Screening for polysaccharide-degrading micro-organisms

New Frontiers in Screening for Microbial Biocatalysts Edited by K. Kieslich, C.P. van der Beek, J.A.M. de Bont and W.JJ. van den Tweel © 1998 Elsevier Science B.V. All rights reserved.

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Screening for polysaccharide-degrading micro-organisms H.J. Ruijssenaars, L.G. van de Wiel and S. Hartmans Division of Industrial Microbiology, Department of Food Science, Wageningen Agricultural University, P.O. Box 8129, 6700 EV Wageningen, The Netherlands

1. INTRODUCTION Polysaccharides are widely applied in the food industry as thickeners, texturisers and stabilisers. Outside the food industry, polysaccharides are used in pharmaceuticals, cosmetics, detergents, textiles, adhesives, paper, paint and oil recovery. Many polysaccharides with different structures are available, but often they do not have the properties that are desired for a certain application. As the functional properties of a polysaccharide depend on its primary structure, specific modification of the structure can result in a polysaccharide with desired properties. Modification of polysaccharides comprises polymerisation, depolymerisation (formation of oligosaccharides) and tailoring (selective adjustment of primary structure) (1). Polysaccharides can be modified chemically or with enzymes. Chemical modification is usually easy and cheap but not very specific. In addition, consumers increasingly dissent from chemically processed food additives. Enzymatic modification is more specific and 'natural'. Oxidases, epimerases and hydrolases or polysaccharases are examples of enzymes that can be used for modification of polysaccharides. Exo-hydrolases (enzymes that usually cleave the first or second glycosidic bond from the non-reducing end) can be used to remove side chains often resulting in altered physical properties. A modification of this kind is described by Bulpin et al. (2). Guar gum is a cheap galactomannan that contains more galactosyl residues than the more expensive locust bean gum. Due to its lower substitution degree, locust bean gum forms gels with other polysaccharides like xanthan. If guar gum is treated with oc-galactosidase, a polysaccharide is obtained that is similar to locust bean gum both in substitution degree and physical properties. Hydrolases are also useful for elucidation of the primary structure of polysaccharides. Exo-hydrolases can be used as a sequencing tool (3), although their use may be limited to oligosaccharide-sequencing. Endo-hydrolases cleave bonds in the backbone of the polysaccharides. In this way, oligosaccharide fragments can be obtained that are relatively easy to characterise. Furthermore, polysaccharide-hydrolases can be used to obtain valuable mono- and oligosaccharides like L-rhamnose, cyclodextrins and trehalose (4).

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2. SCREENING FOR POLYSACCHARASES 2.1. Screening-methods Micro-organisms that degrade polysaccharides are a potential source of polysaccharide degrading enzymes. The most straightforward method of screening for polysaccharases is to screen for growth on plates with a polysaccharide as the only carbon source. If the micro-organism grows on the polysaccharide, it produces enzymes to degrade it. However, growth is only a good indication for polysaccharide degradation if the polysaccharide is relatively pure. Otherwise the observed growth may be due to the presence of contaminating substances that are used as a substrate. Therefore, screening methods that visualise polysaccharide-degradation are preferred. Most of these 'visual' methods are based on specific features of a polysaccharide that visibly change or disappear upon degradation. These methods generally only detect endo-hydrolase activity. Furthermore, most methods are applicable for only one or a limited group of polysaccharides. Some examples will be discussed below. Gel-forming polysaccharides like K-carrageenan, alginate, agar or gellan, can be used in plates as a solidifying agent. If polysaccharases are produced, depressions will be visible around the colonies (5). Insoluble polysaccharides like cellulose can be suspended and mixed with agar. This will result in turbid plates. If the polysaccharide is degraded clear zones around the colonies will emerge (6). This method can also be used with soluble polysaccharides. However, large amounts may be needed to obtain sufficient turbidity. Furthermore, the presence of protein contaminants can also cause turbidity. Thus, colonies producing proteases instead of polysaccharases may be selected. Polysaccharides like xyloglucan or guar gum form highly viscous, gel-like solutions. Test tubes can be filled with a viscous polysaccharide solution and each tube can be inoculated with a single colony. If the polysaccharide is degraded the viscous paste will liquefy (7). However, large amounts of polysaccharide may be required to obtain sufficient viscosity. If protein is present in the polysaccharide preparation, it should be taken into account that the viscosity may be an effect of polysaccharide-protein interactions. Loss of viscosity may therefore be caused by protease-activity. Furthermore, this method requires pure-culturing of micro-organisms before screening. Some polysaccharides, like pullulan, can be precipitated selectively in plates with ethanol (8). Poly anionic polysaccharides like pectin or carboxy methyl cellulose (CMC) can be precipitated with cetylammonium salts (9, 10). Degradation zones in these plates will not turn opaque upon addition of the precipitant thus revealing the activity of polysaccharases. Some dyes have a specific interaction with certain polysaccharides e.g. Congo red interacts with (l->4) P-D, (l->3) P-D and (l->3)(l->4) p-D glucans and (l->4) p-D xylans (11, 12). Upon flooding the plate with the dye-solution, undegraded polysaccharide will be stained by the dye whereas degradation zones will remain uncoloured. Congo red was also applied directly in the medium with CMC (13). Another classical example is the blue coloured complex that is formed between intact amy lose and Lugol's iodine solution. Rinderknecht et al. (14) designed a liquid assay for measuring a-amylase activity using amy lose that had been covalently linked to a dye, Remazol Brilliant Blue R (RBB). In principle, this dye can be coupled to every soluble polysaccharide. RBB was coupled to

241 locust bean gum, CM-cellulose, CM-amylose and CM-barley-glucan (15) and to beechwood-xylan (16). RBB-labelled starch and inulin have been applied in plate-screening methods (17, 18). Also another dye, Direct Green I, was used to label bagasse-xylan for a plate-screening method (19). If the dyed polysaccharide is degraded, dye-labelled oligosaccharides will be formed. These oligosaccharide-fragments will diffuse through the entire plate, resulting in a loss of colour intensity around a polysaccharase-producing colony. As RBB also binds to proteins (20), protein should be removed from the polysaccharide prior to labelling. 2.2. Screening for polysaccharases Most polysaccharides used today are of plant origin. However, also bacteria produce polysaccharides. Especially extracellular polysaccharides (eps's) produced by lactic acid bacteria may find application in foods. Lactic acid bacteria are food-grade organisms and the eps's produced offer a wide variety of structures. The presence of eps is considered to contribute greatly to texture and structure of fermented milk products. An exopolysaccharide produced by Lactococcus lactis ssp. cremoris B40 was chosen as a subject of study. The eps was a gift from the Dutch Institute of Dairy Research (NIZO), Ede, the Netherlands. The eps had no gelling properties, could not be precipitated in plates by ethanol or cetylpyridinium chloride and did not show interaction with Congo red. 80

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Figure 1. Percentage of total carbon converted to C02 from different polysaccharides by soil organisms after 120 hrs (pH 7, 30°C). In an initial experiment, the biodegradability of the eps by a mixed culture was compared to the biodegradability of several commercially available polysaccharides: guar gum, locust bean gum, xanthan and soluble starch. The eps contained approximately 30%

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non-eps carbon; the other polysaccharides were assumed to be pure. Buffered mineral salts medium (pH 7) containing approximately 3 g/1 of polysaccharide was inoculated with a suspension of soil and compost. C02-evolution was measured during incubation at 30°C The amount of total carbon that is converted to C0 2 indicates the extent of polysaccharidedegradation. The amount of C0 2 produced from the inoculum material was neglegible. The results are shown in Figure 1. With the eps, only about 35% of total carbon was recovered as C0 2 , whereas with the other polysaccharides 60-70% of total carbon was converted to C0 2 . This indicates that the eps is less easily biodegradable than the other polysaccharides tested. Nevertheless, attempts were made to isolate eps-degrading micro-organisms from various enrichment cultures with crude eps (20-50% pure) as the carbon source. Nine strains were selected based on difference in growth on plates with and without crude eps (25% pure). As pointed out above, however, growth alone is insufficient proof of the presence of polysaccharases. Therefore, a plate screening method was required that could visualise eps-degradation to demonstrate the ability of these strains to degrade the eps. As the eps had no characteristic properties that could be exploited in a screening method, the generally applicable RBB-method was chosen for screening experiments. The method was also tested with the commercially available polysaccharides guar gum and locust bean gum (both galactomannans). Protein was removed from the eps by treatment with proteinase K and subsequent extraction with hot phenol as described by Navon-Venezia et al. (21). After dialysis and freeze-drying the eps was labelled, like the galactomannans, as described by McCleary (15). Unbound RBB was removed by repeated ethanol precipitation and dialysis. The dyed polysaccharides were incorporated in plates. Table 1. Galactomannan-hydrolysing activities in the culture supernatants of isolated strains, 24 hours after inoculation. The strains were cultured in buffered mineral salts medium supplemented with 2 g/1 of the substrate gum and 0.02 g/1 of yeast extract. Activity (mU/ml) Substrate Growth temperature (°C) Strain 92 locust bean gum 30 "Tl locust bean gum 30 L2 78 115 locust bean gum 30 L3 37 53 locust bean gum LD1 1252 locust bean gum 37 LD2 56 locust bean gum 37 LD3 171 guar gum 30 30-1 33 guar gum 30 30-2 25 guar gum 30 30-3 11 guar gum 30 30-4 45 guar gum 30 30-5 8 guar gum 37 30-6 44 guar gum 37 37-1 25 guar gum 37 37-2

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Six out of the nine strains isolated from the eps-enrichment cultures produced haloes on the RBB-exopolysaccharide plates. Dilutions of enrichment cultures on guar gum or locust bean gum or soil suspensions were plated on RBB-galactomannan plates. Colonies with and without haloes were observed with both labelled polysaccharides. Haloes differed in size and appearance. Halo-producing micro-organisms were isolated from the RBBgalactomannan plates and streaked to purity. The halo-producing strains were cultivated in liquid medium (mineral salts with polysaccharide as the only C-source) and the polysaccharide-hydrolysing enzyme-activity was measured in the culture supernatant 24 hrs after inoculation. Activity was calculated from the increase in reducing sugars during incubation of the culture supernatant with polysaccharide-solution (5 g/1). One unit of enzyme activity releases 1 umole of reducing end groups as glucose equivalents per minute. The strains producing haloes on the RBB-eps plates grew well on mineral salts medium with 3 g/1 crude eps (65% pure). However, no eps-hydrolysing activity could be observed. Samples of culture liquid taken during growth were also analysed by High Performance Size Exclusion Chromatography (HPSEC). The amount of eps and its molecular weight distribution remained unchanged throughout the growth experiment indicating that the exopolysaccharide was not at all degraded.

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Figure 2. Degradation of locust bean gum by cell-free culture liquid from strain LD2 cultivated on mineral salts medium containing 2 g/1 locust bean gum and 0.02 g/1 yeast extract. • = reducing endgroups released from RBB-locust bean gum (mM glucose equivalents); • = reducing endgroups released from locust bean gum (mM glucose equivalents); A = RBB-labelled degradation products (OD590). RBB-labelled gum was incubated with culture supernatant and undegraded RBB-gum was precipitated with 3 volumes of absolute ethanol. The resulting supernatant was analysed spectrophotometrically at 590 nm. Initial gum concentration: 2.25 g/1.

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The majority of the strains that had been isolated from the RBB-galactomannan plates grew well in liquid polysaccharide-medium. However, some halo-producing strains could not be cultured at all in liquid medium with either galactomannan as the C-source. The gum-hydrolysing enzyme actvities in the liquid cultures are given in Table 1. Activities were generally less than 100 mU/ml although one strain, LD2, showed a high activity. In Figure 2, the time course of gum degradation by culture supernatant of strain LD2 is presented. Both RBB-labelled and unlabelled locust bean gum were used as a substrate. The increase in reducing sugars as well as the formation of blue-coloured degradation products were measured. The assays correlated well.

3. DISCUSSION With all three dye-labelled polysaccharides halo-producing colonies were observed. The isolates producing haloes on RBB-galactomannan plates generally produced gumhydrolysing enzymes. This could be demonstrated by measuring both the increase in reducing sugars when gum was incubated with culture supernatant and the increase of RBBlabelled degradation products when supernatant was incubated with RBB-labelled gum. This indicates that the RBB-plate screening method is a suitable method for selecting polysaccharide-degrading micro-organisms. However, none of the strains that produced haloes on RBB-eps were capable of degrading the eps. A possible explanation for this phenomenon would be that an enzyme is produced that either releases the dye from the polysaccharide or degrades or decolourises the dye leaving the polysaccharide intact. Also from the RBB-galactomannan plates, some halo-producing strains were isolated that could not be cultivated in liquid medium with galactomannan indicating that these strains could not degrade the gum. The biodegradability of the labelled polysaccharide may influence the frequency with which polysaccharase-negative, halo-producing colonies are observed. If the polysaccharide is easily degradable, relatively few false positives may be selected; even if the dye would be affected, there is a substantial chance that the polysaccharide is degraded. If the polysaccharide is difficult to degrade -as appears to be the case with the Lactococcus-epsthe chance to find halo-producing strains that do not degrade the polysaccharide is apparently very big. Therefore, in conclusion, although the RBB-method proved to be successful for selection of galactomannan-degrading micro-organisms, it is clear that not all haloproducing colonies exhibit polysaccharase activity. The possibility that the RBB-label is removed or modified enzymatically by the polysaccharase-negative halo-producing isolates is currently under investigation.

ACKNOWLEDGEMENTS This work was financially supported by the Ministry of Economic Affairs, the Ministry of Education, Culture and Science and the Ministry of Agriculture, Nature Management and Fishery in the framework of an industrially relevant research programme of the Association

245 of Biotechnology Centres in the Netherlands (ABON). The authors wish to thank Willemiek van Casteren of the Division of Food Chemistry, Department of Food Science, Wageningen Agricultural University for her assistance with the HPSEC analyses.

REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21.

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