physiological and molecular understanding of bacterial … · all bacterial pmos belong to the aa10...
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Physiological and MolecularUnderstanding of BacterialPolysaccharide Monooxygenases
Marco Agostoni,a John A. Hangasky,a Michael A. Marlettaa,b,c
California Institute for Quantitative Biosciences (QB3), University of California, Berkeley, Berkeley, California,USAa; Department of Chemistry, University of California, Berkeley, Berkeley, California, USAb; Department ofMolecular and Cell Biology, University of California, Berkeley, Berkeley, California, USAc
SUMMARY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2MOLECULAR ASPECTS OF POLYSACCHARIDE MONOOXYGENASES . . . . . . . . . . . . . . . . . . . . . . 2
Classification of Polysaccharide Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2Evolution of Bacterial PMOs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3Structure of Bacterial PMOs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4Domain Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
PHYSIOLOGICAL ASPECTS OF POLYSACCHARIDE MONOOXYGENASES . . . . . . . . . . . . . . . . . . 5Genomic Comparison of Bacterial PMOs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5Environmental Factors: Regulator Elements of Cellulose and Chitin Metabolism . . . . . . . . 5
FUNCTIONS OF BACTERIAL PMOs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7Host-Microbe Interactions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7
Endosymbiosis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7Antifungal properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8Bacterial PMOs as virulence factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9
Role of PMOs in Human Infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9Vibrio cholerae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9Pseudomonas aeruginosa. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10Listeria monocytogenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10
PERSPECTIVES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11ACKNOWLEGMENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11REFERENCES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11AUTHOR BIOS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16
SUMMARY Bacteria have long been known to secrete enzymes that degradecellulose and chitin. The degradation of these two polymers predominantly in-volves two enzyme families that work synergistically with one another: glycosidehydrolases (GHs) and polysaccharide monooxygenases (PMOs). Although bacterialPMOs are a relatively recent addition to the known biopolymer degradation ma-chinery, there is an extensive amount of literature implicating PMO in numerousphysiological roles. This review focuses on these diverse and physiological as-pects of bacterial PMOs, including facilitating endosymbiosis, conferring a nutri-tional advantage, and enhancing virulence in pathogenic organisms. We also dis-cuss the correlation between the presence of PMOs and bacterial lifestyle andspeculate on the advantages conferred by PMOs under these conditions. In addi-tion, the molecular aspects of bacterial PMOs, as well as the mechanisms regu-lating PMO expression and the function of additional domains associated withPMOs, are described. We anticipate that increasing research efforts in this fieldwill continue to expand our understanding of the molecular and physiologicalroles of bacterial PMOs.
KEYWORDS Listeria monocytogenes, Pseudomonas, cellulose, cellulolytic enzymes,chitin, endosymbionts, infectious disease, monooxygenases, polysaccharides
Published 28 June 2017
Citation Agostoni M, Hangasky JA, MarlettaMA. 2017. Physiological and molecularunderstanding of bacterial polysaccharidemonooxygenases. Microbiol Mol Biol Rev 81:e00015-17. https://doi.org/10.1128/MMBR.00015-17.
Copyright © 2017 American Society forMicrobiology. All Rights Reserved.
Address correspondence to Michael A.Marletta, [email protected].
M.A. and J.A.H. contributed equally to thiswork.
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INTRODUCTION
Cellulose (C6H10O5)n and chitin (C8H13O5N)n are the two most abundant biopolymerson Earth (1, 2) and are composed of �(1-4)-linked D-glucose (Glc) and �(1-4)-linked
N-acetyl-D-glucosamine (GlcNAc), respectively. These biopolymers are widely distrib-uted due primarily to their high resiliency and versatility in combining with proteinsand other compounds to form hybrid complexes (3, 4). Cellulose is the primarycomponent of plant cell walls, but it is also found in some bacteria, fungi, and protozoa(5). Cellulose has twice the tensile strength of chitin, conferring an advantage as astructural component for land plants over chitin (6). On the other hand, chitin hasplayed a widespread role in numerous living organisms since the Cambrian life explo-sion (3, 7). In crustaceans and insects, chitin supports the exoskeleton and the attach-ment of muscles to joints and defends against pathogens and predators (8–10). Inyeasts, amoebas, fungi, and sponges, chitin provides rigidity and strength to the cellwall (11–14). Also, chitin confers buoyancy in some marine photosynthetic microor-ganisms by increasing the cell surface area (15).
Bacteria have developed specialized strategies to efficiently degrade chitin andcellulose. Cellulose-degrading bacteria are prevalent in terrestrial and submarine soil(16), whereas chitin-degrading bacteria are prevalent in both terrestrial and aquaticenvironments (17). The degradation of cellulose and chitin predominantly involves twosecreted-enzyme families that work synergistically with one another: glycoside hydro-lases (GHs) and polysaccharide monooxygenases (PMOs) (18–23). GHs, which arecommon in bacteria (17, 24, 25), have been known for a relatively long period of timeand have been the subject of recent reviews (26, 27). Conversely, PMOs are a recentaddition to the known biopolymer degradation machinery; their oxidative chemistrywas first reported in 2010 (19), and since then, they have been identified in eukaryotic,bacterial, and viral genomes (28). PMOs are commonly associated with fungal metab-olism and biomass degradation, and the potential for these proteins in the biorefineryindustry has already been extensively reviewed (20, 29, 30). Bacterial PMOs have alsobeen implicated in having numerous physiological roles. Thus, here we focus onPMO-containing bacteria and their diverse physiological and functional roles.
MOLECULAR ASPECTS OF POLYSACCHARIDE MONOOXYGENASES
The crystalline structures of cellulose and chitin make these biopolymers resistant tohydrolysis. PMOs use an oxidative process leading to the cleavage of �-1,4 glycosidiclinkages, creating new chain ends for glucanase action (19, 31, 32). PMOs are intriguingenzymes, not only as they oxidize C-H bonds but also because this chemistry does notrequire separation of the polysaccharide chain from the crystalline matrix backbone forbond cleavage. Since their discovery, significant contributions to the biochemicalcharacterization of PMOs have been made, and other reviews have thoroughly coveredPMO structure and the copper active site relating to catalysis (28, 33–35); therefore,only a brief summary is given here.
Classification of Polysaccharide Monooxygenases
The Carbohydrate-Active enZYmes (CAZy) database (36), classifies PMOs as auxiliaryactivity (AA) enzymes and places all PMOs into four families based on sequencehomology: AA9 (formerly GH61), AA10 (formerly CBM33), AA11, and AA13. The AA9,AA11, and AA13 families are found in fungi and oxidize cellulose, chitin, and starch,respectively (37–40). All bacterial PMOs belong to the AA10 family and are active oneither cellulose or chitin (19, 32, 41). To date, only approximately 20 out of the over2,400 putative bacterial PMOs present in the CAZy database have been biochemicallycharacterized, leaving many unanswered questions about this large group.
Evolution of Bacterial PMOs
Bacterial PMOs most likely share a distant common ancestor with fungal PMOsbelonging to the AA9 PMO families (42); however, no AA9 or AA11 PMOs are found inbacterial genomes. Phylogenetic clustering of bacterial PMO sequences separates AA10
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PMOs into two major clades: clade I and clade II. Clade I consists of chitin-active PMOs,whereas clade II consists of cellulose-active PMOs (23, 40, 42). Further selection for morefavorable binding interactions with substrates and other proteins likely influenced theorigin of additional subclades (23, 42). An interesting question arises as to whetherbacterial PMOs initially evolved to utilize chitin or cellulose substrates. One couldspeculate that since chitin was present before cellulose, PMOs could have originated todegrade chitin. The first chitin fossil record dates to 505 million years ago (middleCambrian) and was isolated from a skeletal component of early sponges (43), althoughchitin probably originated in protozoans 1,400 million years ago (44). Cellulose origi-nated in land-adapted plants later, around 450 to 470 million years ago (45). Indeed,recent phylogenetic analyses suggested that bacterial PMOs first originated to degradechitin and then evolved for cellulose degradation (42).
Mechanism
PMOs are mononuclear copper-dependent monooxygenases that hydroxylate eitherthe C-1 or C-4 position of the glycosidic bond, forming an unstable intermediate, whichdecomposes with concomitant bond cleavage. Following bond cleavage, C-1 hydroxy-lation ultimately produces aldonic acids through a hemiacetal intermediate, whereasC-4 hydroxylation ultimately leads to ketoaldoses through a hemiketal intermediate(Fig. 1). Loop flexibility, particularly the “loop 2” region, has been reported to have a rolein conveying oxidative regioselectivity, although the molecular basis for regioselectivityremains unanswered (46). The chemical mechanism of PMOs, including the catalyticsteps and the active-site oxidant, still remains to be determined (28, 47). PMOs are alsocalled lytic PMOs (LPMOs) since C-1/C-4 hydroxylation ultimately results in the cleavageof the glycosidic bond. We have commented on this nomenclature previously (28).
Two one-electron reductions and two proton transfers are necessary to activate O2
for the oxidation of the glycosidic bond (Fig. 1). Small-molecule reductants andphotosynthetic pigments (48) have been shown to directly serve as electron donors forbacterial PMOs. Recently, it was shown that phenols coupled with members of theglucose-methanol-choline (GMC) family of oxidoreductases were effective electrondonors for fungal PMOs (49). Also, cellobiose dehydrogenase (CDH), a known electrondonor for fungal PMOs (50, 51), has also been shown to serve as an electron donor forbacterial PMOs (52). Thus, although not yet identified, extracellular oxidoreductases orredox-active enzymes likely serve as biological redox partners for bacterial PMOs. Dueto the range of environments in which bacterial PMOs are found, as well as their diversefunctions (see below), the biologically relevant electron donor may vary based on thebacterial environment.
FIG 1 The reaction catalyzed by polysaccharide monooxygenases. Oxidation at the C-1 or C-4 position of the glycosidic linkage producesaldonolactones and 4-ketoaldoses, respectively (19, 152). Aldonolactone products are most commonly observed as aldonic acids, and4-ketoaldoses are most commonly observed as gemdiols. For cellulose, R indicates Glc and R= indicates OH; for chitin, R indicates GlcNAc,and R= indicates NHCH3CO.
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Structure of Bacterial PMOs
The first crystal structure of a bacterial PMO, that of the Gram-negative bacteriumSerratia marcescens, was reported in 2005 (53). Since then, multiple crystal structuresand one nuclear magnetic resonance (NMR) structure (54) have been reported forvarious organisms, including Serratia marcescens, Enterococcus faecalis (55, 56), Vibriocholerae (57), Streptomyces coelicolor (41), Jonesia denitrificans (58), and Bacillus amy-loliquefaciens (34). These structures show that the catalytic domains of bacterial PMOshave a �-sandwich fold similar to that of the other PMO families. The active sitecoordinates a type II mononuclear copper with two histidines and the N-terminal aminein a T-shaped histidine brace (Fig. 2). The active site is found on a flat substrate bindingsurface ideal for interactions with crystalline substrates (19, 37–40, 54–56, 59, 60), whichallows PMOs to cleave the glycosidic bond without the need to separate the polysac-charide chain from the crystalline matrix. Chitin-active bacterial PMOs have a smallpocket near the copper active site that might serve to accommodate the acetyl groupfound on the glucosamine moiety (41).
The first crystal structures of a PMO with bound cello-oligosaccharides were recentlyreported for the fungus Lentinus similis (61). These structures provide insight to themolecular aspects involved in substrate binding for bacterial PMOs. Hydrogen bondcontacts, a lone pair-aromatic interaction involving His1, and a CH-� interaction help tobind the cello-oligosaccharide (61). The CH-� interaction forms between the pyranosering of the carbohydrate and a highly conserved Tyr on the surface of the PMO. Thehydrogen-bonding network that forms between the oligosaccharide, a water molecule,and N-terminal His1 may serve as a proton transfer pathway needed to stabilize areactive Cu-O2 intermediate (61, 62).
Domain Structure
PMOs are active as single-domain proteins; however, in some cases, the full-lengthpolypeptide has been found to contain additional domains. These additional domainsinclude carbohydrate binding modules (CBMs), fibronectin type III-like domains (FnIIIs),hydrolase domains, and polycystic kidney disease domains (PKDs) (20). Domains withunknown function have also been found fused to PMOs in the AA11 family (39). CBMsare classified into numerous families, based on amino acid sequence. These domainscontain aromatic residues proposed to increase binding to various polysaccharides (63).Likewise, CBMs fused to PMOs have been proposed to increase polysaccharide speci-ficity and binding (64, 65) and to modulate activity (18, 41). CBMs have also beenimplicated in aiding in substrate processivity by GHs (66), but to date, processivity hasyet to be shown for PMOs. FnIIIs and PKDs are typically associated with mediating celladhesion and protein-protein interactions (67–73). Therefore, these domains mayfacilitate substrate binding; however, exactly how they do so is not known, especially
FIG 2 Crystal structure of ScLPMO10B from Streptomyces coelicolor (PDB accession number 4OY6) (153).The histidine brace motif, consisting of the two copper-coordinating histidine residues, is shown in theinset. The surface of the protein is shown in light gray and illustrates the flat substrate binding surface.
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since the PMO catalytic domain is capable of binding the substrate. Thus, a physiolog-ical perspective is necessary to achieve a deeper understanding of the role of multi-domain PMOs.
Studies on the V. cholerae PMO GbpA (VCA0811), a PMO with four domains, providesome insight into the physiological role of PMO domain architecture. GbpA is com-posed of four domains: a catalytic PMO domain, a chitin binding domain, and twobacterial flagellin-like domains (57). Binding of V. cholerae to mucin in the intestinalepithelium of the host is mediated by the GbpA catalytic domain and the two surfaceflagellin-like domains (57); it is unclear if the catalytic domain has an oxidative role inthis process. On the other hand, in an aquatic habitat, the catalytic domain and thechitin binding domain of GbpA attach to the exoskeleton of crustaceans and cleavechitin (74). PMO-associated domains can contribute to our understanding of thephysiological roles of these enzymes. Thus, a detailed understanding of PMO-associated domains found in other bacteria as well as the substrates with which thesePMOs interact is needed not only to advance the field but also to have a betterunderstanding of the physiological roles of these domains.
PHYSIOLOGICAL ASPECTS OF POLYSACCHARIDE MONOOXYGENASESGenomic Comparison of Bacterial PMOs
The Pfam hidden Markov model for bacterial PMOs can be found under accessionnumber PF03067 (75, 76), and it can also be accessed under InterPro database acces-sion number IPR004302. This database identifies PMO families based on sequencehomology. However, this database is not comprehensive, and putative PMOs are stillbeing discovered, creating new Pfams (77, 78). Through the PF03067 database, putativePMOs seem to be extensively present in the phyla Proteobacteria, Actinobacteria, andFirmicutes. The phylum Chloroflexi possesses only three species with PMOs, the phylumBacteroidetes possesses two species, and the phyla Chlamydiae and Verrucomicrobiapossess one species each, indicating that these phyla have a low number of PMOs. TheArmatimonadetes, Cyanobacteria, Fibrobacteres, Planctomycetes, Spirochaetae, and Ther-motogae phyla did not contain any putative PMOs. These results are in agreement withdata from two previous reports (17, 40).
The presence of PMOs in bacterial genomes appears to be influenced by numerousvariables, including habitat and lifestyle. Since PMOs are oxygen dependent, it is notsurprising that anaerobic microbial communities lack PMOs (17). Likewise, it is notsurprising that many bacteria isolated from soil or decomposing biomass possessPMOs. Xylanimonas cellulosilytica, a Gram-positive bacterium of the actinobacterialfamily possessing two PMOs (40), was isolated from a decaying tree and shown todegrade both cellulose and xylan (79). Similarly, Saccharophagus degradans, a Gram-negative proteobacterium possessing one PMO (40), is a carbohydrate-degradingmarine bacterium that can degrade chitin and cellulose, among other polymers (80).However, there are examples of aerobic cellulolytic bacteria that do not possess PMOs.Both Acidothermus cellulolyticus, a Gram-positive bacterium of the actinobacterial familyisolated from an acidic hot spring (81), and the saprophytic Gram-negative proteobac-terium Agrobacterium radiobacter, which grows on decayed plant matter (82), lackPMOs. These examples suggest that other variables beyond lifestyle could dictate thepresence of PMOs in bacterial genomes.
Environmental Factors: Regulator Elements of Cellulose and Chitin Metabolism
Some bacteria use cellulose and chitin as signaling molecules (Fig. 3) (83) and asmarkers for nutrient availability and developmental control (84). In polymer-enrichedhabitats, bacteria secrete PMOs as well as cellulases and chitinases (18–23, 85) todegrade cellulose and chitin into smaller soluble oligosaccharides; this could possiblyprovide the bacterium with simpler carbon and nitrogen sources (Fig. 3). PMOs can beupregulated when bacteria are grown in the presence of cellulose or chitin (86, 87),indicating that bacteria have developed mechanisms to regulate the expression ofthese PMOs in response to these potential substrates (86–88) (Fig. 3 and see below). For
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example, the thermophilic actinobacterium Thermobifida fusca secretes the PMOs E7and E8 when grown on cellulolytic polymers; these PMOs work synergistically with T.fusca GHs to form cellobiose, a sugar utilized for growth (89). Genes that are in the sameoperon are coexpressed and can provide valuable insight into gene function andprotein interactions. However, PMOs do not appear to be organized into operons butinstead are distributed throughout the whole genome close to regulatory systemsinvolved in sensing cellulose or chitin (88, 90).
The genus Streptomyces illustrates very well the complex network of the transcrip-tional control of both cellulose- and chitin-active PMOs. Streptomyces sp. SirexAA-E hassix predicted PMOs. During growth on cellulose, Streptomyces sp. SirexAA-E secretesthree putative PMOs, SACTE_3159, SACTE_6428, and SACTE_2313, whereas the putativePMOs SACTE_2313, SACTE_0080, and SACTE_6493 are secreted when grown with chitin(88). SACTE_2313 was the only putative PMO secreted under conditions of bothcellulose and chitin growth, suggesting substrate-specific responses for the other fourPMOs.
CebR, the master regulator of cellulose/cello-oligosaccharide catabolism, is involvedin sensing cellulose and cellooligosaccharide following their uptake in Streptomyces(86). In the presence of cello-oligosaccharides, including cellobiose, binding of CebR toDNA is weakened, allowing downstream gene transcription to be enhanced (91) (Fig. 3).Genes regulated by CebR include cellobiose/cellotriose ABC transporters, cellulose-active PMOs, and other cellulases (86). In Streptomyces, these genes are found close toa palindromic CebR binding element, which regulates their expression under cellulosegrowth (86, 88).
Chitin and chito-oligosaccharide catabolism, on the other hand, is regulated by theGntR/HutC family regulator DasR, a global transcriptional regulator in Streptomyces (84).In response to the uptake of the chitin monomer N-acetylglucosamine (GlcNAc), DasRregulates numerous genes, including those involved in the phosphotransferase system,the chitobiose ABC transporter system, as well as chitin-active PMOs and other chiti-nases. Interesting, the DasR-DNA complex does not dissociate in the presence ofGlcNAc; rather, glucosamine-6-phosphate, a intermediate in GlcNAc metabolism, is theeffector molecule (84) (Fig. 3). In Streptomyces coelicolor, the SCO0481, SCO2833,SCO6345, and SCO7225 genes were upregulated by DasR (87). Based on sequencealignments, these genes encode putative chitin-active PMOs (see Fig. 5). When Strep-tomyces coelicolor A3 was grown in soil, SCO7225 (chiM) was the only putative PMO thatwas upregulated (92). Microarray analyses confirmed that there were 21- and 3-foldupregulations of the putative PMOs SCO7225 and SCO2833, respectively, in soil withand without chitin; there was no change in SCO6345 under these conditions (93). Thesechanges in PMO transcripts are consistent with a role for chitin in gene regulation.
FIG 3 Bacterial sensing of cellulose and chitin. Bacteria can sense cellulose and chitin through CebR and DasR, respectively. In turn, CebRand DasR regulate the expression of PMOs and other GHs. PMOs cleave cellulose and chitin chains, generating new chain ends morereadily accessible by GHs for further degradation.
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Besides cellulose- and chitin-rich environments, other environmental factors mayinfluence the expression levels of PMOs. The ability of the Gram-negative bacteriumVibrio cholerae to survive under diverse environmental conditions is enhanced throughattachment to the chitinous exoskeletons of zooplankton (94, 95), and the PMO GbpA(VCA0811) plays an important role in chitin degradation for growth (96). Ecologicalstudies have determined that GbpA and the ability to bind to chitin are influenced byenvironmental factors such as increases in temperature (97) and quorum sensing (98).The detailed functions of GbpA are described below in the section on V. cholerae.
FUNCTIONS OF BACTERIAL PMOs
Bacterial PMOs have been implicated in various functions, including nutrition,endosymbiosis, and virulence in pathogenic organisms (88, 96, 99–101) (Fig. 4). It is stilltoo early to definitively know the function of bacterial PMOs, and it is highly likely thatthe physiological role of PMOs varies from organism to organism. In this section, wediscuss the advantages that PMOs confer when organisms rely on carbon sources otherthan glucose, the gene regulatory network involved in sensing cellulose and chitin, theprimary role of PMOs in facilitating host-microbe interactions, and, finally, the proper-ties of PMOs as antifungal agents and virulence factors.
Host-Microbe InteractionsEndosymbiosis. Symbiosis between host organisms and cellulolytic bacteria led to
the evolution of new feeding strategies. Host organisms have taken advantage ofbacterially secreted enzymes, including PMOs, GHs, and other cellulases, to deconstructcell walls and degrade plant biomass. These symbiotic relationships modulate theinteractions of the host organisms with their environment. The first documentedacquisition of symbiotic cellulolytic microorganisms was by lower termites originatingaround 150 million years ago, leading to enormous evolutionary and ecological success(102, 103).
FIG 4 Putative functions of bacterial PMOs. Following secretion, bacterial PMOs can oxidize a range of polysaccharide substrates, resulting in PMOs beingimplicated as having various functions, including degrading biomass, being involved in endosymbiotic relationships, and serving as virulence factors ofpathogenic bacteria and as antifungal agents, in addition to providing a nutritional source for bacteria.
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Some strains of Streptomyces sp. are symbiotic with bark beetles and wood wasps,which rely on the degradation of plant biomass (104). These insects take advantage ofsecreted biomass-degrading enzymes to facilitate energy accumulation inside plant cellwalls for larva development (104). Similarly, the giant snail Achatina fulica possesseshigh cellulolytic activity, mainly due to the abundance of resident microbiota in thegastrointestinal tract (105). Streptomyces sp. strain I1.2 was isolated from this herbivoreinvertebrate and displayed high growth rates on cellulose as the sole carbon source(105). Among genes that putatively encode enzymes involved in plant cell wall decon-struction, there are four genes encoding putative PMOs (105). The authors of that studyconcluded that Streptomyces is involved in symbiotic associations with A. fulica and thatthis association is important for lignocellulose degradation (105). In the marine envi-ronment, host-proteobacterium symbiotic relationships potentially contributed to theevolutionary success of bivalves in a cellulose-enriched habitat (106). Unlike terrestrialsymbiotic relationships, where cellulolytic bacteria are found in the gut, these micro-biotas are found in the gills of the bivalves (107–110). The gammaproteobacteriumTeredinibacter turnerae, isolated from the gills of bivalves, contains numerous GHs andother carbohydrate-active enzymes as well as one predicted PMO (TERTU_0046, whichcontains the two histidines in the active site critical for PMO activity but lacks theconserved regions of the bacterial PMOs depicted in Fig. 5). Additional studies areneeded to know when the genes encoding these putative PMOs are induced and tobiochemically characterize the enzyme as a PMO.
Antifungal properties. As the cell wall of fungi is composed primarily of chitin (13),it is not surprising that some bacterial PMOs possess antifungal activity (100, 111–113).Cbp50 (annotated CBM33), from the Gram-positive soil-dwelling bacterium Bacillusthuringiensis, was the first PMO definitively shown to have antifungal properties (100).Cpb50 strongly binds �-chitin but can also bind colloidal chitin and cellulose (100). Bybinding �-chitin present in the fungal cell wall, Cpb50 may prevent chitin biosynthesisduring cell division (100). Other studies have suggested that other PMOs, such as CHB1and CHB2, may interact with the fungal cell wall. CHB1 and CHB2 are two chitin binding
FIG 5 Sequence alignment of conserved regions of bacterial PMOs. Green highlighting shows copper-coordinating residues, gray highlighting shows conserved residues on the putative substrate bindingsurface, and purple highlighting shows a conserved aromatic residue by the active site. Numbers indicatethe residue of the reference sequence (italics). Shown are bacterial PMOs predicted to oxidize C-1 ofcellulose (A), C-1/C-4 of cellulose (B), and chitin (C).
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proteins with sequence homology to known PMOs (Fig. 5), are secreted by Streptomycesin the presence of �-chitin, and could bind to mycelia from fungi (111, 112). AnotherPMO in Streptomyces, CHB3, was later shown to interact with �-chitin, �-chitin, andchitosan, possibly allowing Streptomyces to bind various polysaccharide substratesfound in fungal cell walls (114). Subsequently, PMOs similar to CHB1 and CHB2 fromBacillus amyloliquefaciens were shown to bind chitin from fungi (113). However, thosestudies did not conclude whether CHB1 and CHB2 have antifungal proprieties. It is alsoimportant to note that not all PMOs enhance antifungal proprieties of bacteria. It hasbeen suggested that CBP21 in Serratia marcescens, BtCBP in B. thuringiensis, and BliCBPin Bacillus licheniformis are able to bind and degrade chitin but do not have any effecton inhibiting fungal growth (115). However, these PMOs were not suspended in asolution containing reducing agents, complicating interpretations of the results.
Bacterial PMOs as virulence factors. The biological roles of bacterial chitinases andPMOs and their interactions with insects and fungi can be understood from anenvironmental perspective where these interactions often result in pathogenesis. Chitinis present in the midgut of most invertebrates, where it serves as a major structuralcomponent (116), functioning as a mechanical barrier for protection and compartmen-talization of digestive processes (117). Pathogenic bacteria must breach this layer toinvade the host organism, and there is growing evidence that PMOs are involved in thisprocess.
Paenibacillus larvae, a Gram-positive bacterium pathogenic to honeybee larvae, isable to degrade the chitin-containing gut during infection (118, 119). Once this layer isbreached, P. larvae can initiate the invasive growth phase (99). PlCBP49, a chitin-activePMO (Fig. 5), was shown to be key for honeybee larva colonization, as its absenceabolished P. larvae virulence (99).
Role of PMOs in Human Infection
In recent years, attention has been drawn to the role that bacterial chitinases andPMOs play in humans, despite the fact that chitin is not an endogenous component ofmammals (120, 121). The bacterial pathogens that show a correlation between PMOactivity and infectibility are V. cholerae, Pseudomonas aeruginosa, and Listeria monocy-togenes. Unfortunately, too little is currently known to review the pathogenic role thatPMOs may play in Enterococcus faecalis infections, where the putative PMO EF0362 isupregulated in the presence of blood and urine (122, 123), and in other pathogenicbacteria with predicted PMOs, such as Serratia marcescens, Bacillus anthracis, andLegionella pneumophila.
Vibrio cholerae. As mentioned above, V. cholerae can survive under low-nutrientenvironmental conditions by attaching to the chitinous exoskeletons of zooplankton(94). Chitin is a substrate for the PMO GbpA (124), which is induced along withchitinases when grown in the presence of chitin and chito-oligosaccharides (125). Notonly is V. cholerae able to use free chitin particles or the chitinous exoskeleton ofcopepods as the sole carbon and nitrogen source for growth (96), but also chitin is usedfor the production of ammonia to increase toxicity to heterotrophic grazers (126).
Kirn et al. hypothesized that the colonization factors promoting adhesion in thenatural environment of V. cholerae could be the same for intestinal mucosal surfacesand identified GbpA as the protein responsible for attachment to epithelial cells (101).GbpA may have originated to function in adhesion with environmental biotic sub-strates and then evolved to be a virulence factor in pathogens (74). V. choleraecolonization of the human intestine is mediated, in part, by GbpA; GbpA interacts withmouse intestinal mucus, with the principal component being the glycoprotein mucin(127). GbpA and mucin mutually enhance expression: GbpA stimulates mucin secretionin a concentration-dependent manner, and mucus secretion increases the level ofGbpA (127).
Quorum sensing and cyclic di-GMP (c-di-GMP) signaling are connected pathwaysthat sense and integrate environmental cues, controlling a broad variety of physiolog-ical functions in V. cholerae (128). Recently, both of these pathways have been shown
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to regulate GbpA. At a high cell density, GbpA expression is repressed by HapR, acentral regulator of cell density-dependent quorum sensing in V. cholerae (129). Fur-thermore, GbpA is degraded by two known quorum sensing proteases, HapA and PrtV(98), strongly indicating that GbpA is regulated through quorum sensing. Genomeneighborhood analysis revealed that gbpA is near two c-di-GMP binding riboswitchescalled Vc1 and Vc2 (130). A recent study confirmed that the second messengersc-di-GMP and cyclic AMP (cAMP) regulate gbpA transcription (131). The levels ofc-di-GMP and cAMP are regulated by numerous abiotic signals, confirming that GbpAexpression is fine-tuned in response to environmental signals (131).
Interestingly, GbpA is not the only PMO predicted to be present in V. cholerae.Although uncharacterized, VCA0140 (which has the two histidines in the active sitecritical for PMO activity but does not possess the conserved regions of the bacterialPMOs present in Fig. 5) was also strongly expressed when V. cholerae was associatedwith living copepods (125). Additional studies are needed to characterize and helpunderstand the function of VCA0140.
Pseudomonas aeruginosa. Pseudomonas aeruginosa is a pathogenic Gram-negativebacterium that inhabits soil and freshwater ecosystems. P. aeruginosa and the patho-genic yeast Candida albicans often form polymicrobial biofilms, especially in the lungsof patients with cystic fibrosis (132). The interaction of these two pathogenic organismsis mainly antagonistic (132). P. aeruginosa can grow on hyphae and kill fungal C.albicans but cannot interact with or kill yeast-form C. albicans (133), likely due todifferences in the proteins and carbohydrates present in the cell walls (134).
Although P. aeruginosa is unable to grow on chitin as a sole carbon source (135), theputative PMO CbpD (PSPA7_4667) is secreted when grown with colloidal chitin-enriched medium. Research into the antagonistic interaction between P. aeruginosaand C. albicans provided evidence that CbpD mediates binding and adhesion to thechitin present in C. albicans hyphae (136). Thus, P. aeruginosa cannot kill the yeast formof C. albicans, as CbpD probably cannot interact with the outermost glycoprotein layerof C. albicans yeast cells (136).
There is growing evidence that CbpD has a virulence role in this opportunisticpathogen. First, there is a strong association between CbpD and cystic fibrosis. Cysticfibrosis patients have been found to have high levels of CbpD in their lungs (137), andmicroarray and protein analyses showed that CbpD was upregulated in strains associ-ated with cystic fibrosis compared to other laboratory strains (137). In addition, in astrain isolated from an acute transmissible cystic fibrosis case, CbpD was abundant inthe secretome compared with the laboratory-adapted strain when grown with artificialsputum medium containing mucin (138). Notably, CbpD has been found only in clinicalisolates of P. aeruginosa but not in nonpathogenic strains isolated from soil (139).
Second, microarray analysis showed that CbpD is important during the early for-mation of biofilm in cystic fibrosis lung rather than maintaining the biofilm (140). Thetranscription of cbpD and its secretion through the type II system are probably underthe control of the quorum sensing system (141). Taken together, these results suggestthat CbpD plays a potentially pathogenic role in distinct and specific areas of inflam-mation and disease and is important for the attachment of the surface leading tobiofilm formation (137, 138).
It is important to note that P. aeruginosa also possesses the putative PMOs CbpA andCbpE (142, 143). Continued studies will shed light on functional homologies anddifferences between CbpD, CbpA, and CbpE and their function in P. aeruginosa.
Listeria monocytogenes. L. monocytogenes is a saprophytic Gram-positive bacteriumfound in water, soil, and decaying plants (144) that causes foodborne infection leadingto septicemia, meningoencephalitis, gastroenteritis, and perinatal infections (145). Thegene lmo2467 encoding a PMO (LmPMO10) in L. monocytogenes is not essential foreither chitinolytic activity (146) or growth in media with different carbon sources asalternatives to glucose (147). Another study confirmed that LmPMO10 was not presentin the secretome, nor was it upregulated when cells were grown in the presence ofdifferent types of chitin (148). Since LmPMO10 is not essential for growth in basic chitin
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medium, it was suggested that it could have a role during pathogenesis. To date, theonly transcriptional study of LmPMO10 under different growth conditions revealed thatthis gene was upregulated when subjected to low temperature or hypoxia, or duringstationary-phase growth, but not in blood from healthy human donors or in thepresence of the intestinal lumen of mice (149). In addition, LmPMO10 was upregulatedat the initial stage of biofilm formation (150), similar to the PMO in P. aeruginosa. Amutant lacking LmPMO10 did not influence bacterial invasion or replication in tissueculture cell lines; however, this mutant exhibited a defect in bacterial colonization inthe spleen and liver of infected mice, and LmPMO10 contributed to virulence in thebloodstream of infected mice (151). These studies unfortunately are not conclusive withregard to the possible role of LmPMO10 in virulence, and the exact mechanism ofaction must still be clarified.
PERSPECTIVES
PMOs in pathogens have been shown to be active on numerous polysaccharides,including cellulose and chitin, yet definitive physiological roles remain unclear. Theseproteins have been implicated in diverse functions, including facilitating endosymbio-sis, providing a source of nutrition, enhancing virulence, and serving as virulencefactors in pathogenic organisms (Fig. 4). With the assumption that the putative PMOactive sites are functional, the key question is, what are the substrates? A starting pointfor any putative PMO would be a carbohydrate moiety; however, this opens the doorto a great many possibilities. The diversity of carbohydrates beginning with themonosaccharide building blocks generates a long list of potential structures. Thesebuilding blocks can then be linked into oligo- and then further to polysaccharides withpotential branch points leading to diverse structures. Glycan attachment to proteinscreates a further diverse array of structure and, correspondingly, potential PMO sub-strates. It is also possible that saccharide-containing glycans found on mammalian cellsand other �(1-4)-linked polysaccharides encountered in pathogenic bacterial biofilmsmay be the targets of bacterial PMOs. Strategies to screen these extensive substrateclasses are clearly needed. An illustrative example is GbpA in V. cholerae, which hasbeen shown to degrade chitin. It was also shown to interact with the glycoproteinmucin, and thus, a role in adhesion was proposed, but mucin as a substrate was notinvestigated. The physiological roles of PMOs will certainly be connected to theenvironmental habitat of the bacteria, and clues to function will no doubt be found inthese habitats as well as in interactions with other organisms in these natural habitatsor with hosts. Mechanisms of host-microbe interactions and pathogenesis will certainlyemerge from these studies. Ultimately, a better understanding of PMO expression andfunction could lead to a deeper understanding of the molecular and physiologicalmechanisms of bacteria for biomass conversion, feeding strategies, as well as symbioticrelationships.
ACKNOWLEGMENTS
We thank Tyler Detomasi, Yirui Guo, Ben Horst, Christopher Lemon, ElizabethNdontsa, Minxi Rao, and Elise Span for critical reading of the manuscript.
This review was prepared by M.A., J.A.H., and M.A.M. M.A. and J.A.H. conducted theprimary literature search and contributed to the development, organization, andwriting of the manuscript. M.A.M. contributed to the organization, writing, and editingof the manuscript. All authors read and approved the final manuscript.
We declare no conflict of interest.
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Marco Agostoni obtained his B.Sc. in marinebiology with an emphasis on microalgalphysiology from the Marche PolytechnicUniversity in Italy. He then moved to theMarine Science Institute at the University ofTexas at Austin, where he received his M.Sc.in Marine Science, focusing on harmful algalblooms. During his Ph.D. in the Molecularand Cell Biology Program and the MSU/DOEPlant Research Lab at Michigan State Univer-sity, he worked on identifying the regulatoryroles of the second messenger cyclic di-GMP in cyanobacteria. After heobtained his Ph.D. in May 2015, he began his postdoctoral fellowship atthe University of California, Berkeley, where he is unraveling the phys-iological role of PMOs during host-pathogen interactions and in theenvironment. The knowledge gained will help expand the understand-ing of the possible functional roles of bacterial PMOs. His broaderinterests lie in leveraging microorganisms as effective tools for practicalapplications.
John A. Hangasky obtained bachelor’s ofscience degrees in chemistry and forensicscience from the University of New Haven inNew Haven, CT. He then pursued graduatestudies at the University of Massachusetts,Amherst, where he received his Ph.D. in chem-istry in 2014, based on his work characterizingalpha-ketoglutarate-dependent oxygenasesresponsible for mediating the hypoxic re-sponse of the transcription factor HIF (hyp-oxia inducible factor). Following his Ph.D., hejoined the laboratory of Michael A. Marletta at the University of Cali-fornia, Berkeley, where he is currently a postdoctoral research associate.His current research interests range from identifying and characterizingnovel PMOs and the polysaccharides that they degrade to studying themolecular mechanisms of PMOs.
Michael A. Marletta earned an A.B. in chem-istry and biology from Fredonia, State Uni-versity of New York. After completion of aPh.D. at UCSF with George Kenyon and apostdoctoral fellowship at MIT with ChrisWalsh, Dr. Marletta joined the faculty at MIT,where investigations into the mammalianbiosynthesis of nitrate led to the laboratory’sfindings on nitric oxide (NO). He continuedstructure-function studies involving NO atthe University of Michigan and now at theUniversity of California, Berkeley. The emergence of the Energy Biosci-ences Institute at Berkeley led the Marletta group toward studies intothe enzymology of cellulose degradation, subsequently leading to thejoint discovery of the polysaccharide monooxygenases. Those studiescontinue and have been expanded to include PMOs in pathogens.
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