Enzymatic depolymerization of chitin and chitosan involves chitinases and chitosanases, respectively (Figure 1). These enzymes hydrolyze the glycoside bonds between the sugars and are thus glycoside hydrolases (GH). Such GH are classified in the Carbohydrate-Active enZYmes database (CAZy) [45–48] that provides a continuously updated list of the GH families and, since a few years, also other families of carbohydrate-active enzymes such as glycosyl transferases and carbohydrate esterases. The CAZy classification is based on the amino acid sequence, which gives very useful information since sequence and structure, and hence mechanism, are related. It should be noted that the CAZy system does not take substrate and product activities into account a priori. The same applies to enzyme properties such as exo- versus endo-binding, processivity, and the presence of additional modules that may improve substrate-binding. All these properties are not taken into account either. The CAZy classification is purely based on amino acid sequence similarities and, indeed, several GH classes contain enzymes acting on a variety of substrates. Many GH enzymes have one or more carbohydrate binding domains in addition to their catalytic domain (Figures 1 and 2). These are referred to as carbohydrate-binding modules (CBMs) and are also classified in the CAZy database.

Chitinases occur in families GH18 and GH19 and both these classes almost exclusively contain these enzymes. Chitinases have the unique ability to hydrolyze A-A bonds and this property discriminates these enzymes from chitosanases. However, as described below, chitinases are perfectly capable of hydrolyzing chitosan, albeit to different extents. Chitinases do not hydrolyze D-D bonds.

Enzymes with chitosanase activity have been found in GH families 5, 7, 8, 46, 75 and 80. GH7 is a cellulase family and in a very few cases chitosanase activity has been detected as a side activity of these enzymes. GH5 contains a variety of enzymatic activities, including chitosanases, cellulases, licheninases, mannanase and xylanases. Again, chitosanase activity has been detected in only a very few cases, and the activity seems to be a side activity of cellulases. In GH8, enzymes annotated as chitosanases occur more frequently (next to e.g., cellulases and xylanases), and this family seems to contain a few true chitosanases [49]. The other three families, GH46, GH75 and GH80, exclusively contain chitosanases. Families GH75 and GH80 have only a few members and there is no structural and only very little functional information available for these enzymes. The best studied chitosanases, by far, are those belonging to family 46 [50–52] (Figure 1).

Based on their substrate specificity towards chitosan, chitosanases have been classified into subclasses I, II and III [53]. Chitosanases in subclass I can hydrolyze A-D and D-D linkages, subclass II enzymes can hydrolyze D-D linkages only, whereas subclass III enzymes can hydrolyze D-A and D-D linkages. Subclass I enzymes have been found in both families 46 and 75 [54,55]. In family 46, subclass III enzymes have also been found [52].

The hydrolysis of the glycosidic linkage is a nucleophilic substitution at the anomeric carbon, and can lead to either retention or inversion of the anomeric configuration [57,58]. Both hydrolysis reactions take place through general acid catalysis, and require a pair of carboxylic acids at the enzyme’s active site. One carboxylic acid is acting as a proton donor, facilitating leaving group departure, and the other acts as a base (inverting mechanism) or as a nucleophile (retaining mechanism). In both mechanisms, the position of the proton donor is within hydrogen-bonding distance of the glycosidic oxygen. The inverting mechanism (also called the single displacement mechanism) is a “one-step” reaction, where the protonation of the glycosidic oxygen occurs simultaneously with a nucleophilic attack on the anomeric carbon by an activated water molecule. This water molecule is located between a carboxylic group and the anomeric carbon and it is activated by the carboxylic group that acts as a base. Since the water molecule approaches the anomeric carbon from the side of the catalytic base, this mechanism leads to inversion of the anomeric configuration. Chitosanases belonging to families GH46, GH75 and GH80 and family 19 chitinases use the inverting mechanism [45,47,49,54,59–61].

The retaining mechanism (also referred to as the double displacement mechanism) is a two-step reaction, were the first step involves the protonation of the glycosidic oxygen (by the catalytic acid) and a congruent nucleophilic attack on the anomeric carbon atom by the nucleophile (the second carboxylic acid). This attack leads to breakage of the glycosidic linkage and the formation of a covalent linkage between the anomeric carbon and the catalytic nucleophile [62]. Subsequently, this intermediate is hydrolyzed by a water molecule that approaches the anomeric carbon from a position close to that of the original glycosidic oxygen, leading to retention of the anomeric carbon configuration.

Family 18 chitinases use a special variant of the double displacement mechanism, referred to as the substrate-assisted double displacement mechanism. Here, the carbonyl oxygen atom from the N-acetylgroup of the sugar bound in subsite -1 act as the nucleophile, leading to formation of an oxazolinium ion intermediate. Because of this involvement of the N-acetylgroup in catalysis, productive substrate-binding of chitosan and chitosan oligomers to family 18 chitinases requires that a GlcNAc is bound in the -1 subsite [63–67].

Chitinases and chitosanases can have endo- or exo-activity, where both the endo- and exo-mode of action can be combined with processivity. Processive enzymes will not release the substrate after one cleavage, but remain associated with the substrate so that a new cleavage can take place as the polymer substrate slides through the substrate-binding cleft (Figure 2). Processivity is difficult to analyze when degrading insoluble substrates such as chitin, but can be studied when using chitosan as substrate [56,64,68,69]. Whereas processivity generally is considered to be favorable for the hydrolysis of crystalline substrates, processivity has been shown to reduce enzyme efficiency towards soluble and more accessible polymeric substrates such as chitosan [69,70]. Thus, for the industrial production of CHOS, the use of non-processive enzyme variants may be beneficial in some cases.

Even though chitin and chitin synthases have not been found in humans, we produce two family 18 chitinases with chitinolytic activity [71]. In addition, two highly homologous proteins named chi-lectins and a homologous protein called oviductin are expressed. These proteins lack enzymatic activity while having retained the typical features of family 18 enzymes, including carbohydrate binding to the active site [71].

One of the human chitinases, called chitotriosidase (HCHT), is synthesized and secreted as a 50-kDa two-domain protein in human macrophages [72]. A significant portion of produced enzyme is routed to lysosomes and processed into a 39-kDa isoform, lacking the C-terminal chitin-binding domain [73]. HCHT was first discovered as a marker for Gaucher disease [72], but has later been associated with several diseases like malaria [74], fungal infections such as candidosis [75], sarcoidosis [76,77], cardiovascular risk [78], coronary artery disease [79], primary prostate cancer and benign prostatic hyperplasia [80], nonalcoholic steatohepatitis [81], multiple sclerosis [82], and Niemann-Pick disease [83]. The other chitinase, acidic mammalian chitinase (AMCase), is also synthesized as a two-domain 50 kDa protein containing a 39 kDa N-terminal catalytic domain and a C-terminal chitin-binding domain. AMCase is found in the stomach [84], in tears [85], sinus mucosa [86], and lungs [12,84]. Chitinases play important roles in the antiparasite responses of lower life forms [87–89]. Both HCHT and AMCase are believed to play similar roles in the human immune defense system, being a part of antiparasitic responses [10,87,90].

The two chi-lectins are associated with numerous diseases. YKL-40 (alternatively called HCgp39 or CHI3L1) is observed in elevated levels for patients with severe asthma [91], cardiovascular disease and diabetes [92], cancer [93], peritoneal endometriosis [94], morbid obesity [95], osteoarthritis [96], and liver fibrosis [97]. The other chi-lectin YKL-39 (or CHI3L2) has been observed up-regulated in osteoarthritic chondrocytes [98] and osteoarthritic cartilage [99]. The loss of hydrolytic activity in these chi-lectins is due to replacement of the catalytic acid (E) and the adjacent aspartic acid (D) in the conserved DxxDxDxE motif with Ala and Leu or Ser and Ile for YKL-40 and YKL-39, respectively. The chi-lectins have retained their ability to bind CHOS [100] and this may be a feature underlying some of the alleged biological effects of CHOS.

Family 18 chitinases are of particular interest, because they are abundant in nature, are crucial in the life cycles of numerous plague and pest organisms, and because they occur in humans themselves. Because of the substrate-assisted catalytic mechanism of GH18 enzymes, catalysis requires that a GlcNAc is bound to the -1 subsite. CHOS that preferably bind in such a way that a GlcN ends up in the crucial -1 subsite will act as an inhibitor. Sugar binding to the -1 subsite leads to an energetically unfavorable distortion [101] which involves the N-acetylgroup [66,67] and which may amount to an unfavorable ΔG as high as ~8 kcal/mol [102]. Although this has not yet been substantiated by experimental data, it seems plausible that binding of GlcN in the -1 subsite in fact could be energetically less unfavourable than binding of a GlcNAc. Thus, binding of a GlcN would be non-productive, but perhaps stronger than binding of a GlcNAc. This shows that the idea of developing partially deacetylated CHOS as inhibitors for family 18 chitinases is worth pursuing. Indeed, the validity of this idea has been substantiated by an early study by Peter and co-workers [103] as well as by more recent work [104,105].

For chitinase B of Serratia marcescens, the -2 subsite has a strong preference for an GlcNAc [67]. The oxygen atom in the acetamidogroup of the -2 sugar forms a bifurcated hydrogen bond with Trp ⁴⁰³ and Gln ⁴⁰⁷, whereas the methyl group packs tightly in an apolar environment provided by the side chains of Tyr ²⁹² and Ile ³³⁷. Thus, the -2 subsite of this enzyme seems optimized for strong binding of an GlcNAc, which is not surprising taking into account that the positive effects of binding sugars in subsites adjacent to the -1 subsite is needed to “pull” the -1 sugar in its distorted conformation [106]. Indeed, one has observed non-productive binding by a DADAA oligomer bound from subsites -3 to +2 in chitinase B [104].

CHOS being based on the substrate, hold a tremendous advantage in being very specific inhibitors towards chitinases, and hence not likely to interfere with other enzymatic systems. Moreover, the binding strength of the CHOS based inhibitor, an important parameter, can be tuned simply by increasing the chain length of the CHOS or by coupling additional groups to the reducing end.

In addition to its natural substrate, the glycosidic linkage of certain bacterial cell wall peptidoglycans, lysozyme may also hydrolyze chitin and chitosans [107]. In very early work, Amano and Ito [107] studied oligomers formed upon lysozyme degradation of an F_A = 0.32 chitosan, and identified the fully N-acetylated trimer and tetramer together with the partially N-acetylated oligomers AAD, DAA, AAAD, ADAA and ADAD among the oligomeric products. Later, Vårum et al. [108] studied lysozyme degradation of a highly N-acetylated chitosan (F_A = 0.65). In this study, NMR-spectroscopy of the degradation products was used to determine the identities of the newly formed reducing and non-reducing ends. This methodology, which has later been used to characterize chitinases (see below), provided insight into the cleavage specificities of the enzyme, i.e., its preference for cleaving A-A, A-D, D-A and/or D-D linkages in chitosans. Some information on the identity of the nearest neighbors to the new reducing and non-reducing ends could also be obtained.

The degradation of chitosan by the family 18 chitinases, ChiA, ChiB and ChiC, from Serratia marcescens has been studied in much detail [56,64,69,118]. Figure 3 shows the size-distribution of oligomers obtained upon degradation of a highly N-acetylated chitosan (F_A = 0.65) to various extents of degradation (α). For ChiA and ChiB, the product profiles obtained during the initial phase of the degradation show a dominance of even-numbered oligomers, which is indicative of processive action [56,64,68]. This product pattern is due to the fact that enzyme-ligand complexes where there is a GlcN bound to the -1 subsite is not productive (in the case of family 18 chitinases). If the enzyme is processive, the enzyme will slide by two sugar units at the time, until a productive complex is formed [the primary condition being that there is an GlcNAc bound in the -1 subsite (see Sørbotten et al. [64] and Eijsink et al. [68] for a more detailed discussion)]. Consequently, while the first product of an enzyme-substrate association may have any length, every subsequent product will be even-numbered. In the case of chitin, all these products would be dimers; in the case of chitosan, these products may be longer even-numbered oligomers. Later during the reaction, the dominance of even-numbered oligomers disappears because there are no longer substrate molecules left and the enzyme is primarily involved in rebinding and further cleavage of oligomers from the preceding processive phase (see Figure 3). For example, an oligomer such as ADADAA that could emerge during processive degradation by ChiB can be converted by this same enzyme to ADA and DAA upon rebinding in a mode that was not explored during the processive phase, where the substrate moves by two sugars at the time (see Horn et al. [56] and Eijsink et al. [68] for a more extensive discussion and explanation).

Figure 3 shows a totally different product pattern for ChiC that is characteristic for an endo-acting, non-processive enzyme. ChiC converts chitosan to a continuum of oligomers of different sizes and the polymer peak disappears early in the degradation reaction. Also, there is initially no accumulation of dimers or other even numbered oligomers. This all indicates that ChiC attacks the polymeric substrate chains in random positions, without processivity. Indeed, this “endo” activity could be confirmed by viscosity measurements during the hydrolysis reaction [118]. The contrast with ChiA and ChiB is perhaps best illustrated by the void peak, which disappears much more slowly in the case of processive enzymes. These latter enzymes perform many cuts per chain instead of a few cuts in every chain as in the case of ChiC.

Analysis of the sequences of the products (Table 2) showed considerable differences between the enzymes as well as differences over the time course of the degradation reactions. In all three enzymes, productive binding requires a GlcNAc in subsite -1, explaining why all products have a GlcNAc at their reducing ends. The enzymes did not have any detectable preferences for GlcNAc versus GlcN in the +1 subsite, and this is reflected in the oligomers having both N-acetylated and deacetylated non-reducing ends. All three chitinases showed a strong, but not absolute, preference for GlcNAc in subsite -2, meaning that oligomers preferentially have an N-acetylated unit next to the reducing end. The kinetics of the reaction with ChiB illustrate this (Figure 4; Table 2): in the initial rapid phase of the reaction almost all oligomeric producs have −AA at their reducing ends; during the second, much slower phase, oligomeric products ending at −DA appear to a larger extent [56, 64]. The preference for a GlcNAc in subsite -2 was strongest and in fact almost absolute for ChiC; all oligomeric products end with −AA, at any point during the reaction (Table 2).

Figure 5 shows how the outcome in terms of the length distribution of products can be manipulated by varying the F_A of the chitosan. Obviously, since we are working with chitinases with clear preferences for GlcNAcs at certain positions in the substrate, the products get longer as the F_A goes down. It is quite remarkable that a chitinase such as ChiB works well on chitosans with F_A close to only 10%.

Sikorski et al. [114,118] have produced a model for the degradation of different chitosans with ChiB, which is capable of accurately predicting the outcome of hydrolysis reactions in terms of the length distributions of the products at varying α. This model can be used to predict how reactions need to be set up in order to maximize the production of CHOS of certain lengths (Figure 6).

ChiG, a bacterial family 19 chitinase from Streptomyces coelicolor A3(2), produces quite different CHOS as compared to the family 18 chitinases, reflecting the very different binding preferences in the active sites of the enzymes [115]. Since ChiG, which operates according to a non-processive endo-mode of action (Figure 7), uses the inverting mechanism, there is no absolute requirement for GlcNAc in subsite -1. This means that the reducing ends of the oligomeric products could be both N-acetylated and deacetylated, as was indeed observed. The non-reducing ends of the oligomers were found to be exclusively N-acetylated, and the sugar units in the neighboring position to reducing ends were also found to be exclusively N-acetylated. Thus, ChiG has an absolute preference for a GlcNAc in subsites -2 and +1. The enzyme also has considerable preference for GlcNAc in -1 and kinetics were clearly biphasic: In the first fast phase cleavage occurred in A-A^↓A sequences, whereas A-D^↓A sequences were cleaved in the slower second phase [115]. So, while the chromatograms of Figure 7 may look somewhat similar to the chromatogram for the family 18 enzyme ChiC in Figure 3, the sequences of the produced oligomers show considerable differences, which may affect bioactivity. More generally, it is clear that ChiG will yield oligomers of different P_A as compared to those obtained by ChiA, ChiB and ChiC. For example, degrading a chitosan with F_A = 0.65 with ChiB and ChiG gives the trimers DAA and AAD, respectively [64,115].

Figure 8 shows that varying the F_A of the chitosan had a major effect on the size distribution of the products and the extent of cleavage. Interestingly, ChiG efficiency is much more sensitive for deacetylation than the efficiency of ChiB (compare Figure 8 with Figure 5).

Other studies on the degradation of chitosan with family 19 chitinases confirm the findings for ChiG. In one study, the family 19 chitinase ChiC from Streptomyces griseus HUT 6037 was used to (extensively) degrade a chitosan with a degree of N-acetylation of 47% and the products were isolated and characterized [116]. The identities of the isolated CHOS were AD, AAD, ADAD, ADDAA and AADDAA, and it was concluded that this enzyme has an absolute specificity for N-acetylated units in the -2 and +1 subsites, in agreement with the results for ChiG from Streptomyces coelicolor. A family 19 chitinase from rice [117] has been found to operate according to a non-processive endo-mode of action, with strong preferences for N-acetylated units in subsites -2 and +1, and with less strong preference for N-acetylated units in the -1 subsite.

The GH 46 family of chitosanases comprises enzymes classified as subclass I (cleaving A-D and D-D linkages) [60] as well as subclass III (cleaving D-A and D-D linkages) [120]. In an early study, a chitosan with F_A = 0.25–0.35 was extensively degraded with CsnN174 from Streptomyces sp. N174 and the oligomeric products were isolated and characterized. In addition to the fully deacetylated oligomers D, DD and DDD, several hetero-oligomers were identified (the dimer DA, the trimers DDA and DAA, the tetramers DDAA and DAAA and the pentamer DDAAA) [60]. These results suggest that CsnN174 has a high specificity for a GlcN in the +1 subsite (new non-reducing ends), whereas both GlcNAc and GlcN seem to be acceptable in the -1 subsite (new reducing ends). Further insight into the properties of CsnN174 would require studies of oligomer production over time, as well as studies using chitosans of varying F_A.

Recent work in our own laboratories on a family 46 chitosanase, Csn88, from Streptomyces coelicolor has shown that this enzyme is capable of degrading chitosans with varying F_A, producing oligomers with both N-acetylated and deacetylated reducing ends [121]. ¹H NMR spectroscopy analysis showed that when the new reducing ends were N-acetylated, the sugar binding in the neighbouring position (i.e., binding in subsite -2) always was deacetylated. However, in oligomers with deacetylated sugars at the new reducing end, both N-acetylated and deacetylated sugar occurred in the neighbouring position (revealed by mass spectrometry). The identity of the new non-reducing ends was studied using ¹³C NMR spectroscopy and this analysis showed that initial products exclusively had deacetylated non-reducing ends, whereas N-acetylated non-reducing ends appeared later during the reaction. All in all, these preliminary data indicate that Csn88 can cleave D-A, A-D and D-D linkages. As expected maximum α values were high, for example 0.59 for a highly deacetylated chitosan (F_A 0.008) and 0.44 for a F_A 0.32 chitosan.

Several authors have employed unspecific enzymes such as papaine and cellulases to degrade chitosans (e.g., [122–124]). Since the enzyme preparations used tend to be rather crude and derived from sources (fungi, plants) that are known to produce chitinolytic enzymes, there remains some doubt concerning which enzymes actually catalyze the hydrolysis reactions. However, for the practical purpose of producing CHOS, the use of (cheap) unspecific enzymes may be quite useful.

Of chemical methods for hydrolysis of chitosan [32–35], acid hydrolysis is probably the best known. Early studies on acid hydrolysis of chitosans had shown that it is possible to convert fully deacetylated chitosan to CHOS in concentrated hydrochloric acid [32]. In later studies [33], using a variety of chitosans, the acid-catalyzed degradation rates of chitosans were shown to depend on F_A, and the initial degradation rate constant was found to increase in direct proportion to F_A. Acid hydrolysis was found to be highly specific to cleavage of A-A and A-D glycosidic linkages, which were hydrolyzed with two to three orders of magnitude higher rates than the D-D and D-A linkages. This preference is probably due to a combination of two factors: (1) the presence of a positively charged amino group (as in GlcN) close to the glycosidic linkage is inhibitory, and (2) the presence of an acetamidogroup (as in GlcNAc) close to the glycosidic linkage may yield some degree of substrateassistance to the hydrolytic mechanism. In the same study it was shown that the rate of deacetylation was less than one-tenth of the rate of depolymerization in concentrated acid, whereas the two rates were found to be equal in dilute acid. It was suggested that this is due to these two processes having different reaction mechanisms with different rate-limiting steps [33].

Chemical synthesis of CHOS requires multiple protection and deprotection steps, and is today not a routine procedure. Chemical synthesis of CHOS gives rise to pure compounds, but most methods existing today are time consuming and require extensive use of organic solvents and/or are not capable of producing anything else than homo-oligomers. There are in fact only a few examples of chemically synthesized CHOS in the literature. Kuyama et al. [125] performed synthesis of fully deacetylated chitosan dodecamers starting with glucosamine monomers using phthalimido as the amino protective group. Aly et al. [126] reported a method for synthesis of fully N-acetylated CHOS from GlcN monomers using dimethylmaleoyl as an amino protective group for synthesis of chitotetraose and chitohexaose. Removal of the amino protective group and N-acetylation was performed in a one-pot reaction to give the desired products [127]. In principle it would be possible to combine the use of these two described protection methods to synthesize partly deacetylated CHOS, but this has so far not been reported (to our knowledge).

Trombotto et al. [128] have reported a method for chemical preparation of partly deacetylated CHOS from fully deacetylated high molecular weight chitosan. The starting chitosan was partially depolymerized using HCl to produce fully deacetylated oligomers that were fractionated by selective precipitation and ultrafiltration to yield a mixture of DP 2–DP 12. The oligomers were then partly N-acetylated using stochiometric amounts of acetic anhydride to reach the decided F_A. In this way, CHOS fractions of DP between 2 and 12 were successfully prepared. The drawback of this method, as for the enzymatic preparation of CHOS, is the heterogeneity of the prepared CHOS.

In an early study, Letzel et al. [103] used a similar approach: chitosan with F_A 0.02 was hydrolyzed with HCl, oligomers were separated by DP using gel permeation chromatography and the resulting CHOS fractions were N-acetylated using substoichiometric amounts of acetic anhydride to control the F_A. Interestingly, some of the fractions produced in this study inhibited the family 18 chitinase ChiB from Serratia marcescens.

In principle, chemoenzymatic synthesis provides opportunities to produce pure CHOS of defined DP, F_A and P_A without the use of extensive protection of the functional groups at the sugar unit. So far this has been done by allowing an oxazoline, imitating the intermediate of chitin hydrolysis, to act as a glycosyl donor in an enzyme-catalyzed reaction where another GlcN/GlcNAc unit acts as a glycosyl acceptor [129]. By using oxazoline oligomeric building blocks of specific DP, F_A, and P_A, longer specific CHOS can be made using this approach. The main disadvantage using the chemo-enzymatic approach is that the product is necessarily also a substrate for the enzyme, which can result in poor yields. To avoid this problem, modified enzymes with reduced hydrolytic activity may be employed. The enzyme modifications would need to be aimed at reducing hydrolytic power, while increasing binding strength for the glycosyl donor in the glycon subsites [130].

It has been known since the 1970s that CHOS have anti-tumor effects [18], and there is also evidence for positive effects of CHOS in reducing metastasis from tumors [19,20]. It was first suggested that the anti-tumor activity was due to the cationic properties of CHOS [18]. Later, M_W also was proposed to play a major role [136]. Maeda and Kimura [137] found that CHOS enhanced the natural killer activity in intestinal intraepitelial lymphocytes at the same time as they reduced tumor growth in mice, and suggested that this CHOS-activation of intestinal immune functions could be useful in treating tumors.

There are indications that apoptosis is involved in the tumor-reducing effects of CHOS. Harish Prashanth and Tharanathan [138] discovered that DNA from Ehrlich ascites tumor cells was fragmented after incubation with CHOS, an indication of apoptosis. CHOS have also been shown to induce apoptosis of human hepatocellular carcinoma cell via upregulation of the pro-apoptotic protein Bax [139].

In recent years, the hypothesis that the anti-tumor effects of CHOS are related to their inhibitory effects on angiogenesis has received some attention [138,140,141]. Angiogenesis is the formation of new capillary blood vessels from already existing blood vessels. This process is important for tumor formation, since tumor growth and metastasis require angiogenesis when the tumor reaches a certain size. Xiong et al. [142] compared effects of dimers to hexamers of fully deacetylated CHOS on angiogenesis and found that the hexamers were the most effective inhibitors, whereas Wang et al. [140] showed that N-acetylated CHOS were more effective in preventing angiogenesis than fully deacetylated CHOS, both in vitro and in vivo.

AMCase is induced during T_H2 inflammation through an interleukin (IL)-13 dependent mechanism and has been demonstrated to be heavily over-expressed in human asthmatic tissue [10,12]. Inhibition of the AMCase with the well known chitinase inhibitor allosamidin reduced the inflammation [12]. The fact that chitinases are a factor in host antiparasite responses and in asthmatic T_H2 inflammation led to the hypothesis that asthma may be a parasite-independent antiparasite response [10], which again suggests that inhibition of AMCase is a potential target for asthma therapy [9–12]. It has been shown that partially deacetylated CHOS can function as inhibitors of family 18 chitinases [103–105]. There is therefore a great potential for CHOS as an anti-inflammatory drug in patients with asthma. For a more detailed description of this and related topics, see the review by Muzzarelli in this special issue of Marine Drugs [143].

Mesenchymal stem cells from the bone marrow are able to differentiate into chondrocytes (cartilage), adipocytes (fat) and osteoblasts (bone). Osteoblasts produce osteoid; which is further mineralized to produce the bone matrix. Bone-tissue is mainly composed of bone matrix and osteoblasts. Chitosan and CHOS are known to increase the differentiation of mesenchymal stem cells to osteoblasts and to consequently facilitate the formation of bone-tissue [21,22].

The mineralization process and bone strength are dependent on Ca²⁺, which helps to support the structure. There is evidence that CHOS increase calcium-deposition in bone [22,144,145]. Jung et al. [144] found that CHOS could efficiently inhibit the formation of insoluble calcium-phosphate salts and consequently increase Ca²⁺ bioavailability and bone strength. They also found that CHOS (<5 kDa) gave increased calcium retention and decreased bone turnover in a rat osteoporosis model. This indicates that CHOS may have beneficial effects as a calcium fortifier in conditions of Ca²⁺ deficiency, such as in osteoporosis.

Chitosans have been successfully used as vectors for delivery of genes (gene therapy) since the first report about 15 years ago [146,147]. Chitosan forms stable complexes with plasmid DNA and can be used as a vector for the administration of genes to mucosal tissues such as the lungs [148] and intestinal epithelium [149,150]. There are, however, certain drawbacks connected to the use of high molecular weight chitosans because of the low solubility at physiological pH, the high viscosity and the fact that the chitosan complexes often tend to form aggregates. By using CHOS instead of chitosan these drawbacks may be overcome [16]. Köping-Höggård et al. [16] showed that fully deacetylated CHOS (DP 24) formed stable complexes with plasmid DNA, and in vitro and in vivo experiments proved that these CHOS were effective vectors for delivery of genes [16,17]. It has been speculated that a delicate balance between the stability of the CHOS-DNA-complexes at lower pH-values (around pH 6) and their instability at higher pH-values (above pH 7) could be the reason for their efficiency [151]. This has recently been confirmed in detailed studies of how chitosan chemistry can be used to create an optimal balance between the stability of the complexes and their unpacking [152].

Some pathogens can initiate disease in humans by using carbohydrate binding proteins (lectins) to attach to complementary membrane-bound oligosaccharides on host cells [153,154]. Observed antibacterial and anti-infective effects of CHOS [13] are thought to be due to their ability to bind to the lectins on human pathogens and, consequently, prevent adhesion to human cells. A priori, one would expect the sequence of GlcNAc and GlcN units in CHOS to be important for binding affinity, and for ensuring selectivity for pathogens (i.e., the CHOS should preferably not bind to lectins of the normal bacterial flora). In a recent study, it was shown that a mixture of 97% deacetylated tetramers significantly inhibited adhesion of certain enteropathogenic Escherichia coli strains to human colon adenocarcinoma epithelial (HT29) cells in tissue culture, whereas the binding of other E. coli strains was not inhibited [13]. Since pathogens vary in terms of their lectins, identification of both the target-lectins and the possible complementary CHOS are important when pursuing this application route of CHOS.

Malaria is caused by several species of the parasite Plasmodium. P. falciparum causes the most serious forms of malaria in humans, whereas P. vivax, P. ovale and P. malarie give a milder disease that is not generally fatal [155]. Each year, 350–500 million cases of malaria occur worldwide, and over one million people die, most of them young children in sub-Saharan Africa. There is no vaccine available, and the identification of molecular targets for vaccine development is of great importance. Anopheles mosquitoes transmit the malaria-parasite from one infected person to another and the most important control strategy for malaria today is to interfere with different stages in the life cycle of the Plasmodium parasite.

During its life cycle, the Plasmodium parasite must be capable of penetrating the chitin-containing periotrophic matrix surrounding the mosquito midgut, to make the mosquito infective. To do so, Plasmodium species secrete family 18 chitinases capable of degrading the periotrophic matrix [23,156–160]. The P. falciparum chitinase (PfCHT1), the P. vivax chitinase (PvCHT1) and the P. gallinaceum chitinase (PgCHT1) have been characterized [161–163]. P. gallinaceum is the only malaria parasite species where more than one chitinase gene has been identified, and PgCHT1 and PgCHT2 are both family 18 chitinases. One approach to vector control might be inhibition of the secreted Plasmodium chitinases by chitinase inhibitors that are taken up by the mosquito via the blood meal [23,156–160]. Several studies have convincingly shown that inhibition of Plasmodium chitinases indeed reduces infectivity [23,163,164]. CHOS may perhaps be developed as nontoxic, inexpensive small-molecule inhibitors of these chitinases.

The use of chitosan in wound dressings has been explored to a certain extent and positive effects have been documented in several studies [14,165–169]. Similar positive effects have been documented for CHOS, which were shown to accelerate the wound healing process [14,15]. It is quite likely that chitosan is converted to CHOS by naturally occurring enzymes and that the activity observed for chitosan might in fact be caused by CHOS. It may thus be advantageous to use CHOS in wound dressings to get a more immediate effect.

CHOS are thought to accelerate wound healing by enhancing the functions of inflammatory and repairing cells [170–172]. For example, it has been shown that subcutaneous injection of hexamers of CHOS enhanced migration of polymorphonuclear cells in dogs [171]. Hexamers of N-acetylated and fully deacetylated CHOS were shown to induce persistent release of IL-8, a potent activator and chemoattractant of polymorphonuclear cells, from fibroblasts from rats in vitro [170]. It must be noted that most authors ascribe the beneficial effect of longer CHOS and (polymeric) chitosan on wound healing to the ability of these compounds to form biocompatible ordered tissue-like structures (see [167] and references therein).

Hemostatic effects may also contribute to the beneficial effects of chitosan/CHOS in wound dressings. Chitosan enhances platelet adhesion and aggregation [165,173] and increases the release of the platelet derived growth factor-AB (PDGF-AB) and the transforming growth factor-β1 (TGF-β1) from platelets in canine blood [173]. These two factors retract inflammatory cells which are thought to be important in wound healing. Chitosan also has the ability of causing erythrocytes to aggregate [169].

Minagawa et al. [166] compared wound break strength and the increase in collagenase activity in wounds in rats after exposure to monomers, oligomers and polymers of the chitin-group (GlcNAc/N-acetylated CHOS/chitin) and the chitosan-group (GlcN/fully deacetylated CHOS/chitosan). They found that all six compounds increased both the wound break strength and the collagenase activity. Overall, the non-acetylated compounds were most effective compared to the corresponding N-acetylated compounds. The oligomers of fully deacetylated CHOS were most effective for wound break strength and GlcN gave the highest activity of collagenase. The enzyme collagenase is produced mainly by fibroblasts and inflammatory cells and its activity is related to remodeling in wound healing [174].

The antifungal activity of chitosan was discovered already in 1979 [175], and has been utilized to inhibit fungal growth in crops [176]. The antifungal potential of CHOS has to a lesser extent been investigated. In 1984 Kendra and Hadwiger [177] tested the antifungal activity of monomers– heptamers of deacetylated CHOS on Fusarium solani, which is infectious to pea crops, and found that the heptamer was most effective.

Subsequent research has revealed that longer oligomers of CHOS (also called low molecular weight chitosan, LMWC) are more effective. LMWC (4.6 kDa, average DP of 23) shows antifungal activity against Candida krusei and inhibits spore germination in Fusarium oxysporum [178]. Seyfarth et al. [26] found antifungal effects of LMWC on different Candida species. Both DP and F_A of the chitosan/CHOS are of great importance for the antifungal potential and LMWC with low F_A so far seems to be the most promising type of compound [179].

The anti-fungal effect of LMWC seems to be caused by its interaction with lipids in the plasma membrane, leading to morphological changes and cell surface disruptions [180,181]. The composition of the fungal plasma-membrane seems to be important for the sensitivity against chitosan, and a higher content of polyunsaturated fatty acids makes the fungi more sensitive [182].

From literature studies, as well as from own unpublished work on non-medical use of LMWC as anti-fungals, it is clear that LMWC indeed have a considerable potential in this area. This is a good reason for giving the application of LMWC, to combat fungal infections in humans, more research attention than it has received so far.