The microbial degradation of tensile test pieces made of poly(3-hydroxybutyrate) [P(3HB)J or a copolymer of 90%o 3-hydroxybutyric acid and 10%o 3-hydroxyvaleric acid was studied in soils incubated at a constant temperature of 15, 28, or 40°C for up to 200 days. In addition, hydrolytic degradation in sterile buffer at temperatures ranging from 4 to 55°C was monitored for 98 days. Degradation was measured through loss of weight (surface erosion), molecular weight, and mechanical strength. While no weight loss was recorded in sterile buffer, samples incubated in soils were degraded at an erosion rate of 0.03 to 0.64% weight loss per day, depending on the polymer, the soil, and the incubation temperature. The erosion rate was enhanced by incubation at higher temperatures, and in most cases the copolymer lost weight at a higher rate than the homopolymer. The molecular weights of samples incubated at 40°C in soils and those incubated at 40°C in sterile buffer decreased at similar rates, while the molecular weights of samples incubated at lower temperatures remained almost unaffected, indicating that molecular weight decrease is due to simple hydrolysis and not to the action of biodegrading microorganisms. The degradation resulted in loss of mechanical properties. From the samples used in the biodegradation studies, 295 dominant microbial strains capable of degrading P(3HB) and the poly(3-hydroxybutyrate-co-3-hydroxyvalerate) copolymer in vitro were isolated and identified. They consisted of 105 gram-negative bacteria, mostly belonging toAcidovorarfacilis and Variovoraxparadoxus, 36 Bacilus strains, 68 Streptomyces strains, and 86 mold strains, mainly belonging to Aspergillusfumigatus and species of the genus PeniciUium.

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The biodegradation of samples of poly(3-hydroxybutyrate)(P(3HB)), poly(3-hydroxybutyrate-co-10%-3-hydroxyvalerate)(P(3HB-co-10 %-3HV)), and poly 3-hydroxybutyrate-co-20%-3-hydroxyvalerate)(P(3HB-co-20%- 3HV)) was investigated in situ in natural waters. The degradation was studied by decrease in mass, molecular weight, and tensile strength. In two freshwater ponds the polymers were slowly degraded. After half a year of submersion the mass loss was less than 7%. After 358 days in a freshwater canal, 34% mass loss was recorded for the homopolymer, and 77% for the P(3HB-co-10%-3-HV) samples, while the P(3HB-co-20%-3HV) samples had completely disappeared. In seawater in the harbour of Zeebrugge, P(3HB) samples lost 31% of their initial mass, and the copolymers 49-52%, within 270 days. In all of these environments, the degradation rate was faster during the summer, when the temperature of the water was higher. No relevant changes in molecular weight could be detected, indicating that the degradation took place only at the surface of the samples. The degradation resulted in considerable loss of tensile strength of the copolymer samples. Ninety-two microorganisms, mainly bacteria, able to degrade P(3HB) in polymer overlayer plates, were isolated and identified by fatty acid analysis. The isolates from one freshwater pond belonged mainly to the bacterial genus Acidovorax, while the microorganisms from the other freshwater pond belonged to various bacterial genera, to Streptomyces, and to the mould genus Penicillium. Most of the 31 bacterial isolates from seawater were identified as Alteromonas haloplanktis. The results demonstrate that P(3HB) and P(3HB-co-3HV) samples are effectively biodegradable in natural waters under real-life conditions and reveal the biodiversity of the microflora responsible for this biodegradation.

Degradation of poly(3‐hydroxybutyrate) and copolymers with 3‐hydroxyvaleric acid was investigated in natural environments, and the microorganisms involved were isolated and identified. The influence of abiotic and biotic factors on the degradation is discussed.

Using a grid pattern, a total of 50 soil borings (DCT 1 through DCT 50) were collected from an abandoned chemical manufacturing facility. Split samples were obtained from six cores collected from a centralized location near a former wastewater lagoon. An additional sample from the 2.1‐ to 2.4‐m depth of core DCT 20 was obtained as representative of the waste present on site. Three subsamples from each of the six cores or a total of 18 samples were sequentially extracted with methylene chloride and methanol for testing in mutagenicity and acute toxicity assays. A separate batch extraction with water was conducted and evaluated, using both chemical and biological test methods. The maximum mutagenic response was 3,237 net revertants induced with metabolic activation by 1 mg of the methanol extract of the 0‐ to 0.6‐m depth at sampling location DCT 32. The water extract of the 0‐ to 0.6‐m depth at sampling location DCT 32 induced 737 net revertants per mg. At sampling location DCT 24, the water extract induced 0.74, 1.32, and 2.5 toxic units in the Microtox® assay and 935, 69, and 39 net revertants per mg in the mutagenicity assay at the 0‐ to 0.3‐, 1.2‐ to 1.4‐, and 2.1‐ to 2.4‐m depths, respectively. The chemical analysis of the samples from location DCT 24 indicated that the concentration of 2,3,6‐trichlorobenzene acetic acid ranged from 16 ppm in the 0‐ to 0.3‐m depth and to 0.95 ppm and below the detection limits at the 1.2‐ to 1.4‐ and 2.1‐ to 2.4‐m depths. The methanol extract of the soil from the 0‐ to 0.3‐m depth induced 19.2 toxic units in the acute toxicity assay and 1,467 net revertants in the mutagenicity assay. These results suggest that the most accurate site assessment is obtained when a water extract is evaluated to measure leaching potential and a solvent extract is evaluated to determine the relative total hazard of the sample. In addition, the combined use of chemical analysis, mutagenicity, and acute toxicity bioassays provides more accurate information from which to make a risk assessment.