sorokiniana XJK cultivation for 6 days. It was found that the characteristic absorption peak of disperse blue 2BLN had its maximum absorption near 560 nm in the visible region.⁵ With the growth of microalgae XJK, the maximum absorption peak disappeared at 560 nm. It was speculated that the chromophore group in the disperse blue 2BLN structure was destroyed or the intensity of the color was reduced by the behavior of Chlorella sorokiniana XJK.

Fig. 4 (a) The UV-vis spectrum of dye degradation: (m) untreated by algae; (n) treated by algae for 6 days; (b) The FTIR spectra of disperse blue 2BLN (pH = 10, biomass density = 9.9 × 10⁶ cell per mL, dye conc. = 60 mg L⁻¹).

3.3.2 FTIR analysis. The group absorption of disperse blue 2BLN and its degradation products were further estimated by FTIR spectroscopy during the growth of XJK (Fig. 4b). It was found that the aromatic ring near 797.1 cm⁻¹ disappeared and some chromophore groups attached to the destruction of aromatic structures, such as the peak at 1043.9 cm⁻¹ attributed to C–NH₂ vibration, were observed.³⁶ In addition, the peak near 2978.5 cm⁻¹ denoted C–H asymmetric stretching and the peak at 3418.7 cm⁻¹ represented the N–H vibration during the decolorization process.^37,38 It was found that some new peaks were observed after 6 days of culture. Furthermore, the anthraquinone rings were fragmented, which was indicated by the peak becoming weaker near 1625.3 cm⁻¹ due to the combination of stretching vibration between C=O and C=C.³⁷ Therefore, the large chromophore structure of disperse blue 2BLN was degraded into smaller molecules.

3.3.3 Ligninolytic enzyme analysis. Table 1 reports the ligninolytic enzyme activities obtained with C. sorokiniana XJK in dye medium. The results showed that laccase and MnP activities worked majorly on the removal of disperse blue 2BLN. Rovena Dosdall et al.³⁹ also reported the MnP activity from M. arcangeliana SBUG 1709 displayed the highest efficiency on the degradation process than other enzymes. During the incubation time, higher extracellular enzyme activities were examined on dye decolorization than that of the intracellular activities, where the laccase and MnP activity increased and reached 3.89 U mL⁻¹ and 4.86 U mL⁻¹, respectively. This was due to the extracellular enzymes involved in the oxidative reaction, which catalyzed the oxidation of phenolic compounds.⁴⁰ However, the LiP activity kept at a low level all the time with an opposite variation trend when compared to laccase and MnP. That is, the enzymes have different contributions to the entire biodegradation process, which was proven by the work of Martorell et al.⁴¹

Table 1 The ligninolytic enzyme activities of Chlorella sorokiniana XJK during the decolorization process

Time (d)	Laccase (U mL^−1)		MnP (U mL^−1)		LiP (U mL^−1)
	Intracellular	Extracellular	Intracellular	Extracellular	Intracellular	Extracellular
2	0.05	0.22	0.87	2.40	3.53	3.53
3	0.08	0.48	1.53	3.27	3.25	2.17
4	0.19	1.99	3.92	4.86	2.99	1.09
5	0.21	3.89	3.27	3.49	1.09	0.27

---

3.3.4 GC-MS analysis. The qualitative results of the GC-MS analysis and some of the identified intermediate products of disperse blue 2BLN during the biological treatment are reported in Table 2. The identified degradation products were as follows: dibutyl phthalate, 7,9-di-tert-butyl-1-oxaspiro[4,5]deca-6,9-diene-2,8-dione, 2,6-di-tert-butyl-p-benzoquinone, 2-(2-aminoanilino)ethanol, ethylbenzene, propylamine and ethyl acetate.

Table 2 GC-MS analysis of disperse blue 2BLN and its degradation products ([pH] = 10, biomass density = 9.9 × 10⁶ cell per mL⁻¹, [dye] = 60 mg L⁻¹)

Number	Product	Chemical structure	Formula	Molecular weight	Incubation time (days)
1	Disperse blue 2BLN		C14H9O4N2Cl	304.3	0
2	Dibuty phthalate		C16H22O4	278.34	1
3	7,9-Di-tert-butyl-1-oxaspiro[4,5]deca-6,9-diene-2,8-dione		C17H24O3	276.37	1
4	2,6-Di-tert-butyl-p-benzoquinone		C14H20O2	220.31	3
5	2-(2-Aminoanilino) ethanol		C8H12N2O	152	2
6	Ethylbenzene		C8H10	106.17	4
7	Ethyl acetate		C4H8O2	88.11	5
8	Propylamine		C3H9N	59	6

---

The degradation pathway was proposed starting from the products numbered 2 and 3, which may be isomers. The m/z value of 191 belonged to 2,6-di-tert-butyl-p-benzoquinone (number 4) generated by an open-loop, with the molecular formula being C₁₇H₂₄O₃ (Fig. 5a); the m/z value of 149 may extrapolate to 2-(2-aminoanilino) ethanol (number 5) (Fig. 5b). The m/z value of 88 and 59 belong to small molecular products. In addition, these products could further be degraded to give products such as CO₂ and H₂O (ref. 42) in other ways. However, numerous peaks were indistinguishable and these peaks may come from the secretion of Chlorella sorokiniana XJK by the degrading enzyme.⁴³

Fig. 5 The GC-MS spectra of the identified compounds in the disperse blue 2BLN solution of the algae-based treatment.

Disperse blue 2BLN was confirmed to be broken down by UV-vis, FTIR, GC-MS and enzymatic analysis. Experimental evidence for dye degradation: the characteristic absorption peak of dye had a dramatic decrease (UV-vis), the chromophore C=O was fractured (FTIR) and some new peaks were found (GC-MS). It is worth mentioning that the ligninolytic enzyme played an important role in dye degradation, particularly laccase and MnP.

4. Conclusions

The microalgae named C. sorokiniana XJK used in this study was exposed to disperse blue 2BLN under various operating conditions. C. sorokiniana XJK played a vital role in dye degradation, in addition to the accumulation of bioenergy. Under the optimal conditions, XJK can remove the dye with a maximum 83% decolorization rate and obtain 570 mg L⁻¹ of biomass and 43% of lipid content at same time. Above all, during the enzyme production function as well dye removal, laccase and MnP activity played essential roles during the decolorization of the dye by C. sorokiniana XJK. Moreover, the results of UV-vis, FTIR and GC-MS analysis observed that the anthraquinone structure was actually broken down into small molecules and some products were formed during the degradation process.

Acknowledgements

This study was financially supported by the National Natural Science Foundation of China (21466032) and Scientific Research Foundation for Changjiang Scholars of Shihezi University.

References

1. L. Liu, J. Zhang, Y. Tan, Y. Jiang, M. Hu, S. Li and Q. Zhai, Chem. Eng. J., 2014, 244, 9–18.
2. D. A. Yaseen and M. Scholz, Ecol. Eng., 2016, 94, 295–305.
3. S. Rangabhashiyam, E. Suganya, N. Selvaraju and L. A. Varghese, World J. Microbiol. Biotechnol., 2014, 30, 1669–1689.
4. N. Colin, A. Maceda-Veiga, N. Flor-Arnau, J. Mora, P. Fortuno, C. Vieira, N. Prat, J. Cambra and A. de Sostoa, Ecotoxicol. Environ. Saf., 2016, 132, 295–303.
5. E. Baldev, D. MubarakAli, A. Ilavarasi, D. Pandiaraj, K. A. Ishack and N. Thajuddin, Colloids Surf., B, 2013, 105, 207–214.
6. S. C. D. Sharma, Q. Sun, J. Li, Y. Wang, F. Suanon, J. Yang and C. P. Yu, Int. Biodeterior. Biodegrad., 2016, 112, 88–97.
7. V. Gopalakrishnan and D. Ramamurthy, BioMed Res. Int., 2014 DOI:10.1155/2014/529560.
8. G. Boduroglu, N. K. Kilic and G. Donmez, Environ. Technol., 2014, 35, 2410–2415.
9. M. Cerboneschi, M. Corsi, R. Bianchini, M. Bonanni and S. Tegli, Appl. Microbiol. Biotechnol., 2015, 99, 8235–8245.
10. C. R. Holkar, A. B. Pandit and D. V. Pinjari, Bioresour. Technol., 2014, 173, 342–351.
11. J. P. G. Rodriguez, D. E. Williams, I. D. Sabater, R. C. Bonugli-Santos, L. D. Sette, R. J. Andersen and R. G. S. Berlinck, RSC Adv., 2015, 5, 66360–66366.
12. R. Lu, L. Ma, F. He, D. Yu, R. Fan, Y. Zhang, Z. Long, X. Zhang and Y. Yang, Bioprocess Biosyst. Eng., 2015, 39, 381–390.
13. X. Yang, J. Zheng, Y. Lu and R. Jia, Environ. Sci. Pollut. Res., 2016, 23, 9585–9597.
14. V. V. Pathak, R. Kothari, A. K. Chopra and D. P. Singh, J. Environ. Manage., 2015, 163, 270–277.
15. H. C. Kim, W. J. Choi, S. K. Maeng, H. J. Kim, H. S. Kim and K. G. Song, Bioresour. Technol., 2014, 159, 128–135.
16. P. S. Kumar, J. Pavithra, S. Suriya, M. Ramesh and K. A. Kumar, Desalin. Water Treat., 2015, 55, 1342–1358.
17. D. G. Ülküye and D. Gönül, Desalin. Water Treat., 2013, 51, 3597–3603.
18. S. Katia, A. L. Lewis, V. Elie, M. Isabella and L. R. Nicoletta, J. Phycol., 2015, 51, 1172–1188.
19. N. Hatvani and I. Mécs, Process Biochem., 2001, 37, 491–496.
20. H. M. N. Iqbal, M. Asgher and H. N. Bhatti, BioResources, 2011, 6, 1273–1287.
21. H. Bermek, İ. Gülseren, K. Li, H. Jung and C. Tamerler, World J. Microbiol. Biotechnol., 2004, 20, 345–349.
22. E. G. Bligh and W. J. Dyer, Can. J. Biochem. Physiol., 1959, 37, 911–917.
23. D. Rajkumar, B. J. Song and J. G. Kim, Dyes Pigm., 2007, 72, 1–7.
24. R. Waqas, M. Arshad, H. N. Asghar and M. Asghar, Int. J. Agric. Biol., 2015, 17, 803–808.
25. Q. He, H. Yang and C. Hu, Bioresour. Technol., 2015, 198, 876–883.
26. M. T. Ale, M. Pinelo and A. S. Meyer, Prep. Biochem. Biotechnol., 2014, 44, 242–256.
27. P. Feng, Z. Deng, Z. Hu and L. Fan, Bioresour. Technol., 2011, 102, 10577–10584.
28. S. Mahalakshmi, D. Lakshmi and U. Menaga, Int. J. Environ. Res., 2015, 9, 735–744.
29. S. V. Mohan, Y. V. Bhaskar and J. Karthikeyan, Int. J. Environ. Pollut., 2004, 21, 211–222.
30. K. V. Kumar, S. Sivanesan and V. Ramamurthi, Process Biochem., 2005, 40, 2865–2872.
31. S. Xue, Z. Su and W. Cong, J. Biotechnol., 2011, 151, 271–277.
32. Y. Xiao, J. Zhang, J. Cui, X. Yao, Z. Sun, Y. Feng and Q. Cui, Algal Res., 2015, 11, 55–62.
33. G. Yildiz and Ş. Dere, Acta Oceanol. Sin., 2015, 34, 125–129.
34. K. G. Schulz, J. Barcelos e Ramos, R. E. Zeebe and U. Riebesell, Biogeosciences Discuss, 2009, 6, 4441–4462.
35. J. Fan, H. Xu, Y. Luo, M. Wan, J. Huang, W. Wang and Y. Li, Appl. Microbiol. Biotechnol., 2015, 99, 2451–2462.
36. J. M. Fanchiang and D. H. Tseng, Chemosphere, 2009, 77, 214–221.
37. C. W. M. Yuen, S. K. A. Ku, P. S. R. Choi, C. W. Kan and S. Y. Tsang, Res. J. Text. Apparel, 2005, 9, 26–38.
38. F. N. Memon and S. Memon, Pak. J. Anal. Environ. Chem., 2012, 13, 148–158.
39. R. Dosdall, V. Hahn, F. Preuß, H. Kreisel, J. Miersch and F. Schauer, Int. Biodeterior. Biodegrad., 2014, 96, 174–180.
40. G. McMullan, C. Meehan, A. Conneely, N. Kirby, T. Robinson, P. Nigam, I. M. Banat, R. Marchant and W. F. Smyth, Appl. Microbiol. Biotechnol., 2001, 56, 81–87.
41. M. M. Martorell, H. F. Pajot and L. I. C. de Figueroa, Int. Biodeterior. Biodegrad., 2012, 66, 25–32.
42. S. Saraf and B. Thomas, Process Saf. Environ. Prot., 2007, 85, 360–364.
43. J. S. Ye, H. Yin, J. Qiang, H. Peng, H. M. Qin, N. Zhang and B. Y. He, J. Hazard. Mater., 2011, 185, 174–181.

---