Inl. J. Biochem. Vol. 24, No. 4, pp. 545-552, 1992 Printed in Great

0020-711X/92$5.00+ 0.00 Copyright 0 1992Pergamon Press plc

Britain. All rights reserved

MINIREVIEW ENZYMIC

DEGRADATION

OF ALGINATES

PETER GACESA Biochemistry Department, University of Wales College of Cardiff, P.O. Box 903, Cardiff CFl IST, U.K. [Tel. (222) 874128; Fnx (222) 8741161 (Received 10 May 1991)

INTRODUCTION

Alginate is a gelling polysaccharide present in the cell walls and intracellular material of the brown seaweeds (Phaeophyceae), and comprises almost 30% of the dry weight of these plants. Approximately 22,000 tonnes/annum of alginate are extracted, mainly from three of the 265 known genera of brown algae, and the polysaccharide is used for a variety of applications by the food, pharmaceutical and other industries (Gacesa, 1988). Certain bacteria, Azotobacter chroococcum, Azotobacter vinelandii, Pseudomonas aeruginosa and other Pseudomonads, also synthesis alginate as an extracellular polysaccharide. The physiological roles of bacterial alginate have not been established although it is considered to be a major virulence factor in chronic infections of P. aeruginosa in the cystic fibrosis lung (Gacesa and Russell, 1990). Alginate is a linear polysaccharide comprised of (l-4)-linked fi-D-mannuronate and a-L-guluronate (Fig. 1). These uranic acids are arranged in block structures which may be homopolymeric (polyguluronate, poly G; polymannuronate, poly M) or heteropolymeric, i.e. essentially random sequences (poly MG), and all three types of block may be present within a single alginate molecule (Haug et al., 1967). The block structure of alginate determines the physical properties of the polysaccharide, and particularly the type of gel formed in the presence of divalent cations. Alginates rich in poly G blocks form strong but brittle gels in the presence of Ca2+ whereas a predominance of poly M or poly MG results in weak but elastic gels (Rees, 1972). Bacterial alginates are usually substituted with 0-acetyl groups on the 2 and/or 3 position of D-mannuronate (Skjak-Braek et al., 1985), which affects the water-binding properties (Skjak-Braek et al., 1989) and ion-binding selectivity of the polymer (Skjak-Braek et al., 1989; Geddie and Sutherland, 1991). Both the block structure and the degree of 0-acetylation determine the susceptibility of alginates to enzymic degradation (see below).

ALGINATE-DEGRADING

ENZYMES

Alginate-degrading enzymes have been isolated from many sources including marine algae, marine molluscs and microorganisms. Extracts from several species of the brown algae, including Laminaria digitata (Madgwick et al., 1973), Pelvetia canaliculata

(Madgwick et al., 1978) and Undaria pinnatifida (Watanabe and Nisizawa, 1982, 1984), have measurable alginase activity, and a cell-wall bound alginase has been implicated in the development of Fucus zygotes (Vreeland and Laetsch, 1990). Some marine molluscs secrete alginases into their gut, presumably to facilitate digestion of brown algal tissues. Enzyme activity has been measured in extracts of the mid-gut gland of Turbo cornutus (Muramatsu et al., 1977; Shun et al., 1984) the hepatopancreas of Littorina sp. (Favorov, 1973) and Dofabella auricula (Nisizawa et al., 1968), the style of the Surf Clam Spin& solidissima (Jacober et al., 1980) and the marine mussels Choromytilis meridionafis and Perna perna (Seiderer et al., 1982). Alginate lyases have also been extracted from the gut juice of Ap/ysia spp. (Kloareg et al., 1989; Kloareg and Quatrano, 1987a). Microorganisms have proved to be a prolific source of alginases, and enzyme activity has been detected in three species of alginate-producing bacteria, Pseudomonas aeruginosa (Dunne and Buckmire, 1985; Linker and Evans, 1984), Azotobacter vinelandii and Azotobacter chroococcum (McDowell et al., 1991). Many other bacteria also produce inducible alginases; these include Alginovibrio aquatiiis (Stevens and Levin, 1977) Alteromonas sp. (Vilter, 1986) Bacillus circuluns (Hansen and Nakamura, 1985) Beneckea pelugia (Sutherland and Keen, 1981) KlebsieNa aerogenes (pneumoniae) (Boyd and Turvey, 1977; Lange et al., 1989), Pseudomonas alginovora (Boyen et al., 1990), Pseudomonas spp. (Kashiwabara et al., 1969; Min et al., 1977a; Sutherland and Keen, 1981) and various unidentified isolates of marine (Davidson et al., 1976; Doubet and Quatrano, 1984; Preston et al., 1985) and soil bacteria (Kaneko et al., 1990). The gene for an alginate lyase from Kfebsiella pneumoniue has been cloned (Caswell et al., 1989) and over-expressed in Escherichiu coli under the control of the luc promoter (Gacesa and Caswell, 1990). Other microbial sources of alginases include four species of marine fungi (Schaumann and Weide, 1990; Wainwright and Sherbrock-Cox, 1981) and a bacteriophage which infects A. vinelundii (Davidson et al., 1977). Expression of alginase activity in bacteria is invariably induced by alginate, and one example of catabolite repression has also been reported (Macauley and Preston, 1990). In contrast, the cloned alginase of K. aerogenes (pneumoniue) is expressed at a basal level

545

546

PETERGACESA

HO-,,

-oo&f!+ HO

Fig. 1. The structure of alginate. The uranic acids p-o-mannuronate right) are (I A)-linked to form blocks of polymannuronate (centre), sequences (not shown).

in Escherichia coli in the absence of alginate and may be over-expressed with IPTG (Caswell et al., 1989). This has the advantage that enzyme may be readily obtained free of contamination by alginate. Alginases from bacteria are often considered to be extracellular although the evidence for this is usually based simply on the observation that enzyme activity can be detected in the culture medium. Few attempts have been made to determine whether this is the result of cell lysis or directed export of the protein from the cell. The precise localisation of the enzyme has been determined in only a few cases and results have not always proved to be conclusive. For example, the enzyme from Klebsiella aerogenes (pneumoniae) has been reported as extracellular by one group (Boyd and Turvey, 1977) and mainly intracellular by another (Lange et al., 1989). In practice, both groups are partially correct because the ratio of intra- to extracellular enzyme is dependent on the extent of growth of the bacterial cells (Caswell and Gacesa, 1990a). The indications are that the Klebsiella enzyme is probably periplasmic in origin, as is the case with the alginases of A. chroococcum and A. uinelandii (McDowell er al., 1991). Export of enzyme from the cell into either the periplasm or the medium implies a precursor form of the enzyme. Preliminary analysis of the cloned a1.ygene from K. pneumoniae indicates that a pro-alginase is the primary product of translation and is processed by proteolytic cleavage to the mature form of the enzyme (Caswell and Gacesa, 1990b). It is also of note that the cellular distribution of cloned enzyme is dependent on the host strain of E. coli (Caswell et al.. 1989). All but one of the reported alginases depolymerise alginate by a a-elimination reaction mechanism (Gacesa, 1987). These alginate lyases (EC 4.2.2.3) function by a three-stage reaction similar in mechanism to the alkaline degradation of polyuronides. First, the carboxyl group on the substrate has to be neutralised, possibly by the formation of a salt bridge

(top left) and n-L-guluronate (top polyguluronate (bottom) or random

with a positively charged amino acid side-chain in the active site of the enzyme. Second, a general basecatalysed abstraction of the proton at C5 of the uranic acid occurs with formation of a resonance-stabilised enolate anion intermediate. Third, a transfer of electrons from the carboxyl group to form a double bond between C, and C, results in the elimination of the 4-0-glycosidic bond (Fig. 2). A consequence of this reaction mechanism is that 4-deoxy+erythrohex-4-ene pyranosyluronate is produced at the nonreducing end of the resultant oligosaccharide regardless of whether the j-elimination occurred with either b-D-mannuronate or s-r=guluronate. Therefore it is not possible to identify the uranic acid that was originally on the 4-O-linked side of the cleaved glycosidic bond, which has implications for substrate specificity studies. This mechanism is analogous to that proposed for other polysaccharide lyases such as hyaluronan lyase (Niemann et al., 1976) and the pectate/pectin lyases (Whitaker, 1990). Although the p-elimination reaction mechanism is well understood, little advance has been made in the identification of active site residues in alginate lyases. Lyases from Turbo cornutus are inactivated if cysteine, tryptophan or lysine residues are chemically modified (Muramatsu and Egawa, 1982; Muramatsu and Imasato, 1987). The tentative conclusion from kinetic analysis of the data is that these enzymes require a single cysteine, tryptophan and lysine residue at the active site. There has been only one report (Schaumann and Weide, 1990) of an alginate hydrolase enzyme which is present in extracts of the marine fungi, Asteromyces cruciatus and Dendryphiella salina. It has been proposed that alginate is degraded in a two-stage process by these marine fungi. The first phase of depolymerisation, which is characterised by a rapid decrease in viscosity and an increase in reducing substances, is catalysed by an endo-hydrolase. The second phase, which results in a small further decrease in viscosity

Enzymic degradation of alginates

cooIO

R-O

w

R-OH

Fig. 2. A proposed reaction mechanism for alginate lyases. To simplify the diagram the linkage at C, and the hydroxyl groups at positions 2 and 3 have been omitted. AA1 and AA2 refer to amino acid residues within the active site of the enzyme.

but a high increase in the formation of 4-deoxy-r.erythro -hex-Gene pyranosyluronate and the corresponding oligosaccharides, is catalysed by an alginate lyase. However, the various alginate-degrading enzymes of these organisms will have to be isolated and purified before the existence of this unique alginate hydrolase is confirmed. Whereas other polyuronides are degraded by both lyases and hydrolases, this appears to be the exception for alginate. However, the most likely source of alginate hydrolases may well prove to be the brown algae (Gacesa, 1988), a largely unexplored source, where they could be instrumental in zygote development and tissue remodelling (Vreeland and Laetsch, 1990). Numerous methods have been developed for the detection and quantitation of alginases. The incorporation of alginate into solid growth media allows the detection of alginase-producing bacteria. The localised depolymerisation of alginate can be visualised with dilute HCl (Romeo and Preston, 1986c), calcium chloride (Schaumann and Weide, 1990), cationic detergents, e.g. cetyl pyridinium chloride, or Ruthenium Red (Gacesa and Wusteman, 1990). Fur-

541

thermore, it is possible to determine simultaneously the substrate specificity of the enzyme by use of alginate block structures rather than the intact polysaccharide (Gacesa and Wusteman, 1990). A simple turbidimetric method for detecting bacterial alginate degradation in liquid culture has been described and is based on the co-precipitation of undegraded alginate with an acidic solution of serum albumin (Kitamikado et al., 1990). Although this method is claimed to be more sensitive than plate assays it does have the disadvantage of being less suitable for screening large numbers of isolates. Several enzyme assay methods have been used with success. The most sensitive, but also the most difficult to quantify, is viscometry (Madgwick et al., 1973; Stevens and Levin, 1976). More routine methods rely on the measurement of released reducing end groups (Boyd and Turvey, 1977), the assay of unsaturated sugars using the thiobarbiturate reagent (Preiss and Ashwell, 1962) or direct estimation of the U.V. absorbance at 232nm (Boyd and Turvey, 1977). The thiobarbiturate method has the widest applicability because of its specificity and lack of interference by compounds present in crude enzyme preparations. Substrate overlay techniques have also proved useful for the determination of alginase activity after isoelectric focusing (Caswell et al., 1986) and for the direct determination of enzyme in plunta (Vreeland and Laetsch, 1989). Attempts to assay alginate lyases using alginate covalently modified with Reactive Black 5 have proved unsuccessful (Ragan, 1990) although the use of substrate labelled with customsynthesised dyes shows promise for the determination of enzyme activity (Gacesa and Wusteman, unpublished results). Changes in the circular dichroism spectrum of alginate have also been reported as a result of lyase activity, although this is unlikely to become a routine assay method (Muramatsu, 1986). Alginate lyases have been purified using a variety of conventional procedures including ammonium sulphate precipitation, ion exchange chromatography and gel permeation chromatography. Affinity chromatography on columns of immobilised alginate has facilitated the purification of alginate lyases from Littorina sp. (Favorov, 1973) and A. chroococcum and A. oinelandii (McDowell et al., 1991). The ability to determine the isoelectric point of alginases in crude extracts (Caswell et al., 1986) allows a rational rather than an empirical approach to be taken to the design of purification protocols. For example, with knowledge of the isoelectric point, chromatofocusing may be used as a rapid method of obtaining purified alginate lyase (Caswell, 1988). Alginate lyases are single-subunit enzymes with molecular sizes ranging from 25 to 100 kDa (Table 1) as determined by gel permeation chromatography or sodium dodecyl sulphate (SDStpolyacrylamide gel electrophoresis (PAGE). However, a recent report indicates that two alginate lyases isolated from a mixed culture of soil bacteria may be of more complex structure (Kaneko et al., 1990). Analysis of these enzymes indicates that each is comprised of two subunits of different molecular size (enzyme El, 35 kDa + 20 kDa; enzyme E2, 50 kDa + 38 kDa). However, there is no evidence to show that both subunits are essential for activity and there are

548

PETER GACESA

Table OrgdtllSm

K. urro,qerm Photobacterium? Littorina sp. A. rin&vufii phage Pseudomonas? Ha/i&s

pH optimum 7.0 7.8 5.&7.8h 7.7 7.5 7.0

(nk) 0.1

I .6

0.2 1.0 6.0 0.1

I, Properties

of Some alginate lyases

Mol. wt (kD4

Isoelectric point

28 29

8.9 4.2-5.0

3542 50 43

Sequence specificitya GVX MMIM MVM MIXM GIXG MVG, MIM,

Major end-product Ttimer Trimer Trimer? Trimer? DimeriTrimer

GVM

Mechanism Endo Endo Endo Endo Endo Endo

“G, Guluronate; M, mannuronate; X, either residue. The triangle shows the point of cleavage bThe value is dependent on the ionic strength of the buffer. Data have been compiled from a variety of references clted in this review.

significant discrepancies between the estimates of molecular size obtained by gel permeation chromatography and electrophoresis. The most likely explanation for these results is that the enzymes have not been purified to homogeneity. Physical properties other than molecular size vary markedly depending on the source of the enzyme. Values for the isoelectric point range from 4.2 to 8.9, with some preparations containing several protein bands, indicating either multiple activities or post-translational processing (Caswell et al.. 1986; Romeo and Preston, 1986~). On the basis of a limited amount of information from circular dichroism spectra, it appears that the secondary structure of alginate lyases also varies widely. The poly M specific lyase from a marine bacterium has 74% helical structure (Romeo and Preston, 1986~). whereas analogous enzymes from Turbo cornutus contain predominately b-pleated sheet (Muramatsu et al., 1984). Further evidence of the lack of structural homology has been obtained by Southern blotting and hybridisation studies. A probe derived from the Klebsiella pneumoniae alginate lyase gene hybridised only with the DNA from one other bacterium, Beneckea pelugia; no hybridisation was evident with DNA from five other bacterial isolates (Gacesa et u/.. 1989). Alginate lyases from diverse sources function most efficiently around neutral pH and typically display optima of pH 7.0-8.0 (Table 1). Exceptions include the lyase from II. sulinu. which has optimum activity at pH 5.5-6.0, and the lyase from B. circulans JBH2, which has an optimum of pH 5.8 (Hansen et al., 1984); other B. circuluns strains produce enzymes with optima of pH 7.4 (Hansen and Nakamura, 1985). The pH optimum of some alginate lyases varies markedly with the ionic strength of the medium (Favorov et ul., 1979). This phenomenon is also seen with other enzymes that degrade anionic substrates, and is due to a microenvironmental pH effect at low ionic strength (Gacesa et al., 1981). The ionic composition of the incubation medium also influences catalysis, with most alginate lyases requiring low concentrations (l-10 mM) of divalent cations for maximal activity. For example, the alginate lyase from P. cunuliculatrr is activated by Ca*+ but is inhibited by Mn’+, especially when the substrate concentration is low (Madgwick el al., 1978). The enzymes from Littorina sp. (Favorov et al., 1979), Pseudomonas spp. (Min et al., 1977b), P. aeruginosa (Dunne and Buckmire, 1985), a marine bacterium (Romeo and Preston, 1986c), D. salina and D. arenariu (Wainwright and Sherbrock-Cox, 1981) also require Ca” for maximal activity, whereas the lyase

from B. circuluns JBH2 requires Mg*+ (Hansen et al., 1984). Most of the extracellular bacterial alginases require moderate ionic strength for maximal activity, which presumably reflects the fact that these enzymes operate in a marine environment. The definition of alginase specificity is difficult as the substrate is co-polymeric with a non-regular distribution of monomeric units and in most cases a rather superficial but pragmatic approach has been taken to the problem. Alginate can be partially hydrolysed and fractionated to yield preparations enriched in each of the three types of block structure (Haug et al., 1967). The activity of the enzyme is tested against these enriched preparations and substrate specificity is defined in terms of a preference for poly M or poly G blocks. However, as the enriched fractions still contain significant proportions of other linkages, this is an unsatisfactory method of analysis. Mechanistically, the crucial catalytic step in the /?elimination reaction is probably stabilisation of the enolate anion (Gacesa, 1987). Therefore, the conformation of the monomer that forms the unsaturated product should be of paramount importance in determining reaction rates. However, currently there is no direct evidence to support a concept of specificity in these terms. The problem of defining specificity has been considered in some detail by Haugen et aI. (1990). Using a combination of kinetic measurements with wellcharacterised substrates and end-group analysis of the reaction products, they concluded that the “poly G specific” enzyme from Klebsiellu will cleave both G-G and G-M linkages at similar rates. In contrast, the “poly M specific” enzyme from Haliotis preferentially attacks M-M linkages but also degrades M-G and G-M diads at significant rates. Therefore, neither of these enzymes appears to be specific for a single type of diad, although the authors recognised that the same results could have been obtained if the enzyme preparations contained more than one alginase, a not uncommon phenomenon (Caswell et al., 1986). Nevertheless, this study does represent the most successful attempt to delineate alginase specificity. However, until more studies of this type have been undertaken, the specificity of alginases will continue to be described in terms of preferential degradation of fractions of enriched block structures. Modification of alginates, either by 0-acetylation of mannuronate residues or the formation of propylene glycol esters, results in diminished activity with all alginate lyases tested (see, for example, Romeo and Preston, 1986a; Sutherland and Keen, 1981; Lange et al., 1989). This implies that unmodified hydroxyl

Enzymic degradation of alginates groups on mannuronate residues and free carboxyl groups are essential for enzyme activity regardless of whether the lyase is mannuronate or guluronate “‘specific”. Kinetic analysis of the enzyme reaction is complicated by the copolymeric nature of alginate. Expression of K, values in conventional terms is virtually meaningless when the enzymes have different rates of transelimination for each of the four diads units G-G, G-M, M-G and M-M. A more useful estimate of lu, is obtained if results are expressed in terms of either mannuronate or guluronate concentration depending on the “specificity” of the enzyme. This value tends to be relatively constant regardless of the mannuronate to guluronate ratio of the substrate (Davidson et al., 1976). Typically, K,,, values for alginate lyases are in the range 0.1-5.0 mM uranic acid (Table 1). Comparison of estimates of i ‘In,, is of little value, as in most cases the constants are expressed in idiosyncratic units which cannot be related to the work of other laboratories. Examples of both substrate (Min et al., 1977b) and product inhibition (Boyd and Turvey. 1978) of alginate lyases have been reported, although this does not occur with preparations of enzyme from other sources (Romeo and Preston, 1986a). Most alginate lyases are endo-acting enzymes producing a range of unsaturated oligosaccharide products. Reaction products have been fractionated using a variety of techniques including paper chromatography (Boyd and Turvey, 1978; Schaumann and Weide, 1990), high-perfo~ance liquid chromatography (HPLC) (Romeo and Preston, 1986b) and urea polyacrylamide g,el electrophoresis (PAGE) (Hansen er al., 1984). A major advantage of the HPLC method is that individual oligosaccharides can be quantified and therefore estimates made of preferential bond cleavage (Romeo and Preston, 1986a). The absolute structure of reaction products has been determined by chemical analysis and nuclear magnetic resonance (NMR) (Boyd and Turvey, 1978). Also, NMR has been used to determine which uranic acid predominates at the reducing end of oligosaccharide fragments, thus allowing detailed analysis of substrate specificity (Haugen et al., 1990). The major endproduct of the endo-acting alginate lyases appears to be the unsaturated triuronide (Table 1); however, this will be dependent on the block structure of the substrate. Although only one exo-acting alginate Iyase has been purified (Doubet and Quatrano, 1984) it is apparent that others exist as several microorganisms are able to utitise alginate as their sole carbon source (Davidson et al., 1976; Preston et al., 1985; Wainwright, 1980). This implies a role for both endo- and exo-acting enzymes in these organisms to ensure that the polysaccharide is degraded to monomers. The carbon skeleton can then enter the central metabolic pathways via a well-defined route which involves the cleavage of the six-carbon sugars into three-carbon units (Preiss and Ashwell, 1962). Microbial degradation of alginates is not just an academic curiosity but presents a significant problem to the seaweed-cultivation industry. Several countries in the Far East rely on a supply of cultivated seaweeds for the specialist food market. Several phytopathogenic microorganisms synthesise alginate lyases

549

as virulence factors causing frond damage to the plants and thus reducing crop yields (Tseng, 1984). The activity of endogenous alginate lyases also has important consequences in the post-harvest processing of seaweeds. Und~rj~ ~inn~tl~~ is a popular seafood in Japan and some 200,000 tonnes wet wt/annum are dried to produce suboshi wakame (Nisizawa et al., 1987). However, an endogenous alginate lyase often causes frond softening during storage, resulting in an inferior product. Pretreatment of the fronds with alkaline briquet ash completely inhibits enzyme activity, although the nature of the product, haiboshi wakame, is somewhat altered in taste and appearance (Watanabe and Nisizawa, 1984). APPLICATIONS

OF ALCINATE

LYASES

Alginate lyases have been used in a variety of applications, including the analysis of the tine structure of alginates and the production of brown-algal protoplasts. Attempts at defining alginate structure enzymically have been hampered by a lack of information on enzyme specificity. Welt-~haracte~sed substrates are required to define enzyme specificity before the lyases can be used to analyse alginate structure. Nevertheless, significant progress has been made despite these problems, Alginate lyases have been used to determine block lengths in alginates (Boyd and Turvey, 1978), and the results are in reasonable agreement with independent end-group analyses. The ratio of mannuronate to guluronate in alginates may be determined by correlation of K, estimates with standard samples (Davidson et al., 1976). Some success has also been achieved in the determination of diad frequencies in samples of alginate (Min et aI., 1977c) although this technique has now been surpassed by the use of NMR (Grasdalen, 1983). Crude enzyme extracts containing alginate lyases have proved useful for the preparation of brown-algal protoplasts. The cell walls of brown algae consist of a complex mix of polysaccharides including alginate and cellulose (Kloareg and Quatrano, 1988). The combined or sequential use of cellulases and an alginate lyase preparation from either A&ssia punttata, Haliotis tuberculata or f&the&a tunicata results in almost complete removal of cell walls of Fucus distichus with high yields of viable protoplasts (Kloareg and Quatrano, 1987b). With some species of seaweed, e.g. Sargassum muticum, protoplasts have been obtained most successfully by treatment with crude enzyme extracts from an unidentified amoeba. This organism produces alginate lyases, as shown by its ability to grow on alginate as its sole carbon source (Polne-Fuller and Gibor, 1987). Alginate lyases have also been used to generate protoplasts from Laminaria spp. (Boyen et ~1.~ 1990; Butler et al., 1989), Macrocysris p_vr$ra {Kloareg et al., 1989) and SphoceZur~u (Ducreux and Kloareg, 1988). This is an important first step towards being able to modify the brown algae genetically. A potential clinical application of alginate lyases is the treatment of P. neruginosa infections in patients with cystic fibrosis (Russell and Gacesa, 1988). These patients suffer from chronic lung infections

PETERGACESA

550

caused by the mucoid, alginate-producing form of P. aeruginosa, which is often the major cause of morbidity and mortality (Dinwiddie, 1990). It is now well established that alginate is a major virulence factor of this bacterium which complicates treatment of the infection (Gacesa and Russell, 1990). Experiments in vitro have demonstrated that phagocytosis of mucoid P. aeruginosa by human monocyte-derived macrophages is enhanced by treatment of the cells with an alginate lyase (Eftekhar and Speert, 1988). Therefore, provided that a suitable method can be devised for the delivery of the enzyme, there is evidence to suggest that treatment of patients with alginate lyases will be efficacious. Selective enzymic degradation may also prove useful for the preparation of alginates with well-defined structures and physical/chemical properties in applications where there is a high added value, e.g. products for the pharmaceutical industry. Alginates are already used in a variety of pharmaceutical applications, including calcium alginate wound dressings, and anti-reflux and anti-ulcer agents (Gacesa, 1988), and lyase-modified alginates with novel properties will allow the development of new products for specific applications. CONCLUSION

In the past, the study of alginate lyases has often been hampered by a lack of well-defined substrates and the use of crude or partially purified preparations of enzyme. However, the advent of more modern methods of analysis, together with recent advances in the cloning of alginate lyase genes (Caswell et al., 1989), will allow new insights to be gained into the mechanism of action of these enzymes. The availability of nucleotide sequence data (Gacesa and Hicks, unpublished results) also opens the way for site-directed mutagenesis of the alginate lyases. These advances will ensure that significant progress is made in our understanding of alginate lyases over the next few years. Ackno~ledgemenfs-The author is grateful to the Cystic Fibrosis Research Trust. NATO (Grant CRG 910163). the SERC and the Wellcome Trust f;or research support.’ REFERENCES

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gene from Klebsiella pneumoniae which encodes alginate lyase. Biochem. Sot. Trans. 18, 929-930. Caswell R. C. and Gacesa P. (1990b) In vitro transcription translation of the alginate lyase gene from Kiebsiella pneumoniae and detection of a precursor form of the enzyme. Biochem. Sot. Trans. 18, 927-928. Caswell R. C., Gacesa P., Lutrell K. E. and Weightman A. J. (1989) Molecular cloning and heterologous expression of a Klebsiella pneumoniae gene encoding alginate lyase. Gene 75, 127-134. Caswell R. C., Gacesa P. and Weightman A. J. (1986) Detection of alginate lyases by isoelectric focusing and activity staining. Int. J. Biol. Macromol. 8, 337-341. Davidson I. W., Lawson C. J. and Sutherland I. W. (1977) An alginate lyase from Azotobacter vinelandii phage. J. gen. Microbial. 98, 223-229.

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Favorov V. V. (1973) Purification of alginases by affinity chromatography on a Bio-Gel alginate column. Int. J. Biochem. 4, 107-I 10. Favorov V. V., Vozhova E. I., Denisenko V. A. and Elyakova L. A. (1979) A study of the reaction catalysed by alginate lyase VI from the sea mollusc, Litforina sp. Biochim. biophys. Acta 569, 259-266.

Gacesa P. (1987) Alginate-modifying enzymes. A proposed unified mechanism of action for the lyases and epimerases. FEBS Letf. 212, 199-202. Gacesa P. (1988) Alginates. Carbohydr. Polym. 8, 161-182. Gacesa P. and Caswell R. C. (1990) Control and heterologous expression in Escher&hia ‘coli of the Klebsielia pneumoniae gene encoding alginate lyase. Hydrobiologia 2041205, 661-666.

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