The microstructure of polymers enzymatically synthesized in a self-assembling environment

E L S E V I E R Materials Science and Engineering C 4 (1996) 161-168

M A T E R I A L S SCIENCE &

ENGINEERING

C

The microstructure of polymers enzymatically synthesized in a self-assembling environment

Xiaodong Xu a, N. Kommareddi a, M. McCormick a, T. Baumgartner a, V.T. John a,*, G.L. McPherson u, J.A. Akkara c, D.L. Kaplan c

Department of Chemical Engineering, Tulane University, New Orleans, LA 70118, USA b Department of Chemistry, Tulane University, New Orleans, LA 70118, USA

c US Army Soldier Systems Command, Natick, MA 01760, USA

Abstract

Polyphenols are synthesized in the self-assembling environment of reversed micelles using an oxidative enzyme, horseradish peroxidase (HRP). Hydrogen-bonding interactions between the monomer and the surfactant lead to monomer partitioning to the micellar interface. The polymer has the morphology of distinct and interconnected micron-sized spherical particles. The internal microstructure of the particles appears to be made up of nanometer-sized polymer patches. By controlling the extent of reaction it is possible to modify the internal density of these polymer mierospheres. Intramicellar selutes such as enzymes and nanoparticles are entrapped within the microspheres and confer specific properties to the resulting composite. This leads to a method of microencapsulation and the preparation of novel polymer-protein and/or polymer--ceramic composites.

Keywords: Reverse micelles; Microencapsulation; Polymer microspheres

1. Introduction

Lignification is a peroxidase-catalyzed reaction involving a one-electron oxidation ofp-hydroxy, methoxy-substituted cinnamyl alcoh0t units [ 1]. In recent years, this abilility of peroxidases to oxidatively couple phenolic units has led to research in the enzyme-based synthesis of polyphenolics [2,3 ] as alternatives to the chemically synthesized phenol- formaldehyde polymers widely used in the coatings and resins industry [4]. Fig. 1 illustrates a simplified schematic of the oxidative coupling of phenols using peroxidase type enzymes. Phenoxy radical centers that are first formed, migrate to ortho (or para) positions following which coupling occurs. The enzymaticaily synthesized material is envi- ronmentally more acceptable than chemically synthesized polyphenolics due to the avoidance of formaldehyde in the process. However, the enzymaticaily synthesized material is chemically dissimilar to phenol-formaldehyde polymers, since these polymers lack the intervening methylene units between the rings. The direct ring-to-ring coupling shown in Fig. 1 leads to a polymer that is conjugated and therefore electro-optically active. These polymers therefore have appli-

* Corresponding author.

0928-4931/96/$15.00 © 1996 Elsevier Science S.A. All rights reserved PHS0928-4931 (96) 00148-8

H H 0 0

(Poroxldase)

L .J

Fig. 1. A simplified representation of HRP catalyzed polymerization of p-substituted alkylphenols.

cations as conductive polymers and in applications to nonlinear optics [ 3,5 ].

In our earlier research, we have shown the feasibility of enzymatic synthesis of polyphenolics in the environment of water-in-oil microemulsions commonly known as reversed micelles [6,7]. These are water droplets suspended in an organic bulk phase and stabilized by a surfactant, typically an anionic such as bis(2-ethylhexyl) sodium sulfosuccinate, whose structure is shown in Fig. 2(a). The enzyme, horseradish peroxidase (HRP) is encapsulated in the water pools and maintains eatalytic activity. An interesting aspect of such synthesis is the fact that the monomer, being amphiphilic, partitions to the oil-water interface. The situation is shown schematically in Fig. 2(b); the head of the arrow represents

162 X. Xu et aL / Materials Science and Engineering C 4 (1996) 161-168

~)

BIs(2..ethylhexyl) sodium suli'osucclnate (AOT)

Nii:

~'/Surfa~nt i ~ Monomer'

Fig. 2. (a) Structure of the anionic surfactant, AOT. (b) Schematic of a reversed micelle illustrating enzyme solubilization and monomer partitioning to the interface.

polar hydroxyl groups of the phenolic species. As discussed in the text, evidence of monomer penetration to the micellar interface can be clearly seen from the perturbations to the surfactant C=O vibrations.

An interesting aspect of enzymatic polymerization in these systems is the observation that the polymer precipitates out of solution during synthesis in the form of both distinct and interconnected microspheres [7]. In earlier work, we have found that a 3:1 molar ratio of surfactant to monomer in the system reproduceably results in polymer microsphere formation [7]. Since polymer microspheres have many useful applications in coatings technologies due to their high dis- persibility, we have continued work on morphology genera- tion, seeking to reach a better control of polymer particle characteristics. The present paper details our observations on an interesting transition from diffuse to dense microspheres. The implications of being able to encapsulate enzymes in these diffuse microspheres with retention of catalytic activity is also discussed.

2. Materi=ds and methods

2.1. Materials

The anionic surfactant, bis(2-ethylhexyl) sodium sulfosuccinate (AOT) and isooctane (HPLC grade) were obtained from Aldrich Chemicals, Milwaukee, WI. All enzymes, peroxidase (type II: from horseradish), cytochrome-c (from horseheart, type VI) and phosphodiesterase I (from Crotalus atrox), and the buffer, HEPES [N-(2-hydroxyethyl)- piperazine-.N'-(2-ethanesulfonic acid)] were purchased from Sigma Chemicals, St Louis, MO. HzOz, the monomer 4-ethylphenol, and the enzyme substrate bis (4-nitrophenyl) - phosphate were obtained from Aldrich. Polystyrene standards (Aldrich) were used for polymer molecular weight deter- minations. Doubly distilled and deionized water was used in all buffer preparations.

2.2. Methods

2.2.1. Polymer synthesis The synthesis can be visualized through the schematic

shown in Figs. 1 and 2. In a typical experiment, the enzyme (HRP) dissolved in 0.01 M HEPES (pH 7.5) was added to a dry reversed micellar solution of AOT in isooctane, followed by the addition of the monomer, 4-ethylphenol (EP). The enzyme concentration in the buffer was adjusted so that the final enzyme concentration in the micellar solution was 0.5 mg ml- 1. The reaction mixture typically has the overall composition 0.5M AOT, 0.15M 4-ethylphenol, 0.5 mg ml- 1 HRP, and a Wo of 15 (Wo is the water to surfactant molar ratio). The reaction is initiated by the addition of H202 in aliquots to minimize enzyme deactivation. Varying levels of HzOz were added as is explained in the section on results. Polymer rapidly forms and precipitates out of solution. The recovered polymer is sonicated mildly, and thor- oughly washed with isooctane to remove adsorbed AOT and monomer.

2.2.2. Characterizations The polymer morphology was characterized by scanning

electron microscopy (SEM) and transmission electron microscopy (TEM). For SEM analysis a drop of the sample dispersed in iso-octane was placed on an aluminum stub, dried and coated first with 20 nm thickness carbon black and then with gold. The micrograph was taken at an acceleration voltage of 15-30 kV in a Jeol JSM-820 scanning electron microscope. For TEM analysis, a Phillips 410 transmission electron microscope with a L a B 6 (lanthanum hexaboride) crystal electron source was used. The particles were dispersed on carbon-coated grids and directly imaged using an accler- ation voltage of 60 kV.

An 'ATI-Mattson Galaxy 6021 FTIR spectrometer was used for FTIR measurements. A liquid sample cell (SpectraTech) with CaF 2 windows and path length 0.3 mm was used for liquid samples. Molecular weight distributions of the polymers were measured using gel permeation chro- matography (GPC). The set-up consisted of a 25 cm Jordi Gel DVB mixed bed column, a Perkin Elmer bit-compatible binary pump (model 250), and a Perkin Elmer Diode Array detector (model 235) interfaced with a personal computer. The eluent was THF, and the column operated under the eluent head pressure adjusted to maintain the flow rate of 0.75 ml rain- 1. The dry polymer was dissolved in THF at a c o ncentration of 0.1% ( w / v ) and 20/M of this solution was used for the measurements. Polystyrene samples in the molecular weight range of 60(O200 000 were used as calibration standards.

2.2.3. Encapsulated enzyme activity measurements The activity of phosphodiesterase (PD) both in reversed

micelles and encapsulated in the polymer, was assayed using the hydrolysis of bis(p-nitrophenyl)phosphate as the model reaction. The release ofp-nitrophenol from the hydrolysis is

X. Xu et at./Materials Science and Engineering C 4 (1996) 161-168 163

measured through its absorbance at405 nm; the solution turns yellow upon release of p-nitrophenol. Prior to absorbance measurements, the sample was passed through 0.2/zm filters to remove interference from the oligomer/polymer traces in solution.

3. Results and discussion

3.1. Polymerization characteristics

FTIR spectra of the AOT carbonyl vibrations both in the absence and in the presence of varying amounts of the monomer are illustrated in Fig. 3. The perturbation in the vibrational characteristics and the low frequency shift are indicative of hydrogen bonding interactions between the phenol and the surfactant. The low frequency component increases with increasing amounts of the monomer. This ability of the monomer to interact with surfactmlt head groups in the micelle is evidence that the monomer penetrates to the interracial region. In fact, AOT-phenol interactions are so strong that at higher AOT levels, the system becomes an organogel [ 8]. It is to be noted, however, that the concentra- tions used to obtain the spectra of Fig. 3 are not fully repre- sentative of the polymerization reaction and are used primarily to emphasize the C=O perturbations. In the presence of water, which also hydrogen bonds with the surfactant, it is not possible to fully separate out water effects from phenol effects, although the monomer adds to perturbations of the C=O vibrations.

It is therefore reasonable to assume that initial chain formation occurs at the micellar interface. The macroscopic observation is that the solution darkens and a polymer precipitate is formed. The precipitate, when collected and imaged through scanning electron microscopy, shows the morphology of both distinct and interconnected micros-

0.25 i , I AOT 0.05 M

a. 0 E P

b. 0.025 M EP 0.20 e. 0.05 M EP -

d. 0.40 M EP

0.15

8 ¢3

.~ 0.10

.<

0.05

0.00 I r i 1760 1740 1720 1700 1680

Wavenumbers (cm "1) Fig. 3. Perturbation of AOT C=O vibrational frequencies by p-ethylphenol (EP) hydrogen bonding. Spectra taken with water-free reversed micelles.

o e

t

Fig. 4. Scanning electron micrographs of polymer rnicrospheres: (a) a large collection of spheres; (b) showing both clusters and single spheres.

pheres, as reported in our earlier work [7]. Fig. 4(a) is a scanning electron micrograph of large number of polymer particles, and Fig. 4(b) illustrates that both single particles and interconnected particle clusters are formed. In order to understand the morphology development better, the following experiments were carried out.

3.2. Effect of reaction time and H202 levels

Horseradish peroxidase (HRP) catalysis in the presence of H2Oa can be written in the simplified form [ 1 ] :

HRP + H202 ~ compound I

compound I + SH ~ S* + compound II

compoundII + SH ~ S* + HRP

Compounds I and II are intermediate states of the enzyme, compound I resulting from a two-electron oxidation of the heme group and compound II resulting from the one-electron reduction of compound I. S-H is the substrate, either the phenolic monomer or the polymer, and S* is the phenoxy radical species. The overall coupling reaction can be expressed as

HRP

2S - H + HzOa ~ 2S - S + 2H20

and it is noted that each coupling step requires one molecule of HzOz. If it is assumed that most coupling steps involve monomer addition to a chain unit, then there is a 1:1 corre-

164 X. Xu et al. / Materials Science and Engineering C 4 (1996) 161-168

i(a) .

, 2 : .,

° 2 ; ~ : . . . . . . . . . , , . :

:x4414oo ' ..... . . . . . . ?

m . . . . .

Fig. 5. Transmission electron micrographs of polymer microspheres at different stages of densification: (a) 15 rain sample; (b) 4 h sample; (e) 24 h sample.

spondence between the H202 usage and monomer conversion, as has been experimentally observed [ 6].

Our typical synthesis of poly(p-ethylphenol) involves addition of H202 30% in excess of the stoichiometric require- ment for complete conversion of the monomer. The H202 is added in three to four atiquots in succession. Polymer for- marion is rapid and after 5-10 min of reaction time, analysis of the micellar supernatant reveals that over 95% of the initial monomer content has been lost. The typical procedure is to leave the reaction mixture stirred for about 24 h to complete reaction. When the polymer particles are recovered and imaged through transmission electron microscopy they appear internally dense, as shown in Fig. 5 (c). Interestingly, however, when the particles are sampled after 4 h of reaction time, many of them appear to have a diffuse interior (Fig. 5 (b) ) . Inspection of the diffuse particles reveals that they seem to have a dark ring or shell on the outside while the interior seems to be made up of small patches. If the polymer is sampled 15 min after reaction initiation, almost all particles have the characteristic patchy diffuse interior (Fig. 5(a) ) . A count of a few hundred particles imaged through T]EM at different reaction times indicates that about

4% of the particles are filled at 15 min reaction time, 75% are filled in 4 h and 96% are filled over 24 h.

It is thus evident that the polymer particles have an internal microstructure that evolves with reaction time. The overall particle size and spherical morphology develops very rapidly, and it is the interior that gradually densities with time. Fig. 6 is a higher resolution TEM of a single particle where the patchy, diffuse interior can be clearly observed.

The origin of the patchy interior can be understood by further control of particle growth through manipulation of monomer conversion levels. This is accomplished by adjust- ing the H202 addition level. Fig. 7 represents the transmission electron micrograph of the polymer precipitate sampled very soon after adding 0.04 M I-I202, just sufficient for 25% monomer conversion. In addition to the patches, there is initial evidence of the organization of the patches to spheres. A rough estimate of patch size is in the range 5-20 nm, which is not significantly larger than the micelle diameter. It may then be possible that the patches are the consequence of initial growth and nucleation at the micellar interface and that micellar collisions lead to linking and attachments between the patches. The polymer is ' 'sticky" due to hydrogen bonding

X. Xu et aL /Materials Science and Engineering C 4 (1996) 161-168 165

Fig. 6. A higher resolution TEM of a single particle at an early stage of densification.

4

x 75_:~90 - . . . . . . . . . . . . . . . . . . . . . . . . I I . . . . . . . .

Fig. 7. TEM of polymer at an early growth stage when just sufficient HzO2 for 25% monomer conversion is added.

between phenolic hydroxyls, which should also contribute to patch attachments and attachments of the clusters of patches. Fig. 7 thus provides an insight into the formation of polymer particles that are two orders of magnitude larger than the micelle size.

We have observed that the internally diffuse but fully formed microspheres of Fig. 6 are obtained when I-IzO2 is added to approximately 80% monomer conversion. Upon keeping the solution stirred for 24 h, no further increase in internal density is observed. We have found that microspheres with a diffuse interior can be reproduceably obtained through two techniques: (1) by addition of H202 to levels slightly less than total monomer conversion; (2) by recovery of the polymer within 15-30 rain of reaction time (when excess H2Oa is added), following which the particles are thorougly washed with isooctane and water to remove monomer and H202 trapped in the matrix.

The densification of the particle interior is therefore attrib- uted to reaction involving residual monomer, H202 and

Fig. 8. High resolution TEM of gold-labeled peroxidase in diffuse polymer microspheres. The dar k spots in the center of the micrograph are the gold particles.

enzyme that may be entrapped in the polymer matrix. To prove that HRP is entrapped in the polymer particles, a trace of gold-labeled HRP was added to the reaction system prior to synthesis. The electron-dense gold particles are observed in the high resolution TEM of an internally diffuse particle in Fig. 8. Further qualitative evidence of the presence of entrapped enzyme is the fact that the internally diffuse polymer particles, when separated out of the reaction mixture, are capable of carrying out the reaction when contacted with a fresh batch of monomer in reversed micelles. In other words, the diffuse polymer particles contain enzyme with at least partial retention of catalytic activity. Additional evidence for the densification being caused by residual reaction over a 24 h period comes from a measurement of monomer levels in the polymer particle, when the polymer particles are recovered and analyzed for monomer without any washing steps. The monomer levels were measured by dissolving the recovered polymer particles in benzene followed by gas chroma- tography analysis. Early and incomplete H202 addition experiments showed trapped monomer levels around 10% of initial monomer levels. On the other hand, 24 h incubation, excess H202 addition experiments, showed that the levels had fallen to 3% or less of initial monomer content.

The question arises as to whether the residual reaction that leads to densification, results in increased molecular weights with connections between existing chains being created through monomer bridging. The molecular weight data shown in the gel permeation chromatograrns of Fig. 9 indicates that the molecular weight distribution remains virtually constant between an early time sample (the internally diffuse particles) and a 24 h sample (the dense particles). If there is a change with increased reaction time at all, it is to slightly lower molecular weights, and the peak molecular weight changes from about 1300 to 1200 Da. The molecular weight information indicates that particle densification is not the consequence of significant internal chain linkages. It is more

166 X. Xu et aL /Materials Science and Engineering C 4 (1996) 161-168

I I I I I I l L.. 1o5 l - ~ g p ~o,o~e, '= l a. 180%, 5 rNn. J I_~ / " ~ standards o / . . . . . b. 180O/o, 24,r. q

I ' / 1o2f- , , , , , , q .

Retention Time ~" p-ethylphenol

1.3ao.ooo ] z 9o,0oo I: a. 12,ooo i: 4. 4.0oo F.

le. eoo / /

8 10 12 14 16 18 20 22 Retention Time (rain)

Fig. 9. Gel permeation chromatograms illustrating no significant change in molecular weight between diffuse and dense microspheres; 130% H202 added (30% in excess of the stoichiometric level needed for complete monomer conversion).

likely that the densification is the result of new chain growth with unreacted monomer.

Additionally, we wish to correct an error in the molecular weight data reported in our earlier paper on polymer morphology in this journal, where an average molecular weight of 90 kDa was reported [7]. This apparently high molecular weight is the result of rapidly eluting aggregates in the highly hydrogen bonded "sticky" polymer. When the aggregates are broken, the apparent molecular weights are significantly smaller [9]. Similar results have now been obtained in the labororatories of both sets of researchers.

3.3. Enzyme encapsulation

The observations that the internal density of the polymer microspheres can be controlled, and that HRP encapsulated in the interaally diffuse polymer remains catalytically active leads to the possibility that other enzymes may also be encapsulated within the polymer with retention of activity. In order to determine if other enzymes cosolubilized in reversed micelles with HRP are also encapsulated, experiments have been carried out with a model enzyme, cytochrome-c. This is a relatively small heme-containing protein with molecular weight, 12 800 Da. The protein has an absorption peak at 410 nm with minimal interference at this wavelength from phenol oligomers in solution. Fig. 10 illustrates the results of polymerization in the presence ofcytochrome-c cosolubilized in reversed'~ micelles. As observed, the micellar supernatent shows almost completely removal of cytochrome-c, indicat- ing that thi,; protein has become entrapped in the precipitating polymer. At this point, we do not know if the entrapment happens on a micellar scale, when initial chains are nucleated at the micelle interface, or if the entrapment occurs during the secondary stage when the chains coalesce to form larger particles.

I I I I

1.0 t Supernatant (after reaction)

"~. 0.8 • Control (before reaction)

0.6

<

0.4

0'21

- - I - T - v - - - - ' t ' v - - w

350 400 450 500 550

Wavelength (nm)

Fig, 10. Cytochrome-c removal from reversed micelles and entrapment in the polymer matrix. The control sample had all reaction constituents (0.5 mg ml- 1HRP, 0.5 M AOT, wo t5, 0,15 M H202 and 1 mg ml- 1 cytochrome-c) with the exception of the monomer. Absorbances are measured after five-fold dilution to bring the absorbance (410 nm) down to calibration levels and to minimize turbidity effects after polymerization,

The next step is to determine whether these cosolubilized enzymes, when entrapped, also retain catalytic activity. In principle, enzyme encapsulation in microparticles with activity retention has several potential benefits. Such polymer- enzyme composites could be used for enzyme delivery and catalytic coatings. There is a possibility that such encapsulation could improve enzyme stability by minimizing denaturation and preventing enzyme aggregation. The encapsulation could also protect enzymes from protease catalyzed lysis.

We have used phosphodiesterase as an illustrative and useful example for its ability to degrade specific organophosphorus compounds. Organophosphorus compounds range from relatively innocuous insecticides and pesticides to highly potent nerve agents. The general mechanistic scheme for the enzymatic hydrolysis of organophosphorus corn- pounds can be written as [ 10]

X X II Pholphoestera=e II

R - P - Z + H 2 0 ~ R - P - O H + ZH 1 I

R' R'

where X=oxygen or sulfur, R is an alkoxy (or hydroxyl)group, R' is an alkoxy, phenoxy, or phenyl group, and Z is a phenoxy group or a fluorine atom.

In this work, initial experiments have been carried out using the hydrolysis of bis(p-nitrophenyl) phosphate in reversed micelles as the model reaction. Both enzyme and substrate are readily available commercially (Sigma- Aldrich). The enzymatic hydrolysis reaction releasesp-nitrophenol which can be analyzed through its absorbance at 410 nm (the solution turns yellow). Fig. 11 compares the activity of free phosphodiesterase solublized in reversed micelles to the activity of entrapped phosphodiesterase, when

X. Xu et al. /Materials Science and Engineering C 4 (1996) 161-168 167

4.0

3.5 q

" 3.0

2.5

:~ 2.0 g

:~ 1.5 <

1.0 I

t.U

0.5

0.0

[ . -O - free enzyme 1 [ ~ immob'~'zed enzyme I

, u r

I I I I v

1 2 3 4 5 6 7 Time (days)

Fig. 11. Activity of free and polymer-entrapped phosphodiesterase (PD). Reaction carried out in reversed micelles of w o 15, 0.5 M AOT using i mM substrate [bis(p-nitrophenyl)phosphate]. 1 mg m1-1 phosphodiesterase used in the free enzyme studies and in encapsulation. Encapsulated enzyme activity based on the assumption of complete entrapment. Each data point is a separate sample incubated for the prescribed time period at ambient conditions.

contacted with reversed micellar solution containing the substrate. As observed, the initial activity of the encapsulated enzyme is only a third of the activity of the free enzyme, but the encapsulated enzyme exhibits an enhanced stability. It must be noted that the experiments are initial and the encapsulation procedure has not been optimized for maximum activity. The potential for enzyme microencapsulation can only be promoted when a complete study is made of activity and stability in various environments and when a variety of enzymes consistently exhibit reasonable activity and enhanced stabilities. Nevertheless, the fact that the polymer particles with entrapped enzyme are catalytically active provides a motive for further study.

An eventual objective is to use highly efficient phospho- triesterases such as the phosphotriesterase from Pseudo-

monas diminuta [ 10]. Incorporation of these enzymes into microparticles with activity retention may have some novel applications related to chemical decontamination. These polymers could be used in the development of protective coatings on fabrics, as alternatives to the bulky, activated- charcoal based protective suits used presently. The particles could be sprayed together with water, using traditional fire-fighting equipment, to the site of nerve agent decontamination.

The internally dense microparticles are also useful in composite development. It is possible to encapsulate inorganic nanoparticles (ferrites for example), by first synthesizing the nanoparticles in reversed micelles and subsequently con- ducting enzymatic polymerization to encapsulate the nanoparticles. Allowing the polymer to densify leads to full immobilization of the nanoparticles, and the inorganic component then confers its specific properties to the composite.

For example, we have shown that polymer-ferrite composite exhibits superparamagnetism and low temperature spin-glass properties characteristic of the ferrite component. Full details of the composite magnetic properties are listed in an accom- panying publication [ 11 ].

4. Conclusions

The synthesis of polyphenols in a reversed micellar environment leads to polymer microspheres whose internal microstructure can, to a degree, be controlled through manipulation of the extent of reaction. During the course of polymerization and precipitation, the polymer entraps and encapsulates intramicellar solutes such as proteins and or nanoparticles. These encapsulates appear to retain their specific properties in the composite. Enzymes encapsulated in the polymer matrix confer catalytic properties to the composite. Similarly, encapsulated nanoparticles confer their unique properties to the composite (e.g. superparamagnetism). Thus, this is a sort of ship-in-a-bottle approach to the synthesis of polymer-protein and polymer-nanoparticle composites. The fact that these composites can be prepared as microspheres implies an ease in processing for coatings or dispersions technologies.

The work described above is an example of biocatalysis operating in a microstructured fluid environment. The polymer produced as a result of such enzymatic reaction is itself microstructured, and the synthesis environment may provide a template for synthesis. While reversed micelles have often been studied as an analogue to biological membranes [ 12], it is perhaps inaccurate to classify this system and reaction as an example of biomimetics. However, it is clear that the polyphenol produced is of use not just for its chemical properties, but also for its microstructural characteristics. As has been pointed out by Mann and coworkers [ 13], biological materials synthesis is unique for its ability to replicate properties with high fidelity. Polymer synthesis using biological methods may be a route to new and useful microstructures.

Acknowledgements

Financial support from the National Science Foundation, the US Army, and the Tulane/DoD Center for Bioenviron- mental Research are gratefully acknowledged. We are also grateful to Ms Zuzannah Hruska of the Tulane Coordinated Instrumentation Facility for help with electron micrography.

References

[1] B. Halliwell and J.M.C. Gutteridge, Free Radicals in Biology and Medicine, Clarendon Press, Oxford, 1989.

[2] J.L. Popp, T.K. Kirk and J.S. Dordick, Enzyme Microb. TechnoL, 13 (1991) 964.

[3] J.A. Akkara, K.J. Senecal and D.L. Kaplan, J. Polym. Sci., 29 ( 1991 ) 1561.

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[4] P.W. Kopf, Phenolic resins, in Encyclopedia of Polymer Science, Vol. 11, Wiley, New York, 1985.

[5] J.A. Akkara, M. Ayyagari, F. Bruno, L. Samuelson, V.T. John, C. Karayigitoglu, S. Tripathy, K.A. Marx, D.V.G.L.N. Rao and D.L. Kaplan, Biomimetics, 2 (1994) 331.

[6] A.M. Rao, V.T. John, R.D. Gonzalez, J.A. Akkara and D.L. Kaplan, BiotechnoL Bioengng, 41 (1993) 531.

[7] C.F. Karayigitoglu, N. Kommareddi, R.D. Gonzalez, V.T. John, G.L. McPherson, J.A. Akkara and D.L. Kaplan, Mater. Sci. Engng, C2, (1995) 165.

[8] M. Tara, V.T. John, Y.Y. Waguespack and G.L. McPherson, 9'. Phys. Chem., 98 (1994) 3809.

[9] M.S. Ayyagari, K.A. Marx, S.K. Tripathy, J.A. Akkara and D.L. Kaplan, Macromolecules, 28 (1995) 5192.

[ 10] D.P. Dumas, R. Caldwell, J.R. Wild and F.M. Raushel, J. Biol. Chem., 264 (1989) 19659.

[ 11] N. Kommareddi, M. Tata, G. McPherson, V.T. John, C.J. O'Connor, Y.S. Lee, J. Akkara and D.L. Kaplan, Chem. Mater., 8 (1996) 801.

[12] M. Vacher, M. Waks and C. Nicot, J. Neurochem., 52 (1989) 117. [ 13] D.D. Archibald and S. Mann, Nature, 364 (1993) 430.

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The microstructure of polymers enzymatically synthesized in a self-assembling environment