Journal of Polymers and the Environment, Vol. 8, No. 2, 2000

Polymer Mineralization in Soils: Effects of Cold Storage on Microbial Populations and Biodegradation Potential Jason S. Lee,1 Belinda L. Daniels,1 David T. Eberiel,1 and Richard E. Farrell1–4

Soil retrieval, processing and storage procedures can have a profound effect on soil microorganisms. In particular, changes in soil microbial populations may adversely affect the biological activity of a soil and drastically alter the soil’s potential to mineralize added substrates. The effects of cold storage on the biodegradation of a series of test polymers was investigated using two soils—a synthetic soil mix (SM-L8) and a field soil (Bridgehampton silt loam) from Rhode Island (RI-1). Biodegradation tests were conducted using freshly prepared/ collected soil and again following storage at 48 C for 3 to 8 months. Prior to each biodegradation test, the soils were incubated at 60% water-holding capacity (WHC) and 258 C to rejuvenate the microbial populations; the soils were incubated for periods of 48 h (freshly collected soil) or 25 days (soils stored at 48 C). Soil microbial populations were assessed by enumerating different segments of the population on agar plates containing different selective media. Mineralization of the test polymers (cellulose, poly-3-hydroxybutyrate, and starch acetate, d.s. 1.5) was monitored using standard respirometric techniques. Our results demonstrated that cold storage had a generally negative effect on the soil microbial populations themselves but that its effect on the capacity of the soil microorganisms to degrade the test polymers varied between soils and polymer type. Whereas cold storage resulted in dramatic shifts in the community structure of the soil microbial populations, substantial restoration of these populations was possible by first conditioning the soils at 60% WHC and ambient temperatures for 25 days. Likewise, although the effects of cold storage on polymer mineralization varied with the test polymer and soil, these effects could be largely offset by including an initial 25-day stabilization period in the test. KEY WORDS: Microbial degradation; biodegradable polymers; cellulose; starch acetate; poly(3hydroxybutyrate).

1. INTRODUCTION

known that the procedures used to retrieve, process, and store soils can dramatically affect the microbial population of a soil [14] and, hence, its potential to degrade an added substrate (e.g., a polymer or plastic). Although it is best to use a soil immediately after collection, this is not always feasible. Thus, if biodegradation tests in a soil are to be repeated, or if comparative tests are to be made several months apart, we are left with two options: (1) collect soil from the same site each time it is needed or (2) collect a large amount of soil—enough for multiple tests—and store it until needed. To ensure that environmentally relevant data are obtained, it is essential that the soils used in polymer biodegradation tests contain a microbial population that is comparable to that found in the field. Multiple

Microorganisms are the most important segment of the soil biota. Indeed, soil microorganisms are responsible for nutrient cycling and the decomposition of organic materials (both natural and synthetic). It is well 1 NSF–Biodegradable

Polymer Research Center, Department of Biology, University of Massachusetts Lowell, One University Avenue, Lowell Massachusetts 01854 2 NSF–Biodegradable Polymer Research Center, Department of Chemistry, University of Massachusetts Lowell, One University Avenue, Lowell, Massachusetts 01854. 3 Present address: University of Saskatchewan, Department of Soil Science, 51 Campus Drive, Saskatoon, Saskatchewan S7N 5A8 Canada. 4 To whom all correspondence should be addressed. Telephone: (306) 966–2772; email: [email protected]

81 1566-2543/ 00/ 0400-0081$18.000/ 0  2000 Plenum Publishing Corporation

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retrievals from the same site are confounded by seasonal changes, which alter the microenvironment and, in turn, the soil microbial community, and may have a significant effect on mineralization potential [5]. Likewise, the conditions used to store soils can have a significant effect on the soil microorganisms. Stotzky et al. [12] have shown that methods of storage, such as freezing (< 08 C) and air drying, can result in drastic changes in the soil biota and, in turn, may effect the capacity of the soil to mineralize organic matter. Cold storage (4–58 C) at a moisture content of 40 to 50% water-holding capacity (WHC), on the other hand, is usually considered to have less of an effect on the soil microorganisms [1, 14]. Furthermore, it has been suggested that a short stabilization period—with the soil maintained at optimum moisture and temperature status—can be used to stabilize soil microbial populations following cold storage [1, 7]. Consequently, cold storage followed by a stabilization period at room temperature is deemed to be the most practical means of storing a soil and maintaining its microbial activity. Nevertheless, several questions remain to be answered; e.g., Does cold storage affect the soil biota to the extent that polymer mineralization studies can not be repeated? And, if so, is there a way to negate these effects so that tests performed months apart will yield comparable results? In addition, storage and handling procedures should be standardized to ensure the biological integrity of the soil and facilitate interlaboratory comparisons of polymer biodegradation.

MATERIALS AND METHODS Soils A standard soil mix (SM-L8) was prepared from potting soil (Peters Professional Potting Soil; GraceSierra Horticultural Products Co., Milpitas, CA), acid washed sand (construction grade sand washed with 5% HNO3 ), and dehydrated and composted cow manure (1881 Select; Earthgro Inc., Lebanon, CT) in a ratio of 1 : 1 : 0.1 (w/ w/ w). Biodegradation tests also were conducted using a field soil (Bridgehampton silt loam; South Kingston, RI) collected in late August, 1995. The composition of the standard soil mix is similar to that described by McCassie et al. [9], but with a smaller proportion of added manure so that the total organic carbon content of the standard mix was comparable to that of the field soil (see Table II). Both soils were mixed thoroughly and sieved through a 2-mm screen prior to analysis for pH, nutrient status, water content, and WHC [4, 13].

Lee, Daniels, Eberiel, and Farrell Polymer biodegradation tests were carried out immediately after collection/ processing—using a portion of each soil. The remaining (unamended) soil was stored at a temperature of 48 C and the moisture content maintained at 40–50% WHC. The soils were stored in thin (1 mil) polyethylene bags, placed inside plastic buckets, and covered loosely to allow for some aeration while minimizing water loss. The soils were then placed in the cold room and stored for up to 8 months.

Characterization of Soil Microbial Populations Enumeration on Selective Media Microbial populations are a key component of soils, and differences in both population size and community structure are likely to have a significant effect on polymer mineralization. Thus, to truly understand how polymers biodegrade in soils, it is necessary to assess the size and structure of the soil microbial community. One method of doing so involves enumerating the soil microbes using selective media [6, 16]. The media used in this study were: Tryptic Soy agar (TSA), Soil Extract agar (SEA), Czapek-Dox agar (CDA), Rose Bengal agar (RBA), TSB-Vancomycin agar (TSB-V), and Starch Casein agar (SCA). TSA was used to enumerate populations of total heterotrophs, and thus provide an estimate of the overall bacterial population. However, because TSA supplies high concentrations of essential nutrients and other growth factors, this medium may favor fast-growing microorganisms and yield population estimates that are too high [2]. SEA, on the other hand, is more representative of the soil environment in which the native microorganisms evolved and, therefore, is more likely to yield better estimates of the total population of heterotrophs. Actinomycetes were enumerated using SCA. Gram-negative (GM) bacteria were enumerated using TSB-V agar. Yeasts and molds were enumerated using CDA; fungi were enumerated using RBA. The microorganisms were released from the soil and enumerated using standard serial dilution techniques, i.e., a 10.0-g sample of moist (60% WHC) soil was added to 90-mL of phosphate-buffered saline (PBS) in a milk dilution bottle and shaken vigorously for 20 min (10 − 1 dilution); a 10-mL aliquot of this suspension was then pipetted into another bottle containing 90mL of PBS (10 − 2 dilution). This procedure was repeated three more times to yield dilutions of 10 − 3 , 10 − 4 , and 10 − 5 . Finally, 0.10 mL from the various dilutions was

Polymer Mineralization in Soils pipetted onto the agar plates (5 reps per dilution) and spread using a sterile glass “hockey stick.” The agar plates were then inverted and placed in an incubator at room temperature (i.e., 208 –228 C). Microbial populations were enumerated by counting the number of colonies on each plate after 3 and 7 days. Polymer Mineralization Studies Standard Test Respirometric studies of polymer mineralization (i.e., the conversion of polymer-C to CO2 ) were conducted using a method based on that described by Bartha and Pramer [3]. Freshly prepared/ collected soils were adjusted to 60% of their WHC and preincubated for 48 h to eliminate the flush of CO2 that normally accompanies the rewetting of a soil. The test polymer was then added to the soil at a loading of 5-mg polymer-C gm − 1 soil (oven-dry weight basis), i.e., 250 mg polymer-C was mixed with 50-g (oven-dry weight) of soil and placed in a 250-mL reactor vessel (i.e., biometer flask; Belco Glass Inc., Vineland, NJ). Blanks (i.e., empty reactors) were set-up to determine the amount of CO2 in the headspace of the reactor flasks; reactors containing unamended soil were used as the controls. All systems were run in triplicate. The side-arm chamber of each biometer flask was charged with 20 mL of 0.5 M KOH, the flasks were then stoppered and placed in a warm room at 308 C. The CO2 evolved during mineralization was trapped in the KOH and the CO2 traps were sampled daily for the first 7 to 10 days and at 2–3-day intervals thereafter. The soils were aerated by allowing the systems to equilibrate with atmospheric air for 20 min after the CO2 traps were sampled. Potassium hydroxide in the traps was quantitatively transferred to 25-mm × 250-mm screw top test tubes containing 3.5 mL of 1.5 M BaCl2 and were allowed to sit for approximately 1 hour to precipitate the carbonates formed by reaction of the CO2 with the hydroxide. A known volume (4.0 ml) of the remaining (unreacted) KOH was titrated using standardized 0.10 M HCl. Net mineralization was then determined relative to the blanks and controls. Note that biological, chemical, and physical analyses of the soils were carried out while the mineralization studies were underway. Modified Tests Additional biodegradation tests were conducted with the standard soil mix after it had been in cold stor-

83 Table I. Combinations of Test Factors Storage temperature (8 C) Test IDa 22/ 02 22/ 25 04/ 02 04/ 25 a ID

4

√ √

Stabilization time (days)

22

2

√ √

√ √

25

√ √

code, storage temperature/ stabilization time.

age (48 C) for 3 months. Likewise, the effects of cold storage on polymer mineralization also were determined using the Bridgehampton silt loam soil (RI-1). In this case, the biodegradation tests were carried out within 72 h of the time the soil was brought to the lab and sieved, and then again after 8 months in cold storage (48 C). In both cases, prior to testing, the soil was adjusted to a moisture content of 60% WHC and incubated for 2 or 25 days at ambient temperature. The various combinations of test factors studied are listed in Table I. Test Polymers Biodegradation tests were carried out using the following polymers: cellulose (TLC grade, microcrystalline; Sigma Chemical Corp., St. Louis, MO), poly(3hydroxybutyrate), P3HB (Grade c 5100, M N ≈ 130, 000; Monsanto, St. Louis, MO), and starch acetate (d.s. 1.5, M N ≈ 150,000; National Starch & Chemical Co., Bridgewater, NJ). All polymers were supplied in powdered form and were hand mixed with the soils. Data Analysis Biodegradation data were first plotted in the form of polymer mineralization versus time curves (see Figs. 3–8). Nonlinear regression techniques were then used to fit a “three-parameter, single exponential rise to a maximum” model (SPSS, 1997): y c y0 + a(1 − exp − kt )

(1 )

where y is amount of substrate carbon (expressed as mg ThCO2 ) mineralized at time t, y0 is the intercept, a is the amplitude, and k is the empirical rate coefficient. (Note: Eq. 1 is fitted to the data beginning at the end of the lag period; in all cases R2 ≥ 0.95). Results of the mineralization studies also were interpreted using the following response parameters: (1) MAX-CO2 , defined as

84

the maximum amount of CO2 -C evolved during mineralization of the test substrate—calculated using Eq. (1); (2) LAG, defined as the time required for net CO2 –C evolution to reach 10% of the MAX-CO2 ; (3) PDP, the primary degradation phase of the test exposure—defined as the time from the end of the lag period to the start of the plateau region of the net mineralization curve (i.e., the point where net mineralization c 23 MAX-CO2 ); (4) r pdp , defined as average rate of mineralization during the primary degradation phase—calculated as the slope of the linear least-squares regression line plotted over this region of the net mineralization curve; and (5) the relative biodegradation index (RBI), defined as the ratio of cumulative net mineralization of the test polymer following cold storage to cumulative net mineralization of the test polymer prior to cold storage. In addition, the time required for mineralization of the test polymers to reach 50% ThCO2 (t 50 ) was calculated from the mineralization equation (Eq. 1). Analysis of variance (ANOVA) and Tukey’s test for significant differences were conducted using SigmaStat ver. 2.0 [11]. Exploratory data analysis revealed that most of the plate count data were log-normally distributed; consequently, all statistical analyses were carried out using the log-transformed data.

RESULTS AND DISCUSSION Microbial Characterizations of the SM-L8 and RI-1 Soils Air drying a soil produces considerable changes in its biological activity and is not recommended for studies of microbial processes. Cold storage at 28 –58 C and 40–50% WHC, on the other hand, is generally considered to produce relatively small perturbations in soil microbial activity and is usually recommended as the preferred method of storing soils for studies of microbial processes [14]. Nevertheless, even under these storage conditions, biological activity may not accurately reflect that in a freshly collected soil [15]. Indeed, we found that cold storage had a significant ( p c 0.05) negative impact on indigenous microbial populations in both the SM-L8 and RI-1 soils (Figs. 1 and 2). However, whereas it was possible to rejuvenate most segments of the microbial population by incubating the SM-L8 soil at 55–60% WHC and ambient temperatures for 25 days (Fig. 1), the same was not true for the RI-1 soil (Fig. 2). In particular, populations of total heterotrophs (SEA only), GM-bacteria, yeasts and molds, and actin-

Lee, Daniels, Eberiel, and Farrell

Fig. 1. Effects of cold storage and length of the stabilization period on microbial populations in the standard soil mix (SM-L8). Populations enumerated using selective media. Note: Plate counts were obtained on the last day of the stabilization period. Legend, storage temperature (8 C)/ stabilization time (days). Error bars represent 1 SD above the mean.

omycetes in the SM-L8 soil exhibited significant ( p c 0.05) increases in number following the 25-day stabilization period. Fungi, on the other hand, exhibited only a very limited rejuvenation. These results are similar to those of Zelles et al. [15] who found that indicators of bacterial activity generally increased more rapidly than indicators of fungal activity following soil storage under various conditions—including cold storage at 48 C. On

Fig. 2. Effects of cold storage and length of the stabilization period on microbial populations in the Bridgehampton silt loam soil (RI-1): Populations enumerated using selective media. Note: Plate counts were obtained on the last day of the stabilization period. Legend, storage temperature (8 C)/ stabilization time (days). Error bars represent 1 SD above the mean.

Polymer Mineralization in Soils

85 Table II. Soil Chemical Analyses Soil variablea

Soil

pH

TOC (%)

SOC (mg g − 1 )

TN (%)

C:N

Inorganic N (mg g − 1 )

P (mg g − 1 )

K (mg g − 1 )

SM-L8 RI-1

7.2 5.2

4.80 4.29

520 44

0.29 0.40

16.6 10.8

87 10

615 6

1729 70

a TOC,

total organic carbon; SOC, soluble organic carbon; TN, total kjeldhal nitrogen; C : N, TOC/ TN; inorganic N, NO3 -N + NH4 -N.

the other hand, in the RI-1 soil only the total heterotrophs (SEA only) and GM-bacteria exhibited any significant ( p c 0.05) rejuvenation during the 25-day stabilization period (Fig. 2). Poor recovery of most segments of the microbial population in the RI-1 soil are most likely a reflection of the lower organic matter and nutrient concentrations in this soil (Table II). In particular, there is considerably less soluble organic carbon (SOC) in the RI-1 soil. Given that this fraction of the total soil organic matter pool represents the most readily bioavailable form of carbon in the soil, it seems likely that rejuvenation of the microbial populations in the RI-1 soil following cold storage was limited by the relatively low supply of available carbon. Indeed, it has been reported that, under adverse environmental conditions, microbial survival is related to both the amount and biodegradability of the organic matter present in the growth medium [8]. Likewise, Anderson [1] reported that the primary reason for microbial biomass loss following soil storage was carbon starvation. Clearly, storage at 48 C for extended periods (i.e., 3–8 months) can produce significant changes in soil microbial populations, even when followed by a relatively lengthy conditioning/ stabilization period. The impact of these population changes on mineralization of the test polymers was determined next.

28-day biodegradation test exposure (Table III). Likewise, there was a 30% decrease in the overall rate coefficient (k) and a fivefold decrease in the average rate of mineralization during the primary degradation phase (r pdp ). Incubating the soil at room temperature (228 C), while maintaining optimal moisture conditions, for 25 days prior to initiating the biodegradation test resulted in significant increases in the r pdp and overall rate coefficient, but only a 9% increase in net cumulative mineralization. Given that cellulose degradation is mediated primarily by fungi, these results presumably reflect a decrease in fungal populations in the soil following cold storage (see Fig. 1).

Poly(3-hydroxybutyrate) As with cellulose, cold storage of the soil resulted in significant decreases in cumulative net mineralization of the P3HB ( − 28%), as well as the overall rate coefficient ( − 24%) and r pdp (twofold reduction). However, unlike the cellulose degraders, the P3HB degraders appear to

Polymer Mineralization in the Standard Soil Mix (SM-L8) Cellulose The effects of cold storage and length of the stabilization period on the net mineralization of cellulose in the standard soil mix (SM-L8) are illustrated in Fig. 3. Storage of the soil at 48 C for 3 months had a significant negative effect on the ability of the soil microorganisms to degrade cellulose; i.e., there was a 33% decrease in cumulative net CO2 evolution following a

Fig. 3. Effects of cold storage and length of the stabilization period on net mineralization of cellulose in the standard soil mix (SM-L8). Soil maintained at 308 C and 60% WHC throughout the biodegradation test exposure. Legend, storage temperature (8 C)/ stabilization time (days).

Lee, Daniels, Eberiel, and Farrell

86

Table III. Effects of Cold Storage and Preconditioning on Polymer Mineralization in the Standard Soil Mix (SM-L8) Mineralization Treatment %ThCO Polymer substrate Cellulose

P3HB

SA (d.s. 1.5)

Storage (temp/ time) — 4 8 C/ 3 4 8 C/ 3 — 4 8 C/ 3 4 8 C/ 3 — 4 8 C/ 3 8 4 C/ 3

mo mo mo mo mo mo

Stblza (days)

LAGb (days)

PDPc (days)

r pdp d (mg C d − 1 )

25 2 25 25 2 25 25 2 25

3 2 3 2 3 4 2 3 3

6 8 7 9 11 8 25 IND j 32

22.0 4.4 11.6 16.0 7.0 17.6 3.2 2.0 3.1

28 dayse 85.7 57.2 64.0 91.8 66.2 89.7 37.4 21.5 32.0

a b b a b a a b a

MAX-CO2 f

RBIg

kh (day − 1 )

t 50 i (days)

85.3 59.4 64.6 96.5 75.0 99.6 57.5 IND j 58.5

1.00 0.67 0.75 1.00 0.72 0.98 1.00 0.57 0.86

0.149 0.103 0.139 0.101 0.077 0.092 0.038 IND j 0.030

6.3 4.8 9.0 7.0 9.5 11.5 17.1 IND j 21.8

a Duration of the stabilization period (ambient temperature and 60% WHC). b Time required for net mineralization to reach 10% of the MAX-CO . Note:

In those cases where the net mineralization curve did not reach a 2 plateau, the LAG was defined as the time required for net CO2 –C evolution to reach 10% ThCO2 . c Primary degradation phase is defined as the time from the end of the lag period to start of the plateau region (i.e., the point where net mineralization c two-thirds MAX-CO2 ). d Average rate of mineralization during the primary degradation phase is defined as the slope of the linear least-squares regression line plotted between the end of the lag period and start of the plateau region. Note: In those cases where the net mineralization curve did not reach a plateau, the r pdp was calculated for the period from the end of the lag period to the end of the test exposure. e The total amount of CO –C evolved during the test exposure. For each polymer type, values followed by the same letter are not significantly 2 different ( p c 0.05). f The maximum amount of CO –C evolved during mineralization of the test substrate, calculated using Eq. (1). 2 g Relative biodegradation index c (net mineralization following cold storage ÷ net mineralization prior to cold storage). h Calculated using Eq. (1). i Time required for substrate mineralization to reach 50% ThCO . 2 j Indeterminate.

have been successfully rejuvenated during the 25-day stabilization period. That is, following the 25-day stabilization period, values for cumulative (28-day) net mineralization, MAX-CO2 , r pdp , and k were comparable to those obtained prior to cold storage (see Fig. 4 and Table III). These results suggest that there was an increase in the population(s) of P3HB degraders during the extended stabilization period. Mergaert and Swings [10] reported that there is considerable biodiversity of P3HB degrading microorganisms in most environments—particularly in soils and composts. Thus, it should not be surprising that one or more P3HB degraders (perhaps a spore-forming bacterium or actinomycete) would survive the cold storage treatment and thrive under the more favorable conditions of the polymer mineralization test.

exhibit slower degradation kinetics than cellulose [9]. These results notwithstanding, however, the effects of cold storage on mineralization of the starch acetate were

Starch Acetate (d.s. 1.5) Net mineralization of the starch acetate proceeded at much slower rates and to a much lesser extent than either cellulose or P3HB (Fig. 5; Table III). These results are presumably a reflection of chemical substitutions in the starch—in much the same way that cellulose acetates

Fig. 4. Effects of cold storage and length of the stabilization period on net mineralization of poly(3-hydroxybutyrate) in the standard soil mix (SM-L8). Soil maintained at 308 C and 60% WHC throughout the biodegradation test exposure. Legend, storage temperature (8 C)/ stabilization time (days).

Polymer Mineralization in Soils

87

Fig. 5. Effects of cold storage and length of the stabilization period on net mineralization of starch acetate (d.s. 1.5) in the standard soil mix (SM-L8). Soil maintained at 308 C and 60% WHC throughout the biodegradation test exposure. Legend, storage temperature (8 C)/ stabilization time (days).

Fig. 6. Effects of cold storage and length of the stabilization period on net mineralization of cellulose in the Bridgehampton silt loam soil (RI-1). Soil maintained at 308 C and 60% WHC throughout the biodegradation test exposure. Legend, storage temperature (8 C)/ stabilization time (days).

similar to those observed for P3HB. That is, cold storage resulted in significant decreases in both cumulative net mineralization ( − 43%) and r pdp ( − 38%). Moreover, following cold storage, the mineralization curve no longer fit the exponential model (Eq. 1) so that neither an overall rate coefficient or MAX-CO2 could be calculated. As with the P3HB system, the effects of cold storage on mineralization of the starch acetate were largely offset by preconditioning the soils at 60% WHC and ambient temperature for 25 days (Fig. 5; Table III)—presumably reflecting an increase in the population(s) of starch acetate degraders during the extended stabilization period.

light of the relatively low SOC content of the soil, the possibility exists that the added substrates (i.e., test polymers) may have preferentially stimulated the growth of polymer-degrading segments of the microbial population. Regardless, these results demonstrate that the polymer-degrading microorganisms native to the RI-1 soil are more resilient than those in the artificial soil mix (SM-L8). Whether this is a reflection of biological adaptation, environmental factors, or some combination of both remains to be determined.

Polymer Mineralization in the Bridgehampton Silt Loam Soil (RI-1) In general, the potential of the RI-1 soil to mineralization the test polymers was not impaired by the effects of cold storage (Figs. 6–8). On the contrary, two of the polymers (cellulose and P3HB) exhibited increased rates of mineralization and significantly greater amounts of net CO2 production during the 30-day test exposure following cold storage than when the biodegradation tests were performed using fresh soil (Table IV). Given that the temperatures at which the soils were stored (48 and 228 C) are typical of temperate regions, such as Rhode Island, it can be expected that the indigenous microorganisms in this soil are adapted to temperature changes such as those experienced in this study. Moreover, in

Fig. 7. Effects of cold storage and length of the stabilization period on net mineralization of poly(3-hydroxybutyrate) in the Bridgehampton silt loam soil (RI-1): Soil maintained at 308 C and 60% WHC throughout the biodegradation test exposure. Legend, storage temperature (8 C)/ stabilization time (days).

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88

a short (2-day) stabilization period was insufficient to rejuvenate the soil microorganisms—resulting in smaller microbial populations, decreased rates of net mineralization, and less cumulative net CO2 –C production from the polymers (i.e., mineralization) during the test exposure. All of these factors can confound interlaboratory comparisons. Our data indicate, however, that these effects may be negated (at least partially) by extending the stabilization period prior to biodegradation testing. In nearly all cases, the extended (25-day) stabilization period resulted in increased rates of polymer mineralization, as well as in significant increases in cumulative net mineralization. In fact, significant increases (40 ± 2%) in cumulative net mineralization of both cellulose and P3HB were observed for the RI-1 soil. In general, plate counts on selective media provide only a gross indication of microbial populations and, hence, are of only limited use in assessing the effects of environmental variables (e.g., temperature) on the structure of the soil microbial community. Thus, understanding how environmental factors affect soil microbial populations and activity will require the use of more sophisticated methods of analyzing both microbial community structure and population dynamics. Our data also suggest

Fig. 8. Effects of cold storage and length of the stabilization period on net mineralization of starch acetate (d.s. 1.5) in the Bridgehampton silt loam soil (RI-1). Soil maintained at 308 C and 60% WHC throughout the biodegradation test exposure. Legend, storage temperature (8 C)/ stabilization time (days).

SUMMARY AND CONCLUSIONS In general, cold storage had a dramatic effect on both the population of soil microorganisms and their capacity to degrade the test polymers. It was demonstrated that for soils stored for up to 8 months at 48 C,

Table IV. Effects of Cold Storage and Preconditioning on Polymer Mineralization in the Bridgehampton Silt Loam Soil (RI-1) Mineralization Treatment %ThCO2 Polymer substrate

Storage (temp/ time)

Stblza (days)

LAGb (days)

PDPc (days)

r pdp d (mg C d − 1 )

Cellulose

— 48 C/ 8 mo — 48 C/ 8 mo — 48 C/ 8 mo

25 25 25 25 25 25

11 8 15 11 4 2

13 15 9 10 15 12

3.4 4.7 4.2 5.9 1.4 1.0

P3HB SA (d.s. 1.5)

a Duration of the stabilization period (ambient temperature and 60% WHC). b Time required for net mineralization to reach 10% of the MAX-CO . Note:

30 dayse 34.8 49.3 38.7 53.4 17.3 15.5

b a b a a a

MAX-CO2 f

RBIg

kh (day − 1 )

t 50 i (days)

IND j 91.5 84.2 IND j 24.8 17.3

1.00 1.42 1.00 1.38 1.00 0.89

IND j 0.012 0.045 IND j 0.040 0.064

IND j 39.3 22.6 IND j 17.2 10.9

In those cases where the net mineralization curve did not reach a 2 plateau, the LAG was defined as the time required for net CO2 –C evolution to reach 10% ThCO2 . c Primary degradation phase is defined as the time from the end of the lag period to start of the plateau region (i.e., the point where net mineralization c two-thirds MAX-CO2 ). d Average rate of mineralization during the primary degradation phase is defined as the slope of the linear least-squares regression line plotted between the end of the lag period and start of the plateau region. Note: In those cases where the net mineralization curve did not reach a plateau, the r pdp was calculated for the period from the end of the lag period to the end of the test exposure. e The total amount of CO –C evolved during the test exposure. For each polymer type, values followed by the same letter are not significantly 2 different ( p c 0.05). f The maximum amount of CO –C evolved during mineralization of the test substrate, calculated using Eq. (1). 2 g Relative biodegradation index c (net mineralization following cold storage ÷ net mineralization prior to cold storage). h Calculated using Eq. (1). i Time required for substrate mineralization to reach 50% ThCO . 2 j Indeterminate.

Polymer Mineralization in Soils an apparent connection between the supply of available carbon in the soil and the potential of the soil microorganisms to mineralize added polymeric substrates. However, additional research will be required to more fully elucidate the nature of this relationship. It is recommended that: (1) whenever possible, biodegradation tests be conducted within 2 to 3 days of soil collection; (2) for both short- and long-term storage, soils should be maintained at 40–50% WHC and kept in the dark at a temperature of about 48 C; and (3) following storage, the soil should be preincubated for 3–4 weeks prior to initiating the biodegradation test to rejuvenate the soil microbial populations as much as possible. These procedures can help to lessen the adverse effects of cold storage and facilitate interlaboratory comparisons. ACKNOWLEDGMENTS Financial support for this study was provided by the National Science Foundation-Industry/ University Cooperative Research Center for Biodegradable Polymer Research at the University of Massachusetts Lowell. The support of the Center’s co-directors (Drs. R. A. Gross and S. P. McCarthy), and the technical assistance of Thomas Adamczyk are gratefully acknowledged. REFERENCES 1. J. P. E. Anderson (1987) in L. Somerville and M. P. Greaves (Eds.), Pesticide Effects on Soil Microflora, Taylor & Francis, Philadelphia, PA, pp. 45–60.

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