This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Oceguera-Cervantes, A.
Right arrow Articles by Loza-Tavera, H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Oceguera-Cervantes, A.
Right arrow Articles by Loza-Tavera, H.
Agricola
Right arrow Articles by Oceguera-Cervantes, A.
Right arrow Articles by Loza-Tavera, H.

 Previous Article  |  Next Article 

Applied and Environmental Microbiology, October 2007, p. 6214-6223, Vol. 73, No. 19
0099-2240/07/$08.00+0     doi:10.1128/AEM.01230-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Characterization of the Polyurethanolytic Activity of Two Alicycliphilus sp. Strains Able To Degrade Polyurethane and N-Methylpyrrolidone{triangledown}

Alejandro Oceguera-Cervantes,1 Agustín Carrillo-García,1 Néstor López,2 Sandra Bolaños-Nuñez,3 M. Javier Cruz-Gómez,2 Carmen Wacher,3 and Herminia Loza-Tavera1*

Departamento de Bioquímica, Facultad de Química, Universidad Nacional Autónoma de México, Cd. Universitaria, 04510 México, D.F. México,1 Departamento de Ingeniería Química, Facultad de Química, Universidad Nacional Autónoma de México, Cd. Universitaria, 04510 México, D.F. México,2 Departamento de Alimentos y Biotecnología, Facultad de Química, Universidad Nacional Autónoma de México, Cd. Universitaria, 04510 México, D.F. México3

Received 1 June 2007/ Accepted 1 August 2007


arrow
ABSTRACT
 
Two bacterial strains (BQ1 and BQ8) were isolated from decomposed soft foam. These were selected for their capacity to grow in a minimal medium (MM) supplemented with a commercial surface-coating polyurethane (PU) (Hydroform) as the carbon source (MM-PUh). Both bacterial strains were identified as Alicycliphilus sp. by comparative 16S rRNA gene sequence analysis. Growth in MM-PUh showed hyperbolic behavior, with BQ1 producing higher maximum growth (17.8 ± 0.6 mg·ml–1) than BQ8 (14.0 ± 0.6 mg·ml–1) after 100 h of culture. Nuclear magnetic resonance, Fourier transform infrared (IR) spectroscopy, and gas chromatography-mass spectrometry analyses of Hydroform showed that it was a polyester PU type which also contained N-methylpyrrolidone (NMP) as an additive. Alicycliphilus sp. utilizes NMP during the first stage of growth and was able to use it as the sole carbon and nitrogen source, with calculated Ks values of about 8 mg·ml–1. Enzymatic activities related to PU degradation (esterase, protease, and urease activities) were tested by using differential media and activity assays in cell-free supernatants of bacterial cultures in MM-PUh. Induction of esterase activity in inoculated MM-PUh, but not that of protease or urease activities, was observed at 12 h of culture. Esterase activity reached its maximum at 18 h and was maintained at 50% of its maximal activity until the end of the analysis (120 h). The capacity of Alicycliphilus sp. to degrade PU was demonstrated by changes in the PU IR spectrum and by the numerous holes produced in solid PU observed by scanning electron microscopy after bacterial culture. Changes in the PU IR spectra indicate that an esterase activity is involved in PU degradation.


arrow
INTRODUCTION
 
Polyurethane (PU) was developed by Otto Bayer as a substitute for rubber at the beginning of World War II (1938). Due to its range of properties, the polymer is widely used, for example, in liquid coatings and paints, adhesives, sealants, flexible and rigid foams, and elastomers (7). Since PU is such a versatile polymer, PU production has increased, but this has brought with it the problem of safe disposal. Each year, more than 5 million tons of shredder residue containing different plastics and PU foams is generated in the United States and Canada (6). Several mechanical processes, such as regrinding, flexible foam bonding, adhesive pressing, and compression molding, as well as chemical techniques, such as glycolysis, hydrolysis, pyrolysis, and hydrogenation, are used for the recovery of the starting materials or in the production of other PU types (8). Recently, the development of new strategies based on the utilization of biopolymers such as poly[(R)-hydroxyalkanoic acids] (21), the enzymatic polymerization of polyesters and degradation of PU (28), and the discovery of microorganisms (fungi and bacteria) able to utilize PU as a source of carbon and nitrogen (12, 31, 32) is leading the move to a greener chemical industry.

A number of bacterial strains, such as Corynebacterium sp., Pseudomonas fluorescens, P. chlororaphis, and Bacillus subtilis, have been reported to grow in PU media supplemented with yeast extract or glucose (18, 19, 20, 22, 35). However, only Comamonas acidovorans TB-35 has the ability to attack solid PU and use it as a carbon source (31). Protease, urease, and esterase activities have been associated with the degradation of polyester PU by fungi and bacteria (12, 18, 31, 32, 34, 39). Polyurethanase protease activities have been reported for Pseudomonas fluorescens and P. chlororaphis (19, 36), polyurethanase lipase activity has been detected in Bacillus subtilis strains (35), and polyurethanase esterase activities have been reported for Corynebacterium sp., Comamonas acidovorans TB-35, and P. chlororaphis (20, 22, 30). Membrane-bound esterase activity produced by the Comamonas acidovorans TB-35 strain, which is able to attack solid PU (1, 30), is the best characterized of these.

N-Methylpyrrolidone (NMP), the lactam of 4-methylaminobutyric acid, is an organic solvent for natural (waxes and resins) and synthetic (polystyrene, polyesters, and polyvinyl chloride) polymers. It is chemically stable, and it is used in a variety of industrial chemical reactions and in paints, surface coatings, paint strippers, and cleaners, as well as for the recovery of pure hydrocarbons in petrochemical processing, desulfurization of gases, and manufacture of electronic equipment. It has been used as an additive in the manufacture of PU finishes for the last 25 years. Additionally, it is used in agrochemicals, such as insecticides, fungicides, herbicides, seed treatment products, and bioregulators. It has been regarded as having a favorable toxicological and environmental profile in comparison to other, more toxic solvents (5), for which reason it has been used extensively. However, recent findings indicate that NMP has aquatic toxicity for Daphnia magna, with a 50% lethal concentration at 24 h of 1.23 mg·liter–1, and only certain pesticides have greater toxicity than this compound (23). NMP has been reported to be teratogenic in rats when administered orally (38) but not by inhalation (16, 25, 37), although a reduction in fetal weight has been observed to occur in the absence of obvious maternal effects at certain doses (37, 43). It has also been correlated with stillbirth in a woman working in a laboratory where NMP was used routinely (42) and with the poisoning of an agricultural worker by an insecticide in which it was a major component (44). Although NMP was listed in 1994 in the Environmental Protection Agency Emergency Planning and Community Right-to-Know Act, section 313(d)(2)(B) (14), as having serious or irreversible chronic health effects, it is still used broadly. Its extensive use in different industries, its excellent miscibility with water and most organic solvents, and its high percutaneous absorption (4) make the use of NMP potentially hazardous to humans (23). Even though NMP has been reported as being biodegradable (5, 10), there have been no reports on the identification of microorganisms with such a capacity.

In this paper we describe the isolation and characterization of two Alicycliphilus sp. strains featuring two remarkable capacities: to use NMP as a carbon and nitrogen source and to degrade PU with an esterase activity. We also identified an inducible extracellular esterase activity which might be responsible for the polyurethanolytic activity.


arrow
MATERIALS AND METHODS
 
Isolation of bacterial strains able to grow in Hydroform as the only carbon source.
Samples of deteriorated PU were collected from an open-air refuse dump (Bordo de Xochiaca, Nezahualcóyotl, Edo. de México, México). The samples were flexible PU foam pieces collected from the ground or deteriorated upholstery. They were carried in plastic bags to the laboratory, and portions were introduced to 125-ml Erlenmeyer flasks containing 50 ml minimal medium (MM) at pH 7.2 (composition in g·liter–1: KH2PO4, 2.0; K2HPO4, 7.0; NH4NO3, 1.0; MgSO4·H2O, 0.1; ZnSO4·7H2O, 0.001; CuSO4·7H2O, 1 x 10–4; FeSO4·7H2O, 0.01; MnSO4·6H2O, 0.002) plus Hydroform (1.0%, vol/vol; 10.5 mg·ml–1) as the carbon source (MM-PUh). Hydroform (PolyForm, México) is an aliphatic PU water-based coating similar to Impranil DLN (Bayer Corp.), which has been employed in other experimental work to characterize polyurethanolytic bacteria (2, 19, 20, 35, 36). The inoculated media were incubated for 5 days at 37°C and at 200 rpm. Aliquots (100 µl) were taken at different incubation times, inoculated into MM-PUh agar plates, and incubated at 37°C for 48 h, after which colonies were isolated and subcultured in the same medium. In order to obtain axenic cultures, subculturing was performed alternately in MM-PUh agar and Luria-Bertani (LB) agar. Bacterial stocks were prepared in glycerol (30%) and stored at –70°C.

Identification of the isolated strains.
Bacterial strains able to grow in MM-PUh were characterized using biochemical tests (MacConkey and potato agars, nitrate broth, Salmonella/Shigella, cetrimide, sulfate-indol-motility, motility-indol-ornitine, urea, arginine, and oxide fermentation glucose media) (9, 27) and an API 2ONE test (bioMérieux GmbH, Nurtingen, Germany). The two bacterial strains that showed the most growth in MM-PUh were selected for study and identified by 16S rRNA gene sequence comparison. DNA was extracted from colonies growing in LB agar (24), except that after the phenol-chloroform extraction procedure, polysaccharides were eliminated by adding 110.7 µl of 5 M NaCl plus 88.5 µl of cetyltrimethylammonium bromide (10%, wt/vol) in solution in 0.7 M NaCl. The mixture was shaken thoroughly and incubated at 65°C for 10 min (3). DNA integrity was analyzed by agarose electrophoresis and the amount and purity estimated by spectrophotometry. A fragment of the 16S rRNA gene (corresponding to positions 30 to 750 of the Escherichia coli 16S rRNA gene) was amplified by PCR with the following conserved primers close to the 3' and 5' ends of the fragment, respectively: pA (5' AGAGTTTGATCCTGGCTCAG 3') and pH (5' AAGGAGGTGATCCAGCCGCA 3'). A single strand of the PCR product was directly sequenced using an automatic DNA sequencer (ABI Prism 3100 genetic analyzer; Foster City, CA) with primers gamma (5' ACTGCTGCCTCCCGTAGGAG 3'), pD* (5' GTATTACCGCGGCTGCTG 3'), ANTIgamma (5' CTCCTACGGGAGGCAGCAGT 3'), and ANTIKK (5' CGTGCCAGCAGCCGCGGTAAT 3'). Identification was based on partial sequence comparisons (720 bp). Closely matching sequences were found in the GenBank database using the BLAST algorithm (http://www.ncbi.nlm.nih.gov). Sequences were aligned using BioEdit sequence alignment editor and visually inspected. Phylogenetic analysis was performed using the MEGA program, version 3.1. The Kimura two-parameter method was used to calculate evolutionary distances, and dendrograms were constructed according to the neighbor-joining method. Bootstrap values were calculated using 500 replicates.

Bacterial growth.
A portion of glycerol stock was inoculated into LB agar. After 2 days of incubation at 37°C, one isolated colony was inoculated into 5 ml LB broth and incubated at 37°C and 200 rpm. After 12 h, 5 ml of fresh LB was inoculated with cells of this culture to reach an optical density at 660 nm (OD660) of 0.1. The new culture was incubated at 37°C and 200 rpm for the time needed to reach exponential growth, approximately 6 h. This culture was used to start growth in MM-PUh by inoculating fresh medium with the quantity of cells needed to reach an OD660 of 0.02. Incubation was conducted at 37°C and 200 rpm. Similar cultures were established to monitor the increase in wet weight. For this, 50 ml of MM-PUh inoculated with cells to give an OD660 of 0.02 was introduced into glass tubes (16 by 150 mm) in 5-ml aliquots and incubated at 37°C and 200 rpm. At various times over 5 days, the contents of one tube were poured into a preweighed tube and centrifuged at 5,000 x g for 5 min. The wet weight of the cell pellet at each time interval was measured, and each experiment was performed in triplicate. To determine the dependence of bacterial growth on Hydroform concentration, the wet weights of bacterial cultures growing in MM with different amounts of Hydroform (from 10.5 to 105 mg·ml–1) were measured. The growth kinetic parameters maximum growth rate (µmax) and Ks were calculated from the exponential phase of growth (between 0 and 12 h) and a Monod plot produced. Growth kinetic parameters were also calculated for bacteria growing in MM with various concentrations of NMP (from 0.5 to 25 mg·ml–1) as the sole carbon and nitrogen source. All experiments were performed in triplicate, and comparisons between measured values were performed using the Student t test.

Hydroform characterization.
The components present in Hydroform were separated by vacuum distillation. The system was heated to 120°C, and two liquid fractions, at 30°C and 60°C, and a residual solid were obtained. The liquid fractions were analyzed by gas chromatography coupled to a mass spectrometry detector (GC-MS) (HP 6890 series and HP 5973, respectively) with a 5% phenyl methyl siloxane capillary column (30 m by 0.25 mm by 20 µm). The oven was heated from 50°C to 100°C at 20°C·min–1. Helium was used as the carrier gas at a flow rate of 1.8 ml·min–1, and the injection volume was 1 µl. The residual solid was analyzed by nuclear magnetic resonance (1H NMR and 13C NMR) (Unity Inova model, 300 MHz; Varian) and by infrared spectroscopy (IRS) (4,000 to 400 cm–1) in a Fourier transform infrared (FTIR) 1605 Perkin-Elmer apparatus.

Identification of bacterial enzymatic activities related to PU degradation. (i) Microbiological tests.
Three different microbiological media were used to determine which enzymatic activity (protease, urease, or esterase activity) related to PU degradation Alycicliphilus sp. strains were able to produce. To detect protease activity, the strains were subcultured in YES medium plates (composition in g·liter–1: K2HPO4, 1.0; KH2PO4, 0.5; MgSO4·7H2O, 0.5; MnCl2·4H2O, 0.001; CuCl2·2H2O, 1.4 x 10–5; ZnCl2, 1.1 x 10–5; CoCl·6H2O, 2 x 10–5; Na2MoO4·2H2O, 1.3 x 10–5; FeCl3·6H2O, 7.5 x 10–5; agar, 15; and gelatin, 0.02; pH 7). After 48 h of incubation at 37°C, the plates were stained with Coomassie blue R-250 (0.1% in acetic acid:methanol, 3:1, vol/vol) for 30 min and washed with acetic acid:methanol (3:1, vol/vol). A clear, nonstained zone of degradation was observed around colonies exhibiting proteolytic activity (19). To detect urease activity, the strains were subcultured in Christensen's urea agar (11) (composition in g·liter–1: urea, 20; agar, 15; NaCl, 5; KH2PO4, 2; peptone, 1; glucose, 1; and phenol red, 0.012). After 48 h of incubation at 37°C, a pink coloration was observed around colonies showing urease activity. To identify esterase activity, the strains were subcultured in Tween 80 agar (41) (composition in g·liter–1: peptone, 10; NaCl, 5; CaCl2, 0.1; and agar, 12 [plus Tween 80, 10.0 ml·liter–1]) and incubated for 120 h at 37°C. A white precipitation halo was observed around colonies showing esterase activity. A Pseudomonas sp. culture was used as a positive control for the three tests.

(ii) Enzymatic assays.
To determine the presence of extracellular enzymatic activities related to PU degradation, protease, urease, and esterase activities were measured in cell-free supernatant (SN) portions of MM-PUh bacterial cultures, after different times of incubation, obtained by centrifugation at 5,000 x g for 5 min (Sorval RC5 PLUS). Linearity of the enzymatic assays had been tested previously by using different reaction times and enzyme amounts. Protease activity was determined spectrophotometrically by measuring casein hydrolysis (26). Casein buffer solution (400 µl) (1% casein in 50 mM potassium phosphate buffer, pH 7) was preincubated at 37°C for 3 min. The reaction was started with 100 µl of enzyme extract, and after 10 min of incubation, 500 µl of 100% trichloroacetic acid was added. The tubes were centrifuged at 5,000 x g for 5 min. The SNs were collected and the absorbance measured at 280 nm with a UV-VIS Ultrospec 2000 instrument (Pharmacia-Biotech). For blank tubes, the enzyme extract was added after trichloroacetic acid. A protease activity standard curve was constructed by incubating different amounts (1 to 20 mU) of fungal proteinase K (product no. 5530UA; Gibco BRL) under similar conditions. Urease activity was determined by a phenol hypochloride assay (45). Potassium phosphate buffer (180 µl) (50 mM, pH 7) plus 100 µl of 5 M urea was preincubated at 37°C for 3 min. The reaction was started by the addition of 20 µl of enzyme extract and incubated for 3 min. After incubation, 100 µl of phenol solution {7% phenol, 0.34% sodium nitroprussiate [Na2Fe(CN)5NO·2H2O]} was added, followed by 200 µl of NaClO solution (0.37 M NaOH, 1 M Na2HPO4·12H2O, 1% NaClO; pH 12). Immediately after the addition of these solutions, the absorbance was read at 639 nm with a UV-visible Ultrospec 2000 instrument (Pharmacia-Biotech). Blank tubes were prepared for each sample by addition of the enzyme after the NaClO solution. Proteus mirabilis culture SN was used as the enzyme source for positive controls. To determine the amount of NH4+ released in the reaction, a standard curve was performed by using NH4Cl and phenol and NaClO solutions as described above. Esterase activity was measured spectrophotometrically by a p-nitrophenyl acetate (p-NPA) hydrolysis method (13). The reaction was carried out at 37°C at 1 ml final volume containing 250 µl reaction buffer (200 mM phosphate, pH 7.5), 100 µl enzyme extract, 400 µl of water, and 250 µl of a 20 mM solution of p-NPA in acetonitrile. The reaction was started by adding the substrate and the A405 recorded 30 s after the start of the reaction and at further 1-min intervals for 5 min. p-NPA chemical hydrolysis was calculated from blank experiments performed with SN from a noninoculated MM-PUh. The measurements of chemical hydrolysis were subtracted from those from experiments with enzyme preparation. A standard curve was constructed to measure the hydrolysis of p-NPA by incubating different concentrations of the reagent under conditions similar to those used for the enzymatic assay but using 100 µl 1 M NaOH.

GC-MS analysis of the MM-PUh SN after bacterial culture.
Cell-free SNs of MM-PUh cultures (2 ml) taken at different incubation times were extracted with chloroform (1 ml). The organic phase was recovered using a Pasteur pipette and transferred to a fresh tube. A small amount of sodium sulfate was added to eliminate water. The sample was analyzed in a GC-MS system as described in "Hydroform characterization" above. The concentration of NMP in the samples was quantified based on a standard curve (0.6 to 24.6 mM) constructed with pure NMP (catalog no. M6762; Sigma). The measurements were performed in triplicate.

IRS-FTIR analysis of PU after bacterial growth.
Three samples of solid PU were obtained from the SNs of 5-day MM-PUh cultures inoculated with the bacterial strains by using noninoculated MM-PUh cultures as the control. The solid obtained from SNs by drying in vacuo was pelleted and analyzed by IRS-FTIR as described in "Hydroform characterization" above.

SEM.
Solid PU sheets 3 mm thick were prepared by pouring Hydroform into a petri dish, drying in an oven at 65°C, and then cutting into 1-cm2 pieces. Five pieces were incubated in MM-PUh inoculated with each bacterial strain for 15 days at 37°C. The same number of polymerized PU pieces was incubated in noninoculated MM-PUh under similar conditions. After incubation, sheets were recovered, rinsed with water, gold coated by evaporation, and analyzed by scanning electron microscopy (SEM) using a JEOL 5900 LB microscope at x500 magnification.


arrow
RESULTS
 
Two Alicycliphilus sp. strains were isolated by using MM-PUh.
Inoculation of deteriorated flexible PU foam pieces, collected from a refuse dump, in MM-PUh and subsequent rounds of selection in MM-PUh agar allowed us to obtain several bacterial isolates. The two bacterial strains (BQ1 and BQ8) which showed the best growth in MM-PUh were selected for study. According to biochemical characterization, both strains were identified as members of the Comamonadaceae family. However, although both were positive for reduction of nitrate, adipic and malic acid assimilation, and cytochrome oxidase, small differences, such as the arginine dihydolase test positive for strain BQI and negative for BQ8 and BQ8 positive but BQ1 negative for assimilation of potassium gluconate, were observed. Analysis of the partial 16S rRNA gene sequence confirmed the observation that strains BQ1 (GenBank accession no. EF463077) and BQ8 (GenBank accession no. EF463078) cluster with the family Comamonadaceae of Betaproteobacteria and showed that, although they are closely related to the members of this family Acidovorax sp. and Comamonas sp., they cluster with the genus Alicycliphilus (Fig. 1). Strain BQ1 had 100% identity and strain BQ8 99% identity with Alicycliphilus sp. strain R-24611 (GenBank accession no. AM084014) and 99% and 98%, respectively, with strain K601 (GenBank accession no. AJ418042). Strain K601, Alicycliphilus denitrificans, is the type species, a new genus and new species (29). Strain R-24611 was isolated from activated sludge and cultivated in growth media with ethanol or succinate as the carbon source and with nitrate as the nitrogen source (17) (Fig. 1). Some branching patterns (those with bootstrap values lower than 75%) cannot be considered stable. It is possible that higher bootstrap values would be obtained if more members of this species were sequenced and included in the analysis.


Figure 1
View larger version (23K):
[in this window]
[in a new window]

  		FIG. 1. Dendrogram showing the phylogenetic relationships of strains BQ1 and BQ8 to Alicycliphilus sp. A phylogenetic analysis of a 720-bp fragment (corresponding to positions 30 to 750 of the E. coli 16S rRNA gene) of the 16S rRNA gene was performed with the MEGA program, version 3.1. The Kimura two-parameter method was used to calculate evolutionary distances, and dendrograms were constructed according to the neighbor-joining method. Bootstrap values were calculated by using 500 replicates. Numbers at branching points refer to bootstrap values (500 resamplings).

Growth in MM-PUh showed a hyperbolic behavior, with an exponential growth phase between 4 and 12 h and the stationary phase starting around 60 h of incubation (Fig. 2A). BQ1 showed a higher maximum growth (17.8 ± 0.6 mg·ml–1) than BQ8 (14.0 ± 0.6 mg·ml–1) after 100 h of culture. The dependence of bacterial growth on the Hydroform concentration (from 10.5 to 105 mg·ml–1) showed a hyperbolic pattern (Fig. 2B), with µmax values of 3.94 ± 0.2 and 4.55 ± 0.6 h–1 for BQ1 and BQ8, respectively, and Ks values of 16.9 ± 3.5 and 27.7 ± 1.9 mg·ml–1 for BQ1 and BQ8, respectively. Statistically significant differences (P = 0.05) between Ks values were observed, suggesting that BQ1 is more efficient than BQ8, since it reached half of the maximum growth with a lower concentration of Hydroform.


Figure 2
View larger version (15K):
[in this window]
[in a new window]

  		FIG. 2. Growth of Alicycliphilus sp. in MM-PUh. (A) Wet weight was recorded during 115 h of culture in an MM with 1% (vol/vol) (10.5 mg·ml–1) of Hydroform (MM-PUh) as the only carbon source (BQ1, {square}; BQ8, {blacksquare}). The experiments were performed in triplicate. Bars indicate standard deviations. (B) Dependence of Alicycliphilus sp. growth on Hydroform concentration. MM with different concentrations of Hydroform (10.5 to 105 mg·ml–1) as the sole carbon source were used to cultivate BQ1 ({square}) and BQ8 ({blacksquare}) strains. Wet weight values measured at the exponential growth phase (0 to 12 h) were used to construct the Monod plot. The experiments were performed in triplicate. Bars indicate standard deviations.

Hydroform is a polyester PU.
Since the Hydroform chemical composition has not been published, analyses to determine the type of compounds and the nature of the PU present were performed. Hydroform was vacuum distilled and two liquid fractions, corresponding to 71.5% of the total volume, were obtained, together with a solid residue. GC-MS analysis of the distilled fractions showed that the larger of these (50% of the total volume) was water. The remaining 20% comprised a mixture of NMP and two dipropyleneglycol methyl ether isomers (DPGME and DPGME2) (Fig. 3A). NMR analysis from the solid residue (Fig. 3B) showed the presence of ester bonds at peaks 3.6 to 3.9, 7.9, and 8.4 ppm in the 1H NMR spectrum and at 44 to 50, 63, and 66 to 70 ppm in the 13C NMR spectrum (Table 1) . The IRS-FTIR analyses of the solid residue confirmed the ester bond at peak 979 cm–1, the amide group at 1,534 to 1,547 cm–1, and the carbonyl group of the ester bond at 1,722 to 1,735 cm–1, characteristic of PU (data not shown). Data obtained from these analyses allow us to propose that PU in Hydroform is a polyester type and has the structure presented in Fig. 3C.


Figure 3
View larger version (19K):
[in this window]
[in a new window]

  		FIG. 3. Hydroform characterization. (A) Chromatogram obtained from GC-MS analysis of chloroform-extracted cell-free SNs from noninoculated MM-PUh cultures. (B) 1H NMR and 13C NMR spectra of PU present in Hydroform. An analysis of the spectra is presented in Table 1. (C) Proposed chemical structure of PU present in Hydroform.


View this table:
[in this window]
[in a new window]

  		TABLE 1. NMR spectrum analysis of solid Hydroform

Alicycliphilus sp. was able to utilize NMP.
To determine whether the Alicycliphilus sp. strains were able to utilize the other carbon compounds differently than the PU (NMP or DPGME) present in the Hydroform, chloroform extracts of the MM-PUh cell-free SNs were analyzed by GC-MS after different incubation times (0, 3, 6, 9, and 12 h). Analyses showed that NMP, but not the DPGME isomers, was consumed during the growth of the bacteria in inoculated media (Table 2). BQ1 consumed NMP at a higher rate (0.34 µmol·h–1·ml–1) than BQ8 (0.16 µmol·h–1·ml–1). Both NMP and DPGME remained in incubated and noninoculated media during this period.


View this table:
[in this window]
[in a new window]

  		TABLE 2. Concentrations of NMP in the SNs of MM-PUh during the growth of Alicycliphilus sp.

To demonstrate that Alicycliphilus sp. strains were able to utilize NMP as their sole carbon and nitrogen source, the dependence of their growth on NMP concentration (from 0.5 to 25 mg·ml–1) in an MM without other sources of carbon or nitrogen was determined by recording wet weight increases during the exponential growth phase (between 0 and 12 h). µmax values were 0.87 ± 0.04 and 0.94 ± 0.08 h–1 for BQ1 and BQ8, respectively, and Ks values were 7.0 ± 0.9 and 8.9 ± 1.9 mg·ml–1 for BQ1 and BQ8, respectively. No statistically significant differences between them were observed. Calculated values denote that both strains were able to sustain their growth based on NMP. Experiments testing the capacities of BQ1 and BQ8 to sustain their growth by using 2-pyrrolidone (2-P), an NMP-related compound, showed that none of the Alicycliphilus sp. strains were able to do it (data not shown).

Esterase activity, but not protease or urease activity, is induced in Alicycliphilus sp. growing in MM-PUh.
Protease, urease, esterase, and lipase activities identified in polyurethanolytic microorganisms have previously been shown to be associated with PU degradation (18, 19, 20, 30, 35, 36). To determine which activity was displayed by Alicycliphilus sp. strains, the strains were grown in three differential media. Growth in YES-gelatin and Christensen agar indicated that both strains were protease and urease negative. However, growth in Tween 80 medium showed precipitation around the colonies, indicating that the strains were esterase positive (data not shown). To detect the presence of extracellular esterase activity, cell-free SNs of inoculated MM-PUh cultures were used to measure this activity by the p-NPA method. Esterase activity was not detected after 6 and 10 h, even when the SN was concentrated 5x, but was clearly detected after 12 h of incubation. At 18 h, the maximum activity presented by the esterase was 25% higher for BQ8 (52 ± 3.6 nmol p-NP·min–1·ml–1) than for BQ1 (38 ± 2.2 nmol p-NP·min–1·ml–1). Beyond this point, activity began to decrease, remaining at 120 h of culture at approximately at 46% and 68% of the maximum (24 ± 4.0 and 26.5 ± 4.0 nmol p-NP·min–1·ml–1) for BQ8 and BQ1, respectively (Fig. 4). Esterase activity was not detected in SNs of LB broth cultures at 8 or 16 h, even if the SN at 8 h was concentrated 5x (data not shown). These results demonstrate that esterase activity was induced by the presence of compounds having ester linkages, such as Tween 80 or polyester PU. Protease and urease activities were also measured in cell-free SNs of inoculated MM-PUh cultures at the same times and during the same period as for esterase. None such activities were detected even though the assays were performed with 10x-concentrated SN.


Figure 4
View larger version (16K):
[in this window]
[in a new window]

  		FIG. 4. Esterase activity in culture medium SN. Cell-free SNs (100 µl) of BQ1 ({square}) and BQ8 ({blacksquare}) cultures grown in MM-PUh were used to measure esterase activity by the p-NPA method. The experiments were performed in triplicate for each point. Bars indicate standard deviations.

Effect of Alicycliphilus sp. growth on PU.
To determine if these strains were also able to attack the PU component of Hydroform, a number of aspects were analyzed. The effect of Alicycliphilus sp. on the PU present in the liquid MM-PUh was determined by analyzing the polymerization of PU after bacterial growth. Inoculated and noninoculated MM-PUh were incubated for 5 days, after which the cell-free SN was sterilized. After this treatment, the PU present in noninoculated medium was found to have polymerized (white precipitate), while PU from inoculated media had not (Fig. 5A, left three panels). To demonstrate that the absence of NMP in the inoculated media caused by early bacterial growth was not the cause of the loss of polymerizing properties in PU, MM-PUh was prepared using NMP-free Hydroform. The latter was obtained by dialysis and analyzed by GC-MS to confirm the absence of NMP. This medium was incubated for 5 days without inoculation and then sterilized. After incubation, it was found that NMP-free MM-PUh did in fact polymerize, indicating that the absence of NMP was not the cause of the lack of polymerization observed (Fig. 5A, far-right panel). These results indicate that bacterial activity had changed the properties of the PU present in the inoculated media, as a result inhibiting its polymerization.


Figure 5
View larger version (97K):
[in this window]
[in a new window]

  		FIG. 5. Effect of Alicycliphilus sp. culture on PU characteristics. (A) Inhibition of polymerization. MM-PUh noninoculated (NI) or inoculated with BQ1 or BQ8 and NMP-free MM-PUh were incubated at 37°C for 5 days in a rotary shaker incubator. After that time, cell-free SNs were sterilized. NMP-free MM-PUh was prepared using dialyzed Hydroform (analyzed by GC-MS to confirm that no NMP was present). The figure depicts the SNs of each culture after sterilization. (B) IRS-FTIR analysis. MM-PUh NI or inoculated with Alicycliphilus sp. strain BQ1 or BQ8 was incubated at 37°C for 120 h. Cell-free SNs were vacuum dried to obtain the solid component (PU). PU was analyzed by IRS-FTIR. Relevant peaks are indicated. (C) SEM analysis of polymerized Hydroform sheets. MM-PUh containing 1-mm-thick sheets (1 cm by 1 cm), NI or inoculated with BQ1 or BQ8. After 15 days of culture, the sheets were recovered, rinsed with water, gold coated by evaporation, and analyzed by SEM. Five sheets were incubated and analyzed for each culture. Representative results are shown.

The chemical composition of solid PU after bacterial growth was therefore analyzed. For this, bacteria were cultured in MM-PUh for 5 days, after which the SN was removed and vacuum dried and the residual solid analyzed by IRS-FTIR. Analysis revealed that carbonyl (1,734 cm–1) of the ester signal (979 cm–1), observed in solid PU not exposed to bacterial growth (Fig. 5B, top), increased in the solid obtained from the SN after culture with BQ1 (1,727 cm–1); at the same time, the ester signal decreased, and other signals corresponding to ether, methyl, methylene, and terminal alcohols (983 to 1,450 cm–1) appeared (Fig. 5B, middle). These changes indicate that ester bonds were hydrolyzed, producing increases in the signals of the indicated groups, in BQ1-exposed media. The amide signal (1,530 to 1,550 cm–1) increased in BQ1-exposed PU, discarding the hypothesis of hydrolysis of this group by a protease or urease activity. The spectrum presented by BQ8-exposed PU showed less ester hydrolysis than BQ1-exposed PU (Fig. 5B, bottom). These results indicate that BQ1 growth affects PU-ester bonds but not urethane bonds and that BQ8 growth does not affect them to the same extent.

To study the effect of growth of Alicycliphilus strains on polymerized PU, thin polymerized Hydroform sheets were incubated for 15 days in liquid media inoculated with the two bacterial strains. After that time, sheets were viewed by SEM, which showed that sheets exposed to BQ1 had numerous holes on their surface (Fig. 5C, middle), whereas those exposed to BQ8 had few holes (Fig. 5C, bottom). These results clearly indicate that BQ1 can attack solid PU and that, while BQ8 also attacks PU, it does so to a lesser extent.


arrow
DISCUSSION
 
In this work, two strains able to grow in PU were isolated by several rounds of selection in a medium comprising Hydroform, a commercial aliphatic, water-based PU, as the only carbon source. By sequencing of the 16S rRNA gene, the two strains were identified as Alicycliphilus sp. Alicycliphilus denitrificans, the first bacterium classified in this genus, is a new genus and new species reported as a denitrifying and cyclohexanol-degrading bacterium (29) belonging to the Comamonadaceae family in the Betaproteobacteria class. The name Alicycliphilus means alicyclic compound liking, referring to the substrates used for the isolation of this organism (29). This genus is phylogenetically related to Acidovorax defluvii and Comamonas denitrificans, which have previously been isolated from activated sludge (15, 40). Moreover, another bacterium of the Comamonadaceae family, C. acidovorans, has been reported as being able to use various types of PU to sustain its growth (2, 30).

Chemical characterization of Hydroform by NMR, IRS, and GC-MS showed that it was composed of a polyester-type PU, NMP, and two DPGME isomers. We found that Alicycliphilus sp. consumed NMP present in Hydroform during its early phase of growth. Moreover, we were able to confirm that NMP was utilized by the isolated bacteria, since both strains were found to grow in an MM with NMP as the sole carbon and nitrogen source. Pseudomonas aeruginosa was reported as being able to use 2-P, an NMP-related compound, as the nitrogen and carbon source, but it was unable to use NMP (33). On the contrary, Alicycliphilus sp. was able to use NMP but not 2-P. As far as is known, this is the first report of a bacterium being able to utilize NMP to sustain growth. Since the toxicity of NMP has been recognized, our finding that Alicycliphilus sp. is able to consume NMP could be of great relevance in biotechnology research directed towards avoiding the toxic consequences of its release into the environment.

Experiments with differential media and by measurements of protease and urease activities in SNs of MM-PUh inoculated with these bacteria at different growing times demonstrated that the isolated Alicycliphilus sp. strains did not exhibit protease or urease activity. However, esterase activity was detected in Tween 80 media, and by using the p-NPA assay, the esterase activity secreted to the culture medium was measured. A lipase activity has been identified in Bacillus subtilis (35), and extracellular esterase activities have been reported for Pseudomonas chlororaphis and Comamonas acidovorans growing in media with Impranil (2, 20, 36). In C. acidovorans TB-35, an extracellular esterase and a membrane-associated esterase have been identified (30). Analysis of these esterase activities on solid PU and poly(diethylenglycol adipate) has demonstrated that only the membrane-associated esterase was able to degrade poly(diethylenglycol adipate) and PU, this being the one with polyurethanolytic activity (1). Based on the changes observed in the IR spectra of the solid PU obtained from the SN of MM-PUh after culture with strain BQ1 (Fig. 5B), we determined that it was ester groups that were being affected by a bacterial esterase activity. Whether the extracellular esterase activity detected in Alicycliphilus sp. was responsible for the PU attack or whether there was membrane-associated esterase activity will be the subject of further investigation. The absence of an extracellular esterase activity when Alicycliphilus sp. grows in LB broth and detection of this activity at 12 h of culture in MM-PUh, by which time NMP has greatly diminished in the culture medium, suggest that esterase activity is induced in response to the chemical composition of the medium. In previous studies, a medium containing Impranil supplemented with yeast extract has been used to characterize polyurethanolytic bacteria (2, 19, 20, 35, 36). It might be possible that the presence of molecules more easily consumed than PU and which could also be utilized as a carbon source could help in the establishment of polyurethanolytic bacteria in PU media. This early growth might activate the expression of genes encoding proteins involved in the efficient utilization of PU, for example, esterase and other proteins needed for the uptake and metabolism of molecules produced after breakdown of PU-ester bonds. In this work, the presence of NMP and its utilization during early growth of Alicycliphilus sp. might provide an easily used carbon source, but after it has been exhausted, esterase activity has to be induced to allow the bacteria to utilize PU, a more complex carbon source.

Various data demonstrate the capacity of Alicycliphilus sp. to degrade PU, including a loss of polymerizing ability of PU after exposure to bacteria culture, indicating the loss of groups needed for the polymerization reaction. These groups could be lost by cleavage of ester bonds, generating carboxyl groups and short-chain alcohols unsuitable for polymerization. IRS analysis, in which the PU-ester bond peaks disappeared and carbonyl group signals increased in the PU spectrum after culture with the BQ1 strain, supports this suggestion. More work might be done to confirm this reaction. A more graphic demonstration was the observation of the numerous holes produced in polymerized PU after bacteria culture. Despite the fact that both Alicycliphilus strains share certain characteristics, such as the ability to grow in MM-PUh and NMP, the production of extracellular esterase activity when they grow in MM-PUh, and loss of the polymerizing capacity of PU after their growth, the action of BQ1 on PU seems to be more effective than that of BQ8. A more detailed analysis demonstrates that, although BQ8 showed higher extracellular esterase activity (measured by the p-NPA method) than BQ1, it reached a lower maximum growth in MM-PUh (10.5% Hydroform), it had larger Ks values for PU, it almost did not alter ester bond signals in IR spectrum after its growth, and it did not produce as many holes in polymerized PU as BQ1. These observations reflect the distinct genetic and biochemical backgrounds of both strains and suggest that differences, for example, in biochemical mechanisms of the esterase activity over PU, existence of other degradative activities for PU attack, diverse transporters of PU breakdown products, or different catabolic pathways inside the cell, might exist between the two strains. Work is being performed in our laboratory to identify the enzymes and the genes encoding them which participate in the different pathways involved in PU degradation. In conclusion, in this paper we report two important findings: the ability of Alicycliphilus sp. to utilize NMP as a carbon and nitrogen source, which is the first report of a microorganism able to do this, and the capacity of Alicycliphilus sp. to attack PU due to an esterase activity.


arrow
ACKNOWLEDGMENTS
 
NMR, IRS-FTIR, and SEM techniques were performed at Unidad de Servicios y Apoyo a la Investigación (USAI), Facultad de Química, UNAM. Biochemical tests for strain identification were performed by Alejandro Camacho, Facultad de Química, UNAM. We are grateful to Fernando Pérez Lara and Brenda Porta-Briseño for their help with GC-MS, NMR, and IRS-FTIR analyses and to León Patricio Martínez for his assistance with the phylogenetic analysis.

This study was partially supported by a PAIP-FQ-UNAM 6290-04 grant to H.L.-T. A.O.-C., A.C.-G., and S.B.-N. are grateful to CONACYT for scholarships.


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: Departamento de Bioquímica, Facultad de Química, Universidad Nacional Autónoma de México, Cd. Universitaria, 04510 México, D.F. México. Phone: (52) (55) 5622-5280. Fax: (52) (55) 5622-5329. E-mail: hlozat{at}servidor.unam.mx Back

{triangledown} Published ahead of print on 10 August 2007. Back


arrow
REFERENCES
 
    1
  1. Akutsu, Y., T. Nakajima-Kambe, N. Nomura, and T. Nakahara. 1998. Purification and properties of a polyester polyurethane-degrading enzyme from Comamonas acidovorans TB-35. Appl. Environ. Microbiol. 64:62-67.[Abstract/Free Full Text]
    2. 2
  2. Allen, B. A., N. P. Hilliard, and G. T. Howard. 1999. Purification and characterization of a soluble polyurethane degrading enzyme from Comamonas acidovorans. Int. Biodeterior. Biodegrad. 43:37-41.[CrossRef]
    3. 3
  3. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl. 1994. Current protocols in molecular biology, vol. 1, p. 2.4.1-2.4.4. John Wiley & Sons, Inc., New York, NY.
    4. 4
  4. Bader, M., S. A. Keener, and R. Wrbitzky. 2005. Dermal absorption and urinary elimination of N-methyl-2-pyrrolidone. Int. Arch. Occup. Environ. Health 78:673-676.[CrossRef][Medline]
    5. 5
  5. BASF Intermediates. 1994. N-Methylpyrrolidone biodegradability. BASF Intermediates, Ludwigshafen, Germany. http://www.basf.com/diols/pdfs/nmp_brochure_biod.pdf.
    6. 6
  6. Center for the Polyurethanes Industry. 2006. Year in review. Center for the Polyurethanes Industry, Arlington, VA. http://www.polyurethane.org/s_api/bin.asp?CID=820&DID=4985&DOC=FILE.PDF.
    7. 7
  7. Center for the Polyurethanes Industry. 2007. Polyurethane: history. Center for the Polyurethanes Industry, Arlington, VA. http://www.polyurethane.org/s_api/sec.asp?CID=853&DID=3487.
    8. 8
  8. Center for the Polyurethanes Industry. 2007. Recycling and waste reduction. Center for the Polyurethanes Industry, Arlington, VA. http://www.polyurethane.org/s_api/sec.asp?CID=867&DID=3520.
    9. 9
  9. Chapin, K. C., and P. R. Murray. 1995. Media, p. 1687-1707. In P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (ed.), Manual of clinical microbiology, 7th ed. ASM Press, Washington, DC.
    10. 10
  10. Chow, S. T., and T. L. Ng. 1983. The biodegradation of N-methyl-2-pyrrolidone in water by sewage bacteria. Water Res. 17:117-118.
    11. 11
  11. Christensen, W. B. 1946. Urea decomposition as a means of differentiating Proteus and paracolon cultures from each other and from Salmonella and Shigella types. J. Bacteriol. 52:461-466.[Free Full Text]
    12. 12
  12. Darby, R. T., and A. M. Kaplan. 1968. Fungal susceptibility of polyurethanes. Appl. Microbiol. 16:900-905.[Medline]
    13. 13
  13. Desphande, M. V., K. E. Eriksson, and L. G. Pettersson. 1984. An assay for selective determination of exo-1,4-ß-glucanases in a mixture of cellulolytic enzymes. Anal. Biochem. 138:481-487.[CrossRef][Medline]
    14. 14
  14. Federal Register. 1994. Addition of certain chemicals; toxic chemical release reporting; community right-to-know. Fed. Regist. 59:61432-61485.
    15. 15
  15. Gumaelius, L., G. Magnusson, B. Pettersson, and G. Dalhammar. 2001. Comamonas denitrificans sp. nov., an efficient denitrifying bacterium isolated from activated sludge. Int. J. Syst. Evol. Microbiol. 51:999-1006.[Abstract]
    16. 16
  16. Hass, U., B. M. Jakobsen, and S. P. Lund. 1995. Developmental toxicity of inhaled N-methylpyrrolidone. Pharm. Toxicol. 76:406-409.[Medline]
    17. 17
  17. Heylen, K., B. Vanparys, L. Wittebolle, W. Verstraete, N. Boon, and P. de Vosi. 2006. Cultivation of denitrifying bacteria: optimization of isolation conditions and diversity study. Appl. Environ. Microbiol. 72:2637-2643.[Abstract/Free Full Text]
    18. 18
  18. Howard, G. T. 2002. Biodegradation of polyurethane: a review. Int. Biodeterior. Biodegrad. 40:245-252.
    19. 19
  19. Howard, G. T., and R. C. Blake. 1998. Growth of Pseudomonas fluorescens on a polyester-polyurethane and the purification and characterization of a polyurethanase-protease enzyme. Int. Biodeterior. Biodegrad. 42:213-220.[CrossRef]
    20. 20
  20. Howard, G. T., C. Ruiz, and N. P. Hilliard. 1999. Growth of Pseudomonas chlororaphis on a polyester-polyurethane and the purification and characterization of a polyurethanase-esterase enzyme. Int. Biodeterior. Biodegrad. 43:7-12.[CrossRef]
    21. 21
  21. Jendrossek, D. 2001. Microbial degradation of polyesters, p. 293-325. In T. Scheper (ed.), Advances in biochemical engineering/biotechnology, vol. 71. Springer-Verlag, Berlin, Germany.
    22. 22
  22. Kay, M. J., R. W. McCabe, and L. H. G. Morton. 1993. Chemical and physical changes occurring in polyester polyurethane during biodegradation. Int. Biodeterior. Biodegrad. 31:209-225.[CrossRef]
    23. 23
  23. Lan, D.-H., C.-Y. Peng, and T.-S. Lin. 2004. Acute aquatic toxicity of N-methyl-2-pyrrolidinone to Daphnia magna. Bull. Environ. Contam. Toxicol. 73:392-397.[Medline]
    24. 24
  24. Lawson, P. A., P. Llop-Perez, R. A. Hutson, H. Hippe, and M. D. Collins. 1993. Towards a phylogeny of the Clostridia based on 16S rRNA sequences. FEMS Microbiol. Lett. 113:87-92.[CrossRef][Medline]
    25. 25
  25. Lee, K. P., N. C. Chromey, R. Culik, J. R. Barnes, and P. W. Schneider. 1987. Toxicity of N-methyl-2-pyrrolidone (NMP): teratogenic, subchronic, and two-year inhalation studies. Fundam. Appl. Toxicol. 9:222-235.[CrossRef][Medline]
    26. 26
  26. Lovrien, R., and D. Matulis. 1995. Assays for total protein, unit 3.4, p. 3.4.4-3.4.24. In J. E. Coligan, B. M. Dunn, H. L. Ploegh, D. W. Speicher, and P. T. Wingfield (ed.), Current protocols in protein science. John Wiley & Sons, Inc., New York, NY.
    27. 27
  27. Malloy, P. J., and J. J. Bradna. 1992. Interpretation of aerobic bacterial growth on primary culture media, p. 1.6.1-1.7.1. In H. D. Isenberg (ed.), Clinical microbiology procedures handbook. American Society for Microbiology, Washington, DC.
    28. 28
  28. Matsumura, S., Y. Soeda, and K. Toshima. 2006. Perspectives for synthesis and production of polyurethanes and related polymers by enzymes directed toward green and sustainable chemistry. Appl. Microbiol. Biotechnol. 70:12-20.[CrossRef][Medline]
    29. 29
  29. Mechichi, T., E. Stackebrandt, and G. Fuchs. 2003. Alicycliphilus denitrificans gen. nov., sp. nov., a cyclohexanol-degrading, nitrate-reducing ß-proteobacterium. Int. J. Syst. Evol. Microbiol. 53:147-152.[Abstract/Free Full Text]
    30. 30
  30. Nakajima-Kambe, T., F. Onuma, Y. Akutsu, and T. Nakahara. 1997. Determination of the polyester polyurethane breakdown products and distribution of the polyurethane degrading enzyme of Comamonas acidovorans strain TB-35. J. Ferment. Bioeng. 83:456-460.[CrossRef]
    31. 31
  31. Nakajima-Kambe, T., F. Onuma, N. Kimpara, and T. Nakahara. 1995. Isolation and characterization of a bacterium which utilizes polyester polyurethane as a sole carbon and nitrogen source. FEMS Microbiol. Lett. 129:39-42.[Medline]
    32. 32
  32. Nakajima-Kambe, T., Y. Shigeno-Akutsu, N. Nomura, F. Onuma, and T. Nakahara. 1999. Microbial degradation of polyurethane, polyester polyurethanes and polyether polyurethanes. Appl. Microbiol. Biotechnol. 51:134-140.[CrossRef][Medline]
    33. 33
  33. Noe, F. F., and W. J. Nickerson. 1958. Metabolism of 2-pyrrolidone and {gamma}-aminobutyric acid by Pseudomonas aeruginosa. J. Bacteriol. 75:674-681.[Free Full Text]
    34. 34
  34. Pathirana, R. A., and K. J. Seal. 1984. Studies on polyurethane deteriorating fungi. Part 2. An examination of their enzyme activities. Int. Biodeterior. 20:229-235.
    35. 35
  35. Rowe, L., and G. T. Howard. 2002. Growth of Bacillus subtilis on polyurethane and the purification and characterization of a polyurethanase-lipase enzyme. Int. Biodeterior. Biodegrad. 50:33-40.[CrossRef]
    36. 36
  36. Ruiz, C., T. Main, N. P. Hilliard, and G. T. Howard. 1999. Purification and characterization of two polyurethanase enzymes from Pseudomonas chlororaphis. Int. Biodeterior. Biodegrad. 43:43-47.[CrossRef]
    37. 37
  37. Saillenfait, A. M., F. Gallissot, and G. Morel. 2003. Developmental toxicity of N-methyl-2-pyrrolidone in rats following inhalation exposure. Food Chem. Toxicol. 41:583-588.[CrossRef][Medline]
    38. 38
  38. Saillenfait, A. M., F. Gallissot, I. Langonne, and J. P. Sabate. 2002. Developmental toxicity of N-methyl-2-pyrrolidone administered orally to rats. Food Chem. Toxicol. 40:1705-1712.[CrossRef][Medline]
    39. 39
  39. Santerre, J. P., R. S. Labow, D. G. Duguay, D. Erfle, and G. A. Adams. 1994. Biodegradation evaluation of polyether and polyester-urethanes with oxidative and hydrolytic enzymes. J. Biomed. Mater. Res. 28:1187-1199.[CrossRef][Medline]
    40. 40
  40. Schulze, R., S. Spring, R. Amann, I. Huber, W. Ludwing, K. H. Schleifer, and P. Kampfer. 1999. Genotypic diversity of Acidovorax strains isolated from activated sludge and description of Acidovorax defluvii sp. nov. Syst. Appl. Microbiol. 22:205-214.[Medline]
    41. 41
  41. Sierra, G. 1957. A simple method for the detection of lipolytic activity of micro-organisms and some observations on the influence of the contact between cells and fatty substrates. Antonie Leeuwenhoek 23:15-22.[Medline]
    42. 42
  42. Solomon, G. M., E. P. Morse, M. J. Garbo, and D. K. Milton. 1996. Stillbirth after occupational exposure to N-methyl-2-pyrrolidone: a case report and review of the literature. J. Occup. Environ. Med. 38:705-713.[CrossRef][Medline]
    43. 43
  43. Solomon, H. M., B. A. Burgess, G. L. Kennedy, and R. E. Staples. 1995. 1-Methyl-2-pyrrolidone (NMP): reproductive and developmental toxicity study by inhalation in the rat. Drug Chem. Toxicol. 18:271-293.[Medline]
    44. 44
  44. Wen, W.-I., L.-J. Liang, and C.-E. Tsung. 2001. Acute poisoning with the neonicotinoid insecticide imidacloprid in N-methyl pyrrolidone. J. Toxicol. Clin. Toxicol. 39:617-621.[CrossRef][Medline]
    45. 45
  45. Witte, C.-P., and N. Medina-Escobar. 2001. In-gel detection of urease with nitroblue tetrazolium and quantification of the enzyme from different crop plants using the indophenol reaction. Anal. Biochem. 290:102-107.[CrossRef][Medline]

Applied and Environmental Microbiology, October 2007, p. 6214-6223, Vol. 73, No. 19
0099-2240/07/$08.00+0     doi:10.1128/AEM.01230-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Oceguera-Cervantes, A.
Right arrow Articles by Loza-Tavera, H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Oceguera-Cervantes, A.
Right arrow Articles by Loza-Tavera, H.
Agricola
Right arrow Articles by Oceguera-Cervantes, A.
Right arrow Articles by Loza-Tavera, H.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»