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Ability of fungi to degrade synthetic polymer nylon-6
Chemosphere 67 (2007) 2089–2095 www.elsevier.com/locate/chemosphere
Ability of fungi to degrade synthetic polymer nylon-6 Jozˇefa Friedrich b
a,*
, Polona Zalar b, Martina Mohorcˇicˇ a, Ursˇa Klun
a,1
, Andrej Krzˇan
a
a National Institute of Chemistry, Hajdrihova 19, SI-1000 Ljubljana, Slovenia Biotechnical Faculty, Biology Department, University of Ljubljana, Vecˇna pot 111, SI-1000 Ljubljana, Slovenia
Received 15 February 2006; received in revised form 13 September 2006; accepted 17 September 2006 Available online 25 January 2007
Abstract Fifty-eight fungi have been tested for their ability to degrade a recalcitrant synthetic polymer polyamide-6, generally known as nylon6. Most of them were isolated from a factory producing nylon-6. After preliminary screening, 12 strains were selected for submerged culture in a medium with nylon fibres as the only N-source. No degradation was observed with the isolates from the factory. Wood degrading fungi from a culture collection, however, degraded nylon after incubation for several weeks. Bjerkandera adusta disintegrated the fibres most efficiently, starting with the small transverse grooves, which deepened into cracks. The superficial layers crumbled to leave a thin inner core of the fibre, which finally broke down into fragments. The remaining insoluble part of the nylon showed a decrease in number average molecular mass from 16 900 to 5600 during a 60-day incubation. Its thermal properties, such as shifts in melting points and broadening of the melting endotherms, were altered. The reduction of the amount of nylon and the composition of the liquid phase indicated that part of the polymer was degraded into soluble products. After 50 days, the total nitrogen content of the soluble fraction was 10-fold higher than in the control sample. Manganese peroxidase, presumably responsible for the degradation, was detected in the liquid phase. The study shows that only white rot fungi are able to break down nylon-6. For the first time this polymer was shown to be disrupted by B. adusta. The extent of the biodegradation indicates its potential for application in nylon waste reduction. 2006 Elsevier Ltd. All rights reserved. Keywords: Biodegradation; Polyamide-6; White rot fungi; Bjerkandera adusta; Manganese peroxidase
1. Introduction The rapid development of the chemical industry in the last century has led to the production of 140 million tons of various polymers annually (Shimao, 2001). Many of these are not biodegradable and persist almost indefinitely in the environment (Kawai, 1995; Alexander, 1999; Shimao, 2001; Madigan et al., 2003). Their accumulation has triggered research to develop more readily degradable materials and to identify new methods for eliminating existing polymer waste (Kawai, 1995; Shimao, 2001). In general, biodegradation occurs either by hydrolysis or oxidation. Synthetic polyamides are particularly difficult to *
Corresponding author. Tel.: +386 1 476 0200; fax: +386 1 476 0300. E-mail address: [email protected] (J. Friedrich). 1 Present address: Medis, d.o.o., Brncˇicˇeva 1, SI-1000 Ljubljana, Slovenia. 0045-6535/$ - see front matter 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.chemosphere.2006.09.038
degrade. Their monomers are linked by amide bonds, similar to those in peptides and proteins, but which cannot be cleaved by proteolytic enzymes (Alexander, 1999). Aliphatic polyamides, such as nylon-66 and nylon-6, have been produced industrially since the late 1930s and continue to be important materials (Brydson, 1989). Nylon66 is a polymer of adipic acid and hexamethylene diamine, while nylon-6 is a polymer obtained by ring-opening polymerisation of e-caprolactam and has several commercial names including Perlon, Nylon, and Steelon (SzostakKotowa, 2004). Several attempts have been made to degrade these materials using microorganisms. The degree of microbial degradation has been shown to be lower the larger the molecule. A nylon-6 monomer (a cyclic form e-caprolactam or its linear form 6-aminohexanoic acid) is metabolized by numerous microorganisms, including the bacterial genera Pseudomonas, Achromobacter and Corynebacterium
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(Fukumura, 1966a,b; Shama and Wase, 1981) and the fungal genera Absidia, Aspergillus, Byssochlamis, Penicillium, Rhodotorula and Trichosporon (Shama and Wase, 1981). In an oligomer consisting of a small number of monomeric units of nylon-6, biodegradation was observed only with Pseudomonas and Flavobacterium (Negoro et al., 1994; Prijambada et al., 1995; Negoro, 2000). With increased polymerization to over 100 monomers (Prijambada et al., 1995), the material was still more recalcitrant. Although nylon-66 could be degraded by ligninolytic white rot fungi (Deguchi et al., 1997), until recently nylon-6 was generally regarded as a non-biodegradable polymer (Fukumura, 1966a; Gonsalves et al., 1991; Oppermann et al., 1998). The only reported microbial degradation was achieved by a thermophilic bacterium related to Bacillus pallidus (Tomita et al., 2003); enzymatic attack by a manganese peroxidase from an unidentified white rot fungus was also mentioned (Deguchi et al., 1998). Due to scarce information on nylon-6 biodegradability and due to its widespread use we believe that the potential of microorganisms to degrade it has not been adequately investigated. There is a surprising lack of studies using fungi in spite of the fact that they are known as a source of the greatest variety of enzymes (Lowe, 1992). We hypothesized that fungal strains could degrade the highly recalcitrant synthetic nylon-6 when subjected to nutritional pressure. Active strains or their isolated enzymes could then be exploited for waste polymer biodegradation and, potentially, for its recycling. The objectives of the present study were firstly to identify fungi possessing such activity and secondly to investigate the degradation process with the most promising strain. 2. Materials and methods 2.1. Isolation and taxonomic identification of fungi isolated from nylon-6 fibres Samples of nylon-6 fibres were collected from different locations in a factory producing nylon-6 where adapted strains were likely to be found. They were plated onto a selective agar medium (Kelley and Yaghmaie, 1988) in Petri dishes that contained nylon-6 instead of polycaprolactone, with the addition of chloramphenicol to prevent bacterial growth. The composition was as follows: 1.0 g nylon-6 powder with particles <0.5 mm diameter (washed with distilled water to eliminate monomers and soluble oligomers), 1.0 g (NH4)H2PO4, 0.1 g KCl, 0.1 g MgSO4 Æ 7H2O, 0.1 g CaCl2, 50 mg chloramphenicol, 6.0 g agar in 1 l distilled water. The pH was adjusted to 7.0. The medium was autoclaved at 121 C for 20 min and then poured into sterile Petri dishes. After inoculation the plates were incubated at room temperature (20–22 C) for one week. The resulting fungal colonies were purified by multiple transfers of conidia to fresh potato dextrose agar (Merck) plates containing nylon powder and chloramphenicol. Fungal strains were identified based on microscopic observation
of morphological characters (Ellis, 1971; Ellis, 1976; Domsch et al., 1980; Samson et al., 2000). Identification of Aspergillus strains was based on colony and spore morphology and on secondary metabolite profile (Frisvad and Thrane, 1993). 2.2. Screening fungal degradation activity on agar plates Microorganisms isolated in a factory producing nylon-6 and several other strains from the Culture Collection of the National Institute of Chemistry (MZKI), Ljubljana, Slovenia, were tested. All together, 58 fungal strains belonging to 30 genera were included in the study. For screening, the same medium was used as for isolation, but omitting the antibiotic. Inoculated plates were incubated at 30 C, the usual temperature for growing mesophilic fungi, for up to 10 days. The plates were observed each day for the appearance of a clear zone around the colony. When the zone became significant the agar plates were flooded with Lugol’s solution (Merck) for better resolution and the diameter was measured. 2.3. Submerged culture in a medium with nylon-6 fibres In the first experiment of submerged culture the same medium as in agar plate screening was used, but omitting the agar. Since poor growth and no nylon degradation were observed, a glucose-mineral medium (Deguchi et al., 1997) with slight modification was used in all subsequent experiments. The medium contained, per litre of distilled water: 10 g glucose, 1 g KH2PO4, 0.2 g NaH2PO4, 0.5 g MgSO4 Æ 7H2O, 0.1 g CaCl2, and 0.169 g (1 mM) MnSO4 Æ H2O. Fibres (20 lm diameter) of commercial grade nylon-6 (Mn = 16 900) provided by Yulon Aquafil, Ljubljana, Slovenia, were the only source of nitrogen. They were cut into approximately 2 cm long fragments and added in aliquots of 200 mg per 100 ml medium in 500ml Erlenmeyer flasks. The pH of the medium was 6.3. The flasks were autoclaved at 121 C for 20 min. Spores or mycelium scratched from an agar slant were suspended in 20 ml of sterile water and 5 ml of the suspension (approximately 106–107 spores ml 1) was used to inoculate 100 ml of the medium. Fermentations with each fungus were performed with two flasks in parallel. Non-inoculated flasks were used as abiotic controls throughout the study and were treated in the same way as the biotic samples. Cultivation was performed on a rotary shaker at 90 rpm at 30 C. Samples were taken periodically for analysis. Nylon-mycelium clusters were examined microscopically. The broth was filtered (filter paper ‘‘black ribbon’’; Schleicher & Schuell) and the liquid and solid phases analyzed separately. 2.4. Microscopic examination Growth and fibre degradation were observed every 10 days or monthly for the slower growing fungi. The samples
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were examined in optical and scanning electron microscopes (SEM). Samples for SEM were prepared by cutting off a small part (approximately 5 mm in diameter) of a nylon-mycelium cluster, fixing with glutaraldehyde and drying in CO2. For easier observation of the damaged fibres in some samples, mycelia were separated from the nylon fibres by soaking in a 1:1 mixture of 5 M NaOH and concentrated NaOCl (13% active Cl) (Gonsalves et al., 1991) until the mycelium detached (overnight or longer). The fibres were rinsed with distilled water and remaining mycelium was removed in an ultrasound bath (Elma, Darmstadt, Germany) at 100% ultrasound power for 20 min. The samples were air dried overnight. Abiotic controls were treated identically. After drying, all samples were gold-plated by sputtering and observed by SEM (Jeol JSMT220, Tokyo, Japan). An acceleration voltage of 15 kV was applied. 2.5. Viscosity and differential scanning calorimetry (DSC) The nylon was recovered from the filter cakes by soaking the biotic and abiotic samples in 3 ml trifluoroethanol and mixing to solubilize the nylon. The mixture was filtered through a 0.22 lm membrane filter (Durapore GVPP, Millipore) and the filtrate was poured into a Petri dish. The samples were kept overnight at room temperature in a fume hood to allow the solvent to evaporate. The resulting nylon film was weighed and used for analyses of viscosity and DSC. The relative viscosity (grel) of 1% nylon-6 solutions in H2SO4 was measured in an Ubbelhode viscometer (Visco/ Alpha, Middletown, NY) at 25 C using a standard procedure. The number average molecular mass (Mn) was calculated from the equation: Mn = 11 500 (grel 1) (Ciaperoni and Mula, 2001). Melting and crystallization behaviour of the samples was evaluated by DSC analysis using a Perkin Elmer Pyris 1 (Perkin Elmer, Wellesley, MA). A heating rate of 10 C min 1 was used in the range from 20 to 250 C under nitrogen. Melting temperatures (Tm) and enthalpies of fusion (DH) were measured during the first and second heating scans and the first cooling scan. 2.6. Elemental analysis of the liquid phase Soluble substances were concentrated by lyophilizing the filtrates (Micro Modulyo, Edwards, Cambridge, UK). Total nitrogen was determined by the standard method (ISO 11905-1:1997) using oxidative digestion with peroxodisulfate. The contents of carbon and sulphur were determined by the standard methods after dry combustion (ISO 10694:1995 and ISO 15178:2000, respectively). 2.7. HPLC analysis of soluble degradation products A mixture of the following compounds was used as a standard: 6-aminohexanoic acid, e-caprolactam and cyclic
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dimer, each of them at a concentration of 2.5 g l 1 in pure methanol. The same solvent was used for the lyophilized filtrates (17 g l 1) and the solubilized part of the sample was analyzed. The mobile phase consisted of acetonitriledeionized water solution using a concentration gradient from 15% to 50% (v/v). The analyses were performed on a Hewlett Packard 1100 using a 250 · 4.0 mm ID column thermostated at 22 C, packed with Nucleosil 120-5 C18, silicagel, octadecyl-modification, 5 lm particle size, 12 nm pore size, endcapped with HMDS (BIA, Ljubljana, Slovenia). The flow rate of the mobile phase was 1 ml min 1. Detection was at 200 nm. 2.8. Enzyme activity measurement Manganese peroxidase (MnP) and lignin peroxidase (LiP) activities in the liquid phase were determined as described (Mohorcˇicˇ et al., 2004). The substrates were 2,6-dimethoxyphenol (in the presence of Mn2+) and veratryl alcohol, respectively. Reactions were started by the addition of H2O2 and the formation of products was followed spectrophotometrically (Beckman, Fullerton, CA) up to 120 s. One unit of enzymatic activity was defined as 1 lmol of product formed per min. 3. Results 3.1. Fungi from nylon samples and agar plate screening Twenty-five fungi that grew on the medium with nylon-6 as sole carbon source were recovered from the factory producing nylon-6. Growth was restricted to few mycelial clusters, which sporulated and were in close contact with the nylon fibres. With one exception the cultures were asexually reproducing Ascomycetes: five Fusarium spp., four Aspergillus spp. and Penicillium spp., three Cladosporium spp. and Ulocladium spp., two Trichoderma spp., and one strain each of Gliocladium roseum, Pithomyces chartarum, Trichotecium roseum. There was also one Mucor hiemalis as a representative of Zygomycetes. The efficiency of nylon degradation was evaluated by the rapidity of formation and the diameter of the clear zone around the colonies in the opaque agar. Of the 58 fungi tested, 12 strains formed the zone in the first 10 days, seven being from the factory producing nylon-6 and five from the culture collection (Table 1). Surprisingly, clarification was not the most rapid with the strains supposed to be adapted to nylon, but with three ligninolytic wood degrading fungi, two Phanerochaete chrysosporium strains and Bjerkandera adusta. 3.2. Degradation of nylon-6 in submerged cultures Of the 12 most active fungi selected after agar plate screening, nine strains, one from each genus (Table 1, footnote), were tested for their ability to degrade nylon in submerged culture. The use of the same medium as in agar
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Table 1 Fungi selected after screening on agar plates: rapidity of formation of clarification zone and its diameter Fungal strain a
Phanerochaete chrysosporium MZKI B-223 Phanerochaete chrysosporium MZKI B-186 Bjerkandera adusta MZKI G-84a Ulocladium sp. MZKI B-1085a Ulocladium sp. MZKI B-1082 Aspergillus puniceus MZKI A-518a Penicillium sp. MZKI P-261a Mucor hiemalis MZKI B-1078a Aureobasidium pullulans MZKI B-802a Aureobasidium pullulans MZKI B-241 Fusarium sp. MZKI B-1083a Trichotecium roseum MZKI B-1080a a
Source
Time (d)
Diameter ± 2 (mm)
ATCC 24725, Rockville, USA BKM F-1767, Madison, USA MUCL 38680, Louvain la Neuve, B Nylon-producing factory Nylon-producing factory Nylon-producing factory Nylon-producing factory Nylon-producing factory Hypersaline water, saltpans ATCC 15233, Rockville, USA Nylon-producing factory Nylon-producing factory
2 2 3 5 5 5 5 6 8 8 8 9
63 58 50 57 53 44 36 42 62 47 44 36
Selected for nylon-6 degradation in submerged conditions.
plate screening resulted in very little growth and no nylon degradation, even after prolonged cultivation. Therefore, a slightly modified medium, previously found to enable fungal degradation of nylon-66 (Deguchi et al., 1997), was used in further experiments. Nylon-6 was the sole N source in the medium. To enable observation of structural changes it was used in the form of fibres instead of powder. Growth was slow to moderate and some of the fungi could be seen only by microscope. The nylon fibres became clumped in both control and inoculated flasks. The fungal mycelium was tightly interwoven with the nylon fibres to form a small number of clusters of uneven shapes and sizes. In some cultures hyphae were enveloped in a slimy material. Most of the fungi formed spores and the mycelium was often dark coloured. During the prolonged fermentation the volume of the liquid phase was reduced. In two months almost 50% of the liquid evaporated. Physical degradation of nylon-6 fibres could be observed by SEM. Only the white rot fungi, B. adusta and P. chrysosporium, attacked the nylon fibres. No degradation was observed either with the species isolated from the factory producing nylon-6 or with the other strains from the culture collection. 3.3. Nylon-6 degradation by B. adusta The first signs of fibre degradation were observed in cultures of the basidiomycete B. adusta. The initially smooth surface of the fibres was observed in some of them to develop superficial damage in the form of transverse grooves that were visible after 10 days of incubation (Fig. 1A). These grooves were evenly distributed along the fibre. The extent of erosion of the fibres however was not consistent throughout the culture. After 10 days it was estimated that only about 10% of the nylon fibres were damaged. The proportion of attacked fibres and the extent of degradation increased with exposure time. Grooves deepened into cracks that elongated and were densely distributed along the fibre (Fig. 1B). The diameter of the nylon fibres (initially 20 lm) decreased due to the superficial layer peeling off. After 40 days, the strongly eroded
thick outer layer of the fibres began to crumble, leading to a reduced diameter of 10–20 lm. After 50 days, a substantial amount of amorphous material was observed, which was attributed to degraded nylon particles and to mycelial autolysis. As the incubation continued, the diameter was reduced to 5 lm (Fig. 1C). These fibres lost most of their mechanical strength and were easily broken. The degradation of the polymer was reflected in a decrease of the amount of nylon in the filter cake from 0.173 g to 0.097 g per flask after 50-day incubation. The relative viscosity of the remaining nylon began to decrease 10 days after the samples were exposed to the fungus. The largest drop in viscosity was detected in the first 30 days from the initial value of 2.47 to 1.80. After 60 days the relative viscosity was 1.49. In the abiotic samples the viscosity diminished only slightly from 2.47 to 2.29 after 60 days, indicating an active role of the fungus in the degradation. The number average molecular mass (Mn) also decreased with fermentation time (Fig. 2), from 16 900 g mol 1 for the initial nylon-6, to 14 800 g mol 1 for the abiotic control after 60 days and 5600 g mol 1 for the material exposed to the fungus. Physical and chemical changes in the material were also followed by DSC. DSC curves were recorded in the melting and crystallization temperature range between 140 and 230 C for samples exposed to the fungus for 10, 40, and 60 days (Fig. 3). In all three scan series, peak shapes and temperatures changed with time. The crystallization temperature was reduced from 187 to 178 C in parallel with a reduction in the heat of fusion from 67 to 49 J g 1. The crystallization peak also became substantially broader and gradually developed a shoulder on the lower temperature side. The second heating scan initially exhibited a bimodal distribution with peaks at 211 and 219 C, consistent with the presence of two crystal forms of nylon-6 (a and c) (Schwartz, 1966). As the incubation proceeded, the peaks broadened and shifted toward lower temperatures, e.g. from 219 to 211 C for the higher temperature peak. The trend in the heats of fusion for the second heating scans was less consistent than in the case of crystallization. The first cooling and second heating DSC scans of the
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Fig. 2. Decrease in the number average molecular mass of nylon-6 during cultivation of B. adusta (d) compared to control (j).
Fig. 3. DSC curves of nylon-6 samples exposed to B. adusta for 10 (—), 40 (- - -) and 60 (Æ Æ Æ Æ) days.
Table 2 Total nitrogen, carbon and sulphur content in soluble dry matter and enzyme activities in the liquid phase after 50-day incubation of nylon-6 with B. adusta Liquid phase
Fig. 1. SEM micrographs of nylon-6 fibres degraded by B. adusta during submerged culture. Mycelium was removed prior to sample preparation. (A) The starting stages of degradation are seen as transverse grooves on the left-hand side fibre; an intact fibre is seen on the right-hand side (10 days). (B) After 30 days grooves are more numerous and deeper. (C) Strongly eroded fibres showing crumbling of the outer layers to leave a thin central core, which will eventually break down (60 days). In all micrographs the length of the bar is 10 lm.
abiotic control were virtually unchanged throughout the 60-day period. Soluble products in the filtrates were analyzed (Table 2). The total nitrogen content of the biotic sample after 50 days was 10-fold higher than in the corresponding abiotic sample. The content of carbon was at the same level as in the abiotic control while the content of sulphur in the biotic
Sample Control
MnP (U l 1)
LiP (U l 1)
N (mg g 1)
C (%)
S (%)
18.6 ± 0.2 0
0 0
8.01 ± 0.56 0.80 ± 0.06
32.6 ± 1.0 32.1 ± 1.0
0.90 ± 0.13 0.99 ± 0.14
MnP, manganese peroxidase; LiP, lignin peroxidase.
sample was lower. Measurement of enzyme activities showed the presence of MnP but not LiP. HPLC analysis of the liquid phase after 50 days showed that the content of glucose was substantially reduced in the biotic sample, while the content of 6-aminohexanoic acid was about 15% higher than in the abiotic control (Fig. 4). Unidentified new peaks were observed in the retention time interval from 2.25 to 4 min. The amount of cyclic monomer decreased by more than 50%, while cyclic dimer was no longer present.
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Fig. 4. HPLC analysis of nylon degradation products in the soluble dry matter after 50 days cultivation of B. adusta in a medium with nylon-6 fibres as the sole N source. G – glucose; LM – linear monomer (6aminohexanoic acid); CD – cyclic dimer; CM – cyclic monomer (ecaprolactam).
4. Discussion We focused on the polymer degrading ability of several fungi, with the expectation that adapted strains could be found in a factory producing nylon-6. The fungi recovered were anamorphic forms of Ascomycetes and one Zygomycete that are all common inhabitants of soil. Although after the first screening, seven isolates out of 25 appeared to be promising, none of them degraded nylon-6 fibres in submerged culture. It is possible that most of the isolated strains had been carried in from the outside environment and were not under any selective pressure to utilize the nylon. In submerged cultures no polymer degradation was detected in a medium used in agar plate screening. The inconsistency of these results with screening could be explained perhaps with the different cultivation conditions used (surface vs. submerged) or the adapted screening method was not adequate. After changing the medium, we detected nylon degradation by two ligninolytic white rot fungi, B. adusta and P. chrysosporium. Under these conditions, other fungi exhibited some growth but had no detectable effect on the nylon fibres. On the basis of these results and on the literature data we assumed that the reason could lie in different degradation mechanisms involved. The white rot fungi utilize oxidation by MnP (Deguchi et al., 1998), while anamorphic Ascomycetes usually degrade their substrates by hydrolysis. As B. adusta can produce MnP (Ward et al., 2004), whose activity was detected also in our experiments, we assume that non-specific oxidation was involved in this case. Though anamorphic Ascomycetes
and yeasts could metabolize the nylon-6 monomer (Shama and Wase, 1981) and glycine containing polyamides (Gonsalves et al., 1991), they were not effective against the nylon-6 polymer as shown in our experiments. We found that B. adusta was superior to the well-known ligninolytic fungus P. chrysosporium in its ability to degrade nylon fibres. While the common circular pattern of the grooves that developed initially on the fibres was similar to that observed with P. chrysosporium (Klun et al., 2003) and with an MnP from an unidentified white rot fungus (Deguchi et al., 1998), the breakdown of the fibres by B. adusta increased substantially. The incubation period during which physical disintegration of the fibres was observed coincided with a decrease in the relative viscosity and molecular mass of the remaining nylon (Fig. 2). After two months exposure to B. adusta, Mn was only 33% of the initial value, better than the ability of P. chrysosporium, which resulted in 50% of the starting Mn of the polymer (Klun et al., 2003). Structural modifications of nylon-6 are evident in the melting temperature and heat of fusion. These data provide an estimate of molecular stacking and mobility, that are both dependent on the molecular weight distribution in the sample and on the chemical structure, such as functional groups, end groups, etc. The DSC curves (Fig. 3) show that the trend in the heats of fusion for the second heating scans was less consistent than in the case of crystallization. This difference could be due to opposing effects resulting from degradation: less efficient packing due to an increasing number of end groups and increased mobility of the shorter chains. A similar effect of degradation, characterized by a broadening of melting peaks, their shift toward lower melting temperatures, and an inconsistent trend in heats of fusion, was observed by Janigova´ et al. (2002) for polyhydroxybutyrate degradation. The crystallization peak became substantially broader and gradually developed into a shoulder on the lower temperature side. These results are consistent with the progression of degradation in which crystallization is impeded by the increasing number of end groups that are formed during degradation and constitute irregularities. The reduction in viscosity and the change in DSC pattern reflect changes in the insoluble nylon part; the more severely degraded portion, however, was soluble. Analyses of the liquid phase support the assumption that nylon was partly solubilized. The increase in total nitrogen amount was substantial. Not all of it, however, necessarily originates from the nylon degradation products. Part of the nitrogen could be attributed to enzymes and mycelial autolysis. Nevertheless, it can be assumed that mycelial growth and protein production would not be possible without acquisition of nitrogen from the medium in which nylon-6 was the only N-source. HPLC analysis showed differences in composition of liquid phase in the biotic and abiotic samples. The slight increase in monomer content (6-aminohexanoic acid) could be due to partial chemical hydrolysis detected also in the
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abiotic control. The increase in the biotic sample could be ascribed to larger surface area of the damaged fibres exposed to the medium. Due to lack of standards we could not identify several peaks in the interval from 2.25 to 4 min. We assume that these substances are the main degradation products. In the abiotic control, the cyclic monomer and dimer, which are commonly present as impurities in nylon fibres, were attributed to their extraction during autoclaving. Of these two, only cyclic monomer, in reduced amounts, was detected in the biotic sample. 5. Conclusion A large number of fungi have been screened for their ability to degrade nylon-6. Most of the fungi were unable to attack the polymer, but only white rot Basidiomycetes were able to degrade nylon-6 when grown on nylon as the only N-source. Presumably, MnP was the responsible enzyme due to its non-specific oxidative action. The substantial extent of biodegradation of nylon-6 observed with B. adusta is the greatest so far observed. It was assumed that the insoluble polymer was partially solubilized and metabolized by the fungus. These results open up new prospects for the use of fungi for biodegradation of nylon-6 and possibly other environmental pollutants in the form of recalcitrant synthetic polymers. Acknowledgements The authors thank A. Gusˇtin from Yulon Aquafil for viscosity measurements, HPLC analyses and helpful support, and I. Sˇkraba and P. Sever for technical assistance. The authors also thank Prof. Majda Zˇigon for valuable discussions. This work was supported by project grants P4-0176, P20145, and P1-514-104 from The Ministry of Higher Education, Science and Technology of The Republic of Slovenia, and by the COST Action D25. References Alexander, M., 1999. Biodegradation and Bioremediation. Academic Press, San Diego, pp. 393–407. Brydson, J.A., 1989. Plastics Materials. Butterworths, London, p. 448. Ciaperoni, A., Mula, A., 2001. Chimica e tecnologia delle poliammidi. Pacini Editore, Pisa, p. 154. Deguchi, T., Kakezawa, M., Nishida, T., 1997. Nylon biodegradation by lignin-degrading fungi. Appl. Environ. Microb. 63, 329–331. Deguchi, T., Kitaoka, Y., Kakezawa, M., Nishida, T., 1998. Purification and characterization of a nylon-degrading enzyme. Appl. Environ. Microb. 64, 1366–1371. Domsch, K.H., Gams, W., Anderson, T.H., 1980. Compendium of Soil Fungi. Coordinating ed., Academic Press, London, p. 859. Ellis, M.B., 1971. Dematiaceous Hyphomycetes. Commonwealth Mycological Institute, Kew, p. 608.
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Ellis, M.B., 1976. More Dematiaceous Hyphomycetes. Commonwealth Mycological Institute, Kew, p. 507. Frisvad, J.C., Thrane, U., 1993. Liquid column chromatography of mycotoxins. In: Betina, V. (Ed.), Chromatography of Mycotoxins: Techniques and Applications. J. Chromatogr. Library 54, 254–372. Fukumura, T., 1966a. Bacterial breakdown of e-caprolactam and its cyclic oligomers. Plant Cell Physiol. 7, 93–104. Fukumura, T., 1966b. Splitting of e-caprolactam and other lactams by bacteria. Plant Cell Physiol. 7, 105–114. Gonsalves, K.E., Chen, X., Wong, T.K., 1991. Synthesis, characterization and biodegradation test of nylon 2/6 and nylon 2/6/6. J. Mater. Chem. 1, 643–647. Janigova´, I., Lacı´k, I., Choda´k, I., 2002. Thermal degradation of plasticized poly(3-hydroxybutyrate) investigated by DSC. Polym. Degrad. Stabil. 77, 35–41. Kawai, F., 1995. Breakdown of plastics and polymers by microorganisms. Adv. Biochem. Eng. Biot. 52, 151–194. Kelley, J., Yaghmaie, P.A., 1988. Screening of fungal strains employed in the testing of plastics materials. Int. Biodeterior. 24, 289–298, Cited in Paterson, R.R.M., Bridge, P.D., 1994. Biochemical Techniques for Filamentous Fungi, IMI Technical Handbooks No. 1, CAB International, Wallingford, Oxon, pp. 33–34. Klun, U., Friedrich, J., Krzˇan, A., 2003. Polyamide-6 degradation by a lignolytic fungus. Polym. Degrad. Stabil. 79, 99–104. Lowe, D.A., 1992. Fungal enzymes. In: Arora, D.K., Elander, R.P., Mukerji, K.G. (Eds.), Handbook of Applied Mycology, Fungal Biotechnology, vol. 4. Marcel Dekker, New York, USA, pp. 681–706. Madigan, M.T., Martinko, J.M., Parker, J., 2003. Brock Biology of Microorganisms, 10th ed. Prentice Hall, Upper Saddle River, NJ, pp. 676–679. Mohorcˇicˇ, M., Friedrich, J., Pavko, A., 2004. Decoloration of the diazo dye Reactive Black 5 by immobilised Bjerkandera adusta in a stirred tank bioreactor. Acta Chim. Slov. 51, 619–628. Negoro, S., Kato, K., Fujiyama, K., Okada, H., 1994. The nylon oligomer biodegradation system of Flavobacterium and Pseudomonas. Biodegradation 5, 185–194. Negoro, S., 2000. Biodegradation of nylon oligomers. Appl. Microbiol. Biot. 54, 461–466. Oppermann, F.B., Pickartz, S., Steinbu¨chel, A., 1998. Biodegradation of polyamides. Polym. Degrad. Stabil. 59, 337–344. Prijambada, I.D., Negoro, S., Yomo, T., Urabe, I., 1995. Emergence of nylon oligomer degradation enzymes in Pseudomonas aeruginosa PAO through experimental evolution. Appl. Environ. Microb. 61, 2020– 2022. Samson, R.A., Hoekstra, E.S., Frisvad, J.C., Filtenborg, O., 2000. Introduction to Food- and Airborne Fungi, sixth ed. Centraalbureau voor Schimmelcultures, Utrecht, p. 389. Schwartz, E., 1966. Bildung und Verhalten der Polyamide. In: Vieweg, R., Mu¨ller, A. (Eds.), Kunststoff-Handbuch, Band VI, Polyamide. Carl Hanser Verlag, Mu¨nchen, Germany, pp. 75–79. Shama, G., Wase, D.A.J., 1981. The biodegradation of e-caprolactam and some related compounds – a review. Int. Biodeterior. 17, 1–9. Shimao, M., 2001. Biodegradation of plastics. Curr. Opin. Biotech. 12, 242–247. Szostak-Kotowa, J., 2004. Biodeterioration of textiles. Int. Biodeter. Biodegr. 53, 165–170. Tomita, K., Hayashi, N., Ikeda, N., Kikuchi, Y., 2003. Isolation of a thermophilic bacterium degrading some nylons. Polym. Degrad. Stabil. 81, 511–514. Ward, G., Hadar, Y., Dosoretz, C.G., 2004. The biodegradation of lignocellulose by white rot fungi. In: Arora, D.K. (Ed.), Fungal Biotechnology in Agricultural, Food, and Environmental Applications. Marcel Dekker, New York, USA, pp. 393–407.
Ability of fungi to degrade synthetic polymer nylon-6 Jozˇefa Friedrich b
a,*
, Polona Zalar b, Martina Mohorcˇicˇ a, Ursˇa Klun
a,1
, Andrej Krzˇan
a
a National Institute of Chemistry, Hajdrihova 19, SI-1000 Ljubljana, Slovenia Biotechnical Faculty, Biology Department, University of Ljubljana, Vecˇna pot 111, SI-1000 Ljubljana, Slovenia
Received 15 February 2006; received in revised form 13 September 2006; accepted 17 September 2006 Available online 25 January 2007
Abstract Fifty-eight fungi have been tested for their ability to degrade a recalcitrant synthetic polymer polyamide-6, generally known as nylon6. Most of them were isolated from a factory producing nylon-6. After preliminary screening, 12 strains were selected for submerged culture in a medium with nylon fibres as the only N-source. No degradation was observed with the isolates from the factory. Wood degrading fungi from a culture collection, however, degraded nylon after incubation for several weeks. Bjerkandera adusta disintegrated the fibres most efficiently, starting with the small transverse grooves, which deepened into cracks. The superficial layers crumbled to leave a thin inner core of the fibre, which finally broke down into fragments. The remaining insoluble part of the nylon showed a decrease in number average molecular mass from 16 900 to 5600 during a 60-day incubation. Its thermal properties, such as shifts in melting points and broadening of the melting endotherms, were altered. The reduction of the amount of nylon and the composition of the liquid phase indicated that part of the polymer was degraded into soluble products. After 50 days, the total nitrogen content of the soluble fraction was 10-fold higher than in the control sample. Manganese peroxidase, presumably responsible for the degradation, was detected in the liquid phase. The study shows that only white rot fungi are able to break down nylon-6. For the first time this polymer was shown to be disrupted by B. adusta. The extent of the biodegradation indicates its potential for application in nylon waste reduction. 2006 Elsevier Ltd. All rights reserved. Keywords: Biodegradation; Polyamide-6; White rot fungi; Bjerkandera adusta; Manganese peroxidase
1. Introduction The rapid development of the chemical industry in the last century has led to the production of 140 million tons of various polymers annually (Shimao, 2001). Many of these are not biodegradable and persist almost indefinitely in the environment (Kawai, 1995; Alexander, 1999; Shimao, 2001; Madigan et al., 2003). Their accumulation has triggered research to develop more readily degradable materials and to identify new methods for eliminating existing polymer waste (Kawai, 1995; Shimao, 2001). In general, biodegradation occurs either by hydrolysis or oxidation. Synthetic polyamides are particularly difficult to *
Corresponding author. Tel.: +386 1 476 0200; fax: +386 1 476 0300. E-mail address: [email protected] (J. Friedrich). 1 Present address: Medis, d.o.o., Brncˇicˇeva 1, SI-1000 Ljubljana, Slovenia. 0045-6535/$ - see front matter 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.chemosphere.2006.09.038
degrade. Their monomers are linked by amide bonds, similar to those in peptides and proteins, but which cannot be cleaved by proteolytic enzymes (Alexander, 1999). Aliphatic polyamides, such as nylon-66 and nylon-6, have been produced industrially since the late 1930s and continue to be important materials (Brydson, 1989). Nylon66 is a polymer of adipic acid and hexamethylene diamine, while nylon-6 is a polymer obtained by ring-opening polymerisation of e-caprolactam and has several commercial names including Perlon, Nylon, and Steelon (SzostakKotowa, 2004). Several attempts have been made to degrade these materials using microorganisms. The degree of microbial degradation has been shown to be lower the larger the molecule. A nylon-6 monomer (a cyclic form e-caprolactam or its linear form 6-aminohexanoic acid) is metabolized by numerous microorganisms, including the bacterial genera Pseudomonas, Achromobacter and Corynebacterium
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(Fukumura, 1966a,b; Shama and Wase, 1981) and the fungal genera Absidia, Aspergillus, Byssochlamis, Penicillium, Rhodotorula and Trichosporon (Shama and Wase, 1981). In an oligomer consisting of a small number of monomeric units of nylon-6, biodegradation was observed only with Pseudomonas and Flavobacterium (Negoro et al., 1994; Prijambada et al., 1995; Negoro, 2000). With increased polymerization to over 100 monomers (Prijambada et al., 1995), the material was still more recalcitrant. Although nylon-66 could be degraded by ligninolytic white rot fungi (Deguchi et al., 1997), until recently nylon-6 was generally regarded as a non-biodegradable polymer (Fukumura, 1966a; Gonsalves et al., 1991; Oppermann et al., 1998). The only reported microbial degradation was achieved by a thermophilic bacterium related to Bacillus pallidus (Tomita et al., 2003); enzymatic attack by a manganese peroxidase from an unidentified white rot fungus was also mentioned (Deguchi et al., 1998). Due to scarce information on nylon-6 biodegradability and due to its widespread use we believe that the potential of microorganisms to degrade it has not been adequately investigated. There is a surprising lack of studies using fungi in spite of the fact that they are known as a source of the greatest variety of enzymes (Lowe, 1992). We hypothesized that fungal strains could degrade the highly recalcitrant synthetic nylon-6 when subjected to nutritional pressure. Active strains or their isolated enzymes could then be exploited for waste polymer biodegradation and, potentially, for its recycling. The objectives of the present study were firstly to identify fungi possessing such activity and secondly to investigate the degradation process with the most promising strain. 2. Materials and methods 2.1. Isolation and taxonomic identification of fungi isolated from nylon-6 fibres Samples of nylon-6 fibres were collected from different locations in a factory producing nylon-6 where adapted strains were likely to be found. They were plated onto a selective agar medium (Kelley and Yaghmaie, 1988) in Petri dishes that contained nylon-6 instead of polycaprolactone, with the addition of chloramphenicol to prevent bacterial growth. The composition was as follows: 1.0 g nylon-6 powder with particles <0.5 mm diameter (washed with distilled water to eliminate monomers and soluble oligomers), 1.0 g (NH4)H2PO4, 0.1 g KCl, 0.1 g MgSO4 Æ 7H2O, 0.1 g CaCl2, 50 mg chloramphenicol, 6.0 g agar in 1 l distilled water. The pH was adjusted to 7.0. The medium was autoclaved at 121 C for 20 min and then poured into sterile Petri dishes. After inoculation the plates were incubated at room temperature (20–22 C) for one week. The resulting fungal colonies were purified by multiple transfers of conidia to fresh potato dextrose agar (Merck) plates containing nylon powder and chloramphenicol. Fungal strains were identified based on microscopic observation
of morphological characters (Ellis, 1971; Ellis, 1976; Domsch et al., 1980; Samson et al., 2000). Identification of Aspergillus strains was based on colony and spore morphology and on secondary metabolite profile (Frisvad and Thrane, 1993). 2.2. Screening fungal degradation activity on agar plates Microorganisms isolated in a factory producing nylon-6 and several other strains from the Culture Collection of the National Institute of Chemistry (MZKI), Ljubljana, Slovenia, were tested. All together, 58 fungal strains belonging to 30 genera were included in the study. For screening, the same medium was used as for isolation, but omitting the antibiotic. Inoculated plates were incubated at 30 C, the usual temperature for growing mesophilic fungi, for up to 10 days. The plates were observed each day for the appearance of a clear zone around the colony. When the zone became significant the agar plates were flooded with Lugol’s solution (Merck) for better resolution and the diameter was measured. 2.3. Submerged culture in a medium with nylon-6 fibres In the first experiment of submerged culture the same medium as in agar plate screening was used, but omitting the agar. Since poor growth and no nylon degradation were observed, a glucose-mineral medium (Deguchi et al., 1997) with slight modification was used in all subsequent experiments. The medium contained, per litre of distilled water: 10 g glucose, 1 g KH2PO4, 0.2 g NaH2PO4, 0.5 g MgSO4 Æ 7H2O, 0.1 g CaCl2, and 0.169 g (1 mM) MnSO4 Æ H2O. Fibres (20 lm diameter) of commercial grade nylon-6 (Mn = 16 900) provided by Yulon Aquafil, Ljubljana, Slovenia, were the only source of nitrogen. They were cut into approximately 2 cm long fragments and added in aliquots of 200 mg per 100 ml medium in 500ml Erlenmeyer flasks. The pH of the medium was 6.3. The flasks were autoclaved at 121 C for 20 min. Spores or mycelium scratched from an agar slant were suspended in 20 ml of sterile water and 5 ml of the suspension (approximately 106–107 spores ml 1) was used to inoculate 100 ml of the medium. Fermentations with each fungus were performed with two flasks in parallel. Non-inoculated flasks were used as abiotic controls throughout the study and were treated in the same way as the biotic samples. Cultivation was performed on a rotary shaker at 90 rpm at 30 C. Samples were taken periodically for analysis. Nylon-mycelium clusters were examined microscopically. The broth was filtered (filter paper ‘‘black ribbon’’; Schleicher & Schuell) and the liquid and solid phases analyzed separately. 2.4. Microscopic examination Growth and fibre degradation were observed every 10 days or monthly for the slower growing fungi. The samples
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were examined in optical and scanning electron microscopes (SEM). Samples for SEM were prepared by cutting off a small part (approximately 5 mm in diameter) of a nylon-mycelium cluster, fixing with glutaraldehyde and drying in CO2. For easier observation of the damaged fibres in some samples, mycelia were separated from the nylon fibres by soaking in a 1:1 mixture of 5 M NaOH and concentrated NaOCl (13% active Cl) (Gonsalves et al., 1991) until the mycelium detached (overnight or longer). The fibres were rinsed with distilled water and remaining mycelium was removed in an ultrasound bath (Elma, Darmstadt, Germany) at 100% ultrasound power for 20 min. The samples were air dried overnight. Abiotic controls were treated identically. After drying, all samples were gold-plated by sputtering and observed by SEM (Jeol JSMT220, Tokyo, Japan). An acceleration voltage of 15 kV was applied. 2.5. Viscosity and differential scanning calorimetry (DSC) The nylon was recovered from the filter cakes by soaking the biotic and abiotic samples in 3 ml trifluoroethanol and mixing to solubilize the nylon. The mixture was filtered through a 0.22 lm membrane filter (Durapore GVPP, Millipore) and the filtrate was poured into a Petri dish. The samples were kept overnight at room temperature in a fume hood to allow the solvent to evaporate. The resulting nylon film was weighed and used for analyses of viscosity and DSC. The relative viscosity (grel) of 1% nylon-6 solutions in H2SO4 was measured in an Ubbelhode viscometer (Visco/ Alpha, Middletown, NY) at 25 C using a standard procedure. The number average molecular mass (Mn) was calculated from the equation: Mn = 11 500 (grel 1) (Ciaperoni and Mula, 2001). Melting and crystallization behaviour of the samples was evaluated by DSC analysis using a Perkin Elmer Pyris 1 (Perkin Elmer, Wellesley, MA). A heating rate of 10 C min 1 was used in the range from 20 to 250 C under nitrogen. Melting temperatures (Tm) and enthalpies of fusion (DH) were measured during the first and second heating scans and the first cooling scan. 2.6. Elemental analysis of the liquid phase Soluble substances were concentrated by lyophilizing the filtrates (Micro Modulyo, Edwards, Cambridge, UK). Total nitrogen was determined by the standard method (ISO 11905-1:1997) using oxidative digestion with peroxodisulfate. The contents of carbon and sulphur were determined by the standard methods after dry combustion (ISO 10694:1995 and ISO 15178:2000, respectively). 2.7. HPLC analysis of soluble degradation products A mixture of the following compounds was used as a standard: 6-aminohexanoic acid, e-caprolactam and cyclic
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dimer, each of them at a concentration of 2.5 g l 1 in pure methanol. The same solvent was used for the lyophilized filtrates (17 g l 1) and the solubilized part of the sample was analyzed. The mobile phase consisted of acetonitriledeionized water solution using a concentration gradient from 15% to 50% (v/v). The analyses were performed on a Hewlett Packard 1100 using a 250 · 4.0 mm ID column thermostated at 22 C, packed with Nucleosil 120-5 C18, silicagel, octadecyl-modification, 5 lm particle size, 12 nm pore size, endcapped with HMDS (BIA, Ljubljana, Slovenia). The flow rate of the mobile phase was 1 ml min 1. Detection was at 200 nm. 2.8. Enzyme activity measurement Manganese peroxidase (MnP) and lignin peroxidase (LiP) activities in the liquid phase were determined as described (Mohorcˇicˇ et al., 2004). The substrates were 2,6-dimethoxyphenol (in the presence of Mn2+) and veratryl alcohol, respectively. Reactions were started by the addition of H2O2 and the formation of products was followed spectrophotometrically (Beckman, Fullerton, CA) up to 120 s. One unit of enzymatic activity was defined as 1 lmol of product formed per min. 3. Results 3.1. Fungi from nylon samples and agar plate screening Twenty-five fungi that grew on the medium with nylon-6 as sole carbon source were recovered from the factory producing nylon-6. Growth was restricted to few mycelial clusters, which sporulated and were in close contact with the nylon fibres. With one exception the cultures were asexually reproducing Ascomycetes: five Fusarium spp., four Aspergillus spp. and Penicillium spp., three Cladosporium spp. and Ulocladium spp., two Trichoderma spp., and one strain each of Gliocladium roseum, Pithomyces chartarum, Trichotecium roseum. There was also one Mucor hiemalis as a representative of Zygomycetes. The efficiency of nylon degradation was evaluated by the rapidity of formation and the diameter of the clear zone around the colonies in the opaque agar. Of the 58 fungi tested, 12 strains formed the zone in the first 10 days, seven being from the factory producing nylon-6 and five from the culture collection (Table 1). Surprisingly, clarification was not the most rapid with the strains supposed to be adapted to nylon, but with three ligninolytic wood degrading fungi, two Phanerochaete chrysosporium strains and Bjerkandera adusta. 3.2. Degradation of nylon-6 in submerged cultures Of the 12 most active fungi selected after agar plate screening, nine strains, one from each genus (Table 1, footnote), were tested for their ability to degrade nylon in submerged culture. The use of the same medium as in agar
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Table 1 Fungi selected after screening on agar plates: rapidity of formation of clarification zone and its diameter Fungal strain a
Phanerochaete chrysosporium MZKI B-223 Phanerochaete chrysosporium MZKI B-186 Bjerkandera adusta MZKI G-84a Ulocladium sp. MZKI B-1085a Ulocladium sp. MZKI B-1082 Aspergillus puniceus MZKI A-518a Penicillium sp. MZKI P-261a Mucor hiemalis MZKI B-1078a Aureobasidium pullulans MZKI B-802a Aureobasidium pullulans MZKI B-241 Fusarium sp. MZKI B-1083a Trichotecium roseum MZKI B-1080a a
Source
Time (d)
Diameter ± 2 (mm)
ATCC 24725, Rockville, USA BKM F-1767, Madison, USA MUCL 38680, Louvain la Neuve, B Nylon-producing factory Nylon-producing factory Nylon-producing factory Nylon-producing factory Nylon-producing factory Hypersaline water, saltpans ATCC 15233, Rockville, USA Nylon-producing factory Nylon-producing factory
2 2 3 5 5 5 5 6 8 8 8 9
63 58 50 57 53 44 36 42 62 47 44 36
Selected for nylon-6 degradation in submerged conditions.
plate screening resulted in very little growth and no nylon degradation, even after prolonged cultivation. Therefore, a slightly modified medium, previously found to enable fungal degradation of nylon-66 (Deguchi et al., 1997), was used in further experiments. Nylon-6 was the sole N source in the medium. To enable observation of structural changes it was used in the form of fibres instead of powder. Growth was slow to moderate and some of the fungi could be seen only by microscope. The nylon fibres became clumped in both control and inoculated flasks. The fungal mycelium was tightly interwoven with the nylon fibres to form a small number of clusters of uneven shapes and sizes. In some cultures hyphae were enveloped in a slimy material. Most of the fungi formed spores and the mycelium was often dark coloured. During the prolonged fermentation the volume of the liquid phase was reduced. In two months almost 50% of the liquid evaporated. Physical degradation of nylon-6 fibres could be observed by SEM. Only the white rot fungi, B. adusta and P. chrysosporium, attacked the nylon fibres. No degradation was observed either with the species isolated from the factory producing nylon-6 or with the other strains from the culture collection. 3.3. Nylon-6 degradation by B. adusta The first signs of fibre degradation were observed in cultures of the basidiomycete B. adusta. The initially smooth surface of the fibres was observed in some of them to develop superficial damage in the form of transverse grooves that were visible after 10 days of incubation (Fig. 1A). These grooves were evenly distributed along the fibre. The extent of erosion of the fibres however was not consistent throughout the culture. After 10 days it was estimated that only about 10% of the nylon fibres were damaged. The proportion of attacked fibres and the extent of degradation increased with exposure time. Grooves deepened into cracks that elongated and were densely distributed along the fibre (Fig. 1B). The diameter of the nylon fibres (initially 20 lm) decreased due to the superficial layer peeling off. After 40 days, the strongly eroded
thick outer layer of the fibres began to crumble, leading to a reduced diameter of 10–20 lm. After 50 days, a substantial amount of amorphous material was observed, which was attributed to degraded nylon particles and to mycelial autolysis. As the incubation continued, the diameter was reduced to 5 lm (Fig. 1C). These fibres lost most of their mechanical strength and were easily broken. The degradation of the polymer was reflected in a decrease of the amount of nylon in the filter cake from 0.173 g to 0.097 g per flask after 50-day incubation. The relative viscosity of the remaining nylon began to decrease 10 days after the samples were exposed to the fungus. The largest drop in viscosity was detected in the first 30 days from the initial value of 2.47 to 1.80. After 60 days the relative viscosity was 1.49. In the abiotic samples the viscosity diminished only slightly from 2.47 to 2.29 after 60 days, indicating an active role of the fungus in the degradation. The number average molecular mass (Mn) also decreased with fermentation time (Fig. 2), from 16 900 g mol 1 for the initial nylon-6, to 14 800 g mol 1 for the abiotic control after 60 days and 5600 g mol 1 for the material exposed to the fungus. Physical and chemical changes in the material were also followed by DSC. DSC curves were recorded in the melting and crystallization temperature range between 140 and 230 C for samples exposed to the fungus for 10, 40, and 60 days (Fig. 3). In all three scan series, peak shapes and temperatures changed with time. The crystallization temperature was reduced from 187 to 178 C in parallel with a reduction in the heat of fusion from 67 to 49 J g 1. The crystallization peak also became substantially broader and gradually developed a shoulder on the lower temperature side. The second heating scan initially exhibited a bimodal distribution with peaks at 211 and 219 C, consistent with the presence of two crystal forms of nylon-6 (a and c) (Schwartz, 1966). As the incubation proceeded, the peaks broadened and shifted toward lower temperatures, e.g. from 219 to 211 C for the higher temperature peak. The trend in the heats of fusion for the second heating scans was less consistent than in the case of crystallization. The first cooling and second heating DSC scans of the
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Fig. 2. Decrease in the number average molecular mass of nylon-6 during cultivation of B. adusta (d) compared to control (j).
Fig. 3. DSC curves of nylon-6 samples exposed to B. adusta for 10 (—), 40 (- - -) and 60 (Æ Æ Æ Æ) days.
Table 2 Total nitrogen, carbon and sulphur content in soluble dry matter and enzyme activities in the liquid phase after 50-day incubation of nylon-6 with B. adusta Liquid phase
Fig. 1. SEM micrographs of nylon-6 fibres degraded by B. adusta during submerged culture. Mycelium was removed prior to sample preparation. (A) The starting stages of degradation are seen as transverse grooves on the left-hand side fibre; an intact fibre is seen on the right-hand side (10 days). (B) After 30 days grooves are more numerous and deeper. (C) Strongly eroded fibres showing crumbling of the outer layers to leave a thin central core, which will eventually break down (60 days). In all micrographs the length of the bar is 10 lm.
abiotic control were virtually unchanged throughout the 60-day period. Soluble products in the filtrates were analyzed (Table 2). The total nitrogen content of the biotic sample after 50 days was 10-fold higher than in the corresponding abiotic sample. The content of carbon was at the same level as in the abiotic control while the content of sulphur in the biotic
Sample Control
MnP (U l 1)
LiP (U l 1)
N (mg g 1)
C (%)
S (%)
18.6 ± 0.2 0
0 0
8.01 ± 0.56 0.80 ± 0.06
32.6 ± 1.0 32.1 ± 1.0
0.90 ± 0.13 0.99 ± 0.14
MnP, manganese peroxidase; LiP, lignin peroxidase.
sample was lower. Measurement of enzyme activities showed the presence of MnP but not LiP. HPLC analysis of the liquid phase after 50 days showed that the content of glucose was substantially reduced in the biotic sample, while the content of 6-aminohexanoic acid was about 15% higher than in the abiotic control (Fig. 4). Unidentified new peaks were observed in the retention time interval from 2.25 to 4 min. The amount of cyclic monomer decreased by more than 50%, while cyclic dimer was no longer present.
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Fig. 4. HPLC analysis of nylon degradation products in the soluble dry matter after 50 days cultivation of B. adusta in a medium with nylon-6 fibres as the sole N source. G – glucose; LM – linear monomer (6aminohexanoic acid); CD – cyclic dimer; CM – cyclic monomer (ecaprolactam).
4. Discussion We focused on the polymer degrading ability of several fungi, with the expectation that adapted strains could be found in a factory producing nylon-6. The fungi recovered were anamorphic forms of Ascomycetes and one Zygomycete that are all common inhabitants of soil. Although after the first screening, seven isolates out of 25 appeared to be promising, none of them degraded nylon-6 fibres in submerged culture. It is possible that most of the isolated strains had been carried in from the outside environment and were not under any selective pressure to utilize the nylon. In submerged cultures no polymer degradation was detected in a medium used in agar plate screening. The inconsistency of these results with screening could be explained perhaps with the different cultivation conditions used (surface vs. submerged) or the adapted screening method was not adequate. After changing the medium, we detected nylon degradation by two ligninolytic white rot fungi, B. adusta and P. chrysosporium. Under these conditions, other fungi exhibited some growth but had no detectable effect on the nylon fibres. On the basis of these results and on the literature data we assumed that the reason could lie in different degradation mechanisms involved. The white rot fungi utilize oxidation by MnP (Deguchi et al., 1998), while anamorphic Ascomycetes usually degrade their substrates by hydrolysis. As B. adusta can produce MnP (Ward et al., 2004), whose activity was detected also in our experiments, we assume that non-specific oxidation was involved in this case. Though anamorphic Ascomycetes
and yeasts could metabolize the nylon-6 monomer (Shama and Wase, 1981) and glycine containing polyamides (Gonsalves et al., 1991), they were not effective against the nylon-6 polymer as shown in our experiments. We found that B. adusta was superior to the well-known ligninolytic fungus P. chrysosporium in its ability to degrade nylon fibres. While the common circular pattern of the grooves that developed initially on the fibres was similar to that observed with P. chrysosporium (Klun et al., 2003) and with an MnP from an unidentified white rot fungus (Deguchi et al., 1998), the breakdown of the fibres by B. adusta increased substantially. The incubation period during which physical disintegration of the fibres was observed coincided with a decrease in the relative viscosity and molecular mass of the remaining nylon (Fig. 2). After two months exposure to B. adusta, Mn was only 33% of the initial value, better than the ability of P. chrysosporium, which resulted in 50% of the starting Mn of the polymer (Klun et al., 2003). Structural modifications of nylon-6 are evident in the melting temperature and heat of fusion. These data provide an estimate of molecular stacking and mobility, that are both dependent on the molecular weight distribution in the sample and on the chemical structure, such as functional groups, end groups, etc. The DSC curves (Fig. 3) show that the trend in the heats of fusion for the second heating scans was less consistent than in the case of crystallization. This difference could be due to opposing effects resulting from degradation: less efficient packing due to an increasing number of end groups and increased mobility of the shorter chains. A similar effect of degradation, characterized by a broadening of melting peaks, their shift toward lower melting temperatures, and an inconsistent trend in heats of fusion, was observed by Janigova´ et al. (2002) for polyhydroxybutyrate degradation. The crystallization peak became substantially broader and gradually developed into a shoulder on the lower temperature side. These results are consistent with the progression of degradation in which crystallization is impeded by the increasing number of end groups that are formed during degradation and constitute irregularities. The reduction in viscosity and the change in DSC pattern reflect changes in the insoluble nylon part; the more severely degraded portion, however, was soluble. Analyses of the liquid phase support the assumption that nylon was partly solubilized. The increase in total nitrogen amount was substantial. Not all of it, however, necessarily originates from the nylon degradation products. Part of the nitrogen could be attributed to enzymes and mycelial autolysis. Nevertheless, it can be assumed that mycelial growth and protein production would not be possible without acquisition of nitrogen from the medium in which nylon-6 was the only N-source. HPLC analysis showed differences in composition of liquid phase in the biotic and abiotic samples. The slight increase in monomer content (6-aminohexanoic acid) could be due to partial chemical hydrolysis detected also in the
J. Friedrich et al. / Chemosphere 67 (2007) 2089–2095
abiotic control. The increase in the biotic sample could be ascribed to larger surface area of the damaged fibres exposed to the medium. Due to lack of standards we could not identify several peaks in the interval from 2.25 to 4 min. We assume that these substances are the main degradation products. In the abiotic control, the cyclic monomer and dimer, which are commonly present as impurities in nylon fibres, were attributed to their extraction during autoclaving. Of these two, only cyclic monomer, in reduced amounts, was detected in the biotic sample. 5. Conclusion A large number of fungi have been screened for their ability to degrade nylon-6. Most of the fungi were unable to attack the polymer, but only white rot Basidiomycetes were able to degrade nylon-6 when grown on nylon as the only N-source. Presumably, MnP was the responsible enzyme due to its non-specific oxidative action. The substantial extent of biodegradation of nylon-6 observed with B. adusta is the greatest so far observed. It was assumed that the insoluble polymer was partially solubilized and metabolized by the fungus. These results open up new prospects for the use of fungi for biodegradation of nylon-6 and possibly other environmental pollutants in the form of recalcitrant synthetic polymers. Acknowledgements The authors thank A. Gusˇtin from Yulon Aquafil for viscosity measurements, HPLC analyses and helpful support, and I. Sˇkraba and P. Sever for technical assistance. The authors also thank Prof. Majda Zˇigon for valuable discussions. This work was supported by project grants P4-0176, P20145, and P1-514-104 from The Ministry of Higher Education, Science and Technology of The Republic of Slovenia, and by the COST Action D25. References Alexander, M., 1999. Biodegradation and Bioremediation. Academic Press, San Diego, pp. 393–407. Brydson, J.A., 1989. Plastics Materials. Butterworths, London, p. 448. Ciaperoni, A., Mula, A., 2001. Chimica e tecnologia delle poliammidi. Pacini Editore, Pisa, p. 154. Deguchi, T., Kakezawa, M., Nishida, T., 1997. Nylon biodegradation by lignin-degrading fungi. Appl. Environ. Microb. 63, 329–331. Deguchi, T., Kitaoka, Y., Kakezawa, M., Nishida, T., 1998. Purification and characterization of a nylon-degrading enzyme. Appl. Environ. Microb. 64, 1366–1371. Domsch, K.H., Gams, W., Anderson, T.H., 1980. Compendium of Soil Fungi. Coordinating ed., Academic Press, London, p. 859. Ellis, M.B., 1971. Dematiaceous Hyphomycetes. Commonwealth Mycological Institute, Kew, p. 608.
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