Degradation of chitosans with chitinase B from Serratia marcescens

Production of chito-oligosaccharides and insight into enzyme processivity


K. M. Vårum, Norwegian Biopolymer Laboratory (NOBIPOL), Department of Biotechnology, Norwegian University of Science and Technology, 7491 Trondheim, Norway
Fax: +47 73591283
Tel: +47 73593324


Family 18 chitinases such as chitinase B (ChiB) from Serratia marcescens catalyze glycoside hydrolysis via a mechanism involving the N-acetyl group of the sugar bound to the −1 subsite. We have studied the degradation of the soluble heteropolymer chitosan, to obtain further insight into catalysis in ChiB and to experimentally assess the proposed processive action of this enzyme. Degradation of chitosans with varying degrees of acetylation was monitored by following the size-distribution of oligomers, and oligomers were isolated and partly sequenced using 1H-NMR spectroscopy. Degradation of a chitosan with 65% acetylated units showed that ChiB is an exo-enzyme which degrades the polymer chains from their nonreducing ends. The degradation showed biphasic kinetics: the faster phase is dominated by cleavage on the reducing side of two acetylated units (occupying subsites −2 and −1), while the slower kinetic phase reflects cleavage on the reducing side of a deacetylated and an acetylated unit (bound to subsites −2 and −1, respectively). The enzyme did not show preferences with respect to acetylation of the sugar bound in the +1 subsite. Thus, the preference for an acetylated unit is absolute in the −1 subsite, whereas substrate specificity is less stringent in the −2 and +1 subsites. Consequently, even chitosans with low degrees of acetylation could be degraded by ChiB, permitting the production of mixtures of oligosaccharides with different size distributions and chemical composition. Initially, the degradation of the 65% acetylated chitosan almost exclusively yielded oligomers with even-numbered chain lengths. This provides experimental evidence for a processive mode of action, moving the sugar chain two residues at a time. The results show that nonproductive binding events are not necessarily followed by substrate release but rather by consecutive relocations of the sugar chain.


chitinase B


degree of polymerization






degree of scission


fraction of acetylated units

Chitosans are linear, cationic polysaccharides composed of (1→4)-linked units of 2-amino-2-deoxy-β-d-glucopyranose (GlcN, D-unit) which may be N-acetylated to varying extents. Chitosans are soluble in acidic solution, whereas chitin, a structural polysaccharide occurring mainly in the exo-skeleton of arthropods, is insoluble [1]. Chitin is composed of (1→4)-linked units of 2-acetamido-2-deoxy-β-d-glucopyranose (GlcNAc, A-unit). Chitin shares many similarities with cellulose, e.g. the conformation of the monomers, di-equatorial glycosidic linkages, crystalline structure, and lack of solubility in aqueous solvents.

The detailed chemical composition of chitosans will vary, depending on the fraction of acetylated units (FA) and their synthesis or preparation. This variation affects many properties of chitosan, such as solubility as a function of pH [2], binding to lysozyme [3], susceptibility to degradation by lysozyme [4,5], as well as functional properties in drug delivery [6] and gene delivery systems [7]. Previous studies have shown that chitosans obtained by homogeneous de-N-acetylation of chitin, such as those used in the present study, have a random distribution of N-acetyl groups [8–10].

Chitinases and chitosanases are capable of converting chitin and chitosans to low molecular mass products (oligosaccharides) by hydrolyzing the β(1→4) glycosidic linkages between the sugar units. There are several families of chitinases (glycoside hydrolase families 18 and 19) and chitosanases (glycoside hydrolase families 46, 75 and 80) [11]. These enzymes differ with respect to their preference for A- and D-units on each side of the scissile glycosidic bond and, consequently, differ with respect to their activities towards different types of chitosan. Like many other chitinolytic bacteria, the Gram-negative soil bacterium Serratia marcescens produces multiple chitinases (ChiA, ChiB, ChiC1 and ChiC2)[12–14]. These enzymes all belong to family 18 chitinases, which have a characteristic catalytic mechanism which depends on participation of the N-acetyl group of the sugar unit bound to the −1 subsite [15–19]. This substrate-assisted catalytic mechanism implies that family 18 chitinases are expected to have an absolute preference for A-units in the −1 subsite and that the presentation of a deacetylated sugar (D-unit) to the −1 subsite would represent nonproductive binding.

Chitinase B (ChiB; EC from S. marcescens is a two-domain family 18 chitinase [20]. The active site of ChiB has defined subsites from −3 to +3 [16,20], and the substrate-binding cleft is tunnel-like, permitting tight interaction with the polymeric substrate. The aglycon-side of the substrate-binding groove is extended by the aromatic surface of a putative chitin-binding domain. The crystal structure of ChiB further suggests that the active site cleft is partially blocked at the −3 subsite [20]. Together with studies showing that ChiB converts chitin primarily to dimers [14,21], the structure of ChiB suggests that the enzyme exerts exo-activity and degrades chitin chains from their nonreducing ends [16,20].

It is generally accepted that enzymes that degrade carbohydrate polymers can do so by three principally different mechanisms, as first described by Robyt and French [22,23]: (a) A multiple-chain mechanism, where the enzyme-substrate complex dissociates after each reaction. (b) A single-chain mechanism, where the enzyme remains associated with the substrate until every cleavable linkage in the chain has been hydrolyzed. (c) A multiple attack mechanism, where a given average number of attacks are performed after the enzyme-substrate complex has been formed. The latter two mechanisms are often referred to as ‘processive’. Carbohydrate degrading enzymes with tunnel-like active site clefts are often suggested to act processively [24]. For example, cellobiohydrolases are thought to degrade cellulose according to a multiple attack mechanism, where dimers are cleaved off from the end of the polysaccharide chain [24–27]. These cellulose-degrading enzymes convert cellulose to cellobiose, along with small amounts of monomers and trimers, which are thought to be the result of the initial binding/cleaving event. Whereas plausible models exist for cellulase action, mainly based on the results of site-directed mutagenesis and structural studies, less is known about processivity in chitinases. The only available information comes from Uchiyama et al. [28], who used electron microscopy to study the degradation of β-chitin fibers by ChiA from S. marcescens and who concluded that their results were compatible with a processive mode of action.

One goal of the present study was to exploit the unique heteropolymeric nature of chitosan to study the mechanism of action of ChiB. A second goal of this study was to explore possibilities of producing chito-oligosaccharide mixtures by degrading chitosans with ChiB. This latter goal is of interest because chito-oligosaccharides have a number of potential applications [29–32]. To obtain these goals, we have analyzed the degradation of well-characterized, fully water-soluble, high molecular mass chitosans with ChiB, using a newly developed chromatographic method for oligomer separation and previously established NMR-based methods [5,9,10,33] for analysis of oligomer composition. For example, we studied the time course of depolymerization of a highly acetylated but still water-soluble chitosan of high molecular mass, and the resulting oligosaccharides were characterized with respect to their chain length and chemical composition. In addition, high molecular mass chitosans of varying FA were degraded, and the resulting oligosaccharides were characterized in the same way. The results show that ChiB acts as an exo-chitinase/-chitosanase with specific requirements towards acetylated/deacetylated sugar units in subsites −2 and −1, which is reflected in the formation of oligomers with widely varying size distributions and chemical compositions, depending on the type of chitosan used. The results also provide unequivocal experimental evidence for a processive mode of action in ChiB.


Size-exclusion chromatography of chito- oligosaccharides

The size-exclusion chromatographic system was extensively calibrated using fully acetylated and fully de-N-acetylated oligomer standards. Figure 1 shows the degree of polymerization as a function of the distribution coefficient


where Ve is the elution volume of the actual oligomer, and V0 and Vt is the void volume and the total volume of the column, respectively. Although the two homologous oligomer-series do not elute at exactly the same Kav values, the calibration lines are close and nearly parallel. Extrapolation of the calibration lines to Kav = 0 revealed that the corresponding degree of polymerization (DP)-value was 40. The calibration lines of the acetylated- and the de-N-acetylated oligomers did not intercept (an interception of the calibration lines would mean that a fully deacetylated oligomer would co-elute with an acetylated oligomer of different chain length). Thus, the chromatographic system allows the separation of mixtures of partially N-acetylated oligomers according to size (DP) and not according to chemical composition, at least in the separation range between DP 4 (see below for details) and a DP of approximately 20 (Fig. 4; these are the longest oligomers that can be observed separately). It should be noted that the fully acetylated and fully de-N-acetylated standards represent the extreme limits of the possible variation in chemical composition of an oligomer with a given DP, while the enzymatic degradation of chitosans results in oligomer fractions with less variation in chemical composition. As shown in Fig. 1 and by the results presented below, the fully acetylated dimer (AA) and trimer (AAA) are resolved from the corresponding deacetylated dimer (DD) and trimer (DDD).

Figure 1.

Calibration of SEC-columns. Degree of polymerization (DP) plotted as a function of the distribution coefficient (Kav) for fully acetylated and fully de-N-acetylated oligomers (monomer to hexamer). The lines correspond to the following equations: log DP = −1.826 Kav +1.597 (fully acetylated); log DP = −1.714 Kav +1.604 (fully de-N-acetylated).

Figure 4.

Size-distribution of oligomers after ChiB degradation (FA =0.65). Chromatograms showing the size-distribution of oligomers obtained after hydrolysis of chitosan (FA = 0.65) with ChiB to degrees of scission (α) varying from 0.03 to 0.37. Peaks are labelled with DP-values or with the sequence of the oligomer.

ChiB degradation of a high molecular mass chitosan with FA = 0.65

Time course of the reaction

Progress in chitosan degradation can be determined as the degree of scission (α), i.e. the fraction of glycosidic linkages in the chitosan that has been cleaved by the enzyme, by monitoring the increase in reducing end resonances relative to resonances from internal anomer protons in a 1H-NMR spectrum of the reaction mixture. Figure 2 shows the anomer region of the 1H-NMR-spectrum of a high molecular mass chitosan with FA of 0.65 which had been incubated with ChiB for periods from 15 min (α = 0.03) to 168 h (α =0.37). These spectra were assigned from previously published assignments of 1H-NMR spectra of hydrolysed chitosans [33,38]. The β-anomer reducing end resonance appears as a doublet due to the relatively large coupling constant of 8.1 Hz to H-2. Depending on whether the preceding unit is an A-unit or a D-unit, the β-anomer reducing end resonance appears as two doublets at 4.705 p.p.m. (–AA) and 4.742 p.p.m. (–DA). This assignment was made from 1H-NMR data on acid hydrolysis of two chitosans with FA of 0.65 and 0.30. The relative intensities of the α/β-anomers correspond to the equilibrium ratio between the two anomers, which is 60 : 40 at pH 4–5 [39]. The α- and β-anomer reducing end resonances from a D-unit, which would be expected to appear at 5.43 p.p.m. [–D(α)] and 4.92 p.p.m. [–D(β)][33], are completely absent in the spectrum. The internal D- and A-units resonate at around 4.9 p.p.m. (–D-) and 4.6 p.p.m. (–A-) [9]. Using these assignments, the ratio between internal protons and reducing end protons can easily be calculated.

Figure 2.

Anomer region of the 1H-NMR spectrum of chitosan (FA = 0.65) after incubation with ChiB for periods ranging from 15 min (α = 0.03) to 10 080 min (α = 0.37) at 37 °C. The reducing end of the α-anomer of an acetylated unit resonates at 5.19 p.p.m., while the β-anomer from the same unit resonates at 4.7–4.8 p.p.m. [33]. D- and A-units within the chain and at nonreducing ends resonate near 4.9 and 4.6 p.p.m., respectively [9]. -AAβ and -DAβ are the reducing end β-anomer where the preceding unit is an A-unit or a D-unit, respectively.

The time course of hydrolysis of the FA = 0.65 chitosan (Fig. 3) shows that the product formation rate is constant until the degree of scission reaches approximately 0.19 (i.e. 19% of the glycosidic linkages has been cleaved). After this initial phase, a second, much slower kinetic phase becomes apparent in which the rate of the enzymatic reaction is decreased by a factor of 4.1. This slow phase continues until the degree of scission reaches 0.37, that is, a situation where on average more than one out of three glycosidic linkages have been cleaved (note that for an enzyme producing dimers only, the maximum value of α is 0.5 and that the presence of D-units lowers the maximum value of α).

Figure 3.

Time course of ChiB degradation of a chitosan with FA =0.65. The graph shows the degree of scission (α) as a function of time. The slope of the line at low α is 1.8 × 10−3 min−1, and at higher α-values 4.3 × 10−4 min−1. The α -values continue to increase to α = 0.37.

Size distribution of oligomers as a function of α (FA = 0.65)

The size distributions of oligomers resulting from hydrolysis of FA = 0.65 chitosan at different α-values are shown in Fig. 4. Undegraded chitosan, with a number average relative molecular mass of 160 000, elutes in the void volume of the column, as do all chitosan chains with DP > 40. Dimer fractions were found to elute in two peaks, which were identified (by 1H-NMR spectroscopy) as being AA (the dimer with the lowest Kav value) and DA. Similarly, trimer and tetramer fractions were found to elute in two peaks.

During the initial phase of the degradation process, i.e. at α-values lower than 0.2 (Fig. 3), the peak eluting in the void volume disappeared slowly, while short, mainly even-numbered oligomers (DP 2–12) were produced (Fig. 4). Around α = 0.20, a transition took place: as the void peak disappeared, the reaction slowed down (Fig. 3), and the relative amounts of oligomers with an odd number of sugar units began to increase. Thus, it seems that the slow phase of the reaction starts as the polymer eluting in the void volume is used up. In the final product mixture (α = 0.37), oligomer frequencies appear to be determined by oligomer length only, and do not seem to depend on the presence of an odd- or even number of sugar residues. The increased appearance of odd numbered oligomers coincides with the appearance of the second, slower kinetic phase visible in Fig. 3.

Chemical composition of oligomers

To obtain further insight into the chemical composition of the oligomers, we isolated several fractions obtained at α = 0.11 and α = 0.37, and analyzed these by 1H-NMR spectroscopy. The yield of odd-numbered oligomers (α = 0.11) was too low to obtain well-resolved NMR spectra. The results of these analyses are summarized in Table 1.

Table 1.  Composition of isolated oligomers. Chemical composition and sequence of isolated oligomer fractions obtained after degradation of chitosan (FA = 0.65) with ChiB to degrees of scission (α) of 0.11 and 0.37. ND, not determined.
α = 0.11FA = 0.93NDFA = 0.73NDFA = 0.66FA = 0.61
86% AA     
α = 0.37FA = 0.83FA = 0.65FA = 0.51FA = 0.57FA = 0.53ND

Because of an absolute preference for A-units at newly formed reducing ends (i.e. at subsite −1 of the enzyme), only two dimers are formed, AA and DA. In accordance with the NMR analyses shown in Fig. 2, the chromatographic data (Fig. 4) show that, at low α, the dimer mainly consists of AA (86% at α = 0.11). By the end of the reaction, the relative amount of DA starts to increase, finally reaching 34% of the total amount of dimers.

NMR analyses of the tetramer fraction at α = 0.11 (Fig. 5A) revealed a FA of 0.73 and a DP of 4.1. Electrospray ionization mass spectroscopy (ESI-MS) analysis showed that the dominating tetramer was composed of three A-units and 1 D-unit (results not shown). The minor resonances between 4.8 and 4.9 p.p.m. suggest that minor amounts of DDAA are present. The absence of a doublet at 4.742 p.p.m. in Fig. 5A (–DA; Fig. 2) shows that all tetramers has an A-unit next to the reducing end (–AA). The presence of only one major D-unit resonance at 4.865 p.p.m., and the FA of 0.73 show that there is one dominating tetramer which can be ADAA or DAAA.

Figure 5.

1H-NMR spectra (anomer region) of isolated oligomers obtained after hydrolysis of a chitosan with FA = 0.65 with ChiB. (A) Tetramer fraction (α = 0.11). (B) Tetramer fraction (α = 0.37). (C) Trimer fraction (α = 0.37).

1H-NMR analyses of the tetramer fraction at α =0.37 revealed a DP of 4.2 and an FA of 0.51 (Fig. 5B). ESI-MS data showed that the majority of the tetramers were composed of two A-units, whereas minor amounts of tetramers with one and three A-units could be detected (results not shown). There are three possibilities for tetramers containing two A-units with an acetylated reducing end, which are DDAA, DADA and ADDA. The fact that at least three different doublet resonances from internal D-units are observed (at 4.855, 4.895 and 4.905 p.p.m.; Fig. 5B) indicates that indeed several oligomers occur. The relative intensities of the doublets at 4.705 p.p.m. (–AA) and 4.742 p.p.m. (–DA) (Fig. 5B) show that the nearest-neighbour to the reducing ends are 70% acetylated and 30% deacetylated. Thus, one dominating tetramer must be DDAA. The other possible tetramers with an acetylated reducing end are DDDA, ADDA, DADA, DAAA, ADAA and AADA.

NMR analysis of the trimer fraction (α = 0.37; Fig. 5C) showed a DP of 2.90 and an FA of 0.65. The resonance at 4.705 p.p.m. and the absence of a resonance at 4.742 p.p.m. show that the nearest neighbour to the reducing end unit is an A-unit (–AA). This is confirmed by the two doublet resonances at 4.648 and 4.660 p.p.m. with a coupling constant of 8.1 Hz, which are assigned to an A-unit next to the reducing end and where the ratio between the two doublets reflects the α/β-anomer equilibrium at the neighbouring reducing end unit. The resonance at 4.880 p.p.m. is assigned to a deacetylated unit which must be located at the nonreducing end. Thus, the trimer is identified as DAA. Two very small doublet resonances at 4.742 p.p.m. reflect the presence of minor amounts of the trimers with a D-unit next to the acetylated reducing end (i.e. DDA and/or ADA). Based on the FA-value of 0.65 found for the trimer fraction, relative amounts of the trimers with two acetylated units (DAA and ADA) and the trimer with one acetylated unit (DDA) were calculated to be 95 and 5%, respectively (Table 1).

Table 1 also shows FA values for longer oligomers, as determined by 1H-NMR spectroscopy. The results show that the relative amount of D-units in the longer oligomers increases at high α. This indicates that some D-containing sequences are only hydrolyzed as other, more favourable substrates become scarce.

Degradation of chitosans with varying FA

Three high molecular mass chitosans with FA = 0.13, 0.32 and 0.50 (Table 2) were incubated with ChiB, and the chitosans were extensively depolymerized to maximum α. The results (Fig. 6) show that the size distribution of the product mixtures shifts towards higher oligomer lengths for substrates with lower FA values and, thus, longer and noncleavable stretches of consecutive D-units. In addition to a shift to longer oligomers, Fig. 6 shows a reduction in the AA/DA ratio, as would be expected when lowering the FA of the chitosan. A highly deacetylated chitosan (FA < 0.001) was not degraded when incubated with an excess of ChiB (result not shown).

Table 2.  Characterization of chitosans. Fraction of acetylated units (FA), diad frequencies (FAA, FAD, FDA and FDD), number-average block lengths (ND and NA) as determined from 600 MHz 1H-NMR spectroscopy [9]. The intrinsic viscosities ([η]) were determined as described previously [31] and the number-average molecular masses (Mn) were calculated from the intrinsic viscosities [32]. The experimentally determined diad frequencies and block lengths of each chitosan are compared with the calculated diad frequencies and block lengths of a chitosan with a random (Bernoullian) distribution of A- and D-units (numbers given in brackets). ND, not determined.
Chitosan 65%0.65740160 0000.42(0.42)0.23(0.23)0.12(0.12)1.5(1.5)2.8(2.9)
Chitosan 50%0.50850210 0000.27(0.25)0.24(0.25)0.26(0.25)2.1(2.0)2.1(2.0)
Chitosan 32%0.32820250 0000.12(0.10)0.20(0.22)0.48(0.46)3.4(3.1)1.6(1.5)
Chitosan 13%0.13910230 000NDNDNDNDND
Figure 6.

Size-distribution of oligomers after extended hydrolysis of various chitosans with ChiB. Chromatograms showing the size-distribution of oligomers obtained upon extended ChiB-hydrolysis of chitosans with FA of 0.65, 0.50, 0.32 and 0.13 to α-values (corresponding DPn-values in brackets) of 0.37 (2.7), 0.34 (2.9), 0.22 (4.5) and 0.11 (9.5), respectively.

The chemical compositions of selected oligomers (trimer to hexamer) from the experiments with chitosan (FA = 0.32) were investigated by 1H-NMR spectroscopy, using the methods described above (results not shown). The trimer fraction had a DP of 3.04 and a FA of 0.44, and was concluded to consist mainly of DDA and DAA, in addition to minor amounts of ADA. The tetramer fraction showed a DP of 4.1 and a FA of 0.33 and the dominant tetramers were found to be DDDA and DDAA. 1H-NMR analyses of the longer oligomers (pentamers, hexamers and heptamers) revealed two dominating oligomers which both contained A-units at the reducing end but which varied with respect to the neighbouring unit (A or D). All other units were deacetylated.


The NMR analysis of reaction products obtained upon degradation of chitosan by ChiB (Figs 2 and 5) showed that all oligomers had acetylated units at their reducing ends. This is in full agreement with the proposed substrate-assisted catalytic mechanism of ChiB [15,16]. Whereas many details are known about the catalytic mechanism of ChiB and other family 18 chitinases, much less is known about processivity. Some work has been carried out on ChiA, another family 18 chitinases from S. marcescens which, like ChiB, has a deep active site cleft. It was shown that ChiA is an exoenzyme and degrades the polymer from its reducing end [14,21,28]. On the basis of electron microscopy studies, Uchiyama et al. [28] proposed that ChiA acts processively.

In the present study, we have used a new approach to study the properties of ChiB, including possible processivity. When ChiB acts on chitin, it only produces dimers and trimers, indicating that ChiB is an exoenzyme, which can bind either two (subsites −1 and −2) or three (subsites −3, −2 and −1) sugar units in the product binding site [14,21]. The degradation of chitosans yielded products with more than three sugar units very early during the reaction (Fig. 4). This shows that, in the case of a chitosan substrate, the putative physical barrier at the −3 subsite in ChiB [20] does not prevent the enzyme from productive binding events that position more than three sugars on the glycon side of the catalytic centre. The observation of longer oligomers early during the reaction may seem to contrast with the putative exo-activity of the enzyme, but can be explained by processivity, as discussed below. The slow disappearance of the void volume peak and the early appearance of only shorter oligomers (Fig. 4) indicate that ChiB does indeed degrade the chitosan chains from their ends (as opposed to an endo-action), and thus is an exoenzyme.

If ChiB were to be a processive enzyme, one would expect chain movements by an even number of sugar units, as successive glycoside units in chitin/chitosan are rotated by 180° along the chain, meaning that the catalytically important N-acetyl group is positioned correctly in every second sugar only, as proposed for processive cellulases [40–42]. The clear dominance of even-numbered oligomers during the initial phase of the degradation of chitosan with FA 0.65 (Fig. 4) can only be explained by a processive mode of action. If each binding and, for productive binding, cleavage event would be followed by separation and rebinding, the longer initial products would be equally divided between odd-numbered and even-numbered oligomers. The enzyme does not have more than three subsites (−3, −2 and −1) on the glycon side of the catalytic centre, meaning that preferential production of, e.g. hexamers and octamers (as opposed to pentamers and heptamers) cannot be explained by specific binding interactions with the enzyme in a nonprocessive mode of action.

Taking together all information and observations, ChiB is likely to initially bind the chitosan chain with similar chances of having an odd or even number of sugar units positioned on the glycon side (most likely three, occupying subsites −3 to −1, or two, occupying subsites −2 and −1). Depending on the sequence of the bound chain, the first cleavage may then occur directly or after one or more consecutive movements by two sugar units at the time. This would lead to the initial production of an odd- or even-numbered oligomer, respectively. If the enzyme would act processively, all subsequent products coming out of the same binding event will be even-numbered regardless of the initial binding mode, leading to the observed initial dominance of even-numbered oligomers. Interestingly, the initial formation of even-numbered oligomers all the way to at least decamers not only reveals the processive mechanism, but also shows that processivity is not interrupted when a nonproductive or less-preferred enzyme–substrate complex emerges (for example a complex that positions a D-unit in the −1 subsite). In the later stages of enzymatic degradation odd-numbered oligomers appear in larger amounts. This is a consequence of the increased depolymerization of the substrate, which leads to more ‘initial attacks’.

It is generally accepted that cellobiohydrolases, which convert cellulose (which is closely related to and equally insoluble as chitin) mainly to dimers, act processively [40–42]. It is not straightforward to give evidence of a processive mechanism, as the parameter that is often used [the dimer/(trimer + monomer) ratio] strictly speaking does not discriminate between processivity and different initial binding events that occur with particular frequencies, leading to particular dimer/(trimer + monomer) ratios. The availability of the water-soluble, heteropolymeric chitosan, acting as a ‘pseudo-substrate’ for family 18 chitinases and resembling both chitin and cellulose, allowed us to obtain clear experimental data which unequivocally show that the enzyme acts processively.

The current data also provide some insight into the preference for A- or D-units in subsites −2 and +1. Because the nonreducing end sugar of an oligomer must have been bound productively in the +1 subsite in a previous cleavage, the occurrence of both D and A nonreducing ends shows that there is no clear preference for A or D in the +1 subsite. In the initial, faster stage of the depolymerization reaction (α < 0.2), most oligomers formed had an A-unit as nearest neighbour to the reducing end (–AA), indicating that most of the productive enzyme–substrate complexes had an A-unit bound in subsite −2. Upon more extensive hydrolysis, presumably leading to depletion of binding sites with the optimal AA sequence in −2 and −1, the reaction becomes slower, and an increase in oligomers with a D-unit as nearest neighbour to the reducing end (–DA) is observed. Thus, ChiB has a clear preference for A in subsite −2.

Catalysis in family 18 chitinases requires distortion of the sugar in the −1 subsite [16]. In analogy with, e.g. lysozyme, a certain minimum amount of binding energy is likely to be required to achieve this distortion and formation of a productive enzyme–substrate complex. This is reflected by some of the observations made in this study. For example, the trimer AAA is degraded, whereas the trimer DAA (which in principle could bind productively to the −2, −1 and +1 subsites) is not. Likewise, several of the tetramers with an –AA reducing end are not degraded while these two A-units could bind productively to the −1 and +1 subsites. These observations suggest that the ability to cleave a substrate that positions a D in the −2 subsite depends on how many of the subsites on the aglycone side are occupied by sugars.

The distribution of oligomers formed upon extensive ChiB depolymerization of chitosans with varying FA shifted towards higher oligomers with decreasing FA. This was expected because the average chain length of D-blocks that will not be attacked by the chitinase increases with decreasing FA (Table 2). By extensive ChiB-degradation of chitosans with FA from 0.1 to 0.5, we were able to produce longer oligomers (DP > 4) which were composed of only D-units, except for the reducing end unit and in some cases its nearest neighbour. By selecting the chitosan with the appropriate FA and the extent of degradation (α-value), oligomers with predetermined composition and chain length can be prepared.

In conclusion, we have found that chitosans with full water-solubility and known random distribution of A- and D-units are useful substrates for obtaining deeper insight into the mode of action of ChiB, including its processive character. The catalytic mechanism of ChiB requires an acetylated unit in the −1 subsite, but this does not prevent the enzyme from degrading a wide range of chitosans with varying chemical compositions. Thus, ChiB may be used to convert these chitosans to mixtures of oligosaccharides with predictable chain lengths and chemical compositions.

Experimental procedures


Chitin was isolated from shrimp shells by the method of Hackman [34] and milled in a hammer mill to pass through a 1.0-mm sieve. Chitosans, with degrees of N-acetylation of 65, 50, 32 and 13% (FA = 0.65, 0.50, 0.32 and 0.13), were prepared by homogeneous de-N-acetylation of chitin [35]. The chemical properties of the chitosans (FA, diad frequencies, DPn) were characterized by 1H-NMR spectroscopy [5,9], and by intrinsic viscosity measurements [36]. Number-average molecular masses were calculated from the Mark–Houwink–Sakurada equation, as reported by Anthonsen et al. [37]. Characteristic features of the chitosans are given in Table 2, including the fraction of acetylated units (FA), the diad frequencies, the intrinsic viscosities ([η]), the number-average molecular mass (Mn) and the average length of the D-blocks (ND). These characteristics show that the chitosans have a random distribution of acetylated units, i.e. according to Bernoullian distribution.

Chitinase B

ChiB was overexpressed in Escherichia coli and purified by a protocol consisting of a previously described hydrophobic interaction chromatography step [20,21], preceded by ion-exchange chromatography on Q-Sepharose Fast Flow (Amersham Pharmacia Biotech AB, Uppsala, Sweden). The final protein material was collected in an ammonium carbonate buffer [21], and freeze-dried. Before use, the enzyme was dissolved to 1 mg·mL−1 in 20 mm trizma base, pH 7.5. Protein concentrations were determined with the Bio-Rad Protein Assay (Bio-Rad Laboratories Inc., Hercules, CA, USA). The purified protein used in this work displayed the same specific activity and kinetic parameters as ChiB used in previous studies, including the protein used for successful X-ray diffraction studies [20].

Kinetics of chitosan degradation

Samples of 10 mg chitosan (FA = 0.65) were dissolved in 1.0 mL H2O and added to 1.0 mL 0.08 m NaAc buffer, pH 5.5, containing 0.2 m NaCl and 0.2 mg BSA. The samples were placed in a shaking waterbath at 37 °C. The depolymerization reaction was started by adding 5 µg of ChiB and was stopped after between 15 and 10 080 min by adjusting the pH to 2.5 with 1.0 m HCl, and immersing the samples in boiling water for 2 min. The samples were stored at −18 °C until further analysis.

Extended depolymerization of chitosans with varying degree of acetylation (FA)

Four samples of 10 mg chitosan (FA = 0.13, 0.32, 0.50 and 0.65) were each dissolved in 1.0 mL H2O and added to 1.0 mL 0.08 m NaAc buffer, pH 5.5, containing 0.2 m NaCl and 0.2 mg BSA. The samples were immersed in a shaking waterbath at 37 °C. The depolymerization reactions were started by adding 10 µg of ChiB and the reactions were stopped after 1 week as described above.


The samples were dissolved in D2O and the pD was adjusted to 4 with DCl. The deuterium resonance was used as a field-frequency lock, and the chemical shifts were referenced to internal sodium 3-(trimethylsilyl)propionate-d4 (0.00 p.p.m.). The 1H-NMR spectra were obtained at 90 °C at 300.13, 400.13 or 600.13 MHz as previously described [5,9], and FA values were calculated as described by Vårum et al. [9]. The number average degree of polymerization, DPn, of enzyme-degraded chitosans was determined from the anomer (H-1) resonances as follows: DPn = [area of all H-1-resonances (internal and reducing end)]/(area of reducing end resonances). The degree of scission, α, was calculated as α = 1/DPn.

Size-exclusion chromatography (SEC)

Oligomers from enzymatic depolymerization reactions were separated on three columns in series, packed with Superdex™ 30, from Amersham Pharmacia Biotech (overall dimensions 2.60 × 180 cm). The column was eluted with 0.15 m ammonium acetate, pH 4.5 at a flow rate of 0.8 mL·min−1. The effluent was monitored with an online refractive index (RI) detector (Shimadzu RID 6 A, Shimadzu Schweiz GmbH, Reinach, Switzerland), coupled to a datalogger. Fractions of 3.2 mL were collected for analysis. Standard samples contained 10 mg of partially depolymerized chitosan. In experiments where oligomers were collected for further analysis by NMR spectroscopy, samples containing up to 200 mg of partially depolymerized chitosan were injected without loss of resolution. Fully acetylated and fully de-N-acetylated oligomer were used as standards (monomer to hexamer; Seikagaku Corporation, Tokyo, Japan).

To verify the possibility to quantify amounts of oligosaccharides with varying DP and FA from the refractive index detector response, we checked the detector response in the following way: exact amounts (1 mg) of the fully acetylated and fully de-N-acetylated oligomers (dimer, tetramer and hexamer) were dissolved in the mobile phase and injected on the columns, and the refractive index detector signal was used to determine the area of the peaks after elution from the column. These analyses showed that, within the accuracy of the area determination, there was a linear relationship between peak areas and the amount (mass) of injected oligomer, irrespective of DP and degree of acetylation.


This work was supported by grants from the Norwegian Research Council (140497/420 and 134674/I10). We thank Hege Grindheim for carrying out some of the experiments.