The marine ecosystem

Get Complete Project Material File(s) Now! »

Enzymatic transformation of HMW compounds

Proteins and DNA are considered as easily degradable due to their limited number of building blocks (two pyrimidines and two purines in RNA and DNA) (usually 23 L-amino acids in proteins) and the uniform kind of linkages. For example RNA rapidly degrades on the bench, due to the ubiquity of RNAses. In contrast, in polysaccharides and other glycan structures post translational modifications on glycoproteins challenge bacterial degradation systems due to their high diversity and complex structure of assemblage. For instance, for a theoretical hexasaccharide sequence formed by different sugar units, more than 1.05×1012 possible oligosaccharide structures can be calculated. In contrast, a set of six amino acids generates only 46 656 different structures, which is 7 orders of magnitude lower (Laine 1994). This enormous theoretical and observed carbohydrate diversity resides from seven major structural features.
These are i) epimers of the D and L configuration, ii) linear primary structure or branching, iii) ring size, iv) anomeric configuration, v) linkage position, vi) branching positions and vii) reducing terminal attachment, which combine all into various structures with identical mass. This huge chemical variation found in glycans is also the reason for the large abundance of glycoside hydrolases displaying different substrate specificities, and that are classified into families based on their sequence similarity (httpd://www.cazy.org) (Cantarel et al. 2008).

Glycoside hydrolases and their sequence based classification

Glycoside hydrolases (GH) are the enzymes responsible for degradation of carbohydrates to produce mono- and oligosaccharides from substrates with higher degree of polymerization. Glycoside hydrolases are ubiquitous throughout all kingdoms of life where they catalyse the cleavage and formation of glycosidic bonds, reactions which are important in a myriad of biological processes.
The high diversity of their substrates is reflected by the number and diversity of carbohydrate acting enzymes or CAZymes, which are found with highly divergent sequences, different folds, and catalytic mechanisms. However, since the carbohydrate diversity exceeds by far the number of protein folds, CAZymes have evolved from a limited number of ancestors by acquiring novel specificities at substrate and product level (Cantarel et al. 2008).
Carbohydrate active enzymes have been classified into families based on their amino-acid sequence similarity. The carbohydrate active database (CAZy database) merges the present knowledge on 115 glycoside hydrolase, 91 glycosyltransferase, 19 polysaccharide lyase and 52 carbohydrate-binding module families (http://www.cazy.org/).

The catalytic mechanism of glycoside hydrolases

The glycosidic bond is hydrolysed by two critical residues: the acid/base and the base/nucleophile residue that are generally aspartates or glutamates (Davies et al. 1995). The hydrolytic attack occurs via two possible mechanisms which result either in retention or inversion of the configuration of the anomeric carbon (Koshland 1953; Sinnott 1990; Vocadlo et al. 2008). The distance between the two involved catalytic residues differs between these two mechanisms, although exceptions do exist. If the catalytic mechanism involves retention of the anomeric carbon the distance is around 5.5 Å, whereas this distance is generally bigger in enzymes (6.5Å-9Å) of inverting mechanism, since here the activated water molecule requires further space between the nucleophile residue and the anomeric carbon. Inverting glycoside hydrolases proceed via a direct displacement mechanism, in which an oxocarbenium-ion like transition state is formed. The two carboxylic active site residues are placed at an appropriate distance to provide the base catalytic assistance activating a water molecule (nucleophile), while the second catalytic residue (proton donor) provides acid assistance by protonating the glycosidic bond oxygen at the point of cleavage. The catalysis in retaining glycoside hydrolases proceeds via a double-displacement mechanism, in which a covalent glycosyl-enzyme intermediate is formed and subsequently hydrolysed by an oxocarbenium ion-like transition state (Withers 2001) (Figure 12).

Mode of action in glycoside hydrolases

Besides their catalytic mechanism, which separates retaining from inverting enzymes, glycoside hydrolases can be additionally classified by their mode of action. Three modes of action are generally defined by the outcome of the biochemical analysis. Keeping in mind that these definitions are not completely strict in nature, one can define exo-, endo- or a processive mode of action. Exo enzymes specifically attack chain ends of the substrate and then dissociate from their substrate, while endo enzymes cleave anywhere along the chain and dissociate after the reaction. Processive enzymes usually divided into endo- and exo- processive; catalyze several successive rounds of hydrolysis before dissociating from the substrate. The precise characterization of the mode of action of an enzyme is not straightforward, but generally the protein structure analysis can complement the biochemical data. An enzyme with a pocket topology points towards an exo-activity, the presence of a catalytic groove can randomly bind on a polymeric substrate and is indicative of enzymes with an endo-mode of action, but is equally found in processive enzymes. Finally, in the tunnel or tyroid topology that has been described for enzymes with a processive mode of action, large loops or domains close down on the polymer chain that is funnelled through the enzyme, which most probably decreases the dissociation rate.
Within family GH16 examples for all these different topologies have been described. The recent crystal structure of a β-galactosidase from Clostridium perfringens (Tempel et al. 2005) revealed the pocket topology, while both β-agarases display the open-groove catalytic site and the κ-carrageenase CgkA possesses the tunnel topology, which is reminiscent of processive hydrolases (Divne et al. 1994; Divne et al. 1998) (Figure 13).
Figure 13: These three GH16 enzymes present the different active cleft topologies pocket, open cleft and the tyroid. (a) The β-galactosidase with the pocket topology. (b) The β-agarase A with the open cleft. (c) The κ-carrageenase with the tunnel topology. The arrows indicate the substrate binding with the tip pointing towards the reducing end of the substrate.

Subsites in glycoside hydrolases

Within the substrate binding cleft of an enzyme, positive (+) and negative (-) substrate binding subsites can be defined, with respect to the catalytic residues, at the point of cleavage. This nomenclature has been adopted for all glycoside hydrolases, whereby definition, –n subsites represent enzyme/sugar ring interactions with the glycon (new reducing end) and +n subsites represent the interactions with the aglycon (new non-reducing end) (Davies et al. 1997). By definition the point of glycosidic bond cleavage is situated between the -1 and +1 binding sites (Figure 14).
Figure 14: Subsite nomenclature in glycoside hydrolases. Here shown for the -agarases AgaA and AgaB from Z. galactanivorans. Red circles show the L-galactose and white circles the D-galactose rings found in agarose. NR stands for the non reducing end and R for the reducing end of the substrate oligosaccharide.

Agarolytic bacteria and glycoside hydrolases

The first agar-degrading microorganisms were isolated from a Norwegian fiord by Gran in 1902. Since then at least 30 microorganisms with agarolytic activity have been reported. The vast majority of these bacteria are of marine origin belonging to Cytophaga (Duckworth et al. 1968; Duckworth et al. 1969; Duckworth et al. 1969; Van der Meulen et al. 1975), Microbulbifer (Ohta et al. 2004), Pseudomonas (Ha et al. 1997; Kang et al. 2003), Pseudoalteromonas (Belas 1989; Vera et al. 1998; Ivanova et al. 2003; Schroeder et al. 2003), Microscilla (Zhong et al. 2001), Vibrio (Aoki et al. 1990; Sugano et al. 1993; Sugano et al. 1994; Sugano et al. 1994; Araki et al. 1998), Alterococcus (Shieh et al. 1998), Alteromonas (Potin et al. 1993), Thalassomonas (Ohta et al. 2005) , Saccharophagus (Ekborg et al. 2005; Ekborg et al. 2006) and Zobellia (Allouch et al. 2003; Jam et al. 2005). Agarase activity has also been observed in terrestrial organisms, such as Paenibacillus (Hosoda et al. 2003) and Streptomyces (Bibb et al. 1987); genes of agarases have been found in soil by metagenomics (Voget et al. 2003), and interestingly agarase activity was found in an unidentified hospital contaminant (Swartz et al. 1959). Currently there is only one report of an agarase purified from an eukaryote, the mussel Littorina mandshurica (Usov et al. 1975). Because agarolytic bacteria have been isolated from various far eastern mussels, it is likely that the agarase activity reported for L. mandshurica results from an associated bacterial symbiont.
Agars serve as carbon source for these diverse microorganisms, which degrade this class of polysaccharides with glycoside hydrolases specific for the alternating α-1,3 und β-1,4-glycosidic linkages. Based on their specificity, agarases can be classified into two major categories, β- and α-agarases. β-Agarases cleave the β-1,4 linkages to produce oligosaccharides of the neoagarobiose series with D-galactose at the reducing end. The α-agarases cleave the α-1,3 glycosidic linkages and produce oligosaccharides of the agarobiose series with 3,6-anhydro-L-galactose at the reducing end.
One of the most intensively characterized agarolytic systems originates from Pseudoalteromonas atlantica (Day et al. 1975; Groleau et al. 1977). This system consists of an extracellular endo-β-agarase I, which depolymerises agarose to neoagarotetraose. The end-products are subsequently processed by two periplasmic enzymes, the β-agarase II and a neoagarobiose hydrolase, finally yielding 3,6-anhydro-L-galactose and D-galactose (Morrice et al. 1983; Morrice et al. 1983). A similar mode of agar degradation was reported for Pseudomonas elongata (Vattuone et al. 1975) and Flavobacterium flevense (Van der Meulen et al. 1975). In contrast, Vibrio sp. JT0107 secretes two β-agarases that depolymerise agarose to neoagarobiose and neoagarotetraose, respectively which are subsequently degraded by a periplasmic α-L-galactosidase. This enzyme cleaves the α-1,3-linkages of the neoagaro-oligosaccharides from the nonreducing ends (Sugano et al. 1993; Sugano et al. 1994). Recently the agarolytic system from Saccharophagus degradans (Ekborg et al. 2005) has been characterized. This heterotrophic proteobacterium was isolated from decomposing cord grass and was the subject of a genome sequencing project, due to its biotechnological potential. Functional studies have shown that this bacterium degrades a wide array of complex polysaccharides including those originating from algal, fungal and plant sources (Weiner et al. 2008). The agarolytic system present in this microbe has been extensively studied on a genomic and proteomic level (Ekborg et al. 2006), and has been found to be composed of five agarases belonging to three different glycoside hydrolase families, namely GH16, GH50 and GH86.

Family classification of agarases

The sequence based glycoside hydrolase classification has the advantage of relating the structural fold to the function of the enzymes. Consequently, divergent evolution leads to polyspecific enzyme families. On the other hand, this classification will separate enzymes that display the same substrate specificity but that have convergently evolved on different protein folds. As an example, cellulases can be found in 12 different families. In the same manner, the actually known β-agarases are classified into three glycoside hydrolase families, GH16, GH50 and GH86 (http://www.cazy.org) and various β-agarases have been biochemically characterized (for review (Michel et al. 2006).
Interestingly, although active on the same substrate polysaccharide, the three α-agarases known to date fall into their own family, GH96, and are much less abundant, (Flament et al. 2007). Indeed, the environmental abundance of ß-agarases appears much higher when inferred from the available number of -agarase like sequences in the actual databases (NCBI, protein sequence database: 233 β-agarase sequences compared to only three α-agarases). Traditionally, agarases are screened on agar plates and the positive agar lyses from environmental, from expression clone libraries with E. coli as host, or from purified enzymes can be easily detected by the depression created in the gel by enzymatic activity (Figure 15). This method equally detects α-agarase or β-agarase activity and is thus unbiased, compared to the reducing sugar assay that gives negative results with α-agarases. Since the number of reported β-agarases outnumbers the α-agarases, for which the screening method is most probably not the reason, one can assume that α-agarases are less abundant in the marine environment.
Figure 15: Agarolytic activity demonstrated on an agar plate. (a) Alteromonas agaralytica streaked out on an agar plate strongly degrades the agar around the colonies. (b) The recombinantly expressed and purified α-agarase from the same bacterium has the same effect and leads to a hole in the substratum. The arrows indicate the degradation of agar by the enzymatic activity.
The β-agarase family GH16 contains the most agarases in the databases (Tristan Barbeyron personal communication). β-Agarases, and the here presented β-porphyranases provide an interesting model system to investigate the molecular bases of the heterogeneous gel forming polysaccharide degradation by marine heterotrophic microbes.
As mentioned above (The cell wall matrix of marine red algae: Agars and carrageenans) certain components of agar, such as porphyran, may consist mostly of sulfated sugar units. Yet it is unclear whether this type of agarocolloid is degraded by β-agarases displaying very broad substrate specificity, or if specific enzymes have evolved to degrade these agar-variant structures. Until the start of my thesis, no porphyran specific activity had been reported, although various β-agarases from marine bacteria have been used to analyze the primary structure of red algal galactans especially of porphyran from Porphyra spp. (Morrice at al. 1983).

The glycoside hydrolase family GH16

As mentioned previously, the family classification of glycoside hydrolases is sequence based. A major consequence of this is that the catalytic mechanism is in general the same throughout a GH-family of enzymes (Davies et al. 1995; Henrissat et al. 1997). However, recent exceptions have been reported (Gloster et al. 2008). Another consequence is that while some families contain enzymes displaying one and unique specificity, as family GH6 for cellulases or GH11 for xylanases, others including GH1, GH5 or GH13 covers a wide range of substrate specificities. The glycoside hydrolase family 16 (GH16) is such a polyspecific family with eight described specificities and includes more than 900 sequences to date. Numerous structures have been described, two of which are active on red algal galactans, namely the β-agarases and a κ-carrageenase. All GH16 enzymes contain the same catalytic residues and share the retaining reaction mechanism with retention of the configuration at the anomeric carbon (Keitel et al. 1993). The eight enzyme activities that have been described so far cover lichenases, xyloglucan endotransferases, keratan-sulfate endo-1,4-beta-galactosidases, glucan endo-1,3-beta-D-glucosidases, endo-1,3(4)-beta-glucanases, xyloglucanases, β-agarases and κ-carrageenases to which we can add the new activity of β-porphyranases described in my thesis work (Table 2).
Both β-agarases, AgaA and AgaB from Z. galactanivorans as well as the κ-carrageenase CgkA from P. carrageenovora share the jellyroll fold, formed of two β-sheets with of seven β-strands each. These two β-sheets are stacked in a sandwich like manner and this sandwich is twisted around the potential carbohydrate substrate to create an extended binding cleft. This cleft is surrounded by strand connecting surface loops which influence specificity and the mode of action of the enzyme (see below). The catalytic event in retaining enzymes takes place via a double-displacement mechanism that involves two strictly conserved carboxylic residues (Koshland 1953; Sinnott 1990). These catalytic residues, located in the centre of the substrate binding cleft, are characterized by the conserved sequence pattern ExDx(x)E of active GH16.
The first glutamate of this pattern could be identified as the nucleophile and the last glutamate as the acid/base in the 1,3-1,4-β-glucanase from B. macerans (Hahn et al. 1995). The central aspartate residue, which is also strictly conserved, seems to maintain the nucleophile glutamate in the negatively charged state (Kleywegt et al. 1997). Even though the catalytic machinery of different enzyme structures within family GH16 is conserved, local residue substitutions modulate substrate specificity at the active site.

READ  DEVELOPMENT OF SYNTHETIC BIOLOGICAL TOOLS -SYNTHETIC INDUCIBLE PROMOTER

The β-agarases from Z. galactanivorans

The first GH16 enzyme purified from Z. galactanivorans was a κ-carrageenase which was expressed by the bacterium in presence of carrageenan as substrate. This enzyme could be purified to homogeneity from the culture supernatant and was subsequently cloned and characterized (Potin et al. 1991; Barbeyron et al. 1998). When Z. galactanivorans is cultivated in the presence of agar the culture supernatant contains β-agarase activity specific for the β-1,4 glycosidic linkages in the polysaccharide. At least two agarase genes coding for different β-agarases were identified by functional cloning in Z. galactanivorans (Jam et al. 2005). These genes were named agaA and agaB. The product of gene agaA codes for a protein with 539 amino acids and contains a N-terminal sequence of 19 residues which probably targets the gene product into the extracellular medium (von Heijne 1983) and is cleaved of in the mature protein (Figure 16).
Further sequence analysis revealed that the gene product of agaA is modular, and contains three domains which are the catalytic GH16 module coupled to two C-terminal modules of unknown function. The high affinity of the native AgaA enzyme to sepharose beads (cross linked agarose) suggests that these non characterized modules might consist of agarose specific carbohydrate binding modules (Murielle Jam personal communication).
The gene product of agaB is a protein of 40.7 kDa molecular weight and 335 residues and the 20 first residues encode a hydrophobic signal peptide which does not contain the peptidase I specific cleavage site (Nakai et al. 1991) (PSORT) but a possible lipid anchor, suggesting that this protein remains cell wall associated after its expression (Jam et al. 2005). The sequence analysis showed that AgaB is a monomodular enzyme, consisting only of the GH16 catalytic module.
Figure 16: The gene organisation of the two first described β-agarases from Z. galactanivorans copied from (Jam et al. 2005).
The biochemical analysis of these β-agarases revealed that the major reaction products of both are neo-agarohexaose and neo-agarotetraose and that AgaB can further degrade the neo-agarohexaose into neo-agarotetraose and neo-agarobiose. The degradation efficiency of both enzymes was highest on neo-agarooctaose indicating that these enzymes possess a substrate binding cleft consisting of eight subsites, four on the non reducing end (-) and four on the reducing end (+). This interpretation was also confirmed by the crystal structures of both enzymes (Allouch et al. 2003).
Figure 17: The β-agarase AgaB from Z. galactanivorans forms a biological dimer copied from (Jam et al. 2005).
Figure 18: The β-agarase AgaA from Z. galactanivorans contains a second binding on the back of the enzyme possibly involved in the unwinding of the double helical agarose chain as here modelled by extending the two bound oligosaccharides copied from (Allouch et al. 2004).
Furthermore the crystal structure analysis revealed some particular features of both enzymes. AgaB forms a biological dimer induced by an unusual loop in GH16 between residues 282-294 (Figure 17). This dimer may be due to the biological role of AgaB as cell wall bound glycoside hydrolase and it was confirmed by gel filtration experiments.
When the activity of the β-agarases were compared on solid state agarose gel it revealed that AgaA had a significantly higher efficiency in agarose gel degradation than AgaB and the protein crystal structure analysis gave an elegant and surprising explanation for this phenomenon. AgaA has a second agarose binding site which is located at the opposite side of the enzyme in respect to the substrate binding cleft (Figure 18). By extending the oligosaccharides bound to both binding sites the double helical agarose model, proposed by Arnott and Fulmer et al. 1974, could be modelled without disturbing the stereochemistry of the glycosidic linkages and the sugar planes. It was postulated that this new mode of binding could allow AgaA to unwind the double helical agarose which would explain why this enzyme is so active on agarose gel in respect to AgaB (Allouch et al. 2004).

The “knowledge gap” of marine polysaccharides degrading enzymes

In the past, advances in microbiology, including marine microbiology, depended mostly upon culturing. The new age of metagenomics enables the study of the vast majority of microbial species which are as yet unable to be cultivated in the laboratory. These technologies and the analyses they enable (comparative (meta)genomics, (meta)transcriptomics, (meta)proteomics, metabolomics, high throughput gene disruptions, etc) have ushered in a new era of biology with fundamental implications for basic research and biotechnological advances. But, they also pose challenges, especially in marine biology, since this flow of data also highlight the lack of knowledge concerning marine specific metabolisms.
Although projects as the Sorcerer II Global Ocean Sampling Expedition have highlighted the fact that much of the phylogenetic and biochemical diversity of life on Earth is present in its marine microbes, they also highlight that we are way behind in understanding the diversity and functioning in the marine environment. – Up to 60 % of the sequences have no equivalent to previously studied proteins. There are huge numbers of putative genes, the function of which is often unknown and at best only deduced from sequence comparisons. Because more is often known about the genetics and physiology of terrestrial organisms, the number of unknown/putative genes is overwhelming for marine samples because there is so little experimental data on marine model organisms. As an example, despite their ecological importance the number of biochemically and structurally described « marine » glycoside hydrolases is low compared to « terrestrial » glycoside hydrolases (Elifantz, Waidner et al. 2008). Moreover, when identified, these enzymes often constitute completely new protein families: – and -carrageenases (Michel et al. 2003), -agarases (Flament et al. 2007) or fucanases (Colin, 2006). Therefore, these enzymes are only accessible through the application of standard biochemical approaches, since in any genomic approach they would have been annotated as “conserved hypothetical proteins” or given incorrect substrate specificities. The keyword search « glycoside hydrolase » in the Protein Data Bank (21.09.09) gave 413 structure hits of which 170 can be attributed to cellulase or related (CBM) functions. In contrast, a search for marine polysaccharide specific glycoside hydrolases such as agarases results in five structural hits and a search for carrageenase resulted in three hits. Crystal structures for unexploited polysaccharides ulvan or fucoidan do not yet exist, not mentioning enzymes active on unknown substrates from ecologically important unicellular algae. This bias towards biotechnologically important terrestrial enzymes follows an economical rationale but does not reflect the ecological importance of bacterial polysaccharide processing in the sea. The economical interest in marine glycoside hydrolases may change since marine algae currently gain in interest as possible producers of biofuels, increasing the possible applications of new enzymes. Moreover, biochemical characterization of new marine enzymes may help to understand the functioning of the ecosystem.
Most enzymes currently used in biotechnology are of microbial origin. The microbial world contains the greatest fraction of biodiversity in the biosphere. It is thought that the marine environment, covering more than 70 percent of the earth’s surface, contains ~4 x 1030 microorganisms. Commercial expectations are that microbes will deliver the greater part of enzyme diversity and the majority of new applications. However, the well-known dilemma of microbes, whatever their origin , that the majority cannot be cultivated, limits the application of the traditional means of enzyme discovery. Here it was therefore decided to use a different approach and to exploit the available genomic data of one marine bacterium, sequenced because of its algal polysaccharide degrading capacity. Indeed, with this approach that consisted in the combination of knowledge based genome mining, medium throughput cloning and expression, natural substrate screening as well as classical biochemical and structural characterization, it was possible to identify and fully characterize a new glycoside hydrolase function.

Aim: finding new glycoside hydrolases by analysing the agarolytic system of Zobellia galactanivorans

New glycoside hydrolases and enzymes in general can be screened in a number of ways. Expression clone libraries, in which genetic information is linked with an expressed protein, can be screened for a certain substrate. This was demonstrated for the agarases AgaA and AgaB, and numerous other reported agarolytic enzymes. This strategy is especially convenient with agarases since their activity can be directly monitored. An activity based screening approach has the potential to detect new glycoside hydrolase families but depends on available substrates and activity assays. The latter is rarely available for biotechnologically non-exploited polysaccharides from marine algae. If algal polysaccharides were available, marine bacteria could be screened for their ability to grow on these substrates for instance as sole carbon source. The active protein can be purified and the sequence partially determined by classical biochemical approaches such as edman or mass spectrometry protein sequencing. The obtained sequence information can be used to amplify the target gene with degenerated primers to obtain the gene sequence allowing further molecular biological analysis (Hehemann et al. 2008). This approach is laborious but it enables the discovery of new glycoside hydrolases belonging to new families.
Another possibility is to choose sequences of known GH families in genome data bases that display significant divergence from enzymes of known function, hinting at new specificities. Combined with the technical advances (such as medium throughput cloning strategies) that have been reached through structural genomics, one can select a high number of promising targets, analyse the soluble produced enzymes ‘from the genome to the biochemical characterization’ and eventually identify new specificities/functions through this method. This is the knowledge based approach that was applied in this study.

Table of contents :

I. Introduction
I.1 The marine ecosystem
I.1.1 Heterotrophic bacteria as key player in marine carbon cycle
I.1.2 Polysaccharides are an important part of the marine DOM and POM pools The « Sweet Ocean »
I.1.3 Bacterial enzymatic decomposition of marine organic matter
I.1.4 Why enzymatic degradation of gel forming marine galactans impacts the carbon cycle 17 The ocean is a « Sweet Jelly »
I.2 Marine polysaccharides
I.2.1 The cell walls of marine macrophytes
I.2.2 The skeleton component of algal cell walls
I.2.3 The matrix component of algal cell walls
I.2.4 The green algae
I.2.5 The brown algae
I.2.6 The cell wall matrix of marine red algae: agars and carrageenans
I.2.7 Agarose 3D structure
I.3 The marine heterotrophic bacteria
I.3.1 Marine bacteroidetes
I.3.2 Zobellia galactanivorans
I.4 The glycoside hydrolases
I.4.1 Enzymatic transformation of HMW compounds
I.4.2 Glycoside hydrolases and their sequence based classification
I.4.3 The catalytic mechanism of glycoside hydrolases
I.4.4 Mode of action in glycoside hydrolases
I.4.5 Subsites in glycoside hydrolases
I.4.6 Agarolytic bacteria and glycoside hydrolases
I.4.7 Family classification of agarases
I.4.8 The glycoside hydrolase family GH
I.4.9 The β-agarases from Z. galactanivorans
I.5 The “knowledge gap” of marine polysaccharides degrading enzymes
I.6 The aim of the thesis: finding new glycoside hydrolases by analysing the agarolytic system of Zobellia galactanivorans Structural and Functional Organisation of the Agarolytic Enzyme System of the Marine Flavobacterium Zobellia galactanivorans
II. Results
II.1 Medium throughput cloning and expression strategy
II.2 The targets for further characterization: AgaD, PorA and PorB
III. The Structural and Biochemical Characterization of the new -agarase AgaD
III.1 Introduction for manuscript1: Protein crystallization of AgaD
III.2 Manuscript 
III.2.1 Abstract
III.2.2 Introduction
III.2.3 Material and Methods
III.2.4 Results and discussion
III.2.5 Conclusion
III.2.6 Acknowledgments
III.2.7 References
III.3 The crystal structure of AgaD
III.4 Biochemical characterization of AgaD
III.4.1 Catalytic behaviour of AgaD
III.4.2 AgaD is an endo β-agarase cleaving the β-1,4 linkages in agarose
III.4.3 Agarase specificities are different on natural substrates extracted from the agarophytes Gelidium, and Porphyra
III.4.4 PACE and HPLC analysis of porphyran degradation by AgaA, AgaB and AgaD
III.5 Conclusion: The new agarase AgaD together with AgaA,B as part of the agarolytic
IV. The first -porphyranases PorA and PorB
IV.1 Crystallisation of PorA
IV.1.1 3D structure solution of PorA using a gold derivative
IV.1.2 Crystallization of PorB
IV.1.3 The crystal structures of PorA and PorB
IV.2 The discovery of the β-porphyranase activity
IV.3 Introduction for manuscript2: Porphyranases and agarases constitute the first example of a nutrition derived CAZyme update into human gut bacteria
IV.4 Manuscript 
IV.4.1 Abstract
IV.4.2 Discovering a new enzyme activity
IV.4.3 Structural determinants of porphyran active enzymes
IV.4.4 β-Porphyranases are abundant in marine bacteria
IV.4.5 Horizontal gene transfer from marine to human gut bacteria
IV.4.6 Discussion
IV.4.7 Methods summary
IV.4.8 Methods
IV.4.9 References
IV.4.10 Supplementary material
IV.5 Introduction for manuscript3: Production of porphyran oligosaccharides with porphyranase
IV.6 Manuscript 
IV.6.1 Abstract
IV.6.2 Introduction
IV.6.3 Material and Methods
IV.6.4 Results and discussion
IV.6.5 Conclusion
IV.6.6 References
V. Material and Methods
V.1 Expression and purification of PorA and PorB
V.2 DNA techniques and plasmid construction
V.3 The medium throughput cloning strategy
V.4 Screening for crystallization conditions
V.5 Kinetic studies
V.6 Sequences and phylogeny
V.7 Fluorophore-assisted carbohydrate electrophoresis analysis (PAGE)
V.8 Enzyme activity essays
VI. Final discussion and outlook
VI.1 The agarolytic system of Z. galactanivorans
VI.2 Screening for new marine glycoside hydrolases
VI.3 β-porphyranases discovery in marine bacteria
VI.4 Seaweed polysaccharide degrading CAZymes in human gut bacteria
VI.5 Marine glycoside hydrolases as tools to analyse marine POM and DOM
VII. References

GET THE COMPLETE PROJECT

Related Posts