Biomacromolecules 2000, 1, 577-583
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Enzymatic Hydrolysis of Oligomeric Models of Poly-3-hydroxybutyrate Thomas M. Scherer,†,‡ R. Clinton Fuller,§ Steve Goodwin,| and Robert W. Lenz*,‡ Department of Polymer Science and Engineering, Department of Biochemistry and Molecular Biology, and Department of Microbiology, University of Massachusetts, Amherst, Massachusetts 01003 Received February 29, 2000; Revised Manuscript Received September 28, 2000
The mechanism of the enzymatic degradation of poly([R]-3-hydroxybutyrate) (PHB) was investigated by using well-defined model substrates, including both linear and cyclic [R]-3-hydroxybutyrate (3HB) and [R]-3-hydroxyvalerate (3HV) oligomers, with two different PHB depolymerases. The linear and cyclic oligomers containing from 2 to 10 repeating units were hydrolyzed in solutions of the depolymerase isolated from Aspergillus fumigatus and Alcaligenes faecalis, and the rates of hydrolysis and types of products formed were characterized. Both of the depolymerases catalyzed the hydrolysis of the cyclic oligomers (macrolides) which contained more than three 3HB and 3HV repeating units. The degradation reactions of the linear and cyclic 3HB oligomers with the A. fumigatus depolymerase gave similar ratios of monomer-to-dimer products, but PHB itself formed mostly monomer on hydrolysis, indicating that the enzymatic hydrolysis reactions occurred by different mechanisms for these different types of substrates. The results of this study conclusively show that at least the endo mode of polymer hydrolysis occurs with the two enzymes studied, while the A. fumigatus depolymerase was found to utilize both endo and exo modes of hydrolysis to efficiently degrade PHB and 3HB oligomers. Introduction The wide ecological occurrence of bacterial polyhydroxyalkanoates (PHAs) as reserve polymers in many bacteria is accompanied in nature by microorganisms that utilize these polyesters by secreting degradative enzymes. As a result, bacterial PHAs can be readily degraded by many types of microbial populations in marine, active sewage sludge, soil, and compost environments.1,2 The microorganisms in these ecosystems can secrete a class of esterases known collectively as extracellular PHA depolymerases, and many prokaryotic and eukaryotic enzymes of this type have been purified and characterized.3-6 In recent years, the characterization of these PHA depolymerases has contributed information about their biological functions and structural features, and it is known that the ability of these extracellular depolymerases to hydrolyze PHA substrates is determined by the stereochemistry and the types of substituents on the polymers.7-9 Additionally, copolymer content,10 sequence distribution,11 and morphology12,13 were all found to influence the rate and extent of polyester degradation by such depolymerases. The molecular weights of the depolymerases which are particularly active for catalyzing the degradation of poly-3hydroxybutyrate (PHB) are generally between 37 and 60 kDa depending on the microbial source. The primary structures * Corresponding author: Telephone: (413) 545-3060. Fax: (413) 5452595. E-mail address:
[email protected]. † Present address: Wyatt Technologies Corp., Santa Barbara, CA 93117. ‡ Department of Polymer Science and Engineering. § Department of Biochemistry and Molecular Biology. | Department of Microbiology.
of all of these enzymes have been found to include a “lipase box” consensus sequence, as well as conserved regions containing aspartate and histidine residues, indicating that the PHB depolymerases belong to a family of serine esterases.8,14 Primary sequence analysis and limited proteolytic digests have independently provided evidence that a two-domain, tertiary structure organization is common for the different PHB depolymerases.14-16 Despite the well-characterized nature of the enzymes and of the polymers investigated, the process of polymer chain cleavage during these hydrolytic reactions is little understood, especially the nature of the polymer-enzyme interactions. The degradation processes have been investigated primarily through studies of the heterogeneous kinetics of polymer degradation from which several models of enzyme behavior have been proposed for the degradation of PHB.17-19 The types of oligomers formed by hydrolysis have been reported for several different studies.20-22 Results from recent investigations with crude preparations of the A. delafieldii depolymerase on cyclic oligomers of [R]- and [S]-3-hydroxybutyrate ([R]-3HB and [S]-3HB), and also on linear [R]-3HB octamers which were terminated with protecting groups, suggested that endo-type degradation occurred in most cases.23 In contrast, degradation studies of PHB single crystals with the depolymerases from Aspergillus fumigatus and P. lemoignei24 indicated that these enzymes degraded the lamellae from the edges without decreasing the polymer molecular weight, which suggested that an exo-type of chain cleavage could also be involved. The study described in the present report utilized linear and cyclic oligomers as models for PHB to provide direct
10.1021/bm000012c CCC: $19.00 © 2000 American Chemical Society Published on Web 11/22/2000
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evidence for the mechanism of action of the extracellular depolymerases from A. fumigatus and Alcaligenes faecalis in the enzymatic hydrolysis of that polyester. Well-defined cyclic and linear 3HB and 3HV oligomers were used to determine the effects of substrate configuration and size on the kinetics and product distributions of the degradation reactions. Experimental Section Materials. PHB powder with a surface area of 8.0 m2/g, as determined by BET measurement25 was used as obtained from Polysciences Inc. (Warrington, PA). [R]-3HB and [R]-3hydroxyvalerate, [R]-3HV, cyclic oligomers (macrolides) and [R,R,R]-3-hydroxybutyrate and [R,R,S]-3-hydroxybutyrate cyclic trimers (triolides), were synthesized, purified and characterized by procedures according to Seebach and coworkers.26,27 Linear [R]-3-hydroxybutyrate dimer, trimer, tetramer, octamer, and linear octamers of [R,S]-3-hydroxybutyrate with controlled enantiomeric contents were provided Prof. D. Seebach of the Eidgenossische Technische Hochschule in Zurich, Switzerland.28 All solvents and reagents were obtained from J. T. Baker Chemical Company (Phillipsburg, NJ) and Aldrich Chemical Co. (Milwaukee, WI) and were used without further purification. Buffers and biochemical reagents were obtained from Sigma Chemical Co. (St. Louis, MO). PHB Depolymerase. Escherichia coli strain JM109 with plasmid pUC8 carrying the A. faecalis PHB depolymerase gene29 was obtained from Metabolix Inc. (Cambridge, MA). Cultures were grown and maintained on L-broth media (Difco Laboratories) supplemented with 50 mg/L ampicillin. Liquid fermentations of E. coli JM109 utilized L-broth/ ampicillin. Twelve liters of medium was inoculated with a 1.0 L culture of E. coli cells. After 6 h growth at 37 °C, 200 rpm, and aeration of 4 L/min, the cells were harvested and centrifuged at 3500 rpm. The cell pellets (67 g) were suspended in 120 mL of 50 mM tris-HCl pH 7.5 buffer and ruptured by two passes through a French press. The ruptured cell mass was treated with DNAse (0.05 mg/g of cells) at 4 °C and centrifuged to obtain a crude protein solution. Further purification of the crude transgenic A. faecalis PHB depolymerase was conducted according to the procedures of Shirakura et al.30 The depolymerase sample used contained only minor contaminants, which were observed as a second protein band by SDS-PAGE and coomassie blue staining, and had an activity of 1300 AU/mL (see below) as determined by the spectrophotometric PHB assay at 37 °C. The enzyme activity unit, AU, is measured by the change in the intensity of scattered light, or the absorption, of the suspension of the polymer powder in water as a function of time, as described in the following section. PHB depolymerase from A. fumigatus was produced and purified to homogeneity, as determined by SDS-PAGE and coomassie blue staining.6 The purified fungal PHB depolymerase preparation had a protein concentration of 20 µg/ mL and a standardized activity of 3400 AU/mL (see below) by the spectrophotometric PHB assay at 45 °C. Spectrophotometric Assay. PHB powder, at a concentration of 300 µg PHB/mL suspended in 50 mM Tricine-NaOH
Scherer et al.
containing 0.05 mM CaCl2 at pH ) 8.0, was suspended by sonication for 10 min. The resulting suspension was used for determining depolymerase activity as a standard method. The reaction was initiated by the addition of enzyme to 1.0 mL of PHB suspension in a 1.5 mL (10 mm) disposable cuvette at room temperature. After an initial measurement of absorption (A) at 660 nm, the assay mixture was incubated at 45 °C with the A. fumigatus PHBase and at 37 °C for the A. faecalis PHBase. The change in absorption was measured as a function of time with a model 24 UV-vis Beckmann Instruments spectrophotometer (Irvine, CA). A unit of enzyme activity, AU, is defined under these conditions as a change in absorption of 0.001 units per minute (∆mAU/min). Titrimetric Analysis. The contents of the reaction vessel, a stirrer, 2.0 mL of double distilled water, and the substrate, was brought to pH 8.0 with 5.0 mM NaOH under an H2Osaturated nitrogen atmosphere. Substrate hydrolysis was initiated by the addition of 10 µL of purified depolymerase preparation. A heated water bath maintained a constant reaction temperature of 30 °C unless stated otherwise. Reaction pH was maintained at 8.0 by titration of the ester hydrolysis products with 5.0 mM NaOH (degassed) using a recording titristat instrument consisting of a Radiometer pH Meter 26, Titrator 11, Titrigraph SBR2c, Syringe buret SBU1a (Radiometer, Copenhagen, Denmark) fitted with a 2 mL syringe and Teflon/o-ring plunger (Aldrich Chemical Co., Milwaukee, WI) and a combination pH microelectrode M-410 (Microelectrodes Inc., Londonderry, NH). Degradation Kinetics. The determination of oligomer degradation kinetics was conducted using the titristatic method. Approximately 1.0 mg of sample was weighed into a 5.0 mL glass vial. The sample was dissolved in 100-200 µL of dichloromethane to form a solution that was used to coat the interior surface of the vial (approximately 7.5 cm2). Following solvent evaporation, 2.0 mL of distilled H2O was added to the reaction vial, brought to pH 8.0 and allowed to equilibrate to 30 °C in the water bath. The hydrolysis reaction was initiated by the addition of 10 µL purified depolymerase (34 AU A. fumigatus, 13 AU A. faecalis) preparation. All reactions were conducted under the conditions of enzymelimited kinetics, with ratios of enzyme concentration to surface area 20 times lower than those approaching the Vmax of the A. fumigatus PHBase of 2 µg/mL to 8.0 cm2 PHB. The enzymatic degradation of linear and cyclic 3HB and 3HV oligomers was conducted in triplicate. Rates of nonenzymatic hydrolysis for all substrates were determined under identical conditions omiting only the addition of enzyme to the reaction medium. Degradation Product Formation Reactions and Analysis. In a glass reaction vial, 3.0 mg of substrate was added to 150-250 µL of dichloromethane, using the solution to coat the interior surface of the vial. After the solvent had evaporated, 6.0 mL of 50 mM Tricine-NaOH buffer (pH 8.0) and a stirbar were added. The reaction vessel and buffer were maintained at 30 °C in a water bath, and a 0.5 mL aliquot was removed and frozen in liquid nitrogen (t ) 0.0). The addition of 10 µL of A. fumigatus depolymerase preparation initiated the reaction, followed by the removal of 0.5 mL fractions of the reaction medium at 5 min intervals
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that were immediately frozen in liquid nitrogen. Samples were stored at -20 °C until the degradation products were analyzed by HPLC. Oligomer and polyester degradation products were analyzed using a Shimadzu LC-6A HPLC, SPD-6A UV-vis detector (D2 lamp) at 210 nm and a Biorad HPX-87H column with a 5.0 mM H2SO4 mobile phase at a 0.5 mL/min flow rate. Purified samples of 3HB monomer, dimer, trimer, and tetramer were used as product reference standards to calibrate the HPLC detector response to the degradation products. Solution concentrations of 250, 75, and 37.5 µg/mL of the oligomers were used for that purpose. Chromatograph retention times for the linear degradation products were as follows, in minutes: 3HB monomer, 15.1; 3HB dimer, 18.2; 3HB trimer, 22.5; 3HB tetramer, 26.7; 3HB pentamer, 31.6; 3HV monomer, 18.7; and 3HV dimer, 27.9. Results and Discussion Linear Octamer Oligomers of [R]- and [S]-3HB. The degradation, or hydrolysis, reactions of a series of linear octamers containing both [R]- and [S]-3-hydroxybutyrate, 3HB, units were investigated to determine the origin of the stereochemical specificity of PHB depolymerases as reported by Seebach and co-workers.28,31-32 The [R]- and [S]-HB units were arranged in block sequences with segment lengths that varied from one to four units. The oligomers were endcapped with benzyl ether, BzO, groups on the hydroxyl terminus and with tert-butyl alcohol ester, tBuO, groups on the carboxylic acid terminus. The general structures of this series of linear oligomers were as follows: BzO([S]-3HB)n-([R]-3HB)8-n-OtBu I BzO([R]-3HB)n-([S]-3HB)8-n-OtBu II The oligomers are designated in the following discussion according to the sequence lengths of [S]-3HB and [R]-3HB units; that is, as either SnR(8 - n) for series I or RnS(8 n) for series II. For example, the oligomer with a sequence of four successive [S]-3HB units at the benzyl ether terminus followed by four [R]-3HB units with a tert-butyl ether terminus is designated: S4R4; that is, the units adjacent to the benzyl ether terminus are always listed first. Titristatic analysis was used to follow the hydrolysis reactions of these octamers catalyzed by the PHB depolymerase from A. fumigatus. The initial degradation rates (Vo), which are summarized in Table 1, were determined by titration of the carboxyl groups formed during the first 10 min of the hydrolysis reaction. As expected from the results of previous studies, which showed the PHB depolymerases were selective for [R] units, the rates of degradation of the 3HB octamers were found to be strongly influenced by the enantiomeric composition.31,32 The octamers with the longest sequences of S units, S4R4 and R4S4, were the slowest to degrade, and the degradation rates for the linear octamers increased with increasing length of the [R] unit sequences in both series. Surprisingly, however, greater rates of hydrolysis occurred when octamers in Series II were substituted with either one or two [S] segments at the carboxyl terminus. For both series, the results showed that
Table 1. Rates of Hydrolysis of Linear Octamers Catalyzed by the Depolymerase from A. fumigatus at 30 °C octamer oligomera
Vo, µmol/min
extent of enzymatic hydrolysis,d µmol of NaOH
series I: S1R7 S2R6 S3R5 S4R4 series II: R4S4 R5S3 R6S2 R7S1 R8 tBDMSi-R8-Bz R8 unprotectedb PHBc
0.14 0.075 0.058 0.048 0.058 0.078 0.10 0.13 0.080 0.032 0.14 0.008
4.45 2.95 1.98 1.48 1.51 1.90 2.70 3.57 3.99 1.54 4.34 0.46
a See text for designation of oligomers. b Contained free hydroxyl and carboxyl end groups. c High molecular weight poly-3-hydroxybutyrate. d After 50 min of reaction titration of 1 mg samples as a percentage of total 3-hydroxybutyrate ester units theoretically cleavable by PHBase (seven per molecule). Many of the reactions were halted before reaction completion or cessation of enzyamtic activity.
the depolymerase could attack segments of [R]-3HB as short as four units long regardless of the types of units or end groups to which they were attached.32 The hydrolysis reaction rates for the linear octamer samples consisting of eight [R]-3HB units with the different types of protecting groups, and also for the unprotected octamer listed in Table 1, showed that the enzymatic hydrolysis of the stereoblock octamers was not greatly affected by the presence of the protecting groups, which verifies the qualitative observations previously reported by Brandl and co-workers.23 However, the octamer in Table 1, which had tert-butyldimethylsilyl ether, tBDMSi, and a benzyl ester, Bz, protecting groups, degraded more slowly than the other protected octamer and both degraded more slowly than the RS octamers which had ether and ester end groups, which indicated that the presence and type of protecting group can influence the rate of degradation, although the overall degree of enzymatic hydrolysis was not affected by the type of end group. Cyclic Oligomers of [R]-3HB and [R]-3HV. A series of cyclic oligomers containing only [R]-3HB units and varying in size from three to eight units (not including the sevenunit oligomer) was also studied for their rates of hydrolysis by the PHB depolymerase from A. fumigatus. A second series containing only [R]-3-hydroxyvalerate, 3HV, units, varying in size from three to 10 units, was also evaluated for comparison. The rate results obtained for these two series are collected in Table 2. The rates are reported with any chemical or nonenzymatic hydrolysis rate subtracted as baseline. Under the conditions and time scales of these experiments, all subtrates were found to be quite stable to chemical hydrolysis. The cyclic oligomers are referred to in this report as macrolides, a term generally applied to such cyclic esters, and individual oligomers are designated by their ring size. Hence, the three-membered cyclic oligomer (cyclic trimer) is designated as a triolide, and so on. Only the cyclic triolides of both 3HB and 3HV in the macrolide series were not hydrolyzed by the fungal enzyme,
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Table 2. Rates of Hydrolysis of [R]-3HB and [R]-3HV Macrolides Catalyzed by the Depolymerase from A. fumigatus PHB at 30 °C
Scherer et al. Table 4. Rates of Hydrolysis of Linear Oligomers of [R]-3HB by the Hydrolase from A. fumigatus at 30 °C
macrolidea
Vo, µmol/min
relative reactivity,b %
linear oligomer
Vo, µmol/min
3HB:triolide tetralide pentalide hexalide octalide PHB 3HV:triolide tetralide pentalide hexalide heptalide octalide nonalide decalide
0 0.56 0.32 0.29 0.28 0.008 0 0.25 0.037 0.061 0.042 0.055 0.054 0.059
0 7000 4000 3600 3500 100 0 3100 460 760 520 680 680 740
dimer trimer tetramer octamera amorphous PHB crystalline PHB
0 0.27 0.50 0.14 0.054 0.001
a See text for terminology. b Relative reactivity was calculated as a percent of the rate of enzymatic hydrolysis of PHB as the standard.
Table 3. Rates of Hydrolysis of [R]-3HB and [R]-3HV Macrolides Catalyzed by the Depolymerase from A. faecalis at 30 °C macrolidea
Vo, µmol/min
relative reactivity,b %
3HB: triolide tetralide pentalide hexalide PHB 3HV: triolide pentalide hexalide heptalide
0 0.40 0.27 0.37 0.011 0 0.006 0.010 0.008
0 3600 2480 3350 100 0 58 87 76
a
See footnotes in Table 1.
while the tetralides in both series reacted most rapidly, and all of the active cyclic oligomers reacted much faster than PHB itself. The 3HB macrolides were hydrolyzed by the A. fumigatus depolymerase at rates 2-10 times faster than the rates of their 3HV counterparts, as shown by the data in Table 2. In film weight loss studies of polyesters previously carried out in this laboratory it was shown that the A. fumigatus PHB depolymerase degraded PHB at approximately twice the rate of poly-3-hydroxyvalerate, PHV, possibly because of effects of substituted group interactions on the activity of the depolymerase.6 The bacterial PHB depolymerase from A. faecalis was also evaluated, and it too had hydrolytic activity toward the 3HB and 3HV macrolides as shown by the data in Table 3. As with the A. fumigatus depolymerase, the triolides were not hydrolyzed while the tetralide was hydrolyzed somewhat more rapidly in the 3HB series with this depolymerase (the 3HV tetralide was not evaluated). The larger rings again hydrolyzed at similar rates, which is in agreement with the previous observation of the independence of ring size on rate for linear [R]-3HB oligomers that were larger than the tetramer.22 The A. faecalis depolymerase exhibited much less hydrolytic activity toward the 3HV oligomers than 3HB oligomers, in agreement with the previous report of its lower reactivity toward PHV than PHB.15 The observed rate of hydrolysis of the cyclic oligomers by the depolymerase from A. fumigatus and A. faecalis
a
From Table 1.
clearly showed these enzymes could catalyze the hydrolysis of such substrates if they contained more than three 3-hydroxybutyrate units. Therefore, both enzymes must be capable of catalyzing endo-type hydrolysis reactions. A more detailed study of the activity of the 3HB triolide indicated that this cyclic trimer acted as an inhibitor for both types of PHB depolymerases. For example, triolide concentrations of 50 and 250 µg/mL inhibited 67% and 93%, respectively, of the original fungal hydrolase activity, while concentrations of 50 and 250 µg/mL inhibited 88% and 98%, respectively, of the bacterial hydrolase activity toward PHB. 6 Linear Oligomers of [R]-3HB of Varying Size. The enzymatic hydrolysis rates of a series of linear [R]-3HB oligomers, with free hydroxyl and carbonyl end groups, containing two, three, four, and eight units by A. fumigatus depolymerase was also investigated with the results shown in Table 4. This study is very similar to that carried out by Bachmann and Seebach for the A. faecalis depolymerase, and we cooperated closely in our investigations.32 As was the case in the Bachmann-Seebach study, the linear dimer of [R]-3HB was not hydrolyzed, but the 3HB oligomers larger than dimer in Table 4 were rapidly hydrolyzed by the A. fumigatus depolymerase. Indeed, the dimer and monomer of 3-HB are the final products formed in the hydrolysis of PHB by this enzyme, which ultimately degrades the polymer to both types of products. It is interesting to note that the fungal enzyme hydrolyzed the tetramer at a faster rate than either the trimer or octamer, and all three of these linear oligomers were hydrolyzed much faster than PHB. Data for the rates of hydrolysis of both the amorphous and crystalline fractions of PHB, which were obtained in previous studies,6 are included for comparison. It is also of interest to note that while the cyclic trimer of 3HB was not hydrolyzed by the enzyme, the linear trimer was. This result suggests that the conformation of the former is such that it cannot be accommodated by either or both the binding and/or catalytic domains of the enzyme. In sharp contrast, the linear tetramer was hydrolyzed at essentially the same rate as the cyclic tetramer (3HB tetralide in Table 2). Indeed, as noted above, the linear tetramer was the most reactive of the three linear active oligomers in Table 2, so its size or shape corresponds to an optimum for binding to the catalytic site and being hydrolyzed by the enzyme.33 Degradation Products from Macrolides. The enzymatic hydrolysis rates for the degradation of the HB macrolides and linear oligomers by the A. fumigatus depolymerase are shown in Tables 2 and 4, respectively. Both types of substrates gave monomeric and dimeric products in a molar ratio of close to 2:1, as shown in Table 5. In contrast, as
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Enzymatic Hydrolysis of Oligomeric Models Table 5. Products of the Enzymatic Hydrolysis of Cyclic and Linear 3HB Oligomersa by the Hydrolase from A. fumigatus no. of HB units in oligomer 3 4 5 6 8 PHB a
monomer:dimer mole ratio from cyclic oligomera 2.0 ( 0.1 1.8 ( 0.1 1.9 ( 0.4 2.0 ( 0.1
from linear oligomera 1.4 ( 0.1 1.6 ( 0.1
2.1 ( 0.2 6.0 ( 0.1
Oligomers in Tables 2 and 3.
Figure 2. Products from the hydrolysis of 3.0 mg [R]-3HB linear tetramer catalyzed by the A. fumigatus PHB depolymerase at increasing reaction times: (0) monomer; ([) dimer; (O) trimer; (2) tetramer.
Figure 1. Products from the hydrolysis of 3.0 mg [R]-3HB tetralide, catalyzed by the A. fumigatus PHB depolymerase at increasing reaction times: (0) monomer; ([) dimer; (O) trimer; (2) tetramer.
shown by the entry at the bottom of the Table 5, the hydrolysis of PHB by the same enzyme gave a much higher ratio of monomeric-to-dimeric products. Furthermore, the hydrolysis products formed from all of the 3HV macrolides in Table 2 by A. fumigatus PHB depolymerase were found to have monomer-to-dimer ratios closer to 1.5:1 (data not shown) even though the hydrolysis rates followed the same pattern as those of the 3HB macrolides. Apparently, therefore, the active site of the enzyme is very sensitive to the unit structure of the substrate, and in that regard, the degradation product results for the A. fumigatus PHB depolymerase also differed from those obtained with the A. faecalis PHB depolymerase on the same linear oligomers, as reported by Bachmann and Seebach, who observed the formation principally of dimer from the linear 3HB oligomers.32 When degradation product formation was observed over the course of the reaction for the [R]-3HB substrates, the ratio of monomeric-to-dimeric products remained essentially constant at 2:1 as shown in Figure 1 for the [R]-3HB tetralide and at 1.4:1 as shown in Figure 2 for the linear [R]-3HB tetramer. In the hydrolysis products of the cyclic substrates, very small amounts of linear trimer if any, and no higher oligomeric degradation products were observed. As with the cyclic tetralide, the [R]-3-HB hexalide degradation process also did not form measurable amounts of the trimeric product, shown in Figure 3. For the degradation product formation from both the linear tetramer as well as those from the linear [R]-3HB octamer (shown in Figure 4), the amount of trimer
Figure 3. Products from the hydrolysis of 3.0 mg [R]-3HB hexalide catalyzed by the A. fumigatus PHB depolymerase at increasing reaction times: (0): monomer; ([) dimer; (O) trimer; (2) tetramer.
Figure 4. Products from the hydrolysis of the linear [R]-3HB octamer catalyzed by the A. fumigatus PHB depolymerase at increasing reactions times: (0) monomer; ([) dimer; (O) trimer; (2) tetramer.
intermediate formed was clearly observed to pass through a maximum value. These observations are consistent with a model of enzymatic activity that includes both initial attack on the linear oligomers and on the polymer at any point along the chain
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Figure 5. Products from the hydrolysis of PHB by the A. fumigatus PHB depolymerase at increasing reactions times: (0) monomer; ([) dimer.
(endo) and selective attack at the terminal units (exo) to form well-defined degradation products. The cyclic [R]-3HB oligomers (macrolides) form linear substrates after the initial hydrolysis reaction, which must be an endo-type cleavage, and it is surprising that these compounds form much smaller amounts of trimer intermediates than do the linear substrates. The dimer product is presumably formed when the enzyme reaches the end of the linear oligomeric substrate in either case, and it is apparently not an active substrate for further enzymatic cleavage. Dimer product may also be formed when the enzyme innitiates cleavage in endo mode two units away from one end of a linear substrate, while depolymerizing the substrate in the other direction of the chain. Evidence of this exists in the lower ratio of monomer to dimer (higher concentrations of dimer products) and the formation of trimer intermediate products from the linear oligomer substrates. Furthermore, the high ratio of monomeric to dimeric hydrolysis products obtained from PHB indicate that the A. fumigatus depolymerase may remain closely associated to (or bound to) one of the two newly formed terminal units of the hydrolyzed polymer or the cyclic oligomer. After the initial endo-type cleavage, the terminus-associated enzyme may selectively remove longer sequences of monomeric units by exo-type attack before reaching the chain end or dissociating from the remaining polymer chain, as suggested by the product ratio (6:1) of predominantly monomeric [R]-3HB degradation units. Also, during PHB hydrolysis the amount of dimeric product formed appears to reach a maximum, while the concentration of monomeric product formed continues to increase as shown in Figure 5. This result suggests that the primary form of PHB degradation by the A. fumigatus PHBase is exo hydrolysis rather than endo. Conclusions Analysis of the enzymatic hydrolysis of linear and cyclic oligomers containing either [R]-3-HB or [R]-3-HV units reveals several characteristics of the degradation mechanisms of the two depolymerases studied. First, hydrolysis of linear oligomers with a carboxyl and hydroxyl protecting groups at each terminus required blocks of at least four [R]-3HB
Scherer et al.
units, which is consistent with the structure of the binding site of A. faecalis proposed by Bachmann and Seebach.32 Second, purified cyclic oligomers of four or more repeating units were hydrolyzed by both PHB depolymerases investigated, at rates faster than that of linear PHB itself, and that the larger cyclic oligomers hydrolyzed at similar rates. Third, the extracellular depolymerase from A. fumigatus and from A. faecalis hydrolyzed oligomers similarly, with both enzymes initiating degradation of cyclic [R]-3-HB substrates through endo-type hydrolysis and neither enzyme capable of hydrolyzing dimers. On the basis of the observed formation of degradation products from the oligomers, the action pattern of the A. fumigatus PHB depolymerase includes both endo and exo-type hydrolsis modes. Collectively, the results from the enzymatic hydrolysis of linear and cyclic [R]-3HB oligomers provide new insight into the mode of PHB degradation by fungal and bacterial PHB depolymerases. Acknowledgment. Financial support was provided by the Center for Environmentally Appropriate Materials (CEAM) at the University of Massachusetts. Linear and cyclic oligomer compounds were provided by B. Bachmann and Prof. D. Seebach, ETH-Zentrum in Zurich, Switzerland, as part of a cooperative research program. References and Notes (1) Mergaert, J.; Anderson, C.; Wouters, A.; Swings, J.; Kersters, K. FEMS Microbiol. ReV. 1992, 103, 317. (2) Nishioka, M.; Tuzuki, T.; Wanajyo, Y.; Oonami, H.; Horiuchi, T. In Biodegradable Plastics and Polymers; Doi Y., Fukada, K., Eds.; Elsevier Science B.V.: New York, 1994; Vol. 12; pp 584-590. (3) Doi, Y.; Mukai, K.; Kasuya, K.; Yamada, K. In Biodegradable Plastics and Polymers; Doi Y., Fukada, K., Eds.; Elsevier Science B.V.: New York, 1994; pp 39-51. (4) Brandl, H.; Bachofen, R.; Mayer, J.; et al. Can. J. Microbiol. 1995, 41 (Suppl. 1), 143-153. (5) Brucato, C. L.; Wong, S. S. Arch. Biochem. Biophys. 1991, 290, 497-502. (6) Scherer, T. M.; Fuller, R. C.; Lenz, R. W.; Goodwin, S. Polym. Deg. Stab. 1999, 64, 267-275. (7) Jaeger, K. E.; Steinbuchel, A.; Jendrossek, D. Appl. EnViron. Microbiol. 1995, 61, 3113-3118. (8) Schirmer, A.; Matz, C.; Jendrossek, D. Can. J. Microbiol. 1995, 41 (Suppl. 1), 170-179. (9) Abe, H.; Matsumura, I.; Doi, Y.; Hori, Y.; Yamaguchi, A. Macromolecules 1994, 27, 6018-6025. (10) Doi, Y.; Kanesawa, Y.; Kunioka, M. Macromolecules 1990, 23, 2631. (11) Hocking, P. J.; Marchessault, R. H.; Timmins, M. R.; Scherer, T. M.; Lenz, R. W.; Fuller, R. C. Macromol. Rapid Commun. 1994, 15, 447-452. (12) Doi, Y.; Kumagai, Y.; Tanahashi, N.; Mukai, K. In Biodegradation of Polymers and Plastics; Vert, M., Feijen, J., Albertsson, A. C., Scott, G., E. Chiellini, G., Eds.; Royal Society of Chemistry: Cambridge, England, 1992; Vol. 109; pp 139-151. (13) Parikh, M.; Gross, R. A.; McCarthy, S. P. In Biodegradable Polymers and Packaging; Ching, C., Kaplan, D., Eds.; Technomic Publ. Co.: Lancaster, PA, 1993; pp 159-170. (14) Jendrossek, D.; Backhaus, M.; Anderman, M. Can. J. Microbiol. 1995, 41 (Suppl. 1), 160-169. (15) Karuso, Y.; Kohama, K.; Uchida, Y.; Saito, T.; Yukawa, H. In Biodegradable Polymers and Plastics; Doi, Y., Fukada, K., Eds.; Elsevier Science B.V.: Amsterdam, 1994; pp 357-361. (16) Fukui, T.; Narikawa, T.; Miwa, K.; Shirakura, Y.; Saito, T.; Tomita, K. Biochim. Biophys. Acta 1988, 952, 164-171. (17) Mukai, K.; Yamada, K.; Doi, Y. Int. J. Biol. Macromol. 1993, 15, 361-366.
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Biomacromolecules, Vol. 1, No. 4, 2000 583 (27) Seebach, D.; Hoffman, T.; Ku¨hnle, F. N.; Lengweiler, U. D. HelV. Chim. Acta 1994, 77, 2007. (28) Seebach, D.; Brunner, A.; Bachmann, B. M.; Hoffmann, T.; Ku¨hnle, F. N. M.; Lengweiler, U. D. Biopolymers and Oligomers of [R]-3HA: Contributions of Synthetic Organic Chemists; Ernst Schering Research Foundation: Berlin, 1995; Vol. 28. (29) Saito, T.; Suzuki, K.; Yamamoto, J.; Fukui, T.; Miwa, K.; Tomita, K.; Nakanishi, S.; Odani, S.; Suzuki, J.; Ishikawa, K. J. Bacteriol. 1989, 171 (1), 184-189. (30) Shirakura, Y.; Fukui, T.; Tomita, K. Biochim. Biophys. Acta 1983, 784, 331-339. (31) Bachmann, B. M.; Seebach, D. HelV. Chim. Acta 1998, 81, 24302461. (32) Bachmann, B. M.; Seebach, D. Macromolecules 1999, 32, 17771784. (33) Hiromi, K.; Nitta, Y.; Numata, C.; Ono, S. Biochim. Biophys. Acta 1973, 302, 362.
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