Application of cloned monocomponent carbohydrases for modification of plant materials

S.B. Petersen, B. Svensson, and S. Pedersen (Eds), Carbohydrate Bioengineering 9 Elsevier Science B.V. All rights reserved.

321

Application of cloned monocomponent carbohydrases for modification of plant materials L. V. Kofod, T. E. Mathiasen, H. P. Heldt-Hansen and H. Dalb0ge Novo Nordisk A/S, Novo Al16, DK 2880 Bagsv~erd, Denmark

Abstract Several plant cell wall degrading enzymes have been cloned by the expression cloning technique. These enzymes can be used to degrade isolated plant cell wall polysaccharides into oligomers or to extract poly- or oligosaccharides from insoluble and complex plant cell wall material, thereby providing soluble dietary fibre or oligosaccharides with potential beneficial physiological effects. Also the cloned enzymes can be used to control e.g. viscosity in the industrial processing of plant material. This is illustrated by the degradation of various arabinoxylans or arabinoxylan containing plant material with cloned xylanases and by the degradation of rhamnogalacturonans or rhamnogalacturonan containing plant material with cloned rhamnogalacturonases and assessory enzymes.

1. I N T R O D U C T I O N Due to the abundancy of plant material in nature and the diversity of its carbohydrate components the plant cell wall is a rich source of polysaccharides. In the primary wall cellulose microfibrils form a structural network which is embedded in a matrix of hemicelluloses and pectic substances [1]. In grasses, such as wheat, corn and barley, the content of pectic substances is usually very low whereas the content of hemicelluloses is high. The major hemicelluloses are the arabinoxylans which are composed of a backbone of 13-1,4-1inked xylose units with different sidechains attached [ 1, 2]. The sidechains are usually single unit c~-1,2 or 1,3 linked arabinofuranose or o~-1,2 linked 4-O-methylglucuronic acid [2]. Some xylose residues can be substituted at both C-2 and C-3 and the degrees of mono- and disubstitution vary within different populations of arabinoxylans [3, 4]. Xylans can be either soluble or insoluble. The reasons for the insolubility of arabinoxylans have not been fully elucidated, since alkali extractable water insoluble arabinoxylans seem to have the same structures as soluble arabinoxylans [4, 5]. In dicotyledons - and in monocotyledons other than the grasses - xyloglucans are the dominating hemicelluloses [1] and the content of pectic substances is relatively high (10-60 % of the wall polysaccharides). The pectic substances are characterized by a high content of galacturonic acid, which is present in homogalacturonan as well as rhamnogalacturonan

322 polysaccharides [1]. In homogalacturonan long stretches of a-l,4-1inked galacturonic acid residues are only occasionally interrupted by a rhamnose residue whereas rhamnogalacturonan is a polymer of alternating rhamnose and galacturonic acid residues [ 1]. In rhamnogalacturonan the rhamnose residues often carry arabinan, galactan or arabinogalactan sidechains [1]. Because of the abundant sidechains the term "hairy region" is often used to describe the rhamnogalacturonan rich regions of the pectic substances [6]. In the primary wall the cellulose fibrils give the necessary strength for the cell to resist turgor pressure, while the hemicelluloses and pectic substances regulate the flexibility and porosity of the wall, necessary for cell expansion during growth [ 1]. Different models for the interlinkage of the wall polymers have been proposed, but it is generally believed that xyloglucans interlace the cellulose fibrils through strong hydrogen bonds [ 1]. Also, it has been suggested that the arabinogalactan or arabinan sidechains of pectic substances are covalently linked to cellulose or xyloglucan [7, 8] but this is not generally accepted [1, 9]. The rhamnogalacturonan (or "hairy") regions of the pectic substances supposedly alternate with homogalacturonan regions, and the pectic substances are crosslinked due to the ability of the homogalacturonan regions to form interchain "egg box junctions" through Ca2+ ions [1, 9]. Recently it was suggested that the "hairy" regions are not composed solely of rhamnogalacturonan, but that e.g. xylogalacturonan regions are an integral part of the "hairy" region alternating with rhamnogalacturonan [ 10]. The ability of saprophytic filamentous fungi to produce plant cell wall degrading enzymes has been utilized in the production of industrial enzyme products [ 11]. These products usually contain varying amounts of glucanases, xylanases and pectinases [ 11-13]. The glucanase and xylanase enzyme systems have been thoroughly described [14]. Glucanases will not be described any further. Xylanases hydrolyse the 13-1,4-1inkage between unsubstituted xylose residues in arabinoxylans but for complete degradation of arabinoxylans exo-enzymes are necessary in order to remove substituents [2, 15]. An example is arabinofuranosidases which remove the arabinose substituents resulting in more available sites for the xylanase [2, 15]. Pectic enzymes working on homogalacturonan regions as well as arabinanases and galactanases have been studied for years [6, 14, 16-21 ]. In contrast only very recently enzymes cleaving within the rhamnogalacturonan regions of pectic substances have been described. Analogous to the glucanase, xylanase and pectinase enzyme systems a set of endo- and exoenzymes exist which synergistically degrade the rhamnogalacturonan [22]. These enzymes include rhamnogalacturonases [23-26], rhamnogalacturonan acetyl esterase [27, 28] and rhamnopyranohydrolase [22]. The industrial multi-enzyme complexes have found application in wine and juice production, pulp and paper industry, baking, animal feed, textile industry, vegetable oil extraction, and production of undigestible oligo- and polysaccharides [12, 29-31]. A battery of enzymes is often necessary for complete degradation of the plant cell wall material and commercial carbohydrases, e.g. pectinases produced from fungi, can contain an extensive amount of different activities [ 13]. However, in new as well as existing applications it has sometimes been realized that only a few enzyme activities are necessary to achieve the desired effect. At best the additional activities are superfluous but in some applications they are even undesirable [ 11]. Some initiatives have been taken to purify selected enzyme activities, however not on a commercial scale. As an alternative to large scale purification of enzymes cloned monocomponent enzymes have the potential of offering a better use of resources in the

323 fermentation, a better control of the industrial enzyme reaction and a more economical and ecological dosage of enzyme protein. Of particular interest is the controlled degradation or modification of specific components of the plant cell wall, whereby selected functional properties might be encouraged. Application of monocomponent enzymes enables an understanding of the relationship between the structure of plant cell wall components and the functionality, e.g. effect on viscosity, waterbinding capacity, mouthfeel etc. The usual way of obtaining monocomponent enzymes is to identify the enzyme component in the enzyme mixture, purify the enzyme, determine the amino acid sequence, use this information to construct a labeled DNA-probe, isolate by hybridization the gene from a cDNA or genomic library constructed from the fungus in question and finally to transform the gene into an expression host for production of high amounts of monocomponent enzyme [32]. Recently, an alternative method for isolation and expression of fungal genes was introduced. In the expression cloning technique (fig. 1) the gene is isolated by virtue of its expression in yeast into an active enzyme. The yeast harboring the gene is identified by the activity of the enzyme, visualized by a sensitive plate screening assay [32]. By use of expression cloning the steps of enzyme purification, amino acid sequencing, construction of probes and hybridization can be excluded [32]. Additionally, more than one enzyme with the activity in question can be isolated simultaneously [32]. The technique has been shown to be a powerful tool for the isolation of plant cell wall degrading enzymes from filamentous fungi such as Humicola insolens and Aspergillus aculeatus [21, 24, 32, 33]. The cell wall polysacchararides of plants is the main contributor to the intake of dietary fibre by humans [34]. Dietary fibre escapes digestion by human digestive enzymes but is fermented in the large bowel to varying degrees [34, 35]. Insoluble dietary fibre is only slightly fermented and mainly serves the physiological purpose of adding bulk to the faeces and decrease transit time [35]. Soluble types of dietary fibre are fermented more extensively and serve to increase the viscosity of gastrointestinal fluids as well as to regulate lipid metabolism [35]. Especially the beneficial effects of soluble types of dietary fibre in blood glucose and cholesterol regulation and control of intestinal flora has caused an increasing interest in the addition of these types of dietary fibre to foods [36-38]. Also oligosaccharides, e.g. xylooligosaccharides, have received some attention due to their possible beneficial effect on intestinal flora [39, 40]. In the present study the possible applications of cloned enzymes for the production of poly- or oligosaccharide and for the processing of different plant material is described.

2. MATERIALS AND METHODS 2.1. Enzyme isolation A cDNA library from A. aculeatus was constructed and transformed into S. cerevisiae as described [24]. For identification of xylanase producing yeast colonies AZCL-xylan (MegaZyme, Australia) was incorporated into the agar plates. Xylanase activity was visualized by a blue halo surrounding the yeast coloni. Rhamnogalacturonases and galactanase were cloned as described [21, 24]. Arabinanase producing yeast colonies were identified by incorporation of AZCL-arabinan into the plates whereas o~-arabinofuranosidase producing colonies were identified with an overlayer of Methylumbelliferyl-t~-arabinofuranoside giving rise to a fluorescent zone. The genes were isolated and transformed into A. oryzae as described

324 [32, 41]. A.oryzae transformants were fermented as described [24] and recombinant enzymes were purified from the culture supernatant by ionexchange chromatography.

Figure 1. The principle of expression cloning. A cDNA library is constructed in a E. coli/yeast shuttle vector from an enzyme producing fungus. The library is amplified in E. coli and subsequently transformed into yeast. Yeast colonies which produce fungal enzymes are detected by appropriate enzyme assays. Vector DNA is isolated from the positive yeast coloni and the gene encoding the enzyme is inserted into an Aspergillus vector. After transformation of Aspergillus large amounts of essentially monocomponent enzyme can be produced.

2.2. Substrates

Birch xylan was obtained from Roth, soluble wheat arabinoxylan from MegaZyme. Insoluble wheat arabinoxylan was produced by treatment of wheat flour with Termamyl| and Alcalase| and recovery of insolubles by centrifugation and sieving. Corn cell wall material (Corn CWM) was isolated by successive treatments of dehulled corn kernels with Alcalase| and Termamyl| and recovery of the insoluble cell wall material by sieving. Modified hairy regions from apples were isolated according to Schols et al. 1990 [42]. Soy cell wall material (Soy CWM) was isolated by Alcalase| treatment and jet cooking (115 ~ 4 minutes) of soy meal followed by centrifugation and recovery of insolubles.

325 2.3. Small scale enzyme treatments Enzyme reactions were carded out at 30 ~ in 1.5ml Eppendorf| tubes in temperature controlled Eppendorf Thermomixers using varying amounts of enzyme and incubation times. The enzyme reaction was stopped by raising the temperature to 95 ~ for 20 minutes. Insoluble substrates were centrifuged after incubation and the supernatants recovered for analyses. Soluble substrates could be analysed with no further purification.

2.4. Viscosity reduction of wheat flour slurries Suspensions of commercial wheat flour (45 % w/w) in water were treated with enzymes (2.35 mg enzyme protein / g wheat flour) for 1 minute at 35 ~ The viscosity was measured at 40 rpm in a Brookfield viscosimeter. 2.5. Production of cloud stable apple juice Apples (Red Belle de Boskop) were cut and milled. Enzyme preparations (25 mg enzyme protein / kg mash) were added to the mash and incubated for 2 hours at 20 ~ whereafter the mash was pressed. The resulting apple juice was pasteurised to discontinue further enzyme degradation. The cloud was measured as turbidity in EF/F units [43]. The cloud stability was determined by a centrifugation test as the amount of turbidity remaining after centrifugation for 4169 x g for 15 minutes [43]. 2.6. HPLC analysis of enzyme digests The molecular weight distribution of enzyme digests was determined by high pressure size exclusion chromatography (HPSEC) which implied separation on three TSK gelfiltration columns (PW G3500, PW G3000 and PW G2000 obtained from TosoHaas) connected in series followed by refractive index detection (RID) on a RID6A (Shimadzu). The saccharides were eluted with 0.4M Sodium acetate buffer pH 3.0 at a flow rate of 0.8ml/min using a Dionex gradient pump (Dionex Corporation). The chromatograms were processed by Dionex software AI450 and Dextran standards (Serva) were used for estimation of the molecular weight (Mw) and degree of polymerization (DP). The amount of soluble saccharide in the sample could be estimated from the area of the chromatogram. Oligomers obtained from the different substrates after enzyme digestion were separated by High Pressure Anion Exchange Chromatography (HPAEC). Oligomers were eluted from a CarboPac PAl column (Dionex Corporation) with a gradient of sodium acetate in 0.1M NaOH. Gradient mixing was controlled by the Dionex gradient pump. 25ml were injected and eluting saccharides were detected by Pulsed Amperometric Detection (PAD) [44]. Xylooligomers were eluted with 0-10 rain of 0.1M NaOH followed by a linear gradient from 0-0.2M sodium acetate over 40 minutes. Rhamnogalacturonan oligomers were eluted with an acetate gradient according to Schols et al. [45]. For the determination of monosaccharide composition enzyme digests were hydrolysed in 2M triflouroacetic acid (TFA) for 1 hour at 121 ~ followed by evaporation. The hydrolysate was redissolved in water and 25 ml was injected into the CarboPac PAl column. The monosaccharides were eluted with a step gradient of from 0-12 min 5mM NaOH, from 12-28 min water, from 28-35 min 0.1M NaOH and a linear gradient from 35-54 min from 0-300mM sodium acetate in 0.1M NaOH. The column was rinsed from 54-64 min with 0.5M NaOH and equilibrated from 64-70 min in 5mM NaOH. The eluting saccharides were detected by Pulsed

326 Amperometric Detection (PAD). For calibration of the detector response standard solutions of 0.25mM, 0.5mM and lmM rhamnose, fucose, arabinose, galactose, glucose, mannose, xylose, galacturonic acid and glucuronic acid (all obtained from Sigma) were hydrolysed in TFA and analysed as described. The content of the individual monosaccharides in the enzyme digests was calculated from linear regression.

3. RESULTS AND DISCUSSION

3.1. Cloning of plant cell wall degrading enzymes from A. aculeatus When an A.aculeatus cDNA library in yeast was screened for xylanase activity on AZCL-xylan several clones were obtained representing three different xylanases, Xyl I, Xyl II, and Xyl 1111. Thus, by expression cloning three different enzymes sharing the same activity could be cloned simultaneously, which verifies the advantage of this technique. The same library was screened for rhamnogalacturonase activity [24] and a new rhamnogalacturonase (RGase B) was identified, whereas the previously described RGase A [23] was cloned by the PCR technique due to lack of expression in yeast [24]. Also, the rhamnogalacturonan acetyl esterase (RGAE) from A.aculeatus was cloned by the PCR technique [28] because of the lack of a suitable plate screening assay. The galactanase (Gal), arabinanase (Ara) and o~arabinofuranosidase (Ara.f) from A aculeatus were cloned from the cDNA library as described z [21]. After expression in A. oryzae the enzymes were purified by ion chromatographic methods essentially as described [21, 24, 33].

3.2. Composition of xylan substrates The arabinose to xylose ratio of the different xylan substrates used in this study has been determined and the results are shown in table 1. The birch xylan contain no arabinose sidechains whereas the soluble wheat arabinoxylan in average contain an arabinose substituent for every second xylose residue. In the insoluble wheat arabinoxylan the arabinose to xylose ratio is the same as in soluble wheat arabinoxylan in accordance with previous reported results [5]. However, most likely some of the xylose residues will be substituted with arabinose at C-3 as well as at C-2. In the corn cell wall material the arabinose content was higher which indicate a high level of disubstitution of xylose residues with arabinose in this substrate.

Table 1 The arabinose to xylose ratio in arabinoxylan substrates Ara/xyl Substrate 0.0025 Birch xylan 0.51 Soluble wheat arabinoxylan 0.52 Insoluble wheat arabinoxylan 0.78 Corn CWM

327 3.3. Degradation of soluble birch xylan When birch xylan was degraded by xyl I, xyl II or xyl III and analysed by HPAEC the xylooligomers eluted as seen in fig. 2. The hydrolysis of the substrate was followed by time course studies involving different dosages of enzyme. In all time course studies for all three xylanases the oligomers showed a valley point at DP 10 throughout the hydrolysis. This strongly suggests that oligomers of around 10 residues are the preferred substrate for the xylanases because they seem to be degraded as soon as they are produced. From the time course studies it was possible to find a degree of depolymerization of the birch xylan substrate which was identical for the three enzymes. At this identical degree of depolymerization of the xylan the oligomer patterns obtained with the three enzymes were almost identical. The only difference was seen in the amount of xylose and xylobiose produced 1. Xyl I produced no xylose but small amounts of xylobiose. Xyl II, which is shown in fig. 2, produced large amounts of xylose and xylobiose, whereas Xyl III produced smaller amounts of xylose and xylobiose 1. 3.4. Degradation of soluble and insoluble wheat arabinoxylans The chromatograms which result from the HPAEC analysis of soluble wheat arabinoxylan degradation products, fig. 3, were slightly more complex than those obtained for the unsubstituted soluble birch xylan. Some extra peaks emerged when compared to the birch xylan oligomers. The additional peaks are expected to be xylooligomers with arabinose substituents. As for the soluble birch xylan the three xylanases produced exactly identical oligomers from soluble wheat arabinoxylan (except for xylose and xylobiose) indicating that the preferred points of cleavage are identical. With wheat arabinoxylan DP 6-8 were produced in very small amounts and instead DP 9-11 accumulated. These results are in agreement with previously reported results [ 10] and indicate that arabinose substituents prevent the previously preferred degradation of xylooligomers with DP around 10. Therefore, if xylooligomers with high DPs are desirable an arabinoxylan substrate should be chosen instead of an unsubstituted xylan. The HPAEC chromatograms become more complicated when the insoluble wheat arabinoxylan is used as substrate, fig. 4. As opposed to the soluble wheat arabinoxylan the oligomers produced by the three xylanases were no longer identical. The differences in the degradation products have not been identified in this study. An increase in disubstituted xylose residues add yet another factor for variation in arabinoxylooligomer structures which can explain the more complex oligomer pattern. Studies on insoluble wheat arabinoxylan have been carried out with two different xylanases isolated from A.niger [46]. In those studies the oligomer structures were identified by NMR and the xylanases were shown to be different in their sensitivity to arabinose substitution [46, 47]. Thus, the differences in the degradation products obtained with the three xylanases from A. aculeatus probably result from differences in the preferred sites of attack in the highly substituted xylan backbone. The degradation of the insoluble wheat arabinoxylan was also followed by HPSEC. The amount of solubilised material could be estimated from the area under the curve in the chromatogram. In fig. 5 three chromatograms were chosen in which the enzyme dosage and time of hydrolysis would give a degree of depolymerization of soluble birch xylan which was identical for the three xylanases. It is clearly seen that Xyl II was not capable of solubilising the same amount of arabinoxylan as Xyl I and Xyl III and that the solubilised material had a lower

328

DP5 xylobiose

L

xylose

II

1,

uC

10 Ill 24 h 240 min. 120 min. 60 min. 15 min.

5

10

15

20

25

30

35

40 min.

Retention time

Figure 2. HPAEC of birch xylan degradation products. In a time course study 1.5 ml aliqouts of a 1% solution of birch xylan in 0.1M acetate buffer pH 5.0 were added 4mg of Xyl II and incubated at 30 ~ for 15, 60, 120 or 240 minutes or 24 hours. The oligomers produced were eluted from a CarboPac PAl column with an acetate gradient resulting in the chromatograms shown. Similar time course studies were performed with Xyl I and Xyl III.

329

uC

24 h 240 min. 120 min. 60 min. 15 min.

_!~, !!! ! ~ ! ! ! ! ! !! !~ ~ !!-! ! ! ! I!,, !! !!~ !l !!!!! ,~! 5

10

15

20

25

30

35

40

min.

Retention time Figure 3. HPAEC of wheat arabinoxylan degradation products. The experimental conditions were as described in fig. 2, except that the substrate was 1% wheat arabinoxylan.

330

uC Xyl III Xyl II Xyl I

5

10

15

20

25

30

35

40 min.

Retention time

Figure 4. HPAEC of insoluble wheat arabinoxylan degradation products. 3 % suspensions of insoluble wheat arabinoxylan were incubated with each of the three xylanases in time course studies. The three chromatograms shown represent an enzyme dosage and time of hydrolysis which with soluble birch xylan as the substrate would give identical degradation for the three xylanases.

331 molecular weight. This is in accordance with the finding that Xyl II has a very low activity on insoluble wheat arabinoxylan compared to Xyl I and Xyl III]. Prolonged degradation with Xyl II did not increase the amount of solubilised material to the level seen with xylanase I and III and the time course studies showed that at no stage in the hydrolysis chromatograms could be obtained with identical appearances for the three enzymes. This is opposed to the results on soluble birch xylan and soluble wheat arabinoxylan and the HPSEC results verify the differences seen with the three enzymes in the HPAEC oligomer analysis.

mV Xyi III Xyl I I Xyl I

Blank

I I I

>500,000 >3,200

125,000 800

8,000 50

500 3

Mw DP

Figure 5. HPSEC of insoluble wheat arabinoxylan degradation products. The molecular weight distributions of the arabinoxylans released from insoluble wheat arabinoxylan by the action of xylanases were determined by HPSEC. The estimated molecular weight (Mw) and degree of polymerisation (DP) is shown in the X-axis. The chromatograms correspond to the HPAEC chromatograms shown in fig. 4. The amount of material released from the substrate can be estimated from the area: Xyl I: 22 %; Xyl II: 9 %, Xyl III: 20 %.

332

3.5. Degradation of corn CWM The material liberated from corn CWM by the prolonged action of xylanases with and without the addition of arabinofuranosidase was studied. From the HPSEC chromatograms in fig. 6 the amount of solubilised material has been calculated and the results are shown in table 2 together with the arabinose/xylose ratio. For all three xylanases the addition of arabinofuranosidase increased the amount of solubilised material and the molecular weight of the solubilised material is reduced. The monosaccharide composition shows that the solubilised material has a higher ara/xyl ratio than the intact CWM which indicates that the xylan remaining in the wall has a lower degree of substitution than the liberated arabinoxylan. Thus arabinose substitution is not a major determinant of how tightly the polymers are fixed in the cell wall matrix.

Table 2 Amount and composition of solubilised material from corn CWM treated with xylanases and arabinofuranosidase Enzyme Amount released* Ara/xyl ratio % (mol]mol) Xyl I 14 1.04 Xyl I + Ara.f. 18 1.09 Xyl II 5 1.01 Xyl II + Ara.f. 9 1.09 Xy111/ 4 1.13 Xyl III + Ara.f. 10 1.14 A. aculeatus 35 0.92 * Estimated from the areas of the HPSEC chromatograms in fig. 6.

In accordance with the results on insoluble wheat arabinoxylan, Xyl II solubilises less material than Xyl I from the insoluble corn CWM. Xyl II releases only half the amount of material of Xyl I even when arabinofuranosidase is added. In contrast to the results obtained on insoluble wheat arabinoxylan, Xyl III does not release the same amount of material from corn CWM as Xyl I. When compared to the action of the complex A. aculeatus supernatant, the cloned enzymes release less material. Thus, from a solubilisation point of view, the cloned enzymes are inferior to the enzyme complex. The high solubilising power of the enzyme complex is probably due to the presence and action of several exo-enzymes which work in synergy with the xylanases [2, 15]. However, the many side activities result in a degradation of the released material into mainly mono- and dimers. This is not desirable if the extracted material is to be used for incorporation into foods as a functional food ingredient or soluble dietary fibre. Therefore, the intended application of the enzyme degradation products determines whether cloned monocomponent enzymes or the entire enzyme complex is preferable.

333 3.6. Viscosity reduction in w h e a t slurries

The purpose of wheat separation is to separate wheat gluten from wheat starch. Industrially this is accomplished in a wet milling process where a slurry of wheat flour in water is centrifuged by means of hydrocyclones or decanters yielding several fractions. The fractions obtained are enriched in gluten, starch and wheat water solubles, respectively. The viscosity of the slurry determines the capacity of the wheat separations plant as well as the quality of the separation.

A. aculeatus product

Xyl III +Ara.fur

Xyl III Xyl II +Ara.fur

mV

Xyl II XylI +Ara.fur

XylI Blank

>500,000 >3,200

125,000 800

8,000 50

500 Mw 3 DP

Figure 6. HPSEC of corn cell wall degradation products. Suspensions of corn CWM (3 % in 0.1M acetate buffer pH 5.0) were incubated with xylanase (100mg to 1.5 ml of substrate) at 30~ for 24 hours. In some experiments ot-arabinofuranosidase (100mg to 1.5 ml of substrate) was used in combination with xylanase. Also, an experiment was performed in which the cell wall material was degraded by a culture supernatant of A. aculeatus. The molecular weight distribution of the released polysaccharides was determined by HPSEC.

334 In fig. 7 the viscosities in wheat slurries which have been added equal amounts of Xyl I, II or III are seen. Wheat suspensions contain both soluble and insoluble xylan, of which the former contributes the most to viscosity [48] The viscosity reduction obtained with Xyl II is considerably higher than that obtained with Xyl I and Xyl III. As previously described Xyl II has a very low activity on insoluble wheat arabinoxylan, contrary to Xyl I and Xyl III. Thus, Xyl II does not cause a release of more xylan into the soluble phase but instead cause an immediate depolymerisation of the soluble xylan, leading to a reduction in viscosity, which is advantageous for separation of wheat components. The disadvantage of Xyl II for solubilization of insoluble xylan for production of xylooligomers or polymers is turned into an advantage when the xylanases are to be used for wheat separation.

Figure 7. Viscosity reduction in wheat flour slurries treated with xylanases. The viscosity after 1 minute of incubation is measured relative to the viscosity of a wheat flour suspension which was not added enzyme.

3.7. Degradation of rhamnogalacturonan substrates Previously it has been shown that the rhamnogalacturonases, RGase A and RGase B cloned from A. aculeatus, are functionally different [24]. Besides marked differences in pH optima and stability, the enzymes were shown to have different ratios of activity towards rhamnogalacturonan from apples, potatoes, lupins and sugar beets. When rhamnogalacturonan from apples was saponified and degraded with the RGases the degradation products obtained after prolonged incubation were shown by HPSEC to be of identical molecular weight. However, analysis by HPAEC showed that the oligomers produced by the two enzymes eluted

335 very differently from the CarboPac column. Therefore, it was anticipated that the new enzyme RGase B cleaves the linkage between rhamnose and galacturonic acid in the rhamnogalacturonan backbone as opposed to the RGase A, which has previously been shown to hydrolyse the linkage between galacturonic acid and rhamnose [24, 45]. In a very recent study it has also been shown that RGase B as well as RGase A acts in synergy with the cloned rhamnogalacturonan acetylesterase (RGAE) from A. aculeatus in the degradation of apple rhamnogalacturonan in which the acetyl esters have not been removed by saponification [28].

3.8. Degradation of soy CWM In the present study the action of the RGases on soy CWM has been investigated. Soy CWM is known to have a very high content of galactan [49-51] which is present as sidechains in the rhamnogalacturonan polymers. Therefore it was interesting to study the degradation of soy CWM with the RGases in combination with RGAE and galactanase. Also arabinanase and arabinofuranosidase were included in order to obtain as complete degradation of the sidechains as possible. A pH of 5.0 was chosen as a compromise between the acidic RGase A and neutral RGase B. At pH 5.0 both enzymes maintain 25 % of the activity at optimal pH. The results of the HPSEC of solubilised material can be seen in figs. 8 and 9 for RGase A and B, respectively. The amount of solubilised material has been estimated from the area of the chromatograms and the results are presented in table 3 together with the monosaccharide compositions.

Table 3 Amount and composition monocomponent enzymes. Amount Enzyme Soy CWM (untreated) RGase A + RGAE + Gal + Ara + Ara.f RGase B + RGAE + Gal + Ara + Ara.f

of material released from soy CWM by the action of cloned released % 0 7 17 44 46 37 52 48 47

Gal.A 19 6 5 4 4 4 3 4 4

Monosaccharide composition Rha Gal 4 38 6 40 5 51 4 59 4 55 5 55 4 58 3 58 4 56

Ara 19 34 35 30 32 34 33 32 33

The HPSEC analysis revealed that RGase A in combination with RGAE released substantial amounts (17 %) of high molecular weight material from soy CWM. RGase B alone was capable of releasing 37 % high molecular weight material, a yield which could be increased to about 50 % by the addition of RGAE. The high molecular weight material has a DP, estimated from dextran standards, of about 300. The composition of the extracted polymers, seen in table 3, shows an almost 1:1 ratio of rhamnose and galacturonic acid and a very high content of galactose and arabinose. Thus, it must be anticipated that the solubilized material is almost

336 entirely composed of fragments of rhamnogalacturonan backbone with long sidechains of arabinogalactans and arabinans attached. This is verified by the fact that all the released material is degraded completely to rhamnogalacturonanoligomers and galactose and arabinose mono- and dimers by the concerted action of RGase, RGAE, galactanase, arabinanase and arabinofuranosidase (figs. 8 and 9) _

mV

) / V 9 [ ~ RGase A+RGAE +Gal+Ara+Ara.f. RGase A+RGAE +Gal RGase A+RGAE

~

RGase A

Illllll

I >500,000 >3,500

125,000 800

8,000 50

500 3

Mw DP

Figure 8. HPSEC of soy CWM released by RGase A in combination with different cloned monocomponent enzymes. Aliquots of 1% suspensions of soy CWM in 0.1M acetate buffer pH 5.0 were incubated with enzymes (40rag of each to 1.5 ml of substrate) at 30 ~ for 24 hours and the solubilized material was analysed.

337

mV RGase B+RGAE +Gal+Ara+Ara.f. RGase B+RGAE +Gal.

w

~ '

ill

illllll,, >500,000 >3,200

RGase B+RGAE ~1 ~ " - - - ~ ' ~ ' " - - J ~v~ RGase B

llllllllll

125,000 800

8,000 50

500 3

Mw DP

Figure 9. HPSEC of Soy CWM released by RGase B in combination with different cloned monocomponent enzymes. Soy CWM was incubated with RGase B as described in fig. 8 for RGase A.

If, as suggested [7, 8], the sidechains of rhamnogalacturonan were covalently attached to the xyloglucan or cellulose of the plant cell wall, then enzymes cleaving in the rhamnogalacturonan backbone should not alone be able to release large amount of material from the wall. Therefore, soy rhamnogalacturonan does not seem to be attached to other plant cell wall constituents by means of the sidechains, which is in accordance with reports on other plant materials [52-54]. It could be argued, though, that only a few galactan or arabinan chains

338 were involved in covalent crosslinks and that these are not released. Then, the addition of galactanase and/or arabinanase should increase the amount of solubilized polymers. This was not observed. The addition of galactanase, arabinanase and arabinofuranosidase to RGase B combined with RGAE did not increase the solubilization (table 3). The only effect of the addition of side.chain degrading enzymes was to depolymerise extensively the soluble material (fig. 9). The results obtained with RGase A were slightly different. Addition of sidechain degrading activities to RGase A and RGAE increased the solubilization, although not to a level exceeding RGase B combined with RGAE. The most likely explanation for the results with RGase A is that the sidechains sterically hinder the action of RGase A and that the addition of galactanase or arabinanases minimises this hindrance. This explanation is verified by the monosaccharide composition which shows that the galactose ratio is lower and the rhamnose and galacturonic acid ratios are higher in the material released by RGase A (+/- RGAE) compared to RGase B. Thus, RGase A preferentially cleaves in rhamnogalacturonan which is not extensively substituted with sidechains. As for RGase B the addition of sidebranch degrading enzymes had the effect of converting the released high molecular weight fragments into mono-, di- and oligomers. RGases are therefore the enzymes of choice if high molecular weight polysaccharides are desired, whereas the addition of sidebranch degrading enzymes is necessary if small oligosaccharides are the preferred endproducts. Besides a high content of galactan, soy cell wall has also been reported to contain substantial amounts of xylogalacturonan [49, 51 ]. Xylogalacturonan has been suggested to be an integral part of the "hairy" regions of apple pectin, rhamnogalacturonan regions being interspersed with xylogalacturonan regions [10]. If, accordingly, the rhamnogalacturonan regions of soy cell walls were interspersed with xylogalacturonan regions then polymers released by RGases would also contain xylose. However, the content of xylose was negligible in the material released by the RGases and assessory enzymes in this study. Also, no homogalacturonan was released along with the rhamnogalacturonan. This indicates that rhamnogalacturonan in soy cell walls is not either interspersed with homogalacturonan. The results with RGases therefore indicate that rhamnogalacturonan in soy cell walls exists in a matrix separate from a matrix of homogalacturonan or xylogalacturonan. This is in accordance with the finding that no rhamnogalacturonan could be extracted from soy CWM by polygalacturonases (results not shown), which is opposed to results obtained by Schols et al. who used pectinases to extract rhamnogalacturonans from various plant sources [42, 55].

3.8. Cloud stable apple juice In several countries, cloudy fruit juices are produced. The quality of these juices is the pulpy appearance and the stability of the cloud is of paramount importance. The cloud stability is influenced by the size and composition of the particles and the viscosity of the juice [43, 56]. Experiments with various pectic degrading enzymes for production of cloud stable apple juice demonstrated that some enzymes attacking the hairy regions of pectin result in increased cloud stability of the juice. RGase B could not be used for apple juice production because of the low pH. In accordance with the results on soy CWM, RGase A alone had almost no effect (results not shown), but when combined with galactanase and RGAE a cloud stable juice could be obtained (table 4). One explanation for the cloud stability can be the increased viscosity which was found in the juice due to a large solubilization of pectic substances (results not

339 shown). In a study on apple protopectin, RGase alone was shown to solubilise some pectic material (homogalacturonan as well as rhamnogalacturonan) and no synergism was seen with galactanase [57]. The deviating results in the present study can possibly be explained by the use of a differently treated substrate with a different origin. The stabilizing effect of the galactanase used alone (table 4) might be due to modifications of the composition of already soluble pectic material or of the cloud particles rather than to the effect of material solubilized by the galactanase.

Table 4 Production of cloudy apple iuice from Red Belle de Boskop Enzyme Turbidity before Increase in turbidity centrifugation relative to untreated control, % Untreated 1061 + 112 100 RGase A + Gal 1333 + 102 125 + RGAE Galactanase 1212 + 28 114

Cloud Stability ATz, % 56 + 3 86 + 10 77 + 24

4. CONCLUSIONS In the present study it has been shown that cloned monocomponent enzymes used alone or in combination can be used for the production of soluble oligo- or polysaccharides from different plant polysaccharides or complex plant cell wall material. First, the choice of plant polysaccharide or material determines the type of saccharide which can be extracted. Secondly, the choice of enzymes determines the composition and molecular weight of the resulting degradation products. Thus, by careful selection of plant material and enzymes it is possible to obtain a wide range of saccharide products, pectic as well as hemicellulosic, with high or low molecular weight, depending on the preference. It has also been shown that the cloned monocomponent enzymes are valuable tools for control of the processing of plant material such as in the wheat separation or apple juice processes. A simil,'u" regulation and control of enzyme reaction products cannot be obtained with multi-enzyme complexes. Finally, it has become evident that an enzyme with inferior properties for one particular purpose, when compared to enzymes of the same class, can be superior for other purposes.

5. A C K N O W L E D G E M E N T S Thanks are due to Sakari Kauppinen, Lene Nonboe Andersen, Stephan Christgau, Tina S. Jacobsen, Torben Halkier, Kurt Drrreich, Susanne Htittel, Lotte R. Henriksen and Flemming M. Christensen for their contribution to this work. Also, we thank Susanne G. Jacobsen, Marianne Rohde and Margit T. Kjaer for skillful technical assistance.

340 6. FOOTNOTES 1 Sandal, T. et al., manuscript in preparation z Andersen, L. N. et al., manuscript in preparation

7. REFERENCES

1 2 3 4 5 6 7 8 9 10

11 12 13 14 15 16 17 18 19 20 21 22 23 24

N.C. Carpita and D.M. Gibeaut, Plant J., 3 (1993) 1. M.P. Coughlan and G.P. Hazlewood, Biotechnol. Appl. Biochem., 17 (1993) 259. R.A. Hoffmann, B.R. Leeflang, M.M.J. de Barse, J.P. Kamerling and J.F.G. Vliegenthart, Carbohydr. Res., 221 (1991) 63. H. Gruppen, F.J.M. Kormelink and A.G.J. Voragen, J. Cereal Sci., 18 (1993) 111. H. Gruppen, R.J. Hamer and A.G.J. Voragen, J. Cereal Sci., 16 (1992) 41. J.A. de Vries, F.M. Rombouts, A.G.J. Voragen and W. Pilnik, Carbohydr. Pol., 2 (1982) 25. T. Sakai, T. Sakamoto, J. Hallaert and E.J. Vandamme, Adv. Appl. Microbiol., 39 (1993) 213. J. Hwang, Y.R. Pyun and J.L. Kokini, Food Hydrocolloids 7 (1993) 39. S.C. Fry, Ann. Rev. Plant Physiol. 37 (1986) 165. A.G.J. Voragen, H.A. Schols and H. Gruppen, in Plant Polymeric Carbohydrates, F. Meuser, D.J. Manners and W. Seibel (eds.), 3-17, Royal Society of Chemistry, Cambridge (1992). O.P. Ward and M. Moo-Young, CRC Crit. Rev. Biotechnol., 8 (1989) 237. T. Godfrey and J. Reichelt, Industrial Enzymologi. The Applications of Enzymes in Industry, Stockton Press, New York (1983). A. Sch/3nfeld and U. Behnke, Die Nahrung 35 (1991) 395. C.A. White and J.F. Kennedy, in Carbohydrate Chemistry, J.F. Kennedy (ed.), Oxford University Press (1988). F.J.M. Kormelink and A.G.J. Voragen, Appl. Microbiol. Biotechnol. 38 (1993) 688. W. Pilnik and F.M. Rombouts, in Polysaccharides in Food, J.M.V. Blanshard. and J.R. Mitchell (eds.), 109-126, Butterworths, London (1979). G. Beldman, M.J.F. Searle-van-Leeuwen, G.A. De Ruiter, H.A. Siliha, and A.G.J. Voragen, Carbohydr. Polym., 20 (1993) 159. P. Lerouge, M.A. O'Neill, A.G. Darvill and P. Albersheim, Carbohydr. Res. 243 (1993) 373. J.M. Labavitch, L.E. Freeman and P. Albersheim, J. Biol. Chem., 251 (1976) 5904. J.W. Van De Vis, M.J.F. Searle-van-Leeuwen, H.A. Siliha, F.J.M. Kormelink and A.G.J. Voragen, Carbohydr. Polym., 16 (1991) 167. S. Christgau, T. Sandal, L.V. Kofod and H. Dalboge, Curr. Genet., 27 (1995) 135. M. Mutter, G. Beldman, H.A. Schols and A.G.J. Voragen, Plant Physiol., 106 (1994) 241. H.A. Schols, C.C.J.M. Geraeds, M.F. Searle-van-Leeuwen, F.J.M. Kormelink and A.G.J. Voragen, Carbohydr. Res., 206 (1990) 105. L.V. Kofod, S. Kauppinen, S. Christgau, L.N. Andersen, H.P. Heldt-Hansen, K.

341

25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54

D6rreich and H. Dalb0ge, J. Biol. Chem., 269 (1994) 29182. T. Sakamoto and T. Sakai, Carbohydr. Res., 259 (1994) 77. J. An, M.A. O'Neill, P. Albersheim and A.G. Darvill, Carbohydr. Res., 264 (1994) 83. M.J.F. Searle-van Leeuwen, L.A.M. van den Broek, H.A. Schols, G. Beldman and A.G.J. Voragen, Appl. Microbiol. Biotechnol., 38 (1992) 347. S. Kauppinen, S. Christgau, L.V. Kofod, T. Halkier, K. D6rreich and H. DalbCge, (submitted for publication) G.R. Beldman, A.G.J. Voragen and W. Pilnik, Enzyme Microb. Technol., 6 (1984) 503. A.G.J. Voragen, H.A. Schols and G. Beldman, Fruit Processing, 2 (1992) 98. E.-M.B.A.W. Dtisterh6ft, J.C. Venekamp, A.G.J. Voragen, World J. Microbiol. & Biotechnol., 9 (1993) 544. H. Dalbcge and H. Heldt-Hansen, Mol. Gen. Genet., 243 (1994) 253. S. Christgau, S. Kauppinen, J. Vind, L.V. Kofod and H. Dalbc~ge, Biochem. Mol. Biol. Int., 33 (1994) 917. P.S. Selvendran, Amer. J. Clin. Nutri., 39 (1984) 320. M.L. Dreher, Handbook of Dietary Fibre. An Applied Approach, Marcel Dekker Inc., New York (1987). S.A. Andon, Food Technology, Jan., (1987) 74. J. Frank and V. Wheelock, British Food Journal, 90 (1988) 22. M. Glicksman, Food Technology, Oct., (1991) 94. K. Koga and S. Fujikawa, Jap.Technol. Rev. Biotechnol., 3 (1990) 124. A.J. Morgan, A.J. Mul, G. Beldman and A.G.J. Voragen, Agro-Food-Industry Hi-Tech, Nov/Dec., (1992) 35. T. Christensen, H. Wr E. Boel, S,B, Mortensen, K. Hjortshetj, L. Thim, and M.T. Hansen, Bio/Technology, 6 (1988) 1419. H.A. Schols, M.A. Posthumus and A.G.J. Voragen, Carbohydr. Res., 206 (1990) 117. S. Hamatschek, Dissertation Hohenheim, (1989). K. Koizumi, Y. Kubota, T. Tanimoto and Y. Okada, J. Chrom., 464 (1989) 365. H.A. Schols and A.G.J. Voragen, Carbohydr. Res., 256 (1994) 97. H. Gruppen, F.J.M. Kormelink and A.G.J. Voragen, J. Cereal Sci., 18 (1993) 129. H. Gruppen, R.A. Hoffmann, F.J.M. Kormelink, A.G.J. Voragen, J.P. Kamerling and J.F.G. Vliegenthart, Carbohydr. Res., 233 (1992) 45. H. Gruppen, F.J.M. Kormelink and A.G.J. Voragen, Enzymes in Animal Nutrition Symposium, Kantause Ittingen (1993). A.M. Stephen, in The Polysaccharides, G.O. Aspinall (ed.), 97-193, Academic Press (1983). J.-M. Brillouet and B. Carr6, Phytochemistry, 22 (1983) 841. H.A. Schols, G. Lucas-Lokhorst and A.G.J. Voragen, Carbohydrates in the Netherlands, 9 (1993) 7. J.-F. Thibault, R. De Dreu, C.C.J.M. Geraeds and F.M. Rombouts, F.M, Carbohydr. Polym., 9 (1988) 119. C.M.G.C. Renard, M.J.F. Searle-van-Leeuwen, A.G.J. Voragen, J.-F. Thibault, and W. Pilnik, Carbohydr. Polym., 14 (1991) 295. C.M.G.C. Renard, H.A. Schols, A.G.J. Voragen, J.-F. Thibault and W. Pilnik, Carbohydr. Polym., 15 (1991) 13.

342 55 56 57

H.A. Schols and A.G.J. Voragen, Carbohydr. Res., 256 (1994) 83. D.L. McKenzie and T. Beveridge, Food Microstructure, 7 (1988) 195. C.M.G.C. Renard, J.-F. Thibault, A.G.J. Voragen, L.A.M. van den Broek, and W. Pilnik, Carbohydr. Polym., 22 (1993) 203.