Degradation of lignosulfonic and tannic acids by ligninolytic soil fungi cultivated under icroaerobic conditions

FOOD/FEED SCIENCE AND TECHNOLOGY

Degradation of lignosulfonic and tannic acids by ligninolytic soil fungi cultivated under icroaerobic conditions

Isis Serrano Silva ^* ; Cristiano Ragagnin de Menezes; Elisangela Franciscon; Eder da Costa dos Santos; Lucia Regina Durrant

Departamento de Ciência de Alimentos; Faculdade de Engenharia de Alimentos; Universidade Estadual de Campinas; Rua: Monteiro Lobato, 80; C. P.: 6121; 3083-862; Campinas - SP - Brasil

ABSTRACT

Soil fungi were evaluated regarding their ability to degrade lignin-related compounds by producing the ligninolytic enzymes. Lignosulfonic and tannic acids were used as sole carbon sources during 30 days under microaerobic and very-low-oxygen conditions. The fungi produced lignin-peroxidase, manganese-peroxidase and laccase . Expressive degradations was observed by C₁₈ reversed-phase HPLC, indicating the biodegradation potential of these fungi, showing more advantages than obligate anaerobes to decontaminate the environment when present naturally.

Key words: lignosulfonic acid, tannic acid, ligninolytic enzymes, microaerobic conditions, soil fungi

RESUMO

Fungos isolados de solo foram avaliados quanto à habilidade em degradarem compostos derivados de lignina pela produção de enzimas ligninolíticas. Os ácidos lignosulfônicos e tânico foram usados separadamente como única fonte de carobono para cultivo dos fungos em 30 dias sob condições microaeróbias. Os fungos foram capazes de crescer e usar tais compostos como fonte de carbono e mostraram produção de lignina-peroxidase, manganês-peroxidase e lacase. Degradações expressivas dos ácidos lignosulfônico e tânico foram verificadas por Cromatografia Liquida de Alta Eficiência (CLAE), indicando grande potencial de uso em processos de biorremediação de macromoléculas aromáticas similares à lignina em ambientes naturais sob condições baixas de oxigenação.

INTRODUCTION

Lignin is a complex polyphenolic macromolecule and a mechanism of support in vegetable tissues, protecting them against the microorganisms. In the nature, white-rot basidiomycete fungi and actinomycetes have ability to degrade lignin (Kajikawa et al., 2000) during the secondary metabolism or in the absence of nitrogen, carbon or sulphur. However, lignin is not degraded as sole carbon and energy sources, requiring additional co-substrate as cellulose, hemicellulose (de Jong et al., 1994) or glucose (Lee, 1997). Several aromatic compounds such as biphenyl, phenols, anysols, diaryl-ethers present in the lignin structure contribute to their irregular and recalcitrant nature.

Lignosulphonate compounds are highly hydrophilic macromolecules due to sulfonic acid content, and just a minor fraction of lignosulfonic acid is removed from the water phase by absorption onto organic carbon, sediments or water treatment. They cause aquatic unbalance since surface sunlight is not available to the photosynthetic algae and bacteria as a result of an intense brown-color into the water. Lignosulfonic acids contain a great amount of wood-carbohydrates (e.g. glucose, xylose), nitrogen and mineral nutrients that make of this compound a good alternative as substrate for ligninolytic fungi (Aitken and Logan, 1996). White-rot fungi have been extensively studied for their ability to degrade the lignocellulose by production of ligninolytic enzymes (lignin- and manganese-peroxidases, laccase) and cellulases (Kapich et al., 2004).

Tannins are polyphenolic compounds very abundant in the vascular plants and very recalcitrant into the nature presenting great resistance to microbial attack (Hernes et al., 2001) since these compounds are capable of complexing with enzymes involved in the degradation process (Bending and Read, 1996a). Tannic acid belongs to hydrolyzed-tannins group, containing glucose molecule linked to gallic acid, showing anti-nutritional property in herbivorous caused by its ability to associate with proteins, cellulose, hemicellulose, pectin and minerals, retarding digestion (McSweeney et al., 2001) and also forming a complex with other substances containing nitrogen (e.g. aminoacids, peptides, quitin, nucleic acids).

Fungal degradation is considered the main mechanism of biological cycling of lignin, causing oxidation of propyl chains, demethylation of 3- and 5-methoxyl groups and aromatic ring cleavage. Brown-rot fungi generate cathecol derivatives as result of demethylation (Filley et al., 2000). Some white-rot fungi have the ability to selectively remove expressive amount of lignin with slight losses of hemicellulose and cellulose, but many others are not selective for lignin degradation.

Phanerochaete chrysosporium is extensively used as a model for the biodegradation of lignin and chemical pollutants by white-rot fungi. The initial depolymerization of lignin is catalyzed by extracellular peroxidases which are responsible for the degradation of several compounds related to lignin structure (Gold and Alic, 1993). The degradation of lignin and phenolic molecules by selective fungi makes them suitable for industrial applications to alter or remove these compounds (Blanchette, 1988). Non-basidiomycete fungi are also good degraders of wood, producing non-specific extracellular ligninolytic enzymes such as manganese peroxidase and laccase, when grown in media containing lignin-related compounds.

Most fungi need oxygen, at least in small amounts in order to grow; however, some anaerobic phycomycetous fungi capable of degrading lignocellulosic materials are present in the rumen microbiota (Durrant, 1996). Tolerance for low levels of oxygen has also been observed in some aquatic fungi, yeasts and several filamentous fungi such as Fusarium oxysporum, Mucor hiemalis, Aspergillus fumigatus which can fermentate sugars (Singh et al., 1992). The aim of this work was to study the ability of soil fungi to degrade lignosulfonic and tannic acids and produce ligninolytic enzymes under microaerobic conditions.

MATERIALS AND METHODS

Filamentous fungal cultures such as Achremonium sp, Aspergillus sp, Fusarium oxysporum, Trichoderma sp, Verticillium sp, Trichocladium canadense and and a basidiomycetous H2 (Durrant, 1996) were used in this work. Cultures were cultivated in 0.2% lignosulfonic acid (LA) or tannic acids (TA) as sole carbon sources in a defined liquid medium containing per liter: KH₂PO₄ (1 g); (NH₄)₂HPO₄ (0.5 g); CaCl₂ (0.3 g); L- cysteine chloridrate as reducing oxygen agent (1 g); resazurine 0.1% as reduction indicator dye (1 ml); micronutrients solution (1 ml); vitamins solution (0.5 ml), pH 5.5-6.0. Both micronutrients and vitamins solutions were previously sterilized by filtration (Millipore membrane, 0.22 µm). The microelements solution was composed by the following salts per liter: ammonium tartarate (0.22 g; manganese sulphate (0.66 g); iron sulphate (0.15 g); cobalt sulphate (0.10 g); zinc sulphate (0.10 g); cooper sulphate (0.64 g) and aluminium potassium sulphate (10.0 g). The vitamins solution was provided also per liter by thiamine (5.0 g); biotine (2.0 g); nicotinic acid (5.0 g); cianocobalamine (0.1 g); folic acid (2.0 g); riboflavine (5.0 g); pyridoxine (10.0 g); DL- pantetonate Ca⁺² (5.0 g) e thiotic acid (5.0 g). Fungi were grown previously on Malt Extract Agar plates for 7-10 days at 30ºC, and 3 x 1cm² of mycelia were used as inoculum in each experimental flask. Microaerobic and very-low-oxygen conditions were performed using L-cisteyne chloridrate and resazurine into the medium as described previously, together with microaerobac plates (Probac-Brazil) into the jars. The oxygen generation was 5 through 15% O₂, and less than 1% when very-low-oxygen condition was applied. Oxidation-reduction reaction produced hydrogen and carbon dioxide after the addition of distilled water into microaerobac or anaerobac plates and jars were sealed immediately afterwards. Microaerobic and very-low-oxygen conditions were then verified by observing the disappearance of resazurin color into the control non-inoculated liquid medium.

Samples were taken on the 30^th days for both the conditions for the enzymatic activities analyses and to verify LA and TA degradations. Abiotic controls were prepared in the same manner as the samples, except inoculating the mycelia. Supernatants of each sample were centrifuged under refrigeration at 5,541 g for 15 minutes and stored at -20 ºC for subsequent HPLC analyses. Enzymatic assays were carried out immediately after each sampling. All the experiments were run in duplicate, including abiotic controls.

Lignin-peroxidase activity (LiP, EC1.11.1.14) was assayed by the determination of veratryl alcohol oxidation to veratryl aldeide (Tien and Kirk, 1993 - modified). Briefly, in a 1ml- cuvette containing culture supernatant (0.6 ml) was added veratryl alcohol (2 mM) in tartaric buffer (0.4 M - pH 3; 0.2 ml), followed by 2 ml H₂O₂ (2 mM). After 10 minutes of reaction, absorbances of samples were read at 310 nm.

Manganese-peroxidase activity (MnP, EC1.11.1.1) was determined by the phenol red oxidation (Kuwahara et al., 1984 - modified), reacting in a cuvette the culture supernatant (0.5 ml), 0.25 M sodium lactate (0.1 ml), 2mM MgSO₄ (0.05 ml), 5% bovine albumine (0.2 ml), 2mM H₂O₂ in succinate buffer( 0.2 M -pH 4.5; 0.05 ml), with phenol red 1% (0.1 ml). After 5 minutes of reaction, 40µl of 2N NaOH was added to stop the reaction, and samples absorbances were read at 610 nm.

Laccase (Lac, EC1.10.3.2) was measured via H₂O₂-independent oxidation of syringaldazine to its quinone form (Szklarz et al., 1989) by adding to 0.6 ml of culture supernatant in a cuvette, 0.2 ml of 0.05M citrate-phosphate buffer (pH 5), 0.1 ml of distilled water, and 0.1 ml of seringaldazine in 0.1% absolute ethanol. After 10 minutes, the reaction was measured at 525 nm.

Enzyme activities were expressed in U l^-1 that means µmol of substrate oxidized per minute of reaction, according to the followed equation: Enzymatic activity (U l^-1) = (ΔAbs 10⁶) (ε R T)^-1, when ΔAbs is the difference between final and initial absorbencies, ε is the molar extinction coefficient of each enzyme (LiP- ε₃₁₀ = 9300 M^-1.cm^-1; MnP- ε₆₁₀ = 4460 M^-1.cm^-1; Lac- ε₅₂₅ = 65000 M^-1.cm^-1), R is the volume in ml of supernatant, and T is the time of reaction in minutes.

HPLC analyses were performed using LC6A chromatographer (Shimadzu) and column Microsorb MV-100 (0.46 cm x 15 cm x 0.2 mm) on C₁₈ reverse-phase (Varian). Supernatant of samples containing LA were diluted (1:20), filtered in 0.22 µm Millipore membrane and injected (20 µl). TA was extracted from the cultures supernatant using propanol in 5% heptane (v v^-1), followed by centrifugation and solvent evaporation under nitrogen flow and re-suspension of the extract in the mobile phase. LA and TA degradations were determined by isocratic elution in methanol: water (30:70), flow rate of 0.1 ml min^-1 and UV absorbance detector set at 254 nm. Percentages of degradation were calculated as: % degradation = [(PAH control – PAH sample)/PAH control] x 100.

RESULTS AND DISCUSSION

Soil fungi were capable of growing in liquid medium containing LA as sole carbon source during 30 days, followed by acidification of the media, reaching pH values less than 3.0 (data not shown). The ligninolytic enzymes might have been produced in this period, and the drop of pH values suggested the production of acid metabolites by the fungi under microaerobic conditions (Pavarina and Durrant, 2002). Some color changes were also observed possibly due to the production of colored metabolites, or reaction with medium components, or with the respective carbon sources (Silva, 2002). The brown color into the medium might also be a consequence of re-polymerization of degradation products by extracellular phenoloxidases (Bending and Read 1996 a,b).

In the present study, Verticillium sp reached the best rates of degradation when grown in LA (27.5 and 32.5%), as well as higher levels of LiP (18.12 and 8.70 U l^-1) under microaerobic and very-low-oxygen conditions, respectively, as indicated in Figures 1 and ^3a.

Aspergillus sp also showed good degradation of LA (26.5%), and highest level of laccase compared with the other strains studied. The basidiomycetous H2 showed high activities of LiP, MnP and laccase and low LA degradation under microaerobic condition.

The microaerobic fungus Trichocladium canadense reached good degradation under low oxygenation condition (Figures 1 and ^3b); however, only activity of laccase was detected in this period. It occurred probably due to a previous production of more specific enzymes under low rates of oxygen that were able to degrade LA. On other hand, it was possible that ligninolytic enzymes could help this degradation; however, not as a key-hole in the process.

Trichoderma sp and Achremonium sp did not show any activity of ligninolytic enzymes under very-low-oxygen condition; however, LA degradations were detected after 30 days of cultivation (10.1 and 9.3%, respectively).

Filamentous fungi, especially Aspergillus niger and Penicillium ssp were capable of growing in TA as sole carbon source. Some non-white-rot fungi have been described as the very good degraders of TA in static cultures (Bhan et al., 1997; Bhat et al., 1998; Falconi, 1998). In this study, Achremonium sp, Fusarium oxysporum and Verticillium sp showed best visual growths in TA as sole carbon source under microaerobic condition with considerable drop of pH values (data not shown). High levels of MnP were detected in supernatants of Trichoderma sp, Achremonium sp and H2 strain. Nevertheless, these fungi showed the lowest degradation under microaerobic condition (Fig.2).

It is believed that MnP could be produced as part of the fungi constitutive metabolism with no direct relation with TA degradation in this case. Some fungi such as Chaetomium, Fusarium, Rhizoctonia, Cylindrocarpon, Trichoderma and Candida are also able to degrade tannic compounds (Saxena et al., 1995). Fusarium oxysporum, Aspergillus sp (Fig. 4a) and Trichocladium canadense (Fig. 4b) were the best degraders of TA producing ligninolytic enzymes, excepting LiP by Aspergillus sp. The strain H2 showed activities of MnP and laccase under very-low-oxygen; however, did not show any TA degradation.

During the lignin degradation, several simultaneous reactions are involved in the oxidation, polymerization and complete mineralization, producing and accumulating metabolic intermediates (Hernandéz-Péres et al., 1998). Aromatic rings cleavage is the main mechanism to degrade the lignin in sulfate-reducing sediments (Dittmar and Lara 2001). Lignin degradation by anaerobic fungi from rumen of herbivorous resulted in a breakdown of ester- and eter- linkages from phenolic units forming phenolic acids with no degradation of the most abundant linkages between lignin and wood polysaccharides (McSweeney et al., 1994).

Similar results of LA degradation by non-basidiomycete fungi under non-oxygenated condition were also observed in other studies, verifying the degradation under microaerobic condition (Falconi, 1998; Sette, 1997). However, these results were lower than results confirmed in the present study. The microaerobic fungi Trihcocladium canadense and H2 were also able to grow in LA and produce laccase and other peroxidases (Durrant, 1996). Fungi may produce different enzymes depending on genetic characteristics or environment conditions (Eggen, 1999),. Therefore, it is likely that other enzymes are activated under this condition and ligninolytic enzymes may also help in LA degradation.

CONCLUSIONS

From results, it could be concluded that ligninolytic enzymes were produced during the degradation process, acting on metabolic intermediates. Apparently tannase and some other specific enzymes to degrade TA were probably activated to promote the degradation of TA under limitation of oxygen. Further studies are required regarding others enzymatic systems involved in this process.

ACKNOWLEDGEMENTS

The authors would like to thank FAPESP (Brazilian Foundation for Research Support, Sao Paulo, Brazil) for the master degree scholarship to Ms Isis Silva and for the financial support to this work.

Received: September 25, 2007; Revised: June 30, 2008; Accepted: August 13, 2009.

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Publication Dates

• Publication in this collection
  11 June 2010
• Date of issue
  June 2010

History

• Received
  25 Sept 2007
• Reviewed
  30 June 2008
• Accepted
  13 Aug 2009