In Depth Study of a New Highly Efficient Raw Starch Hydrolyzing Î±

34

Biomacromolecules 2011, 12, 34–42

In Depth Study of a New Highly Efficient Raw Starch Hydrolyzing r-Amylase from Rhizomucor sp Georges Tawil,† Anders Viksø-Nielsen,‡ Agne`s Rolland-Sabate´,† Paul Colonna,† and Alain Buleón*,† UR1268 Biopolymères Interactions Assemblages, INRA, F-44300 Nantes, Novozymes A/S, Krogshoejvej 36, 2880 Bagsvaerd, Denmarkand Received August 6, 2010; Revised Manuscript Received November 3, 2010

A new R-amylase from Rhizomucor sp. (RA) was studied in detail due to its very efficient hydrolysis of raw starch granules at low temperature (32 °C). RA contains a starch binding domain (SBD) connected to the core amylase catalytic domain by a O-glycosylated linker. The mode of degradation of native maize starch granules and, in particular, the changes in the starch structure during the hydrolysis, was monitored for hydrolysis of raw starch at concentrations varying between 0.1 and 31%. RA was compared to porcine pancreatic R-amylase (PPA), which has been widely studied either on resistant starch or as a model enzyme in solid starch hydrolysis studies. RA is particularly efficient on native maize starch and release glucose only. The hydrolysis rate reaches 75% for a 31% starch solution and is complete at 0.1% starch concentration. The final hydrolysis rate was dependent on both starch concentration and enzyme amount applied. RA is also very efficient in hydrolyzing the crystalline domains in the maize starch granule. The major A-type crystalline structure is more rapidly degraded than amorphous domains in the first stages of hydrolysis. This is in agreement with the observed preferential hydrolysis of amylopectin, the starch constituent that forms the backbone of the crystalline part of the granule. Amylose-lipid complexes present in most cereal starches are degraded in a second stage, yielding amylose fragments that then reassociate into B-type crystalline structures, forming the final resistant fraction.

Introduction Starch is the major energy reserve for a large variety of higher plants such as cereals, legumes and tuberous plants. Starch is mainly composed of two carbohydrates: amylose and amylopectin. Amylose is defined as a mostly linear molecule of R(1,4)-linked D-glucopyranosyl units with less than 1% R(1,6) branching linkages. Amylopectin is a highly branched macromolecule composed of R(1,4)-D-glucan chains linked by 5-6% R(1,6) linkages.1 Starch is biosynthesized as granules with dimensions ranging from 1 to 100 µm with properties strongly depending on their crystalline ultrastructure.1 The inner architecture of native starch granules is characterized by “growth rings” that correspond to concentric semicrystalline 120-400 nm thick shells separated by amorphous regions.1,2 The crystalline shells consist of a regular alternation of amorphous and crystalline lamellae with a repeat distance of 9-10 nm.3,4 Native starch granules exhibit two main allomorphic types (A- or B-type), which can be detected by X-ray scattering or solid state 13C NMR spectroscopy.1,5 The A-type mainly occurs in cereal starches and the B-type in tubers and high-amylose starches.1,6 The crystallinity varies from 15 to 45% depending on the origin of starch, its hydration level, and the characterization method.1,6,7 The most recent models for A and B structures consist of 6-fold lefthanded double helices packed in monoclinic and hexagonal unit cells for A and B structures, respectively.8-12 Amylose is known to form V-type crystalline inclusion complexes with monoacyl lipids (fatty acids and lysophospholipids) present in cereal starches.1 Such complexes are present * To whom correspondence should be addressed. Fax: +33 2 40 67 50 43. E-mail: [email protected]. † Institut National de la Recherche Agronomique. ‡ Novozymes A/S.

in starches naturally containing lipids1,13 but can also be formed in vitro when heating starch or amylose with lipids in presence of water.14-18 In such a structure, the chain conformation consists of a left-handed six residues per turn single helix. It is assumed that the aliphatic part of the lipid is included inside the amylose helix while the polar group lies outside, being too big to be included.19 In the Vh-type, the most common form obtained by complexation of amylose with lipids, the single helices are packed in an orthorhombic unit cell with 16 water molecules within the unit cell.20 Most uses of starch in food and nonfood (pharmaceutics, papers, adhesives, packaging, biofuels, etc.) applications require the disruption of the starch granules through acid, alkaline, enzymatic, or hydrothermal treatments (gelatinization/melting).15 Enzymatic hydrolysis of native starch is involved in many biological and industrial processes as, for example, starch metabolism in plants, digestion by mammals, malting, fermentation, glucose syrup, or bioethanol production. Starch is specifically hydrolyzed by amylolytic enzymes, which can cut either one or both types of glycosidic bonds. Among these enzymes, R-amylases (E.C.3.2.1.1.) are the main enzymes involved in the hydrolysis of R(1,4) bonds.21,22 These enzymes are able to bypass the R(1,6) branch points without cleaving them. Contrary to homogeneous phase hydrolysis where both substrate and enzyme are in solution, the hydrolysis of solid starchy substrates strongly depends on starch structure and amylase source.21-26 For example, porcine pancreatic R-amylase degrades rice or wheat starches (A-type) over 6 times faster than banana starch and over 20 times faster than potato starch (B-type).26 The morphology and the surface of the granule, the amylose content, the crystalline structure or the presence of amylose lipid complexes were shown to be limiting factors to hydrolysis of the starch granule.23-25,27-31 More generally four main param-

10.1021/bm100913z  2011 American Chemical Society Published on Web 12/15/2010

Starch Hydrolyzing R-Amylase from Rhizomucor sp.

eters govern the enzymatic hydrolysis in heterogeneous phase: (i) the diffusion of enzymes in the medium, (ii) the accessibility of the substrate, (iii) the recognition leading to adsorption and formation of the enzyme-substrate complex, and (iv) the proper catalytic action. Adsorption of amylase onto starch granule was found to be a prerequisite for hydrolysis, but it can be inhibited by the presence of oligosaccharides such as maltose or maltotriose.32 R-Amylase and the barley limit dextrinase33 are also reported to be inhibited by specific proteins. For example, porcine pancreatic R-amylase is inhibited by Tendamistat from Streptomyces tendae34 and by RAI, a lectin-like inhibitor from Phaseolus Vulgaris35 and barley R-amylase by subtilisin inhibitor (BASI).36 The mechanisms involved in the hydrolysis of the crystalline domains and especially the disruption of double helices from the crystallite and their disentanglement are not well-known. As double helices are too wide to enter the catalytic site of R-amylases,37 their disentanglement probably occurs during the adsorption stage. B-type crystalline structure and high-amylose content are known to be more resistant to enzymatic hydrolysis than A-type structure and high-amylopectin content, as well as for native starches, for starch gels, and any type of resistant starch.24,30,38 In the literature, the main studies on heterogeneous phase hydrolysis concern starch digestion in humans by pancreatic R-amylase and especially resistant starch (RS), the starch fraction which bypasses the upper gut38,39 without being completely degraded and passes into the colon. RS is still extensively studied for human nutrition due to its impact on delayed energy release and the health potential of the products resulting from its fermentation in colon.40 A classification of resistant starches has been proposed on the basis of the factors responsible for their resistance,41 including native, embedded, encapsulated, retrograded and chemically modified starches. Rhizomucor is a genus of fungi that contains efficient amylases for raw starch degradation.42-46 The present work describes the mode of action of a recombinant amylase from Rhizomucor sp. (RA). This modified fungal R-amylase, which contains a CBM20 from A. niger glucoamylase attached through a glycosylated linker, has been constructed for starch conversion under SSF conditions during bioethanol production.47 Hydrolysis kinetics, evolution of crystalline and macromolecular structures and composition of the resulting soluble fraction were extensively studied to understand its specific behavior and to identify the structural factors responsible for maize starch resistance to R-amylolysis. This enzyme shows a high potential for both low temperature glucose syrup and bioethanol production. Its behavior was compared to that of porcine pancreatic amylase (PPA), which has been widely studied either on resistant starch or as a model enzyme in raw starch hydrolysis studies.48

Experimental Section Materials. Substrates. Normal maize starch was from Cerestar (Cargill Vilvoorde, Belgium). Amylopectin was prepared from waxy maize starch in the laboratory after dissolution in Me2SO/H2O (90/10 v/v). Amylose from potato was purchased from Sigma. 63-R-D-Glucosylmaltotriose and 63-R-D-glucosyl-maltotriosyl-maltriose were purchased from Megazyme (Wicklow, Ireland). Enzymes. Purified preparation of RA (Acc. No. AEI44738) was provided by Novozymes, Denmark. Crystallized and lyophilized porcine pancreatic R-amylase and proteinase K (from Tritirachium album) were purchased from Sigma. All other reagents were of analytical grade. Sample Preparation. Enzymes. PPA was solubilized in 20 mM phosphate buffer, pH 6.5-7.0, containing 2 mM NaCl, 0.25 mM CaCl2,

Biomacromolecules, Vol. 12, No. 1, 2011

35

and 0.2 g · L-1 NaN3. The solution was centrifuged at 4167g, and the supernatant was used for hydrolysis after protein content determination. RA was provided as 3 mg · mL-1 solution in sodium acetate buffer. Measurement of the Enzyme ActiVity. Enzyme activity was determined according to the Ceralpha procedure.49 The assay was based on hydrolysis of the non-reducing-end blocked p-nitrophenyl maltoheptaoside (BPNPG7) by R-amylase, with release of para-nitrophenol (PNP) in the presence of excess amounts of R-glucosidase. This enzyme has no action on the native substrate due to the presence of the blocking group at the reducing end of BPNPG7. The amount of released PNP was then determined spectrophotometrically at 400 nm wavelength. One unit of activity (U) is defined as the amount of enzyme, in the presence of excess thermostable R-glucosidase, required to release 1 µmol PNP from BPNPG7 in 1 min at 40 °C, pH 7. Activities of RA and PPA were 25 U and 36 U per mg of protein, respectively. Substrate Preparation for Homogeneous Hydrolysis. Amylose, amylopectin, and starch solutions were prepared by suspending 600 mg of each substrate in 40 mL phosphate buffer, pH 7, for PPA or acetate buffer, pH 4.5, for RA using gentle nitrogen stirring. The obtained suspensions were heated in an oil bath at 150 °C for amylose and a water bath at 90 °C for starch and amylopectin, leading to dissolution. Each solution was then filtered through a sieve of 5 µm and kept at room temperature for 30 min before being hydrolyzed by either RA or PPA. The amount of total solubilized sugars in the filtrated solution was determined by the orcinol sulfuric method.30 Sample Preparation for HPSEC-MALLS-DRI. Samples (10 mg) were dispersed in 0.4 mL Me2SO/H2O (90/10 v/v) using gentle magnetic stirring for 3 days at room temperature, precipitated in 80% ethanol, and washed in pure ethanol and, thereafter, acetone before drying.50 They were then solubilized at a concentration of 0.5 g · L-1 in water by microwave heating for 40 s (to reach a maximal temperature of 150 °C) at 900 W. The resulting solutions were filtered through 5 µm Durapore membranes (Millipore, Bedford, MA). Carbohydrate concentration was determined by the orcinol sulfuric method.30 Sample recovery rates were calculated from the ratio of the initial concentration to the final concentration in solution. Kinetics of r-Amylolysis in Heterogeneous Phase. R-Amylases from porcine pancreas (PPA) and Rhizomucor sp. (RA) were used at their pH and temperature optima (37 °C, phosphate buffer pH 7 for PPA; 32 °C, acetate buffer pH 4.5 for RA). Four starch concentrations were used, respectively, 0.1 (usual conditions for routine determination of starch susceptibility to amylolysis), 5, 15, and 31% dry basis (d.b.) The amount of RA and PPA was dosed in order to be at the same activity, that is, 15.5 U per mg dry starch. Enzyme solution was added to the starch suspensions and the final volume adjusted to 20 mL with buffer. The suspension was shaken continuously into a water bath at 200 rpm and 2 mL aliquots were withdrawn at different time intervals until 96 h. For each aliquot, the reaction was stopped by adding 80 µL of KOH 1 M and then centrifuged at 4167g at 4 °C for 10 min. The precipitate was used for structural analysis. Total solubilized sugars were measured in the supernatant by the orcinol sulfuric method,30 and the degree of degradation was expressed as the ratio of soluble sugars from starch hydrolysis to the initial mass of starch (d.b.). Kinetics of r-Amylolysis in Homogeneous Phase. R-Amylolysis of solubilized starch, amylose, and amylopectin was performed on 1% d.b. suspensions using the conditions described above for raw starch hydrolysis to compare the nature of the oligosaccharides produced. RA and PPA were dosed at the same activity, that is, 15.5 U per mg of solubilized substrate. The suspensions were shaken continuously and aliquots of 2 mL were withdrawn at 24, 48, and 96 h. The reaction was stopped by adding 80 µL KOH 1 M. The distribution of the oligomers produced by each enzyme was determined using highperformance anion-exchange chromatography with a pulsed amperometric detector system (HPAEC-PAD). Hydrolysis of r(1,6) Linkages by RA. To determine the possible hydrolysis of R(1,6) linkages by RA, hydrolysis of the branched oligomers, 63-R-D-glucosyl-maltotriose and 63-R-D-glucosyl-maltotrio-

36


Tawil et al.

syl-maltriose, was conducted. Solutions (0.5% d.b.) were hydrolyzed with RA according to the procedure as described above (enzyme activity 15.5 U per mg of solubilized substrate). Sampling was performed at 2 and 48 h, and the released oligosaccharides were analyzed by the HPAEC-PAD system. Determination of the Amount of Adsorbed Protein onto the Residual Starch. The amount of adsorbed R-amylase onto the residual starch was determined after 96 h of hydrolysis. Sample at 96 h was withdrawn and centrifuged. The concentration of the protein in the supernatant was determined according to the Bradford’s procedure51 and the amount of adsorbed protein determined as C ) ((C0 - C)/C0) × 100), with C0 being the initial protein concentration. Potential Inhibition of Amylases by the Reaction End Products. To check the potential inhibition of the two amylases by the oligosaccharides released during hydrolysis, the 96 h residues were washed three times in buffer by centrifugation at 4167g. Then, a new fresh amylase solution was added and a new hydrolysis was performed during 96 h, leading to a total hydrolysis time of 192 h. Potential Ineffective Adsorption of Amylases. To check if the final stages of hydrolysis were limited by amylase molecules adsorbed onto residual starch, proteinase K was applied to 192 h residual starch fragments to hydrolyze any bound amylase molecules prior to a new hydrolysis step. Thus, residual starch obtained after 192 h hydrolysis was washed one time in water and four times in Tris-HCL. Aliquot of proteinase K (0.015 mg per mg of starch) was then added onto each residual starch sample for 30 min. Proteinase K was removed by washing the residue five times with acetate buffer for RA and with a phosphate buffer for PPA followed by centrifugation at 4167g. Subsequent hydrolysis was conducted as described above. X-ray Diffraction Analysis. X-ray diffraction (XRD) analysis was performed on native starch and residual starch withdrawn at different time intervals during hydrolysis. The water content of samples was adjusted by water phase sorption for 10 days in desiccators under partial vacuum at a relative humidity of 90% (using a saturated salt solution of baryum chloride). Hydrated samples (20 mg) were then sealed between two tape foils to prevent any significant change in water content during XRD measurement. XRD diagrams were recorded on a Bruker (Wissembourg, France) D8 Discover diffractometer. Cu KR1 radiation (λ ) 0.15405 nm), produced in a sealed tube at 40 kV and 40 mA, was selected using a Gobe¨l mirror parallel optics system and collimated to produce a beam of 500 µm diameter. The diffracted beam was collected with a two-dimensional GADDS detector and recording time was 600 s. The distance from the sample to the detector was 100 mm. Relative crystallinity was determined after normalization of all recorded diagrams at the same integrated scattering between 3 and 30° (2θ). Aand B-type recrystallized amyloses were used as crystalline standards, after scaled subtraction of an experimental amorphous curve in order to get null intensity in the regions without diffraction peaks. Dry extruded potato starch was used as the amorphous standard. The degree of crystallinity of samples having a pure allomorphic type was determined using the method initially developed by Wakelin and coworkers for cellulose.52 Differential Scanning Calorimetry. The melting behavior of both native and residual starches was analyzed using differential scanning calorimeter (DSC Q100, TA Instruments, U.K.) equipped with a refrigerated cooling system (RCS). Samples of 10 mg were weighed into stainless pans. Distilled water was added and adjusted to get a final water content of 70% (d.b.). The DSC run was performed at a heating rate of 3 °C.min-1 from 5 to 130 °C, against a reference pan containing 20 µL water. The instrument was calibrated using indium (Tm ) 156.6 °C, ∆Hm ) 28.55 J · g-1). The melting enthalpy (∆H) was calculated using the software Universal Analysis (TA Instruments) and normalized to the mass of dry solid. High-Performance Anion-Exchange Chromatography (HPAEC). The composition of soluble products released by enzymatic hydrolysis was determined by using high-performance anion-exchange chromatography with a pulsed amperometric detector (PAD) system.53

Figure 1. Effect of raw starch concentration on RA hydrolysis kinetics: 31% d.b. (+) 15% d.b. (2), 5% d.b. (0), 0.1% d.b. (b). Data obtained with PPA on 31% starch (×) are shown for comparison.

Samples were injected into a CarboPac PA-100 anion exchange column (250 × 4 mm) coupled to a CarboPac PA-100 guard column. Two eluents were used: 150 mM NaOH (eluent A) isocratic, and 1 M NaOAc (eluent B). The flow rate was 1 mL · min-1. The applied gradient of acetate was (1) 0-1 min, 150 mM eluent A; (2) 1-10 min, linear gradient 0-75 mM eluent B; (3) 10-19 min, linear gradient 75-180 mM eluent B; (4) 19-111 min, linear gradient 180-225 mM eluent B; (5) 111-115 min, linear gradient 225-500 mM eluent B; and finally return to 0 mM eluent B for column equilibration. As the HPAECPAD response coefficients decrease with increasing DP, % area of long chains are not representative of the exact weight fraction of each DP. Therefore, standard oligosaccharides were used from G1 to G7 for an exact calibration. High-Performance Size-Exclusion Chromatography (HPSEC). The molecular weight distribution of residues at different hydrolysis times was determined using high-performance size-exclusion chromatography with multiangle laser light scattering (MALLS) detection.50 The equipment was the same as that described previously.50 The used column was Shodex KW-802.5 (8 mm ID × 30 cm) from Showa Denko K.K. (Tokyo, Japan) together with KW-G guard column (6 mm ID × 5 cm) from Showa Denko K.K. The column and guard column were maintained at 30 °C using a Crococil temperature control system from Cluzeau (Bordeaux, France). Before use, the mobile phase (Millipore water containing 0.2 g · L-1 sodium azide) was carefully degassed and filtered through Durapore GV (0.2 µm) membranes from Millipore and eluted at a flow rate of 0.3 mL · min-1. The two online detectors comprised a MALLS instrument (Dawn HELEOS) fitted with a K5 flow cell, a He-Ne laser, (λ ) 658 nm) (Wyatt Technology Corporation, Santa Barbara, CA), and an ERC-7515A refractometer (Erma, Tokyo, Japan). After solubilization and filtration, samples were immediately injected into the HPSEC-MALLS system. Sample recovery was calculated from the ratio of the mass eluted from the column (integration of the refractometric signal) to the injected mass which was determined using the orcinol sulfuric method.30

Results Enzymatic Hydrolysis of Raw Starch at Different Concentrations. Figure 1 shows the evolution of hydrolysis kinetics with increasing starch concentration for RA and Table 1 summarizes the kinetic data obtained for RA and PPA. All hydrolysis curves have a classical two phase shape with a first stage of more rapid hydrolysis (8-10 h) and then a slower stage. By dosing the two enzymes at the same activity per mg starch, their efficiency on raw starch can be strictly compared. No matter at what starch concentration tested, RA is much more effective on raw starch than PPA and remarkably active on the 31% starch suspensions since the final hydrolysis degree (FHD)



37

Table 1. Kinetic Data as a Function of the Starch Concentration for RA and PPA (in Brackets) Time (h)

0.1% d.b.

13.6 ( 2.4 (20.6 ( 6.0) 2 21.8 ( 2.2 (38.0 ( 1.8) 8 65.2 ( 0.8 (69.5 ( 0.1) 24 96 ( 4.8 (82.0 ( 1.6) 48 1.5 ( 3.4 (85.0 ( 1.9) 72 95 ( 1.8 (88.0 ( 3.0) 96 (FHD) 99 ( 2.8 (92.0 ( 0.8) 1

5% d.b.

15% d.b.

31% d.b.

18.1 ( 4.6 (25.0 ( 1.8) 30.6 ( 1.2 (35.0 ( 15.0) 71.1 ( 6.0 (50.0 ( 3.6) 89.4 ( 1.6 (66.0 ( 3.8) 84 ( 9.4 (78.0 ( 9.6) 94.3 ( 1.2 (84.0 ( 6.4) 90.0 ( 7.0 (89.0 ( 10.6)

17.0 ( 12.0 (22.0 ( 2.6) 24.0 ( 11.0 (29.0 ( 4.4) 54.0 ( 5.0 (34 ( 3.0) 75.0 ( 12.2 (75.0 ( 8.0) 79.0 ( 5.6 (48.0 ( 1.4) 84.0 ( 4.0 (54.0 ( 1.6) 81.0 ( 3.2 (67.0 ( 8.0)

13.0 ( 6.0 (19.0 ( 8.0) 20.0 ( 16.0 (23.0 ( 2.8) 36 ( 3.2 (27 ( 3.4) 57.5 ( 4.2 (18.4 ( 1.6) 64.0 ( 2.0 (32 ( 1.2) 68.0 ( 2.0 (31.5 ( 0.8) 75.0 ( 5.2 (32.0 ( 2.8) Figure 2. Evolution of the crystalline structure as a function of time and hydrolysis degree (in brackets) during hydrolysis of a 31% starch suspension by RA.

reaches 75% for RA but only 32% for PPA at 96 h (Figure 1 and Table 1). In general, the hydrolysis degree decreases with increasing starch concentration. Although PPA has an efficiency comparable to that of RA in the first hours of hydrolysis (Table 1) independent of the starch concentration, it yields much lower FHD as soon as starch concentration reaches 15% or higher. Changes in the Crystalline Structure during Hydrolysis. The crystalline structure was essentially assessed by X-ray diffraction (XRD). Crystallinity (degree of crystallinity and crystallinity type) of residual starch and corresponding hydrolysis degree are summarized in Table 2 for 5 and 31% starch suspensions at different times of hydrolysis. Native maize starch has A-type crystallinity with characteristic peaks at Bragg angle (2θ) ) 15, 17, 18 and 23° (Figure 2). The crystallinity decreases from 31 to 20% during the first 2 h of hydrolysis of the 31% starch suspensions by RA. The decrease is less pronounced for PPA which is much less effective on concentrated raw starch. This ability to degrade rapidly the crystalline domains at high starch concentration is characteristic for RA when comparing the crystallinity decrease as the hydrolysis proceeds. Indeed at 2 h of hydrolysis, one-third of initial crystallinity is erased while only 20% of the initial starch has been hydrolyzed. On the contrary, no change of crystallinity was observed with PPA on a 31% starch suspension with a final crystallinity of 30%, that is, while 32% of the starch was hydrolyzed at 96 h. When observed in light microscopy PPA hydrolyzed samples exhibit initial granular shapes with Maltese crosses and no presence of channels. No fragment of granules is detectable. These observations are in agreement with earlier studies on wheat starch where starch granules in heterogeneous phase are attacked granule per granule by PPA.21 No complete disappearance of crystallinity was observed even after 96 h hydrolysis. The intensity of the peak at 2θ around 20°, which corresponds to amylose-lipid complexes formed between amylose and endogeneous fatty acids present in maize starch, is remarkably stable. Vh-type structure is more visible as soon as the hydrolysis

reaches a degree of approximately 80%. It is also present in the 96 h PPA residue obtained from 5% starch suspensions. PPA, unable to hydrolyze more than 32% of the concentrated starch suspension, hydrolyzes 89% of the 5% starch suspensions. Thus, it appears that the Vh-type structure is much more resistant to R-amylases than A-type, as already described by Gernat et al.25 In contrast, analyzing RA samples after 48 h of hydrolysis (hydrolysis degree 84%), a mixture of B-type, with characteristic peaks at 2θ about 5.6, 17, and 24°, and Vh-type is observed. This B-type structure is known to result from retrogradation of starch15 or crystallization of short chain amylose in water.54 This latter structure has been shown to be very resistant to enzymatic hydrolysis.23,30 In this study the B-type structure can originate from rearrangements of linear amylose-like fragments released by the enzyme. This point will be discussed later. Differential scanning calorimetry (DSC) was also used for structural characterization of the residual starch, the melting temperature and enthalpy in excess water being connected to both starch structural type and crystallinity. The thermograms of all starch residues exhibit two endotherms corresponding respectively to melting of the residual A-type structure at temperatures around 72-75 °C and melting of amylose lipid complexes at 97-100 °C. Amylose lipid complexes are present within native maize starch and can form during heating in DSC. DSC is well-known to provoke crystallization of amylose lipid complexes after gelatinization of lipid containing starches such as maize.18 Thus, DSC data cannot be used to assess potential changes in amylose complexing during hydrolysis. The melting enthalpy, relative to A-type native residual structure, decreases rapidly from 12.4 to 5.5 J · g-1 in the first hours of hydrolysis for RA (Table 3), which is consistent with the large decrease in crystallinity determined by XRD. The corresponding melting temperature is not significantly different from that of native maize starch. Very few enthalpy changes were observed on PPA

Table 2. Evolution of the Crystalline Structure CS (Degree of Crystallinity in %, Crystalline Type A, B, or Vh) and Degree of Hydrolysis HD (in %) of Maize Starch during Hydrolysis by RA and PPAa native starch

RA 5% starch RA 31% starch PPA 5% starch PPA 31% starch a

nd: not determined.

hydrolyzed 2 h

hydrolyzed 8 h

hydrolyzed 48 h

hydrolyzed 96 h

CS

CS

HD

CS

HD

CS

HD

CS

HD

31, A 31, A 31, A 31, A

30, A 20, A 30, A 30, A

31 19 34 23

15, A 15, A 25, A 30, A

71 36 49 27

nd, B+Vh 15, A 20, A 30, A

84 72 78 32

nd, B+Vh 15, A+Vh 20, A+Vh 30, A

93 75 89 32

38


Tawil et al.

Table 3. Melting Temperature and Melting Enthalpy of Residual Starch during Hydrolysis of 31% Maize Starch Suspensions by RA and PPA sample

melting temperature (°C)

melting enthalpy (J · g-1 dry matter)

native starch

72.3 ( 2.3

12.4 ( 0.2

RA 31% starch 2h 8h 48 h 96 h

72.0 ( 1.7 72.3 ( 0.9 74.1 ( 0.6 75.0 ( 2.1

8.0 ( 0.6 5.5 ( 0.3 5.8 ( 1.3 3.4 ( 2.7

PPA 31% starch 2h 8h 48 h 96 h

71.9 ( 2.5 72.0 ( 1.2 72.0 ( 1.2 72.0 ( 0.4

12.8 ( 0.1 12.0 ( 1.1 11.0 ( 0.6 11.0 ( 0.4

Figure 3. HPSEC chromatographs of residual starch after hydrolysis of 31% maize starch suspensions by RA and PPA: (1) native starch (blank); (2, 3) residual starch obtained after hydrolysis with PPA at 48 and 96 h, respectively; (4, 5) residual starch obtained after hydrolysis with RA at 48 and 96 h, respectively. Relative scale was obtained by dividing each initial normalized chromatogram (normalized refractometric response) by the amount of residual starch.

residues from hydrolysis of the 31% starch suspensions. These observations are in line with the X-ray diffraction results. Changes in the Macromolecular Structure during Hydrolysis. Figure 3 shows the HPSEC profiles of RA and PPA residual starch at 48 and 96 h, as well as raw starch. Raw starch presents two peaks, which are attributed to amylopectin (elution volume around 5.7 mL) and amylose (elution volume around 6.6 mL) according to previous works.50 Residual starch present after PPA hydrolysis shows the same bimodal shape as for native starch, with no decrease of amylopectin weight average molecular weight (Table 4). The ratio of amylose and amylopectin calculated from the height of the HPSEC peaks is equal to that of native starch, which shows that PPA hydrolyzes progressively and, concomitantly, amylose and amylopectin. On the contrary, RA profiles show a drastic decrease of amylopectin average molecular weight and radius of gyration. The apparent density increases from 3.73 to 21.04, which means that the amylopectin remaining fraction is highly branched. At the same time amylose is only slightly hydrolyzed after 48 h and is more strongly hydrolyzed between 48 and 96 h (Table 4). These results are in agreement with the observed large decrease of crystallinity during hydrolysis, because amylopectin is usually assumed to support the framework of the crystalline domains in the starch granule. Furthermore, the low degradation of amylose is also consistent with the measured amylose-lipid complexes at this stage of hydrolysis, while the smaller molar mass observed at 96 h could be linked to the chains involved

in the B-type recrystallization observed for RA at high hydrolysis degree. The amylopectin to amylose ratio cannot be calculated accurately from the HPSEC profiles due to very high hydrolysis degree resulting in peak shifts, and chain fragments released from the hydrolysis of amylopectin elutes close to the elution volume of amylose. Structure of the Soluble Oligosaccharides Produced. The distribution of oligosaccharides present in the soluble phase after 2, 8, and 48 h hydrolysis of 31% starch suspensions by RA or PPA is shown in Table 5. Only the values obtained for DP < 8 are shown because the HPAEC response is not quantitative above DP 7. The distribution is remarkably different. With RA, starch is completely degraded into glucose (DP1). The amount of glucose increases with hydrolysis time from 20% at 2 h to 50% of the total sugar amount after 48 h (results not shown). No other malto-oligosaccharides (DP 2-7) were observed. The release of malto-oligosaccharides was checked after very short hydrolysis times (5-30 min). Only glucose was observed in 31% starch suspensions (Supporting Information, Figure S1), while a little amount of maltose and other branched oligosaccharides was detected during hydrolysis of 5% starch suspensions at hydrolysis times less than 30 min (Supporting Information, Figure S2).

j Gz), and Apparent Density (dGapp) of Amylose and Amylopectin j w), z-Average Radius of Gyration (R Table 4. Weight Average Molar Mass (M after Hydrolysis of Maize Starch (31% d.b.) by RA and PPAa amylopectin j w* (g · mol-1) M 3.08 × 10 9.93 × 107 5.97 × 107 3.28 × 108 3.36 × 108 8

native starch RA 48 h RA 96 h PPA 48 h PPA 96 h

j Gz* (nm) R 270.0 148.4 87.8 270.0 280.2

amylose dGapp* (g · mol-1 · nm-3)

j w+(g · mol-1) M

amylose/amylopectin ratio

3.73 7.25 21.04 3.98 3.65

5.65 × 10 4.93 × 106 1.94 × 106 3.28 × 106 7.24 × 106

0.5 nd nd 0.5 0.4

6

a nd: not determined; *values obtained over the whole amylopectin peak; +values at the maximum of amylose peak; the experimental uncertainty was 5%; dGapp ) Mw/(4π/3)RG3.

Table 5. DP1-DP7 Composition in the Soluble Fraction during Hydrolysis of 31% Starch Suspensions by RA and PPA

RA PPA

time (h)

DP1 (%)

DP2 (%)

DP3 (%)

DP4 (%)

DP5 (%)

DP6 (%)

DP7 (%)

2 8 48 2 8 48

100.0 100.0 100.0 9.9 11.4 17.1

0.0 0.0 0.0 25.1 25.4 25.0

0.0 0.0 0.0 22.6 20.4 16.4

0.0 0.0 0.0 4.1 4.5 7.8

0.0 0.0 0.0 10.6 10.9 11.1

0.0 0.0 0.0 15.3 15.2 12.3

0.0 0.0 0.0 12.5 12.2 10.4



39

Figure 5. Evolution of the activity of RA ([) and PPA (2) with time at their optimum pH and temperature. Line is given for a better view of the evolution of PPA activity.

Figure 4. Relative amount of glucose (×), maltose (| | |) and isomaltose (0) released at 2 and 48 h during hydrolysis of 63-R-Dglucosyl-maltotriose (a) and 63-R-D-glucosyl-maltotriosyl-maltriose (b) by RA.

In contrast, hydrolysis by PPA leads to a wide range of oligosaccharides with DP between 1 and 15. Such a composition is very comparable to that of classical R-amylases, which yield a broader range of oligomers when acting on solid starch.40,53 The major part of this soluble fraction is maltose (DP2), which is known to induce a potential inhibition of amylases,23,55,56 while maltotriose and maltohexaose are slowly hydrolyzed between 8 and 48 h. The action mode of RA is therefore very specific. A similar behavior has already been described for the fungal R-amylases from A. flaVus.57 Planchot et al.58 have also observed the same behavior of the A. fumigatus R-amylase, which produced only glucose in the R-anomeric configuration when acting on solid substrates. They concluded that, as glucose produced by this R-amylase retains its anomeric configuration, such enzyme is different from glucoamylases, which liberates glucose in the β-anomeric form exclusively. Oligosaccharide Profiles in Homogeneous and Heterogeneous Phase Hydrolysis. The nature of oligomers released from hydrolysis of solubilized starch, amylose, and amylopectin was also studied. The soluble fraction stemming from RA action essentially consists of glucose as for hydrolysis of raw starch, while that of PPA contains a larger range of DP. The relative amount of oligomers produced by PPA is slightly different from those released from raw starch, which is not surprising when considering the higher accessibility and mobility of solubilized starch. Potential Hydrolysis of r(1,6) Linkages by RA. The ability of RA to hydrolyze the R(1,6) linkages was checked through hydrolysis of the branched oligosaccharides, 63-R-D-glucosylmaltotriose and 63-R-D-glucosyl-maltotriosyl-maltotriose, followed by HPAEC analysis of the resulting oligosaccharides. The two branched oligosaccharides were completely hydrolyzed after 48 h releasing glucose, maltose and isomaltose. Figure 4 clearly shows that the amount of glucose increases very rapidly

with time of hydrolysis reaching 87% at 2 h and 97% after 48 h for 63-R-D-glucosyl-maltotriose. The resistance of all R(1,6) linkages should have in theory led to the release of 50% isomaltose and 50% glucose (ratio 1:1) in the case of 63-R-Dglucosyl-maltotriose and 57% isomaltose and 43% glucose (ratio 4:3) for 63-R-D-glucosyl-maltotriosyl-maltotriose. Because glucose was detected as the primary product at 48 h, hydrolysis of the R(1,6) linkages must have taken place. However, it must be noticed that hydrolysis of the isomaltose still present at 2 h is very slow because its concentration at 2 and 48 h, respectively, are more or less similar. This may be explained by RA being better in hydrolyzing R(1,6) linkages when they are in a long substrate rather than in a short one such as isomaltose.

Results and Discussion about the Factors Resulting in a Limited Hydrolysis Decrease of Enzyme Activity As a Function of Hydrolysis Time. Figure 5 shows the relative activity of the two enzymes as a function of hydrolysis time under the buffer and temperature conditions as used for hydrolysis. Under these conditions, RA is stable for more than 8 days. Therefore, the action of RA on raw starch for up to 96 h is not limited by a decrease in the activity of the enzyme but rather by the kinetics. When looking at the curves in Figure 1, no plateau is reached neither at 48 h nor at 96 h. In contrast, the activity of PPA decreases progressively over time to become 3 times lower at 48 h. This may explain why the hydrolysis degree changes weakly after 48 h. Inhibition by Soluble Oligomeric End Products. For both RA and PPA, the final degree of hydrolysis of the 5% starch suspensions does not change significantly after washing the residue and adding a fresh amylase solution (Table 6). Therefore, no obvious inhibition by soluble oligosaccharides of the two enzymes is occurring when they act on 5% starch suspensions. For 31% starch suspensions, a new starch fraction is hydrolyzed with the degree of hydrolysis increasing from 75 to 83% and from 30 to 49% for RA and PPA, respectively. It could indicate a slight inhibition of the enzymes by soluble oligosaccharides present at high starch concentrations, especially for PPA because, as mentioned above, the hydrolysis plateau is most likely not reached at 96 h for RA. Nevertheless, other parameters as non effective adsorption of enzymes onto residual starch or very resistant residual structure can also be involved in the decline in hydrolysis rate.

40


Tawil et al.

Table 6. Evolution of the Final Hydrolysis Degree FHD after Removal of the Soluble Fraction and Adding a New Fresh Amylase Solution FHD (%) RA

FHD (%) PPA

time (h)

5%

31%

5%

31%

96 192

93 94

75 83

89 90

32 49

Table 7. Amount of Adsorbed Enzyme (%) on Residual Starch at 96 h Hydrolysis starch concentration (d.b.)

RA (%)

PPA (%)

5% 31%

98 88

93 93

General Discussion

maize starch. Nevertheless, resistance to hydrolysis could not be completely explained by complexation of amylose because Morrison and co-workers60 show that the rate of complexed amylose is only 15% of the total amylose content. Such resistance of high-amylose starch substrates to enzymatic hydrolysis is well-known24,30 and amylose gels have proven to be much more resistant to hydrolysis than amylopectin gels.61,62 On the contrary, amylose and amylopectin amounts decrease concomitantly during PPA hydrolysis, which is consistent with a mode of attack granule by granule already observed for hydrolysis of wheat starch and other starches in the solid state.21,22 Depending on the rate of hydrolysis, A-type (major part of maize starch), Vh-type (amylose-lipid complexes), and B-type (from rearrangement of linear fragments of amylose) structures are detected in the residues. It is well-known that the allomorphic type influences the rate of hydrolysis. For example, B-type, which is generally found in tubers and high-amylose starch, is usually more resistant to enzymatic hydrolysis compared to A-type.1,27,55,63 Resistant A-type structure has only been found in some substrates originating from acid hydrolysis of starch21 or aggregation of starch crystals53,64 due to the lack of accessible glucan double helices for the amylases. In the present work, maize starch presents the A-type characteristic from cereal starch and is characterized by a high susceptibility to enzymatic hydrolysis, as shown by this work and as reported earlier.1,30,63,65 However, such a rapid and preferential hydrolysis of crystalline domains, as demonstrated for RA, has never been described before. It differs from cellulases, which usually degrade preferentially amorphous cellulose, leading to an increase of the degree of crystallinity during hydrolysis.66 The strong resistance of Vh-type structure, characteristic of amylose-lipid complexes, compared to A-type is more controversial. It has been observed during enzymatic hydrolysis of cereal starch at long hydrolysis times by Gernat and coworkers25 and Lauro and co-workers,28 but Gerard et al.24,67 have shown on maize starch mutants that A- and Vh-types have

RA is highly effective in hydrolysis of raw starch suspensions, being able to degrade 75% of 31% starch at 32 °C. This very high efficiency at high starch concentration, when compared to PPA, which is unable to hydrolyze more than 30-35%, makes it very suitable for low temperature glucose syrup or bioethanol production. Its mode of action is also very specific and quite interesting in comparison to other amylases, such as, for example, PPA. It is remarkably efficient on the crystalline structures at high starch concentration because the crystallinity decreases very rapidly during the first stages of hydrolysis. At 5% starch concentration, the hydrolysis kinetics is closer to that of PPA, that is, a higher final rate of hydrolysis but the same type of action on the crystalline structure: a long resistance of the A-type X-ray diagram and a stable amount of Vh-type structure during the major part of the hydrolysis. Amylose is less rapidly hydrolyzed by RA compared to amylopectin during the first stages of hydrolysis, which is consistent with both the rapid hydrolysis of the crystalline domains, that is, originating from the amylopectin, and the resistance to hydrolysis of amylose lipid complexes present in

Figure 6. Crystalline structure of residual starch at very high degree of hydrolysis by RA: (a) 5% starch 48 h, hydrolysis rate 94%; (b) 31% starch 192 h, hydrolysis rate 83%.

Ineffective Adsorption of Enzymes on the Substrate. Table 7 shows the amount of adsorbed enzymes onto the residual substrate at the end of the hydrolysis. The amount of bound enzyme was calculated from the difference of enzyme concentration in the supernatant between the starting time of hydrolysis and after 96 h. Independent of enzyme and starch concentration at least 88% (for RA on a 31% starch suspension and hydrolysis still in progress) and up to 98% was bound at the end of hydrolysis. Therefore, an irreversible and potentially ineffective adsorption takes place during long hydrolysis periods. To check if it could be responsible for the decline of the hydrolysis rate, amylase was hydrolyzed by proteinase K, and after washing/ centrifugation, a fresh amylase solution was added. No further hydrolysis was observed, which means that addition of fresh enzyme does not lead to further hydrolysis, most likely because of a very resistant structure of the remaining starch residue. Crystalline Structure of the Residue. To determine which type of structure could be resistant and prevent a complete hydrolysis, the residues obtained after 192 h of hydrolysis were analyzed by XRD. As shown in Figure 6, for very high degrees of hydrolysis, a mixture of B- and Vh-type is found in the 5% starch residue and almost pure B-type in the 31% starch residue hydrolyzed by RA. Such a rearrangement into B-type is favored at higher starch concentrations when amylose fragments are released by hydrolysis. The higher resistance to amylolysis of Vh-type when compared to A-type in cereal starches has been observed earlier.25,27,28 The B-type structure formed prevents any progress of hydrolysis once the Vh-type is degraded. Indeed B-type structures have proven to be very resistant to hydrolysis.23,24,28 Such rearrangement during hydrolysis has already been described by Lopez-Rubio and co-workers59 when looking at hydrolysis of high amylose starch.


the same susceptibility to PPA hydrolysis. Moreover, it is wellknown that in vitro formed amylose-lipid complexes are completely degraded by R-amylases.56,65,67,68 In our work, the presence of Vh structure during RA hydrolysis at 48 h and longer hydrolysis times could result from two mechanisms: (i) Vh-type crystalline domains could have been present initially in the native granules but in a relative amount too small to be detected by X-ray diffraction before hydrolysis of A-type structure; (ii) Vh-type could be formed during hydrolysis between amylose fragments released by enzyme and lipid present in maize starch. In this work, the B-type structure appears at very high degrees of hydrolysis. It probably results from recrystallization of linear fragments of amylose released by the amylases when remaining amylose is widely hydrolyzed. It could also be favored by the hydrolysis of R(1,6) linkages leading to release of linear fragments with strong preference for recrystallization. The high initial concentration of starch favors crystallization and precipitation and this type of structure probably constitutes the real resistant part to enzymatic hydrolysis. Such reorganization during enzymatic hydrolysis has already been observed by Lopez-Rubio and co-workers59 on resistant starch and high amylose maize starch. The stronger resistance of B-type compared to Vh-type has already been observed by Gerard and co-workers67 on B- and Vh-type starch from aedu double mutants of maize. Recrystallization of amylose into B-type is also a major way to prepare highly resistant starch.59 The efficiency of RA on concentrated raw starch suspensions at 32 °C, the temperature used for bioethanol production, is surprising. RA is also very specific when considering the nature of oligosaccharides produced. It produces only glucose in contrast to PPA, which yields a classical range of oligosaccharides with DP between 1 and 15. This behavior has already been observed for fungal R-amylases from, for example, A. fumigatus58 or glycosylated amylases69 but under much less concentrated conditions. Surprisingly, the amount of PPA adsorbed onto starch at 48 h hydrolysis is similar to that of RA although RA has a covalently linked starch binding domain. Therefore this starch binding domain seems to have a more important role on hydrolysis kinetics than on adsorption. One role could be involvement in the disentanglement of double helices present in the crystalline domains of starch granules, that is, explaining the efficiency of RA on crystalline structures compared to that of PPA.70

Conclusion RA is shown to be much more efficient than PPA on concentrated raw starch suspensions. Its final degree of starch solubilization reaches 75-80% after 96 h in a 31% starch suspension. Our results suggest that the resistance to hydrolysis is due to a mixture of factors and differs between the two enzymes RA and PPA. The presence of amylose-lipid complexes at extensive hydrolysis times in the case of RA is supposed to be an important factor for the resistance, whereas the crystalline domains play a more important role in the case of PPA. In addition, a small inhibitory effect of the soluble oligosaccharides is seen on the hydrolysis rate at high starch concentrations. Finally, at 31% starch concentration and highest degrees of hydrolysis, rearrangements of amylose fragments into B-type structures is probably the final limiting factor for complete hydrolysis. One of the most surprising results is that RA is able to hydrolyze the crystalline domain easier than the amorphous ones, while PPA hydrolyzes them concomitantly.


41

Acknowledgment. Authors thank S. Guilois, M. de Carvalho, and B. Pontoire for excellent technical assistance. B. Henrissat (AFMB, CNRS Marseille) and G. Veronese (LISBP, INSA Toulouse) are also acknowledged for helpful discussion. Supporting Information Available. HPAEC analysis of the composition in DP1-DP7 oligomers produced at short hydrolysis times by RA on 5 and 31% maize starch suspensions. This material is available free of charge via the Internet at http:// pubs.acs.org.

References and Notes (1) Buleon, A.; Colonna, P.; Planchot, V.; Ball, S. Int. J. Biol. Macromol. 1998, 23, 85–112. (2) Donald, A. M.; Kato, K. L.; Perry, T. A.; Waigh, T. A. Starch/Staerke 2001, 53, 504–510. (3) Cameron, R. E.; Donald, A. M. Carbohydr. Res. 1993, 225, 244. (4) French, D. In Starch, Chemistry and Technology; Whistler, R. L., BeMiller, J. N., Parschall, E. F., Eds.; Academic Press: New York, 1984; pp 183-247. (5) Paris, M.; Bizot, H.; Emery, J.; Buzare´, J. Y.; Buleón, A. Carbohydr. Polym. 1999, 39, 327–339. (6) Zobel, H. F. Starch/Staerke 1988, 40, 1–7. (7) Lemke, H.; Burghammer, M.; Flot, D.; Rossle, M.; Riekel, C. Biomacromolecules 2004, 5, 1316–1324. (8) Imberty, A.; Buleon, A.; Tran, V.; Perez, S. Starch/Staerke 1991, 43, 375–384. (9) Imberty, A.; Chanzy, H.; Perez, S.; Buleon, A.; Tran, V. J. Mol. Biol. 1988, 201, 365–378. (10) Imberty, A.; Perez, S. Carbohydr. Res. 1988, 181, 41–55. (11) Popov, D.; Buleon, A.; Burghammer, M.; Chanzy, H.; Montesanti, N.; Putaux, J. L.; Potocki-Veronese, G.; Riekel, C. Macromolecules 2009, 42, 1167–1174. (12) Takahashi, Y.; Kumano, T.; Nishikawa, S. Macromolecules 2004, 37, 6827–6832. (13) Morrison, W. R.; Law, R. V.; Snape, C. E. J. Cereal Sci. 1993, 18, 107–109. (14) Biliaderis, C. G. Food Technol. 1992, 46, 98–&. (15) Buleon, A.; Colonna, P. The chemical physics of food; Blackwell Publishing: Oxford, U.K., 2007; pp 20-59. (16) Kugimiya, M.; Donovan, J. W. J. Food Sci. 1981, 46, 765–777. (17) Kugimiya, M.; Donovan, J. W.; Wong, R. Y. Starch/Staerke 1980, 32, 256–270. (18) Le Bail, P.; Bizot, H.; Ollivon, M.; Keller, G.; Bourgaux, C.; Buleon, A. Biopolymers 1999, 50, 99–110. (19) Godet, M. C.; Tran, V.; Delage, M. M.; Buleón, A. Int. J. Biol. Macromol. 1993, 15, 11–16. (20) Rappenecker, G.; Zugenmaier, P. Carbohydr. Res. 1981, 89, 11. (21) Colonna, P.; Buleon, A.; Lemarie, F. Biotechnol. Bioeng. 1988, 31, 895–904. (22) Oates, C. G. Trends Food Sci. Technol. 1997, 8, 375–382. (23) Colonna, P.; Leloup, V.; Buleon, A. Eur. J. Clin. Nutr. 1992, 46, S17– S32. (24) Gerard, C.; Colonna, P.; Buleon, A.; Planchot, V. J. Sci. Food Agric. 2001, 81, 1281–1287. (25) Gernat, C.; Radosta, S.; Anger, H.; Damaschun, G. Starch/Staerke 1993, 45, 309–314. (26) Williamson, G.; Belshaw, N. J.; Self, D. J.; Noel, T. R.; Ring, S. G.; Cairns, P.; Morris, V. J.; Clark, S. A.; Parker, M. L. Carbohydr. Polym. 1992, 18, 179–187. (27) Kwasniewska-Karolak, I.; Nebesny, E.; Rosicka-Kaczmarek, J. Food Sci. Technol. Int. 2008, 14, 29–37. (28) Lauro, M.; Forssell, P. M.; Suortti, M. T.; Hulleman, S. H. D.; Poutanen, K. S. Cereal Chem. 1999, 76, 925–930. (29) Manelius, R.; Qin, Z.; Avall, A. K.; Andtfolk, H.; Bertoft, E. Starch/ Staerke 1997, 49, 142–147. (30) Planchot, V.; Colonna, P.; Buleon, A. Carbohydr. Res. 1997, 298, 319–326. (31) Sujka, M.; Jamroz, J. Int. Agrophys. 2007, 21, 107–113. (32) Leloup, V. M.; Colonna, P.; Ring, S. G. Biotechnol. Bioeng. 1991, 38, 127–134. (33) Macri, L. J.; Macgregor, A. W.; Schroeder, S. W.; Bazin, S. L. J. Cereal Sci. 1993, 18, 103–106. (34) Wiegand, G.; Epp, O.; Huber, R. J. Mol. Biol. 1995, 247, 99–110. (35) Bompard, G. C.; Rousseau, P.; Rouge, P.; Payan, F. Structure 1996, 4, 1441–1452.

42


(36) Vallee, F.; Kadziola, A.; Bourne, Y.; Juy, M.; Rodenburg, K. W.; Svensson, B.; Haser, R. Structure 1998, 6, 649–659. (37) Andre, G.; Buleon, A.; Haser, R.; Tran, V. Biopolymers 1999, 50, 751–762. (38) Leloup, V. M.; Colonna, P.; Ring, S. G. J. Cereal Sci. 1992, 16, 253– 266. (39) Asp, N. G. Eur. J. Clin. Nutr. 1992, 46. (40) Sharma, A.; Yadav, B. S.; Ritika, Food ReV. Int. 2008, 24, 193–234. (41) Englyst, H. N.; Kingman, S. M.; Cummings, J. H. Eur. J. Clin. Nutr 1992, 46, S33–S50. (42) Adams, P. R. Mycopathologia 1990, 112, 35–37. (43) Adams, P. R. Mycopathologia 1994, 128, 139–141. (44) Ismail, M. A. Int. Biodeterior. Biodegrad. 1993, 31, 143–157. (45) Silva, T. M.; Attili-Angelis, D.; Carvalho, A. F. A.; Da Silva, R.; Boscolo, M.; Gomes, E. J. Microbiol. 2005, 43, 561–568. (46) Turchi, S. L.; Becker, T. Curr. Microbiol. 1987, 15, 203–205. (47) Viksø-Nielsen, A.; Andersen, C.; Liu, J. Pat. WO2007144424-A2, 2009. (48) Faisant, N.; Champ, M.; Colonna, P.; Buleon, A. Carbohydr. Polym. 1993, 21, 205–209. (49) McCleary, B. V.; McNally, M.; Monaghan, D.; Mugford, D. C.; Black, C.; Broadbent, R.; Chin, M.; Cormack, M.; Fox, R.; Gaines, C.; Gothard, P.; Home, S.; Howes, E.; Johnson, C.; Keeping, R.; Koliatsou, M.; Lindhauer, M.; de Sa, R. M.; Martin, R.; Monaghan, D.; Nees, U.; Nishwitz, R.; Palmer, G.; Panozzo, J.; Recabarren, J.; Roumeliotis, S.; Seddig, S.; Solah, V.; Sonnet, M.; Themeier, H. J. AAOC Int. 2002, 85, 1096–1102. (50) Rolland-Sabate, A.; Amani, N. G.; Dufour, D.; Guilois, S.; Colonna, P. J. Sci. Food Agric. 2003, 83, 927–936. (51) Bradford, M. M. Anal. Biochem. 1976, 72, 248–254. (52) Wakelin, J. H.; Virgin, H. S.; Crystal, E. J. Appl. Phys. 1959, 30, 1654–1662. (53) Pohu, A.; Planchot, V.; Putaux, J. L.; Colonna, P.; Buleon, A. Biomacromolecules 2004, 5, 1792–1798. (54) Buleon, A.; Veronese, G.; Putaux, J. L. Aust. J. Chem. 2007, 60, 706– 718.

Tawil et al. (55) Elodi, P.; Mora, S.; Krysteva, M. Eur. J. Biochem. 1972, 24, 577– 582. (56) Faisant, N.; Champ, M.; Colonna, P.; Buleon, A.; Molis, C.; Langkilde, A. M.; Schweizer, T.; Flourie, B.; Galmiche, J. P. Eur. J. Clin. Nutr. 1993, 47, 285–296. (57) AbouZeid, A. M. Microbios 1997, 89, 55–66. (58) Planchot, V.; Colonna, P.; Gallant, D. J.; Bouchet, B. J. Cereal Sci. 1995, 21, 163–171. (59) Lopez-Rubio, A.; Flanagan, B. M.; Shrestha, A. K.; Gidley, M. J.; Gilbert, E. P. Biomacromolecules 2008, 9, 1951–1958. (60) Morrison, W. R.; Tester, R. F.; Snape, C. E.; Law, R.; Gidley, M. J. Cereal Chem. 1993, 70, 385–391. (61) Leloup, V. M.; Colonna, P.; Buleon, A. J. Cereal Sci. 1991, 13, 1– 13. (62) Leloup, V. M.; Colonna, P.; Ring, S. G.; Roberts, K.; Wells, B. Carbohydr. Polym. 1992, 18, 189–197. (63) Gallant, D. J.; Bouchet, B.; Buleon, A.; Perez, S. Eur. J. Clin. Nutr. 1992, 46 (Suppl 2), S3–16. (64) Pohu, A.; Putaux, J. L.; Planchot, V.; Colonna, P.; Buleon, A. Biomacromolecules 2004, 5, 119–125. (65) Seneviratne, H. D.; Biliaderis, C. G. J. Cereal Sci. 1991, 13, 129– 143. (66) Zhang, Y. H. P.; Lynd, L. R. Biotechnol. Bioeng. 2004, 88, 797–824. (67) Gerard, C.; Colonna, P.; Buleon, A.; Planchot, V. Carbohydr. Polym. 2002, 48, 131–141. (68) Holm, J.; Bjorck, I.; Ostrowska, S.; Eliasson, A. C.; Asp, N. G.; Larsson, K.; Lundquist, I. Starch/Staerke 1983, 35, 294–297. (69) Koyama, I.; Komine, S.; Yakushijin, M.; Hokari, S.; Komoda, T. Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol. 2000, 126, 553–560. (70) Juge, N.; Nohr, J.; Le Gal-Coeffet, M. F.; Kramhoft, B.; Furniss, C. S. M.; Planchot, V.; Archer, D. B.; Williamson, G.; Svensson, B. Biochim. Biophys. Acta, Proteins Proteomics 2006, 1764, 275–284.

BM100913Z

In Depth Study of a New Highly Efficient Raw Starch Hydrolyzing Î±

Recommend Documents