Immobilized Hydrophobins as a New Tool To Study Lipases

Biomacromolecules 2003, 4, 204-210

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Solid-Phase Handling of Hydrophobins: Immobilized Hydrophobins as a New Tool To Study Lipases Jose´ M. Palomo,† Marı´a M. Pen˜as,‡ Gloria Ferna´ ndez-Lorente,† Ce´ sar Mateo,† Antonio G. Pisabarro,‡ Roberto Ferna´ ndez-Lafuente,*,† Lucı´a Ramı´rez,‡ and Jose´ M. Guisa´ n*,† Departamento de Biocata´ lisis, Instituto de Cata´ lisis, CSIC, Campus Universidad Autono´ ma, Cantoblanco, 28049 Madrid, Spain, and Departamento de Produccio´ n Agraria, Universidad Pu´ blica de Navarra, 31006 Pamplona, Spain Received June 17, 2002

Hydrophobins are fungal proteins that self-assemble spontaneously at hydrophilic-hydrophobic interfaces and change the polar nature of the surfaces to which they attach. This attribute can be used to introduce hydrophobic foci on the surface of hydrophilic supports where hydrophobins are attached by covalent binding. In this paper, we report the binding of Pleurotus ostreatus hydrophobins to a hydrophilic matrix (agarose) to construct a support for noncovalent immobilization and activation of lipases from Candida antarctica, Humicola lanuginosa, and Pseudomonas flourescens. Lipase immobilization on agarose-bound hydrophobins proceeded at very low ionic strength and resulted in increased lipase activity and stability. The enzyme could be desorbed from the support using moderate concentrations of Triton X-100, and its enantioselectivity was similar to that of lipases interfacially immobilized on conventional hydrophobic supports. These results suggest that lipase adsorption on hydrophobins follows an “interfacial activation” mechanism; immobilization on hydrophobins offers new possibilities for lipase study and modulation and reveals a new application for fungal hydrophobins. Introduction Hydrophobins are small, highly abundant, cysteine-rich, fungal proteins present in the hyphal cell walls and secreted to the culture medium.1 They are involved in the formation of aerial structures, in conferring hydrophobicity to the aerial surfaces of fungi, and in the attachment of hyphae to hydrophobic supports.2 These functions are a consequence of the ability of hydrophobin to spontaneously self-assemble at hydrophilic-hydrophobic interfaces forming very thin (5-12 nm), highly amphipathic films.3-4 The hydrophobic character and spontaneous self-assembly make very difficult their handling as free monomeric proteins. Class I hydrophobin assemblages are insoluble in hot SDS, and consequently, very drastic conditions (such as performic or trifluoroacetic acid treatment) are required to keep them free in solution. These treatments permit hydrophobin purification from cell walls and its recovery from liquid culture supernatants. The edible basidiomycete Pleurotus ostreatus (Jacq. Ex Fr) Kummer contains, at least, five different hydrophobin genes that have been isolated, cloned, sequenced, and mapped.5-7 Four of them (vmh1, vmh2, vmh3 and POH2) are expressed during vegetative growth, while fbh1 expres* To whom correspondence should be addressed. Mailing address: Instituto de Cata´lisis, CSIC, Campus Universidad Autono´ma, 28049 Madrid, Spain. Fax: (34) 915854760. Tel: (34) 915854809. E-mail addresses: [email protected]; [email protected]. † Campus Universidad Autono ´ ma. ‡ Universidad Pu ´ blica de Navarra.

sion has been found to occur exclusively during fruit body formation.5 Vegetative-growth-specific hydrophobins have been purified from fungal cell walls, as well as isolated from liquid culture supernatant. Time-course experiments of hydrophobin gene expression and protein secretion suggest that the different hydrophobins produced during vegetative growth may perform different roles during the fungal life cycle.7 Furthermore, hydrophobin self-assembly at hydrophilic or hydrophobic surfaces is a potentially useful attribute for use in medical or technical applications, such as the immobilization of diverse proteins, including enzymes.8-9 Lipases are probably the most frequently used enzymes in biocatalysis because they combine a broad substrate range with a high regio- and stereoselectivity. These enzymes have a complex catalytic mechanism, called interfacial activation.10 Lipases can adopt two alternative forms: closed (inactive) in which the substrate access to the active center is blocked by a flat or lid polypeptide, and open (active) in which the lid polypeptide flips, opening the active center pocket and exposing the hydrophobic areas that surround it, that is, the internal face of the lid and the surroundings of the active center. In homogeneous aqueous solutions, the two forms are found in an equilibrium shifted toward the closed state. In the presence of a hydrophobic surface (drop of substrate, hydrophobic solids, gas bubbles, other proteins), however, the equilibrium is shifted toward the open form because this is adsorbed on the hydrophobic surface by hydrophobic interactions established with the areas surrounding the lipase active center.11 This complex mechanism has been used as

10.1021/bm020071l CCC: $25.00 © 2003 American Chemical Society Published on Web 01/16/2003

Immobilized Hydrophobins as a Tool To Study Lipases Scheme 1. Immobilization by Interfacial Adsorption on Glyoxyl-Hydrophobin

a tool to achieve one-step purification, immobilization, and hyperactivation of lipases12-14 by using hydrophobic supports resembling the hydrophobic surface of natural substrates that selectively adsorb the lipases.15 Various hydrophobic supports have been used for lipase immobilization, including octyl-agarose, octadecyl-Sepabeads, hydrophobically coated silicates and glasses, etc.12-20 In the present study, we propose that hydrophobins may represent a useful tool for studying immobilized lipases. The aim of this work was the development of strategy for binding hydrophobins on a solid support to prepare new supports for immobilizing lipases via interfacial activation. These hydrophobin-coated supports have disperse hydrophobic nodes (drops) similar in size to the lipase surface area involved in interfacial adsorption. If the hydrophobicity of the immobilized hydrophobin were high enough to interfacially activate the corresponding lipase, then this substrate could provide interesting new ways to modulate lipase properties via “conformational” engineering (Scheme 1). Furthermore, this interaction may provide new opportunities to perform structural studies on open, active lipases. In this paper, we describe the preparation of a support coated with disperse hydrophobin molecules and the first evaluation of the potential of these molecules to interfacially activate lipases. We used hydrophobins purified from vegetative mycelium of the edible basidiomycete P. ostreatus. The hydrophobin fraction purified from vegetative mycelia comprises two proteins of 9 and 17 kDa, the larger of which corresponds to a glycosylated protein. Experimental Section Materials. The characteristics of the basidiomycete Pleurotus ostreatus (Jacq. ex Fr) Kummer var. florida strain N001 have been described previously.5,21 The two nuclei present in this N001 dikaryotic strain have been previously separated by de-dikaryotization,21 and the two corresponding protoclones (monokaryons carrying only one of the nuclei) have been deposited in the Spanish Type Culture Collection (PC9 [CECT20311] and PC15 [CECT20312]). For large-scale hydrophobin production, dikaryon N001 was cultured in Petri dishes containing SMY agar medium (1% sucrose w/v, 1% malt extract w/v, 1% yeast extract w/v)17 and incubated at 24 °C in the dark. After 7-10 days of culture, the mycelium was collected and homogenized using an Omni-Mixer homogenizer at 5000 rpm for 1 min to increase the number of growing tips and to maximize hydrophobin secretion. The macerated mycelia were transferred to 500 mL of liquid SMY Fernbach flasks and grown for 7-10 days at 24 °C in the dark without shaking before processing for hydrophobin purification.

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A racemic mixture of (R,S)-2-hydroxy-4-phenylbutanoic acid ethyl ester (HPBEt) was kindly supplied by VITA INVEST S. A. (Barcelona, Spain), and a racemic mixture of (R,S)-mandelic acid methyl ester was obtained from Sigma-Aldrich Quı´mica S. A. (Madrid, Spain). Lipases from Candida antarctica (fraction B) (Novozym 525L) and Humicola lanuginosa (Novozym 871) were a kind gift from Novo-Nordisk (Denmark). Octadecyl-Sepabeads were kindly donated by Resindion SRL (Mitsubishi Chemical Corporation, Milan, Italy). Lipase from Pseudomonas fluorescens was purchased from Amano Pharmaceutical Co. (Nagoya, Japan). Glyoxyl-agarose 10BCL was kindly donated by Hispanagar S. A. (Burgos, Spain) and prepared as previously described.22-24 Octyl-agarose 4BCL was purchased from Pharmacia Biotech (Uppsala, Sweden). Triton X-100, pnitrophenyl propionate (pNPP), and ethyl butyrate were obtained from Sigma-Aldrich Quı´mica S. A. (Madrid, Spain). Production and Purification of Hydrophobins. Vegetative mycelium-specific hydrophobins from P. ostreatus were isolated using protocols described elsewhere.5,25 Hydrophobins secreted to the liquid medium were aggregated by aeration using a Heidolf Diax 600 homogenizer and collected by centrifugation. The aggregates were disrupted by sonication in trifluoroacetic acid (TFA) at 0 °C, and subsequently the acid was removed by flushing with a nitrogen stream. The TFA-treated protein extracts were resuspended in 60% ethanol, dialyzed, and lyophilized to produce hydrophobins that could be analyzed by SDS-PAGE. Vegetative mycelium was incubated in a solution containing 10 g of sodium dodecyl sulfate (SDS) in 1 L of 0.1 M sodium phosphate buffer (pH 7.0) at 100 °C during 10 min. The insoluble fraction was then used for hydrophobin extraction as indicated above. The P. ostreatus vegetative myceliumspecific hydrophobins used in this work were vmh1, vmh2 and vmh3. Hydrophobins vmh1 and vmh2 run in SDS-PAGE as a band with an apparent molecular mass of 9 kDa, whereas vmh3 runs as a band with an apparent molecular mass of 17 kDa. Immobilization of Hydrophobins on Glyoxyl-Agarose. To immobilize hydrophobin molecules on the support surface in a very disperse way (Scheme 1), a small quantity of hydrophobins (maximum theoretical loading of this support is over 100 mg of protein per wet gram of support and we added only 0.7 mg of hydrophobins) was mixed with the glyoxyl-agarose beads. The protein (0.1 mg/mL) was dissolved in a mixture of 50% dioxane/40% Triton X-100/ 10% 200 mM sodium bicarbonate at pH 10.5, and immobilization was left to proceed for 16 h. After that time, the composites were reduced by addition of an equal volume of 2 mg/mL sodium borohydride in 100 mM sodium bicarbonate (pH 10.5) and washed with the immobilization solution and with 100 volumes of distilled water. Reduced glyoxyl-agarose (used as reference in the immobilization of lipases) was treated exactly under the same conditions. Immobilization of Lipases Via Interfacial Activation. Lipase (0.1 mg) was offered per milliliter of the different supports (octyl-agarose, octadecyl-Sepabeads or hydrophobin-agarose), using 1 mL of support and 3 mL of enzyme suspension (enzyme dissolved in 5 mM sodium

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phosphate at pH 7). A reference suspension was prepared using reduced glyoxyl-agarose to ensure that the lipase adsorption on glyoxyl-hydrophobin was promoted by the presence of hydrophobin. The activity of supernatants and suspensions was assayed as described below. Immobilization yield was calculated from the difference between the activity in the reference suspension (which was identical to the activity of the suspension supernatant, because no immobilization was ever detected in this fraction) and that of the immobilization solution supernatant. The immobilized lipase preparations were washed with 100 volumes of distilled water. Enzyme Activity Assays. Hydrolysis of pNPP. The assay was performed by monitoring spectrophotometrically (absorbance at 348 nm) the release of p-nitrophenol produced by hydrolysis of 0.4 mM pNPP in 25 mM sodium phosphate buffer at pH 7 and 25 °C. To initialize the reaction, 0.05 mL of lipase solution or suspension was added to 2.5 mL of substrate solution. One international unit (IU) of pNPPhydrolyzing activity was defined as the amount of enzyme needed to hydrolyze 1 µmol of pNPP per minute under the conditions described above. Desorption of the Lipases from the Different Hydrophobic Supports. The lipase derivatives were batchincubated in the presence of increasing concentrations of Triton X-100 (from 0.1% to 1% v/v increasing the detergent concentration in steps of 0.1%). The desorption buffer was 5 mM sodium phosphate (pH 7), and the ratio of volumes of gel and suspension was 1:10. The enzyme derivatives were incubated during 15 min at each detergent concentration before the enzymatic activity present in the suspension and supernatant fractions was measured. A reference solution of soluble enzyme was used to check the effect of the detergent on enzyme activity. Desorption was considered to be complete when the activities of the supernatant and suspension fractions were identical. Assay of Stability of the Different Lipase Preparations. The different preparations were incubated under the conditions described above. Samples of these suspensions were periodically removed and their activity assayed as previously described. Studies of Enantioselectivity. Evaluation of the hydrolytic activity and enantioselectivity of C. antarctica B lipase on the hydrolysis of mandelic acid methyl ester was performed by adding 0.25 g of derivative to 16 mL of 10 mM (R,S)mandelic acid methyl ester in 25 mM sodium phosphate buffer at pH 7.0 and 25 °C under mechanic stirring.26 When the enzyme from P. fluorescens was tested, the experiments were performed by using HPBEt by adding 0.5 g of derivative to 10 mL of 5 mM (R,S)-HPBEt in 25 mM phosphate buffer at pH 7.5 and 25 °C.27-30 During the course of each reaction, pH value was maintained constant by automatic tritation and enzymatic activity (micromole of substrate hydrolyzed per minute per milligram of immobilized protein) was estimated from the NaOH consumption in a pH stat. In both cases, the degree of hydrolysis was confirmed via reverse-phase HPLC (Spectra-Physic SP100 coupled to a Spectra-Physic SP8450 UV detector) recording the absorbance at 254 nm. The HPLC column was

Palomo et al.

a Kromasil C18, 25 cm × 0.4 cm (Analisis Vinicos, Spain), and the mobile phase consisted of a mixture of acetonitrile (30%) and 10 mM amonium phosphate (70%) at pH 2.95. The analyses were performed at a flow rate of 1.5 mL/min. At least three repetitions of each assay were made, and the experimental error was always below 5%. At different conversion degrees, the enantiomeric excesses (ee) of the remaining esters were analyzed by chiral reverse phase according to the formula previously reported HPLC. The column was a Chiracel OD-R, the mobile phase was a mixture of 40% acetonitrile and 60% water, and the analyses were performed at a flow of 0.5 mL/min. Enantioselectivity was expressed as an E value calculated from the enantiomeric excess of the remaining ester and the conversion degree according to the formula previously reported by Chen et al.31 E ) ln[(1 - X)(1 - ee)]/ln[(1 - X)(1 + ee)] X ) conversion ee ) enantiomeric excess Results and Discussion

(1)

Immobilization of Hydrophobins. Hydrophobins are proteins with a high ratio of hydrophobic to hydrophilic amino acids. They self-aggregate easily, even at low concentrations, into precipitable water-insoluble aggregates. Hence, maintenance of purified hydrophobins in solution presents difficulties and requires stringent conditions and denaturing solvents. This, therefore, represented a major difficulty in the handling of vegetative mycelium of P. ostreatus purified hydrophobins, even at concentrations of 0.5 mg/mL. Conventional protocols to keep hydrophobins in solution (treatment with TFA) were not compatible with most of the immobilization protocols, and the use of cosolvents (e.g., dioxane, alcohols) was not sufficient to keep the proteins in solution, even when cosolvents were present at a very high concentration (75%). However, hydrophobins were fully soluble in solutions containing a very high concentration of detergent (e.g., Triton X-100). We therefore decided to use a mixture of Triton X-100 (40%) and dioxane (50%) to keep the protein dispersed at a “moderate” detergent concentration to facilitate protein immobilization. Immobilization on glyoxyl-agarose was monitored by determining the protein binding to the support at different times using Bradford’s method.32 Hydrophobin immobilization proceeded more slowly than was observed for other proteins immobilized on this support, although most of the hydrophobin (around 0.5 mg/mL) was immobilized on the support after 24 h. The slow immobilization rate may have facilitated the dispersion of the hydrophobin molecules over the support surface. This composite was used in all of the other studies described in this paper. Immobilization of Lipases on Different Supports. Lipase immobilization on octyl-agarose proceeded very rapidly; all of the enzyme was immobilized in less than 15 min. A certain degree of lipase hyperactivation was observed: H. lanuginosa lipase activity increased 18-fold, that of P. fluorescens 5-fold, and that of C. antarctica B 2-3-fold (Figures 1-3). Lipase immobilization on the glyoxylhydrophobin proceeded more slowly, reaching 50% immobilization in around 1-2 h, but was also correlated with a significant increase in enzyme activity.

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Immobilized Hydrophobins as a Tool To Study Lipases

Figure 1. H. lanuginosa lipase hyperactivation during its adsorption to octyl-agarose compared with its adsorption to glyoxyl-hydrophobin: (a) immobilization on octyl-agarose support, (2) suspension, (9) supernatant, ([) soluble enzyme; (b) immobilization on glyoxylhydrophobin support, (2) suspension, (9) supernatant, ([) soluble enzyme. The experiments were carried out using crude enzyme preparations.

Reduced glyoxyl-agarose treated under the same experimental conditions as the glyoxyl-hydrophin compound was unable to adsorb lipases indicating that lipase adsorption occurred at the hydrophobin moiety of the composite. Furthermore, covalent immobilization or ionic adsorption of lipases on different supports, such as glutaraldehydeagarose, glyoxyl-agarose, or polyethyleneimine-Sepabeads, failed to promote the increments in enzyme activity observed to occur in the case of lipases adsorbed to hydrophobincoated supports, and in some cases, slight decrements of enzyme activity were promoted under these conditions (Figure 3C). Lipases adsorbed on the glyoxyl-hydrophobin support were desorbed by incubation with Triton X-100 in a similar way to lipase desorption from octyl-agarose or octadecylSepabeads (Table 1). Curiously, H. lanuginosa lipase appeared to be adsorbed more strongly to the hydrophobin compound than to octyl-agarose. The opposite was observed for the other two lipases tested because the concentration of Triton X-100 required for desorbing these enzymes from glyoxyl-hydrophobin supports was lower than that required for desorbing them from octyl-agarose. All of these results suggest that lipase immobilization on hydrophobin compounds follows a mechanism similar to that of immobilization on octyl-agarose or octadecyl-Sepabeads, namely, interfacial activation on hydrophobic surfaces. The slower lipase immobilization rate observed in hydrophobin-coated supports could be due to the small percentage of the support surface that is coated with hydrophobic

Figure 2. Immobilization of the Ps. fluorescens lipase on hydrophobic supports: (a) immobilization on octyl-agarose support, (2) suspension, (9) supernatant, ([) reference suspension; (b) immobilization on glyoxyl-hydrophobin support, (2) suspension, (9) supernatant, ([) soluble enzyme. The experiments were carried out using crude enzyme preparations. Table 1. Strength of Lipase Adsorption to Octyl-Agarose and to Glyoxyl-Hydrophobin lipase

adsorption support

% Triton X-100a

H. lanuginosa

octyl-agarose glyoxyl-hydrophobin octyl-agarose glyoxyl-hydrophobin octyl-agarose glyoxyl-hydrophobin

0.5 1 1 0.05 1 0.05

C. antarctica B P. fluorescens

a Percent (v/v) of Triton X-100 required to completely desorb the lipase from the support.

molecules. This may be a consequence of the small quantity of hydrophobin (0.7 mg) that may be immobilized on the support, which is much lower than the 100 mg of protein that may be theoretically immobilized on the support used for these experiments. Stability of the Different Derivatives. The stability of the C. antarctica B lipase glyoxyl-agarose derivative was significantly higher than that of the soluble enzyme (Figure 4). This is very likely a consequence of a certain stiffness of the lipase. The glyoxyl-agarose derivative, however, was more unstable than the interfacially adsorbed lipase (octadecyl-derivative). It has been suggested that the open form of lipases could be more stable,12-13 perhaps because it is a more ordered structure. The hydrophobin-lipase derivative had stability close to that of the interfacially adsorbed derivative made on conventional supports. Again, this result suggests that lipase immobilization on hydrophobins proceeds by interfacial activation.

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Palomo et al.

Figure 4. Thermal stability of different derivatives of the C. antarctica B lipase. Inactivation was performed at pH ) 7 and 50 °C. The experiments were carried out as described in methods. Symbols indicate the following: (2) octadecyl-Sepabeads; (9) glyoxylhydrophobin; (b) glyoxyl-agarose; ([) soluble enzyme.

higher in this case than that with interfacially activated derivatives (approximately E ) 19). All of the glyoxyl-hydrophobin derivatives produced in these experiments could be reused for 10 reaction cycles with neither a detectable decrease in enzyme activity nor variations in the measured E value. Conclusions

Figure 3. Immobilization of the C. antarctica B lipase on different supports: (a) immobilization on octyl-agarose, (2) suspension, (9) supernatant, ([) soluble enzyme; (b) immobilization on glyoxylhydrophobin, (2) suspension, (9) supernatant, ([) soluble enzyme; (c) immobilization on glyoxyl-agarose, (2) suspension, (9) supernatant, ([) soluble enzyme. The experiments were carried out using crude enzyme preparations.

Catalytic Properties of the Lipase Derivatives. The catalytic properties of P. fluorescens lipase are strongly dependent on the immobilization protocol.24 The enantioselectivity of this lipase immobilized on octyl-agarose, octadecyl-Sepabeads, or glyoxyl-hydrophobin was very similar when catalyzing (R,S)-HPBEt hydrolysis, and very different from that observed using the soluble enzyme or covalently immobilized PFL derivatives (Table 2). The hydrolysis of (R,S)-mandelic acid methyl ester by C. antarctica B lipase was very similar on different hydrophobic supports and the glyoxyl-hydrophobin lipase immobilization support (Table 3) (approximately E ) 10), whereas other immobilization techniques (e.g., covalent immobilization on glyoxyl) yielded very different results; enantioselectivity was

White wood-degrading edible basidiomycete P. ostreatus produces and secretes a considerable amount of hydrophobins during its growth in aqueous culture media. Using conventional hydrophobin purification protocols, we have extracted a hydrophobin fraction containing hydrophobins produced by 10-day-old vegetative mycelium grown in an aqueous medium. This duration of vegetative mycelium growth was chosen because it corresponds to the period of maximal hydrophobin production and secretion.7 Because of the selfassembly capacity of these proteins, hydrophobins are able to change the biophysical properties of surfaces, and these proteins have attracted interest for medical and technical applications.2,8,33 However, hydrophobin self-aggregation makes it difficult to handle these proteins as monomers or as free forms after purification. Protein immobilization may be a simple tool to manipulate proteins such as hydrophobins. After immobilization, the presence of detergents or cosolvents is no longer necessary to keep these proteins in a disperse form, and hence, they may be used for different purposes. Here, we have immobilized hydrophobins on a solid support of glyoxylagarose, and we have then used this composite to immobilize lipases. This lipase immobilization proceeds very likely via a mechanism similar to that observed when more conventional hydrophobic supports (octyl-agarose, octadecylSepabeads) were used, namely, interfacial activation. In this case, the hydrophobin hydrophobic moieties would act as hydrophobic drops triggering lipase activation. Most of the properties of the hydrophobin-immobilized lipases produced in these experiments were similar to those of lipases adsorbed on conventional supports. Lipases immobilized in this way may be used to attempt physical modification of the lipase active center without introducing chemical modifications on the lipase itself. The introduction of different residues in these hydrophobic drops may permit alterations to the lipase

Immobilized Hydrophobins as a Tool To Study Lipases

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Table 2. Enantioselective Hydrolysis of (R,S)-HPBEt Catalyzed by Hydrophobic Derivatives of the P. fluorescens Lipase

a

ee ) enantioeric excess at 50% of conversion. b E ) enantioselectivity.

Table 3. Enantioselective Hydrolysis of (R,S)-Mandelic Acid Methyl Ester Catalyzed by Hydrophobic Derivatives of the C. Antarctica B Lipase

a

ee ) enantioeric excess at 50% of conversion. b E ) enantioselectivity.

catalytic site environment, promoting controlled interactions between the lipase and modified hydrophobins. Examples of this could include the directed introduction of different chemical groups (charged, hydrophobic, polymeric, etc.) on the support that could modulate lipase activity. Site-directed modification of hydrophobin sequence and structure may facilitate this approach to study the activity of immobilized lipases. This paper exemplifies the technological use of natural products. We have used a natural protein (hydrophobin) to

immobilize natural enzymes (lipases) to fulfill a completely different function to that designed by nature because (1) the hydrophobins, which we have used as adsorbent, are surfaceactive macromolecules that fulfill a handful of functions in fungal growth and development, such as formation of hydrophobic aerial structures4,33-36 and attachment of hyphae to hydrophobic surfaces,3 and (2) the lipases, which are enzymes the natural function of which is to hydrolyze oils, have been converted to biocatalysts for use in fine chemistry. The new technology described here has been achieved via

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the development of simple technological tools: immobilization of dispersed hydrophobins and interfacial adsorption of lipases. Future modification of hydrophobins (via molecular engineering techniques) may represent a novel method to achieve significant alterations to lipase properties. In this way, engineered hydrophobins such as SC33,36 have been used to change the functional properties of surfaces.37 Different fungi provide a wide variety of hydrophobins with different solubility characteristics that could be assayed for the study of new ways of lipase activation and modulation. Further studies in this direction will be the subject of forthcoming papers. Acknowledgment. The authors gratefully acknowledge the financial support from Resindion SRL (Mitsubishi Chemical Corporation) and Projects BIO2000-0747-C05-02 and BIO1999-0278 from the CICYTa (Spain). G.F.-L. and J.M.P. hold a postdoctoral and a predoctoral fellowship of the Comunidad Auto´noma de Madrid (Spain), respectively. The authors thank Hispanagar S.A. for the gift of glyoxylagarose, Novo-Nordisk for the kind supply of enzymes, Mr. Daminati (Resindion) and Dr. Martı´nez (Novo-Nordisk) for their interesting suggestions and support, and Dr. T. Williams (Universidad Pu´blica de Navarra) for critical reading of the manuscript. References and Notes (1) Wessels, J. G. H. Int. J. Gen. Mycol. 2000, 14, 153-159. (2) Wo¨sten, H. A. Hydrophobins: multipurpose proteins. Annu. ReV. Microbiol. 2001, 55, 625-646. (3) Wo¨sten, H. A. B.; Asgeirsdo´ttir, S. A.; Krook, J. H.; Drenth, J. H. H.; Wessels, J. G. H. Eur. J. Cell Biol. 1994, 63, 122-129. (4) Wo¨sten, H. A. B.; Richter, M.; Willey, J. M. Fungal Genet. Biol. 1999, 27, 153-160. (5) Pen˜as, M. M.; Asgeirsdo´ttir, S. A.; Lasa, I.; Culian˜ez-Macia`, F. A.; Pisabarro, A. G.; Wessels, J. G. H.; Ramı´rez, L. Appl. EnViron. Microbiol. 1998, 64, 4028-4034. (6) Asgeirsdo´ttir, S. A.; de Vries, O. M. Microbiology 1998, 144, 29612969. (7) Pen˜as, M. M.; Rust, B.; Larraya, L. M.; Ramı´rez, L.; Pisabarro, A. G. Appl. EnViron. Microbiol. 2002, 68, 3891-3898. (8) Scholtmeijer, K.; Wessels, J. G. Appl. Microbiol. Biotechnol. 2001, 56, 1-8. (9) Gregory, G.; Martin, G.; Cannon, C.; McCormick, C. L. Biomacromolecules 2000, 1, 49-60. (10) Sarda, L.; Desnuelle, P. Biochim. Biophys. Acta 1958, 30, 513-521.

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