Plant-Polysaccharide-Degrading Enzymes from Basidiomycetes
Johanna Rytioja,a Kristiina Hildén,a Jennifer Yuzon,b* Annele Hatakka,a Ronald P. de Vries,b,c Miia R. Mäkeläa
Department of Food and Environmental Sciences, Division of Microbiology and Biotechnology, University of Helsinki, Helsinki, Finlanda; Fungal Physiology, CBS-KNAW
Fungal Biodiversity Centre, Utrecht, The Netherlandsb; Fungal Molecular Physiology, Utrecht University, Utrecht, The Netherlandsc
SUMMARY
Basidiomycete fungi subsist on various types of plant material in
diverse environments, from living and dead trees and forest litter
to crops and grasses and to decaying plant matter in soils. Due to
the variation in their natural carbon sources, basidiomycetes have
highly varied plant-polysaccharide-degrading capabilities. This
topic is not as well studied for basidiomycetes as for ascomycete
fungi, which are the main sources of knowledge on fungal plant
polysaccharide degradation. Research on plant-biomass-decaying
fungi has focused on isolating enzymes for current and future
applications, such as for the production of fuels, the food industry,
and waste treatment. More recently, genomic studies of basidiomycete fungi have provided a profound view of the plant-biomass-degrading potential of wood-rotting, litter-decomposing,
plant-pathogenic, and ectomycorrhizal (ECM) basidiomycetes.
This review summarizes the current knowledge on plant polysaccharide depolymerization by basidiomycete species from diverse
614
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habitats. In addition, these data are compared to those for the
most broadly studied ascomycete genus, Aspergillus, to provide
insight into specific features of basidiomycetes with respect to
plant polysaccharide degradation.
INTRODUCTION
P
lant biomass is the most abundant renewable carbon source
on Earth. Many microbes have central roles in the degradation
of this biomass to ensure a global carbon cycle. Fungi are specialAddress correspondence to Miia R. Mäkelä, miia.r.makela@helsinki.fi.
*Present address: Jennifer Yuzon, Phytophthora Genomics Laboratory, University
of California, Davis, California, USA.
Copyright © 2014, American Society for Microbiology. All Rights Reserved.
doi:10.1128/MMBR.00035-14
Microbiology and Molecular Biology Reviews
p. 614 – 649
December 2014 Volume 78 Number 4
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SUMMARY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .614
INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .614
PLANT CELL WALL POLYSACCHARIDES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .615
Cellulose. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .616
Hemicellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .616
Pectin. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .616
ENZYMES MODIFYING PLANT POLYSACCHARIDES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .616
Cellulose Degradation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .616
Hemicellulose Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .618
Pectin Degradation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .618
Debranching Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .619
BASIDIOMYCETE GENOMES AND PLANT POLYSACCHARIDE DEGRADATION. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .619
Wood-Rotting Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .619
White rot fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .619
Brown rot fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .623
Litter- and Straw-Decomposing Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .623
Ectomycorrhizal Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .624
Plant-Pathogenic Fungi and Mycoparasites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .624
Basidiomycete Yeasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .625
Comparison of the Genomes of Basidiomycetes and Aspergillus as a Representative of the Plant-Biomass-Degrading Ascomycetes . . . . . . . . . . . . . . . . . . .625
Genes related to cellulose degradation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .625
Genes related to hemicellulose degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .625
Genes related to pectin degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .625
CHARACTERIZED PLANT CELL WALL POLYSACCHARIDE-DEGRADING ENZYMES IN BASIDIOMYCETES AND ASPERGILLUS. . . . . . . . . . . . . . . . . . . . . . . . . . . .625
Cellulose-Degrading Enzymes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .625
Hemicellulose-Degrading Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .630
Xylan degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .630
Mannan degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .631
Pectin-Degrading Enzymes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .633
Hemicellulose- and Pectin-Debranching Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .634
REGULATION OF PLANT POLYSACCHARIDE DEGRADATION IN BASIDIOMYCETES AND ASPERGILLUS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .636
Repression of Gene Expression in Basidiomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .637
Induction of Gene Expression in Basidiomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .637
CONCLUSIONS AND FUTURE PROSPECTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .637
ACKNOWLEDGMENTS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .638
REFERENCES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .638
AUTHOR BIOS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .648
Plant Polysaccharide Degradation by Basidiomycetes
December 2014 Volume 78 Number 4
FIG 1 Simplified model of plant cell wall structure. (A) The structure consists
of three main layers: the middle lamella and the primary and secondary walls.
(A and B) The main polysaccharides and lignin which form the surrounding
structure for the plasma membrane are presented in the primary (B) and
secondary wall (C). The lignin content in the primary cell wall (not illustrated)
varies considerably depending on the plant species (Table 1). The illustrations
are not to scale.
ies. Finally, the so far poorly addressed regulatory mechanisms of
basidiomycetes in plant cell wall degradation are reviewed.
PLANT CELL WALL POLYSACCHARIDES
The three most important polysaccharide building blocks of plant
cell walls are cellulose, hemicellulose, and pectin. Together with
lignin, an aromatic heteropolymer, they form a degradation-resistant and functional complex that provides rigidity and structure
to the plant and protects the cells from microbial attack. The plant
cell wall consists of three main layers: the middle lamella and the
primary and secondary walls (Fig. 1A) (20, 21). Each of these
layers has a unique structure and chemical composition that also
differ strongly between plant species, tissues, and the growth
phase of the plant (Fig. 1B and C).
The major differences in the chemical compositions of softwood (e.g., pine and spruce) and hardwood (e.g., birch, aspen,
and oak) are in the structure and content of hemicelluloses (Table
1). Hemicelluloses in softwood consist mainly of galactoglucomannans, whereas the majority of hardwood hemicelluloses are
glucuronoxylans (Table 1) (20). On average, softwood has higher
lignin content than hardwood, while the amount of cellulose in
softwood is smaller than that in hardwood (Table 1) (20).
The chemical compositions of cell walls in flowering plants also
vary (Table 1). Monocots, i.e., grasses, are considered the most
important renewable-energy crops, and their primary cell wall
consists mainly of cellulose and hemicelluloses, whereas their secondary walls contain larger amounts of cellulose, a different composition of hemicelluloses, and significant amounts of lignin (Table 1) (22). The primary cell walls of dicots differ from those of
grasses by their low xylan and high xyloglucan and mannan con-
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ized to use plant biomass as a carbon source by producing enzymes that degrade plant cell wall polysaccharides into metabolizable sugars. Plant-polysaccharide-depolymerizing enzymes are of
great interest to biotechnology, as the products of their catalysis
can be used as precursors in the processes that generate bio-based
products, e.g., fuels, paper, food, animal feed, and chemicals (1).
The enzymes degrading or modifying plant polysaccharides are
classified as carbohydrate-active enzymes (CAZymes) and are divided into families according to their amino acid sequence and structural similarity (2). The CAZy database (http://www.cazy.org/) is organized into families of glycoside hydrolases (GHs), carbohydrate
esterases (CEs), polysaccharide lyases (PLs), glycosyltransferases
(GTs), and auxiliary activities (AA) (2).
Basidiomycetes colonize or inhabit a diversity of plant material
in forests, meadows, farmlands, and compost. Different species
have various CAZyme sets to meet the needs of their ecological
roles as saprobes (wood-rotting and litter-decomposing fungi),
symbionts and endophytes (mycorrhizas and lichens), parasites,
and plant and animal pathogens (3, 4). Basidiomycetes are the
most efficient degraders of woody biomass (5) and therefore are
essential for the global carbon cycle. The understanding of the
mechanisms that basidiomycetes use for plant polysaccharide
degradation is in its infancy compared to ascomycete studies,
due largely to the traditional and well-established industrial
relevance of several ascomycetes. Since the enzyme sets of basidiomycetes are likely to reflect adaptation to their unique
natural niches, basidiomycetes contain a huge potential for
applications in various industries, which has so far remained
largely unexplored.
As mentioned above, our knowledge of basidiomycetes regarding their ability to decompose plant polysaccharides is limited
compared to the wealth of information on ascomycetes. Before
the genomics era, functional analyses of purified enzymes and
expression studies of the corresponding genes were the main approaches for characterization of the fungal CAZyme machinery.
However, these methods are laborious and cannot provide a full
overview of a fungal CAZyme arsenal. More detailed insights into
the entire polysaccharide-degrading capability of fungi with interesting ecologies have been obtained through genome sequencing
(6–15) together with transcriptome and proteome analyses (16–
18). However, only by combining these omics data with biochemical characteristics of the enzymes can we complete our understanding of the plant cell wall polysaccharide degradation ability
of basidiomycete fungi.
This review explores the enzymatic potential of basidiomycetes
from different biotopes and focuses on their ability to depolymerize cellulose, hemicelluloses, and pectin. The basidiomycetes are
compared to species belonging to Aspergillus, which is one of the
most extensively studied ascomycete genera, to dissect differences
in their strategies for plant polysaccharide degradation. While
there is also a large diversity among the ascomycete fungi, the
aspergilli are among the few ascomycetes that have been studied
with respect to the degradation of all plant polysaccharides (19).
First, a comparison of the putative CAZyme-encoding genes
found in the genomes of wood- and litter-decomposing basidiomycetes, plant pathogens, and ectomycorrhizal (ECM) fungi gives
insight into their plant cell wall polysaccharide-degrading enzyme
potential. Second, previously characterized CAZymes isolated
from basidiomycetes are compared to those from genomic stud-
Rytioja et al.
TABLE 1 Approximate chemical compositions of softwood, hardwood, monocot, and dicot plant cell wallsa
Chemical composition (% dry wt)b
Hemicelluloses
Plant material
Cellulose
Mannan
Xylan
-Glucan
Xyloglucan
Pectin
Lignin
Softwood
Hardwood
33–42
38–47
10–15
2–5
5–11
15–30
—
—
—
—
—
—
27–32
21–31
Monocots
Primary
Secondary
20–30
35–45
Minor
Minor
20–40
40–50
10–30
Minor
1–5
Minor
5
Minor
Minor
20
Dicots
Primary
Secondary
15–30
45–50
5–10
3–5
5
20–30
ND
ND
20–25
Minor
20–30
Minor
Minor
7–10
b
Data were obtained from references 20 and 22.
—, not reported; ND, not detected.
tents (Table 1) (22). In addition, the amount of pectin is notably
larger in dicots than in grasses (Table 1). The secondary wall of
dicots is composed of cellulose, hemicelluloses, and lignin (Table
1) (22).
Cellulose
Cellulose, found in both the primary and secondary cell walls, is
the most abundant polysaccharide in plant matter (40 to 45% dry
weight) and gives the plant cell wall its rigid structure (20). Repeating units of -1,4-linked D-glucose form linear cellulose
chains, which are held together by intermolecular hydrogen bonds
and create linear crystalline structures (microfibrils) (23) and less
crystalline, amorphous regions. The ratio of crystalline to amorphous regions varies between the layers of primary and secondary
cell walls as well as between plant species. Cellulose microfibrils
are more irregularly ordered in the outer layer than in the inner
layer of the primary cell wall, where they are perpendicularly oriented (Fig. 1). Furthermore, the angles and directions of the cellulose microfibrils vary among the three sublayers (sublayer 1 [S1]
to S3) of the secondary plant cell wall (20, 21).
Hemicellulose
Hemicelluloses (20 to 30% plant dry weight) support the structure
of the cellulose microfibrils in the primary and secondary walls of
plant cells (20). There are four types of amorphous hemicellulose
structures with different main monosaccharide units in their
hemicellulose backbone. Xylan is the most common hemicellulose polymer with a -1,4-linked D-xylose backbone. Other hemicelluloses are xyloglucan (-1,4-linked D-glucose), found mainly
in the primary walls; -glucan (-1,3;1,4-linked D-glucose); and
mannan (-1,4-linked D-mannose) (21). Xylan, xyloglucan, and
mannan backbones are decorated with branched monomers
and short oligomers consisting of D-galactose, D-xylose, L-arabinose, L-fucose, D-glucuronic acid, acetate, ferulic acid, and p-coumaric acid that are cleaved by debranching enzymes (24).
Pectin
Pectin is a noncellulosic polysaccharide containing galacturonic
acid that provides additional cross-links between the cellulose and
hemicellulose polymers. It is found mainly in plant primary cell
walls and middle lamella (25). The pectin concentration in the
middle lamella is high at an early stage of plant growth, but the
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concentration decreases during lignification (20). The simplest
pectin structure is homogalacturonan (HG), which is a linear
polymer of ␣-1,4-linked D-galacturonic acid residues that can be
methylated at the C-6 carboxyl group and acetylated at the O-2 or
O-3 position. Xylogalacturonan (XGA) is a substituted galacturonan that has -1,3-linked D-xylose residues attached to the galacturonic acid backbone. The second substituted galacturonan is
rhamnogalacturonan II (RG-II). The structure of RG-II is more
complex than the structure of XGA. Altogether, 12 different glycosyl residues, e.g., 2-O-methyl xylose, 2-O-methyl fucose, aceric
acid, 2-keto-3-deoxy-D-lyxo heptulosaric acid, and 2-keto-3-deoxy-D-manno-octulosonic acid, can be attached to the galacturonic acid backbone (25). The most complex pectin structure,
rhamnogalacturonan I (RG-I), has a backbone of alternating Dgalacturonic acid and L-rhamnose residues, with branching structures consisting of D-galactose and L-arabinose chains attached to
the L-rhamnose residues.
ENZYMES MODIFYING PLANT POLYSACCHARIDES
An overview of the known fungal plant-polysaccharide-degrading
or -modifying enzymes is presented in Table 2. The enzymes are
divided according to their substrates, and their EC numbers, abbreviations, and corresponding CAZyme families (2) are also
shown.
Cellulose Degradation
The main enzymes that hydrolyze cellulose, so-called classical cellulases, are endoglucanases, exoglucanases, and -glucosidases
(BGLs). -1,4-Endoglucanase (EG) (EC 3.2.1.4) cleaves within
the cellulose chains to release glucooligosaccharides (Fig. 2A).
Exoglucanases or cellobiohydrolases (CBHs) release cellobiose
from the end of the cellulose chains. The two types of cellobiohydrolases, CBHI and CBHII (EC 3.2.1.176 and EC 3.2.1.91, respectively), degrade cellulose from either the reducing or the nonreducing end, respectively, with different processivities, i.e., the
efficiency of the sequential hydrolysis of the -1,4-glycosidic
bonds by the cellulase before the dissociation of the enzyme from
the substrate (26). BGL (EC 3.2.1.21) releases the smallest unit,
glucose, from shorter oligosaccharides.
Recently, oxidoreductive cleavage of the cellulose chain has
been reported. Cellobiose dehydrogenase (CDH) (EC 1.1.99.18)
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Plant Polysaccharide Degradation by Basidiomycetes
TABLE 2 Plant-polysaccharide-degrading enzymes
Substrate
Enzyme activity
EC no.a
Abbreviation
CAZyme family(ies)
Cellulose
-1,4-Endoglucanase
Cellobiohydrolase (reducing end)
Cellobiohydrolase (nonreducing end)
-1,4-Glucosidase
Cellobiose dehydrogenase
Lytic polysaccharide monooxygenase
3.2.1.4
3.2.1.176
3.2.1.91
3.2.1.21
1.1.99.18
NA
EG
CBHI
CBHII
BGL
CDH
LPMO
GH3, -5, -6, -7, -9, -12, -45
GH7
GH6
GH1, -3
AA3_1, AA8
AA9
Xylan
-1,4-Endoxylanase
Xylobiohydrolase
-1,4-Xylosidase
3.2.1.8
3.2.1.–
3.2.1.37
XLN
XBH
BXL
GH10, -11
Galactomannan
-1,4-Endomannanase
-1,4-Mannosidase
-1,4-Galactosidase
␣-1,4-Galactosidase
␣-Arabinofuranosidase
Galactomannan acetyl esterase
3.2.1.78
3.2.1.25
3.2.1.23
3.2.1.22
3.2.1.55
3.1.1.–
MAN
MND
LAC
AGL
ABF
GMAE
GH5, -26
GH2
GH2, -35
GH27, -36
GH51, -54
Xyloglucan
Xyloglucan -1,4-endoglucanase
␣-Arabinofuranosidase
␣-Xylosidase
␣-Fucosidase
␣-1,4-Galactosidase
-1,4-Galactosidase
3.2.1.151
3.2.1.55
3.2.1.177
3.2.1.51
3.2.1.22
3.2.1.23
XEG
ABF
AXL
AFC
AGL
LAC
GH12, -74
GH51, -54
GH31
GH29, -95
GH27, -36
GH2, -35
Arabinoxylan
Arabinoxylan arabinofuranohydrolase/arabinofuranosidase
␣-Glucuronidase
␣-1,4-Galactosidase
-1,4-Galactosidase
Acetyl xylan esterase
Feruloyl esterase
3.2.1.55
3.2.1.139
3.2.1.22
3.2.1.23
3.1.1.72
3.1.1.73
AXH
AGU
AGL
LAC
AXE
FAE
GH62
GH67, -115
GH27, -36
GH2, -35
CE1, -5
CE1
Pectin
Endopolygalacturonases
Exopolygalacturonases
Xylogalacturonan hydrolase
Endorhamnogalacturonase
Exorhamnogalacturonase
Rhamnogalacturonan rhamnohydrolase
␣-Rhamnosidase
␣-Arabinofuranosidase
Endoarabinanase
Exoarabinanase
-1,4-Endogalactanase
Unsaturated glucuronyl hydrolase
Unsaturated rhamnogalacturonan hydrolase
-1,4-Xylosidase
-1,4-Galactosidase
Pectin lyase
Pectate lyase
Rhamnogalacturonan lyase
Pectin methyl esterase
Pectin acetyl esterase
Rhamnogalacturonan acetyl esterase
Feruloyl esterase
3.2.1.15
3.2.1.67
3.2.1.–
3.2.1.171
3.2.1.–
3.2.1.174
3.2.1.40
3.2.1.55
3.2.1.99
3.2.1.–
3.2.1.89
3.2.1.–
3.2.1.172
3.2.1.37
3.2.1.23
4.2.2.10
4.2.2.2
4.2.2.23
3.1.1.11
3.1.1.–
3.1.1.–
3.1.1.73
PGA
PGX
XGH
RHG
RHX
RGXB
RHA
ABF
ABN
ABX
GAL
UGH
URH
BXL
LAC
PEL
PLY
RGL
PME
PAE
RGAE
FAE
GH28
GH28
GH28
GH28
GH28
GH78
GH51, -54, -62
GH43
GH93
GH53
GH88
GH105
GH3, -43
GH2, -35
PL1
PL1, -3, -9
PL4, -11
CE8
CE12
CE1
NA, not categorized by the International Union of Biochemistry and Molecular Biology (IUBMB).
and lytic polysaccharide monooxygenases (LPMOs) participate in
cellulose degradation in combination with cellulases (Fig. 2A) (27,
28). CDH is the only known extracellular flavocytochrome that
oxidizes cellobiose and cellooligosaccharides to the corresponding
lactones (29, 30). The exact role of CDH in lignocellulose degradation is still unclear, although there is evidence of its relevance in
December 2014 Volume 78 Number 4
both the cellulolytic and lignin-modifying machinery of fungi (29,
30). The ability of CDH to produce hydroxyl radicals through
Fenton chemistry supports its role in lignin modification, while
oxidation of cellobiose together with the production of electrons
for LPMO-catalyzed cellulose depolymerization demonstrate the
participation of CDH in the degradation of cellulose (29, 31, 32).
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GH3, -43
Rytioja et al.
(C) -glucan; (D) heteroxylan; (E) heteromannan; (F) pectin. Enzyme abbreviations are presented in Table 2. Polysaccharide structures were drawn by using data
reported previously by Mohnen (203) and Doblin et al. (204).
LPMOs are copper monooxygenases that catalyze the direct oxidation of the cellulose chain leading to cleavage of the glycosidic
bond (28, 31, 32). Moreover, fungal LPMOs can be divided into at
least three classes according to their sequence similarity and specific activities toward cellulose (33). Type 1 LPMOs catalyze oxidation of the glucose unit at the C-1 position, resulting in the
formation of aldonic acids at the reducing end of the cellulose
chain (28, 32). Type 2 LPMOs generate ketosugars at the nonreducing end of the cellulose chain by oxidizing at the C-4 position
(34). LPMOs of type 3 are not as specific as type 1 or 2 enzymes,
and they are able to oxidize both positions (32). Oxidation at C-6
has also been proposed (28). The reaction catalyzed by LPMOs
requires an electron donor to reduce copper II to copper I in the
active site of the enzyme and molecular oxygen to form the copper-oxygen complex, which is capable of oxidizing the glycosidic
bond (35). In addition to the above-mentioned CDH, other naturally occurring electron donors for LPMOs have been proposed,
e.g., gallic acid or lignin (28, 36). Also, several compounds, e.g.,
ascorbic acid, have been shown to act as reductants in LPMO
catalysis in vitro (28, 34).
Hemicellulose Degradation
Due to variable structures, a specific set of CAZymes is needed to
degrade the backbone and branching structures of each hemicellulose (Fig. 2B to E) (37). The xylan backbone is cleaved by -1,4-
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endoxylanase (XLN) (EC 3.2.1.8) into shorter oligomers (Fig.
2D). A xylobiohydrolase that hydrolyzes xylan into xylobiose has
also been described (38). -1,4-Xylosidase (BXL) (EC 3.2.1.37)
hydrolyzes xylobiose into its monomeric units and also releases
D-xylose from larger xylooligosaccharides from the nonreducing
terminus (24, 39). The xyloglucan backbone, the structure of
which is similar to that of cellulose, is hydrolyzed by EGs, CBHs,
and BGLs (Fig. 2B) (24). -Glucan can be degraded by EGs into
oligosaccharides (Fig. 2C). The -1,4-linked D-mannose backbone of mannan is cleaved by -1,4-endomannanase (MAN) (EC
3.2.1.78) to mannooligosaccharides (Fig. 2E). -1,4-Mannosidase
(MND) (EC 3.2.1.25) releases D-mannose from the terminal ends
of mannan (24). In addition, BGL acts on the galactoglucomannan backbone.
The enzymatic oxidative cleavage of hemicelluloses was recently
confirmed (40). First, the ability of CDH to accept electrons from
xylooligosaccharides and interact with various LPMOs was detected, suggesting that these enzymes are able to act on hemicelluloses (41). Recently, LPMO9C of the ascomycete fungus Neurospora crassa was shown to cleave xyloglucan, -glucan, and, to a
lesser extent, glucomannan with ascorbic acid as a reductant (40).
Pectin Degradation
Endopolygalacturonases (PGAs) (EC 3.2.1.15) and exopolygalacturonases (PGXs) (EC 3.2.1.67) act within and at the terminal end
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FIG 2 Schematic representation of plant cell wall polysaccharides and selected corresponding polysaccharide-degrading enzymes. (A) Cellulose; (B) xyloglucan;
Plant Polysaccharide Degradation by Basidiomycetes
Debranching Enzymes
The enzymes described above cleave the main chains of cellulose
and the backbone and branches of hemicellulose and pectin.
However, smaller side branches extending from hemicellulose
and pectin require a different set of CAZymes. The debranching
enzymes (also known as accessory enzymes) ␣-D-xylosidase
(AXL) (EC 3.2.1.177), ␣-L-arabinofuranosidase (ABF) (EC
3.2.1.55), arabinoxylan arabinofuranohydrolase (AXH), endoarabinase (ABN), exoarabinase (ABX), ␣-D-galactosidase (AGL) (EC
3.2.1.22), -D-galactosidase (LAC) (EC 3.2.1.23), endogalactanase (GAL) (EC 3.2.1.89), exogalactanase (EC 3.2.1.–), ␣-glucuronidase (AGU) (EC 3.2.1.139), feruloyl esterase (FAE) (EC
3.1.1.73), p-coumaroyl esterase (pCAE) (EC 3.1.1.–), acetyl xylan
esterase (AXE) (EC 3.1.1.72), galactomannan acetyl esterase
(GMAE) (EC 3.1.1.–), rhamnogalacturonan acetyl esterase
(RGAE) (EC 3.1.1.–), pectin acetyl esterase (PAE) (EC 3.1.1.–),
and pectin methyl esterase (PME) (EC 3.1.1.11) work synergistically with the main-chain-depolymerizing enzymes to degrade
plant polysaccharides (19).
BASIDIOMYCETE GENOMES AND PLANT POLYSACCHARIDE
DEGRADATION
To date, an increasing number of basidiomycete genomes have
been sequenced and annotated to understand fungal physiology
and, in several cases, to search for enzymes of interest that could be
of use in industrial applications (Table 3) (42). These fungi inhabit
a wide range of ecological niches and colonize various growth
substrates, such as conifers, deciduous trees, forest litter, crops,
grassland soils, and roots of plants. Differences in the CAZyme
sets can often be linked to fungal habitat. For example, the wooddecaying white rot fungus Phanerochaete chrysosporium has a
larger repertoire of plant cell wall polysaccharide-degrading enzymes than the biotrophic phytopathogen Ustilago maydis, which
possesses a minimal set of CAZyme-encoding genes in order to
prevent host plant defense responses, as suggested in previous
studies (6, 8). While it cannot be automatically concluded that an
increase in the number of genes related to a particular polysaccharide also means an improved degradation of this polysaccharide,
many studies have revealed such correlations (43–50). However,
there are also clear exceptions to this. The most noteworthy
exception is the ascomycete Hypocrea jecorina (anamorph
Trichoderma reesei), which is a very efficient cellulose degrader but
contains a relatively small number of cellulase-encoding genes in
December 2014 Volume 78 Number 4
each genome. Its strategy appears to have focused on high production levels of a limited set of enzymes rather than expanding its
enzyme repertoire (51). This approach appears to be used by only
a minority of fungi, based on an extensive correlation analysis
between genome content and growth on plant biomass substrates
of ⬎150 fungal species (R. P. de Vries, A. Wiebenga, M. Zhou,
P. M. Coutinho, and B. Henrissat, unpublished data).
Wood-Rotting Fungi
Wood-rotting fungi are traditionally divided into white rot and
brown rot fungi according to the modification that they cause to
wood residue during decay. White rot fungi degrade both lignin
and wood polysaccharides (cellulose and hemicelluloses) so that
the residual wood is white or yellowish, moist, soft, and often
fiber-like. More than 90% of all known wood-rotting basidiomycetes are of the white rot type (52), and they are found more
commonly on angiosperm than on gymnosperm wood species in
nature. Brown rot fungi degrade wood to yield brown, typically
cubical cracks that are easily broken down. Less than 10% of all
known wood-decaying basidiomycete species are classified into
this group, which occurs most often on gymnosperm wood (53).
Interestingly, the analyzed genome sequence data show that many
cellulases of wood-rotting basidiomycetes lack the cellulose binding modules (CBMs) generally considered essential for efficient
cellulose hydrolysis (54). More sequence data are needed to clarify
possible ecological and evolutionary advantages for the occurrence of CBM-less cellulases and other polysaccharide-degrading
enzymes in nature.
Genome information indicates that brown rot fungi evolved
several times from ancestor white rot species (11). Thus, individual brown rot species may have different sets of characteristics left,
which makes this group rather heterogeneous, and some of them
resemble white rot fungi. Genome studies of wood-inhabiting basidiomycetes show that there is a need for a more detailed classification of the rot types, since some fungi, e.g., Botryobasidium
botryosum and Jaapia argillacea, do not fulfill the traditional criteria for dichotomous grouping (55). However, it has been suggested that the definition “white rot” should be reserved for those
fungi that degrade all cell wall polymers through the action of the
lignin-modifying peroxidases and have enzymes capable of attacking crystalline cellulose (55).
White rot fungi. White rot fungi are efficient degraders of the
aromatic polymer lignin and cause a characteristic white appearance on degraded wood (56). White rot fungi also have the most
extensive arsenal of putative CAZymes among the basidiomycetes
(Table 4), allowing them to colonize a wide range of plants, from
pine trees to poplars and grapevines (11). White rot fungi make up
the majority of wood-rotting basidiomycetes, and the most intensively studied species are commonly isolated from hardwoods
(56), which have slightly higher cellulose and hemicellulose (glucomannan and glucuronoxylan) contents than do softwoods (Table 1) (20).
Based on the sequenced genomes (Table 3), the white rot basidiomycetes harbor an extensive set of genes encoding putative
cellulolytic enzymes. Genes encoding GH family 6 (GH6) and
GH7 enzymes, which include mainly cellulose-hydrolyzing CBHs,
are typically present with 1 to 7 copies in all white rot fungal
species sequenced so far (Table 4). As an exception, Pleurotus ostreatus harbors 16 putative GH7-encoding genes (Table 4). Several genes from GH3 and GH5 (6 to 17 and 16 to 43 genes, respec-
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of the ␣-1,4-linked D-galacturonic acid polymer, respectively, releasing D-galacturonic acid from the homogalacturonan backbone (Fig. 2F). Xylogalacturonan is cleaved specifically by xylogalacturonan hydrolases (XGHs) (EC 3.2.1.–). The backbone of
rhamnogalacturonan I is hydrolyzed by exorhamnogalacturonase
(RHX) (EC 3.2.1.–), endorhamnogalacturonase (RHG) (EC
3.2.1.171), rhamnogalacturonan rhamnohydrolase (RGXB) (EC
3.2.1.174), and ␣-rhamnosidase (RHA) (EC 3.2.1.40) (19, 24).
Pectin lyase (PEL) (EC 4.2.2.10), pectate lyase (PLY) (EC
4.2.2.2), and rhamnogalacturonan lyase (RGL) (EC 4.2.2.23) also
cleave the pectin backbone, using a -elimination mechanism.
Lyases have different sensitivities to the acetylations (O-2 or O-3)
or methyl esterifications (O-6) of the D-galacturonic acid backbone. In contrast to pectate lyases, pectin lyases prefer substrates
with a high degree of methyl esterification. Rhamnogalacturonan
lyases favor nonacetylated substrates (19, 24).
Rytioja et al.
TABLE 3 List of basidiomycete species with published genomes and CAZyme annotations
Species
Website(s)
Reference(s)
White rot
Auricularia subglabra
Bjerkandera adusta
Ceriporiopsis (Gelatoporia)
subvermispora
Dichomitus squalens
Fomitiporia mediterranea
Ganoderma lucidum
Ganoderma sp.
Heterobasidion irregulare
Phanerochaete carnosa
Phanerochaete chrysosporium
Phlebia brevispora
Pleurotus ostreatus
Punctularia strigosozonata
Stereum hirsutum
Trametes versicolor
http://genome.jgi.doe.gov/Aurde3_1/Aurde3_1.home.html
http://genome.jgi.doe.gov/Bjead1_1/Bjead1_1.home.html
http://genome.jgi.doe.gov/Cersu1/Cersu1.home.html
11
205
12
http://genome.jgi-psf.org/Dicsq1/Dicsq1.home.html
http://genome.jgi-psf.org/Fomme1/Fomme1.home.html
http://www.herbalgenomics.org/galu/
http://genome.jgi.doe.gov/Gansp1/Gansp1.home.html
http://genome.jgi-psf.org/Hetan2/Hetan2.home.html
http://genome.jgi.doe.gov/Phaca1/Phaca1.home.html
http://genome.jgi-psf.org/Phchr2/Phchr2.home.html
http://genome.jgi.doe.gov/Phlbr1/Phlbr1.home.html
http://genome.jgi.doe.gov/PleosPC15_2/PleosPC15_2.home.html
http://genome.jgi-psf.org/Punst1/Punst1.home.html
http://genome.jgi-psf.org/Stehi1/Stehi1.home.html
http://genome.jgi-psf.org/Trave1/Trave1.home.html
11
11
14
205
66
45
6, 57
205
55
11
11
11
White rot-like
Schizophyllum commune
http://genome.jgi-psf.org/Schco3/Schco3.home.html
15
Uncertain classification
Botryobasidium botryosum
Jaapia argillacea
http://genome.jgi.doe.gov/Botbo1/Botbo1.home.html
http://genome.jgi.doe.gov/Jaaar1/Jaaar1.home.html
55
55
Brown rot
Coniophora puteana
Dacryopinax sp.
Fomitopsis pinicola
Gloeophyllum trabeum
Postia placenta
Serpula lacrymans S7.3
Serpula lacrymans S7.9
Wolfiporia cocos
http://genome.jgi-psf.org/Conpu1/Conpu1.home.html
http://genome.jgi-psf.org/Dacsp1/Dacsp1.home.html
http://genome.jgi-psf.org/Fompi3/Fompi3.home.html
http://genome.jgi-psf.org/Glotr1_1/Glotr1_1.home.html
http://genome.jgi-psf.org/Pospl1/Pospl1.home.html
http://genome.jgi-psf.org/SerlaS7_3_2/SerlaS7_3_2.home.html
http://genome.jgi-psf.org/SerlaS7_9_2/SerlaS7_9_2.home.html
http://genome.jgi-psf.org/Wolco1/Wolco1.home.html
11
11
11
11
18
71
71
11
Litter decomposing
Agaricus bisporus var.
bisporus
Agaricus bisporus var.
burnettii
Galerina marginata
http://genome.jgi-psf.org/Agabi_varbisH97_2/Agabi_varbisH97_2.home.html
9
http://genome.jgi.doe.gov/Agabi_varbur_1/Agabi_varbur_1.home.html
9
http://genome.jgi.doe.gov/Galma1/Galma1.home.html
55
Straw decomposing
Volvariella volvacea
http://www.ncbi.nlm.nih.gov/genome/?term⫽Volvariella⫹volvacea
13
Coprophilic
Coprinopsis cinerea
http://genome.jgi-psf.org/Copci1/Copci1.home.html
206
Plant pathogenic
Melampsora laricis-populina
Puccinia graminis
Ustilago maydis
http://genome.jgi.doe.gov/Mellp1/Mellp1.home.html
http://genome.jgi-psf.org/Pucgr1/Pucgr1.home.html
http://www.broad.mit.edu/annotation/genome/ustilago_maydis/Home.html,
http://mips.gsf.de/genre/proj/ustilago/
10
10
8
Parasitic
Tremella mesenterica
http://genome.jgi-psf.org/Treme1/Treme1.home.html
11
Ectomycorrhiza
Laccaria bicolor
http://genome.jgi-psf.org/Lacbi2/Lacbi2.home.html, http://mycor.nancy.inra
.fr/IMGC/LaccariaGenome/
http://genome.jgi-psf.org/Pirin1/Pirin1.home.html
7
Cryptococcus neoformans var.
grubii
Rhodotorula glutinis
http://genome.jgi.doe.gov/Cryne_H99_1/Cryne_H99_1.home.html
81
http://www.ncbi.nlm.nih.gov/nuccore/AEVR00000000
82
Wallemia sebi
http://genome.jgi.doe.gov/Walse1/Walse1.home.html
207
Piriformospora indica
Yeast
Mold-like
tively) (Table 4), which encode other putative cellulolytic
enzymes, such as BGLs and EGs, occur in all white rot fungi. White
rot fungi also possess a large set of genes encoding putative hemicellulose- and pectin-active enzymes from various CAZyme fam-
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ilies. On average, they have more copies of genes from GH families
10 and 11 (xylan related), 28 (pectin related), 43 (xylan and pectin
related), and 74 (xyloglucan related) and carbohydrate esterase
(CE) families 1 (xylan related) and 12 (pectin related) than other
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Ecology
Plant Polysaccharide Degradation by Basidiomycetes
TABLE 4 Distribution of CAZyme-encoding genes in basidiomycetes and Aspergillus speciesd
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a
No -N-acetylhexosaminidase was included.
-1,4-Endoglucanase and -1,4-endomannanase are included.
c
Can also include models associated with more than one category.
d
Gene numbers are based on previously reported data for the following organisms, and basidiomycete data are updated according to Riley et al. (55): Agaricus bisporus var. bisporus
(9), Aspergillus fumigatus (208), Aspergillus nidulans (49, 209), Aspergillus niger (ATCC 1015) (84, 210), Aspergillus oryzae (211), Auricularia subglabra (11), Bjerkandera adusta
(205), Botryobasidium botryosum (55), Ceriporiopsis subvermispora (12), Coniophora puteana (11), Coprinopsis cinerea (206), Cryptococcus neoformans var. grubii (81), Dacryopinax
sp. (11), Dichomitus squalens (11), Fomitiporia mediterranea (11), Fomitopsis pinicola (11), Galerina marginata (55), Ganoderma lucidum (14), Ganoderma sp. (205), Gloeophyllum
trabeum (11), Heterobasidion irregulare (66), Jaapia argillacea (55), Laccaria bicolor (7), Melampsora laricis-populina (10), Phanerochaete carnosa (45), Phanerochaete chrysosporium
(6), Phlebia brevispora (205), Piriformospora indica (77), Pleurotus ostreatus (55), Postia placenta (18), Puccinia graminis (10), Punctularia strigosozonata (11), Rhodotorula glutinis
(82), Schizophyllum commune (15), Serpula lacrymans 7.9 (71), Stereum hirsutum (11), Trametes versicolor (11), Tremella mesenterica (11), Ustilago maydis (8), Wallemia sebi (207),
Wolfiporia cocos (11), and Volvariella volvacea (13). †, white rot-like; *, ecological classification uncertain; —, not in published papers.
b
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lase-encoding genes by the selective white rot fungus. This shortage and low-level expression of cellulase genes are compensated by
a greater dependence on oxidoreductases, which is in line with the
growth pattern of C. subvermispora showing preference for lignin
depolymerization (12). C. subvermispora grows better on pectin
and guar gum (galactomannan) than on cellulose (12). In fact, C.
subvermispora has more endopolygalacturonase (GH28)-encoding genes (six) than P. chrysosporium (four), but significant differences in the amounts of other pectinolytic genes between these
two white rot species were not detected.
Phanerochaete carnosa, a member of the same genus as P. chrysosporium, is found on softwoods, while most other studied white
rot fungi are typically isolated from hardwood (45). The chemical
compositions of the cell walls of softwoods and hardwoods differ
particularly in their hemicelluloses structures (mainly galactoglucomannans are present in softwood, while glucuronoxylan is the
most abundant hemicellulose in hardwood) and in the slightly
higher lignin contents of softwoods (20). The genome of P. carnosa contains 193 GH gene models, which is higher than the number of gene models in the genome of P. chrysosporium (182 gene
models) (45). When the secretome of P. carnosa grown in cellulose
and spruce wood cultures was analyzed, the fungus produced a
pattern of classical cellulases (GH3 EGs and BGLs and GH6 and -7
CBHs), xylanases (GH10 and -11), debranching hydrolases
(GH43), and glucuronoyl esterases (CE1) together with putative
LPMOs (AA9) that was similar to the pattern produced by P. chrysosporium (59). Interestingly, a GH2 -mannosidase, which was
not detected by proteomic analyses in cellulose or wood cultures
of P. chrysosporium (17, 57), was present in cellulose-containing
cultures of P. carnosa (59). Also, peptides corresponding to a GH5
mannanase were identified in cellulose cultures of P. carnosa. In
addition, P. carnosa grows better (based on radial growth and
mycelium density) on guar gum (galactomannan) than on xylanand pectin-containing substrates (45), thus supporting its preference for softwood bioconversion. Biochemical characterization of
P. carnosa hemicellulases is still needed to confirm a correlation
between growth profiles and enzyme substrate specificities.
Another white rot fungus isolated mainly from softwood,
e.g., western yellow pine (Pinus ponderosa) and old coniferous
trunks (60), Dichomitus squalens, has a CAZyme repertoire typical
of white rot species (11). It is able to grow on cellulose-, pectin-,
and lignin-containing minimal media, and it shows better growth
on galactomannan than on xylan. Together with Fomitiporia
mediterranea, it lacks the CE1 genes encoding putative xylan- and
pectin-debranching enzymes. D. squalens also shows a decreased
ability to grow on pectin than on D-glucose, which is in contrast to
the majority of the species studied so far (11). A recent study
shows that the genes encoding CBHs, LPMOs, and CDH are coexpressed when D. squalens grows on spruce wood and in microcrystalline cellulose (Avicel)-containing cultures. Moreover, the
simultaneous expression of the cdh and lpmo genes emphasizes the
role of oxidative degradation of cellulose together with hydrolytic
cellulases in white rot fungi (61).
Ganoderma lucidum is a wood-decaying white rot species and a
model medicinal fungus traditionally used in Asia. It produces a
large variety of bioactive compounds, thus harboring potential for
medical applications (14). G. lucidum possesses a relatively large
number of genes encoding putative CAZymes, including 288
GHs, compared to other white rot basidiomycetes with all the
major cellulose-, hemicellulose-, and pectin-degrading genes (14,
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wood-rotting and litter-decomposing basidiomycetes (Table 4).
Genes belonging to polysaccharide lyase (PL) families PL3, -9, and
-11 are almost absent, while some species have few representatives
in PL1 and -4. Notably, high numbers of gene copies in PL1 were
annotated for P. ostreatus (Table 4). For the oxidoreductases involved in plant polysaccharide degradation, white rot fungi possess typically 1 copy of a CDH (families AA3_1 and AA8)-encoding gene and up to 29 copies of LPMO (AA9)-encoding genes. In
this respect, J. argillacea resembles white rot fungi, as it harbors
similar numbers of genes encoding CDH and LPMOs (Table 4).
Interestingly, B. botryosum has more genes encoding CDHs and
LPMOs than any white rot fungus sequenced so far (55).
The first basidiomycete genome sequenced is the model white
rot fungus P. chrysosporium (6, 57). Its CAZyme content shows
many similarities to the genomes of other white rot basidiomycetes by carrying, for instance, several genes that encode putative
cellulose-hydrolyzing enzymes (EGs, CBHs, and BGLs) (Table 4),
which enables it to completely degrade cellulose (6). P. chrysosporium secretes CBHI, CBHII, EGs, and BGL when grown on microcrystalline cellulose (Avicel) (58). As these cellulases were not
found in P. chrysosporium under ligninolytic culture conditions,
they do not seem to be constitutively produced (57). In Avicel
cultures of P. chrysosporium, the expression of oxidatively polysaccharide-degrading CDH- and putative LPMO-encoding genes
was detected together with the expression of genes encoding classical cellulases (17). P. chrysosporium is also able to degrade hardwood hemicelluloses into their building blocks (6). Genes encoding hemicellulolytic and pectinolytic enzymes (e.g., GH10
xylanase, a putative GH28 exopolygalacturonase, and a putative
CE1 acetyl xylan esterase) were expressed, and the corresponding
proteins were secreted in both Avicel and carbon-limited liquid
cultures, suggesting constitutive expression of the corresponding
genes (17, 57, 58).
Only a limited number of pectinolytic genes are present in the
genome of P. chrysosporium. For example, pectin/pectate lyase-,
exoarabinanase-, or rhamnogalacturonan hydrolase-encoding
genes were not detected (6). Despite this low pectinolytic potential, P. chrysosporium is able to grow on solid cultures of pectin
substrates with a high degree of methyl esterification, such as
soy, apple, and lemon pectins, possibly producing endopolygalacturonase together with galactan- and arabinan-hydrolyzing 1,4-endogalactanase (GH53), -galactosidase (GH35), and ␣-arabinofuranosidase (GH51) (44). However, poor growth on
rhamnogalacturonan and polygalacturonic acid was observed
(44).
Several studies comparing the plant-polysaccharide-degrading
ability of P. chrysosporium to those of other basidiomycetes have
been conducted. The selective white rot fungus Ceriporiopsis (Gelatoporia) subvermispora, which depolymerizes mainly lignin and
hemicelluloses and leaves cellulose almost intact, has a GH family
distribution similar to that of P. chrysosporium. However, some
key differences between these fungi can be pointed out. C. subvermispora possesses fewer GH3 (including BGL)-encoding genes,
with only six copies in the genome (12), while P. chrysosporium
and the other sequenced white rot species harbor at least 8 genes
(Table 4). Also, modest transcript levels for the genes from
GH5, -6, -7, and -12 were observed during the growth of C.
subvermispora on semisolid aspen wood cultures compared to
those observed during the growth of P. chrysosporium, suggesting
a significant reduction in the expression levels of putative cellu-
Plant Polysaccharide Degradation by Basidiomycetes
December 2014 Volume 78 Number 4
multiple evolutionary steps that have led to these two different
life-styles (11). This can be seen, for example, by the loss of ligninmodifying peroxidases, which has been proposed to have occurred several times, resulting in the divergence of brown rot fungi
in the orders Polyporales (e.g., Fomitopsis pinicola, Postia placenta,
and the plant-parasitic brown rot fungus Wolfiporia cocos) and
Boletales (e.g., Coniophora puteana and Serpula lacrymans) and
species Gloeophyllum trabeum and Dacryopinax sp. (11).
A comparison of the representatives of the different CAZyme
families in each plant-biomass-modifying basidiomycete group
indicates that the brown rot fungi studied up to now possess a
significantly smaller set of plant-polysaccharide-depolymerizing
enzymes than white rot and litter-decomposing fungi (Table 4).
The most obvious reduction in the CAZymes of brown rot fungi
can be seen in the small number of putative CBHs (GH6 and -7)
(18, 70). Only the species of the order Boletales and closely related
to ECM fungi, S. lacrymans and C. puteana, harbor one and four
putative CBH-encoding genes, respectively. Also, the genome of
Postia placenta lacks genes for CBHs and for carbohydrate binding
modules from family 1 (CBM1) and contains only two putative
-1,4-endoglucanase-encoding genes (18). Although the genomes of brown rot fungi contain fewer genes encoding CDHs
(AA3_1 and AA8) and LPMOs (AA9) than those of white rot
fungi, it is possible that these putative oxidoreductases of
brown rot fungi take part in enzymatic cellulose depolymerization (11, 18, 71). However, considering the overall lower number of LPMOs and greater variety in the absence and presence
of CDH in brown rot fungi, this implies that their ability to
utilize oxidized sugars is also more variable than in white rot
fungi. When secretomes from semisolid aspen cultures of
brown rot fungi were analyzed, only C. puteana and G. trabeum
secreted a putative CDH and LPMO, respectively, while none of
these proteins were detected in F. pinicola or W. cocos (11).
The substrate preference of brown rot basidiomycetes for softwoods can also be explained by the characteristics of their hemicellulose-degrading capacity. While hardwoods are known to have
a higher proportion of xylan, softwoods have a higher mannan
content. During the evolution of the brown rot fungal life-style,
the number of genes encoding enzymes assigned to GH10 and -11
(endoxylanases) and CE15 was reduced (18, 70). Therefore,
brown rot fungi have slightly lower numbers of xylanolytic enzymes than white rot fungi. In addition, the genomes of C. puteana, Dacryopinax sp., F. pinicola, P. placenta, and S. lacrymans
lack genes encoding putative acetyl xylan or feruloyl esterases
from CE1. Instead, brown rot fungi grow well on guar gum, which
is a galactomannan similar in structure to softwood cell wall galactomannans (11). Several copies of genes encoding putative
-1,4-endomannanases involved in the degradation of mannan
are present in the genomes of brown rot basidiomycetes, presumably helping them to colonize softwoods.
Litter- and Straw-Decomposing Fungi
Litter- and straw-decomposing basidiomycetes participate significantly in the Earth’s carbon cycle, together with wood-decaying
fungi. The genomes of the litter-decomposing fungus Agaricus
bisporus and the straw-decomposing species Volvariella volvacea
(Table 3) have been sequenced because of their importance as
cultivated mushrooms and in recycling decaying plant matter.
The genomes of two A. bisporus strains show similar gene contents
with respect to plant polysaccharide degradation. The economi-
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62). Similar to most white rot fungi, its genome lacks the genes for
putative pectin lyase, pectate lyase, and rhamnogalacturonan lyase
(PL1, -3, -9, and -11) (14).
Based on morphological features, Auricularia subglabra belongs to a group of so-called jelly fungi. A. subglabra is found on
dead and decaying wood, where it causes white rot (11). Compared to the genomes of other white rot species, the genome of A.
subglabra (formerly deposited as Auricularia delicata in the JGI
database) harbors a large number of GH43 and CE16 genes, which
include putative -1,4-xylosidase-, endoarabinanase-, ␣-L-arabinofuranosidase-, and acetylesterase-encoding activities. Cross
sections of colonized wood demonstrate the ability of A. subglabra
to extensively degrade all the main polymers of the wood cell wall
(11). However, it lacks specific xylan side-chain-hydrolyzing enzymes, such as arabinoxylan arabinofuranohydrolases (11).
Schizophyllum commune is a model basidiomycete for mushroom development (15). It has been classified as a white rot fungus, although it has a limited lignin-degrading capacity and therefore does not correspond to the typical characteristics of white rot
species. Instead, S. commune has one of the most extensive cellulose- and hemicellulose-degrading enzyme sets, and each fungal
CAZyme family related to plant biomass degradation is represented in its genome (Table 4) (15). S. commune is found mainly
on fallen hardwood, but it also colonizes softwood and grass silage. S. commune is rich in GH43 enzyme-encoding genes, which
include -1,4-xylosidase and endoarabinanase, and genes encoding xylan- and pectin-degrading enzymes. Another uncommon
characteristic of S. commune is the wealth of putative pectin-degrading lyases (PL1, -2, and -4), which correlates with high-level
pectinase production (15, 63). This is consistent with the strategy
of S. commune to invade adjacent parenchymatic cells in plant
xylem tissue through pectin-surrounded simple and bordered pits
(15).
The dual life-style of the necrotrophic white rot fungus and
economically important forest pathogen Heterobasidion irregulare
(formerly known as H. annosum, intersterility group P [64]) involves pathogenic and saprobic life-styles, which are reflected in
its genome and transcriptome (65). Similar to saprobes, it has all
the enzymes for digesting cellulose/xyloglucan (GH5, -6, -7, -12,
-27, -29, -45, and -74) and pectin (GH28, -43, -51, -53, -78, and
-105; PL1 and -4; and CE8 and -12). However, the whole CAZyme
arsenal is used only during the saprobic growth phase of H. irregulare, while fewer CAZyme-encoding genes are expressed during
the pathogenic phase (66). This shows that H. irregulare has the
ability to extensively degrade plant material, but the fungus uses
its full CAZyme repertoire only when it becomes less dependent
on its living host (66). Other plant-pathogenic basidiomycetes are
discussed in “Plant-Pathogenic Fungi and Mycoparasites,” below.
Brown rot fungi. Brown rot fungi represent ⬃6 to 7% of the
known wood-rotting basidiomycetes and occur mostly on conifers (gymnosperms), which are softwoods (53). While brown rot
fungi are able to efficiently and rapidly break down wood cellulose
and hemicelluloses, they only modify lignin, mainly by demethoxylation, resulting in a characteristic brown residue of decayed
wood (56). In contrast to the enzymatic approach of white rot
fungi (6, 67), brown rot fungi initiate cellulose breakdown with
highly reactive oxidants, such as low-molecular-weight free radicals, including the hydroxyl radicals formed through the Fenton
reaction (68, 69). The difference in cellulose-depolymerizing abilities between white and brown rot fungi is probably a result of
Rytioja et al.
Ectomycorrhizal Fungi
Mycorrhizal fungi depend largely on their plant symbionts for
their carbon source (76), and thus, they have a less extensive
CAZyme arsenal than the wood-rotting and litter- and straw-decomposing fungi (7, 9, 72, 73). The limited plant-polysaccharidedegrading capability of ECM fungi is a result of evolutionary reduction in CAZyme families (7, 71) to suit their role as root
symbionts. The few CAZymes of ECM fungi are most probably
needed for the modification of cell walls of plant roots in order to
establish contact with their host for nutrient exchange. This is
supported by the tightly controlled expression of putative
CAZyme-encoding genes of Piriformospora indica during the fungal colonization of living plant roots (77). Five LPMO-encoding
genes are upregulated at the prepenetration stage, while GH10-,
GH11-, GH18-, and GH62-encoding genes are induced during
prepenetration, colonization, and postcolonization, thus suggesting a role of GHs in the local secretion of enzymes at the penetration site (77). The reduction in CAZymes is also observed for the
genome of L. bicolor and Paxillus involutus (7, 78). L. bicolor possesses mostly enzymes that modify polysaccharide backbones,
such as -1,4-endoglucanase, polygalacturonases, and -1,4-endomannanses for cellulose, pectin, and galactomannan degradation, respectively, but the number of putative genes encoding accessory enzymes is limited. The most abundant CAZyme family
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acting on the plant cell wall in the genome of L. bicolor is LPMO
(7). P. involutus has a unique enzymatic system, similar to that of
brown rot fungi, to decompose plant biomass (78, 79). Transcriptomic studies of P. involutus have revealed that only one -1,4endoglucanase (GH9) and two LPMO genes are expressed during
growth on plant litter or cellulose (79). The oxidative depolymerization of cellulose in cooperation with CDH or low-molecularweight reducing agents (28, 31) supports the role of LPMOs as
important components of the radical-based cellulose-degrading
mechanism of ECM fungi. We suggest that most CAZyme activities have been lost in ectomycorrhizal fungi as an adaptation to
symbiotic growth on host photosynthate. The CAZyme arsenal of
some ECM basidiomycetes, such as P. indica, reflects their ability
to switch their life-styles from mutualist to saprobe. P. indica associates with living and dead barley roots and a variety of monoand dicotyledonous plants. When exposed to dead plant matter
instead of living plant roots, P. indica upregulates several of its
pectin-related enzymes, thus indicating a switch from a mutualistic to a saprobic life-style (77).
Plant-Pathogenic Fungi and Mycoparasites
Ustilago maydis, Melampsora laricis-populina, and Puccinia
graminis are obligate biotrophic pathogens that derive nutrients
from living plant tissues and are not able to survive without their
hosts. In contrast to the genomes of more aggressive ascomycete
pathogens such as Magnaporthe grisea and Fusarium graminearum,
these basidiomycete pathogens have few genes encoding
CAZymes that are most likely employed for penetrating the cell
surface of the host plant (8, 10, 11, 66). The limited CAZyme set
also reflects the avoidance of extensive damage of the host cell
walls, which can trigger the immune response of the plant (8).
However, the GH5 (including -1,4-endoglucanase and -1,4endomannanase activity)-encoding genes are present in several
copies (up to 29) (Table 4), and they are suggested to modify the
polysaccharide backbones of cellulose and hemicelluloses in order
to loosen the plant cell wall structure and to further facilitate the
entry of fungal hyphae into the host cell. In M. laricis-populinainfected cultures of wheat and barley, cellulose- and hemicellulose-depolymerizing CAZyme-encoding genes were highly upregulated (10). A similar upregulation was detected in poplar
cultures infected with P. graminis (80). This suggests that invading
hyphae of these rust fungi secrete polysaccharide-degrading enzymes to form haustoria on the plant surface (10). However, it is
possible that these obligate biotrophic pathogens possess as-yetunidentified strategies for virulence, such as the unexpected set of
small genes with unknown function detected in the genome of the
corn smut fungus U. maydis (8).
Tremella mesenterica is a wood-degrading fungus and mycoparasite of Peniophora species that is morphologically classified
into the group of jelly fungi. The genome of T. mesenterica has a
limited CAZyme repertoire similar to that of ECM fungi, containing only three genes encoding GH3 (no -N-acetylhexosaminidase included) and no genes encoding GH families 6, 7, 10, 11, 12,
28, 43, and 74 (11). This may reflect the parasitic life-style of T.
mesenterica. However, T. mesenterica and related species might
have an alternative mechanism to degrade plant biomass, but
these species have so far been only scarcely studied.
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cally important white button mushroom A. bisporus var. bisporus
originates from Europe, while A. bisporus var. burnettii grows on
leaf litter in North America (9). Another edible fungus, V. volvacea, is widely cultivated in Asia, where it is grown on rice straw,
cotton waste, and other agricultural by-products (13). In addition, the genome of another litter-decomposing species, Galerina
marginata, was recently reported (55).
A. bisporus, G. marginata, and V. volvacea have a close evolutionary relationship with white rot basidiomycetes and ECM fungi
(9, 55, 72), although their genome content resembles that of white
rot rather than ECM genomes. All these fungi grow on partially
decayed plant matter, have diverse sets of CAZymes, and are able
to cause white rot (Table 4). Although litter- and straw-decomposing fungi and the ECM fungi are taxonomically closely related,
their dissimilar ecological niches have resulted in different
CAZyme repertoires (Table 4). Generally, saprobes are more capable of degrading plant polysaccharides than root symbionts. For
example, the coprophilic fungus Coprinopsis cinerea and the litter
decomposer G. marginata secrete a broader set of plant cell walldegrading enzymes than the ECM fungus Amanita bisporigera
(73).
Nonwoody plant tissues contain relatively large amounts of
pectin (74). In accordance with this, some forest litter-decomposing basidiomycetes have been shown to produce pectinolytic enzymes (75). A. bisporus and G. marginata harbor two putative
pectinolytic enzymes encoding genes from PL1, whereas V. volvacea possesses 11 PL1-encoding genes. Up to 5 CE1, 6 CE5, 3 CE8,
4 CE12, and 11 CE16 genes encoding putative carbohydrate esterases have been found in the genomes of these litter- and strawdecomposing fungi, while the CE1, -5, and -12 genes are missing
from several white and brown rot fungal species (Table 4). While
A. bisporus, G. marginata, and V. volvacea have a wide spectrum of
CE genes, only one gene encoding a putative 4-O-methyl-glucuronoyl methyl esterase (CE15) has been detected in A. bisporus
and G. marginata.
Plant Polysaccharide Degradation by Basidiomycetes
Basidiomycete Yeasts
So far, the genomes of only a few basidiomycete yeast species have
been sequenced and analyzed for CAZymes. These unicellular basidiomycetes usually have a very limited pattern of polysaccharide-degrading enzymes, which has been shown for the genomes
of Cryptococcus neoformans (81) and Rhodotorula glutinis (82).
Similarly, Wallemia sebi, a xerophilic mold-like basidiomycete,
has reduced CAZyme sets (55).
Comparison of the Genomes of Basidiomycetes and
Aspergillus as a Representative of the Plant-BiomassDegrading Ascomycetes
December 2014 Volume 78 Number 4
CHARACTERIZED PLANT CELL WALL POLYSACCHARIDEDEGRADING ENZYMES IN BASIDIOMYCETES AND
ASPERGILLUS
Before the era of genome sequencing, various plant cell wall polysaccharide-degrading enzymes from basidiomycetes were isolated
and characterized at the gene or protein level. Several basidiomycete CAZymes have unique biochemical properties, ranging from
extreme temperature tolerance and pH to bifunctional catalytic
activities. Most of the characterized CAZymes are from the white
rot and litter- or straw-decomposing fungi (Tables 5 to 12). These
fungi have more copies of putative CAZyme-encoding genes than
any other group of basidiomycetes (Table 4). The extensive plantpolysaccharide-degrading ability of white rot fungi stems from
their ecology as the dominant wood-degrading species (56).
Cellulose-Degrading Enzymes
Cellulose, the most abundant plant polymer, is hydrolyzed by the
extensive set of cellulolytic enzymes of basidiomycetes. EGs,
CBHs, and BGLs have been isolated from species that represent
various ecophysiological groups, but most of them belong to
wood-degrading white rot fungi (Fig. 3 and Tables 5 to 7). On
average, the molecular masses of basidiomycete EGs and CBHs are
41 kDa and 53 kDa, respectively (Fig. 3A and Tables 5 and 6).
BGLs may be extracellular or cell wall associated, and their structure can be monomeric or multimeric (88). This is shown by the
high level of variation in their molecular masses, ranging from 36
to 640 kDa (Fig. 3A and Table 7). In general, these cellulases have
acidic pI values, with few exceptions, and acidic pH optima (Tables 5 and 6). The average optimum temperature of the characterized basidiomycete cellulases is between 54°C and 58°C (Fig. 3D
and Tables 5 to 7).
Generally, white rot fungi produce more isoenzymes for plant
polysaccharide degradation than do other basidiomycetes. Isoenzymes of EGs have been isolated from several white rot species and
characterized (Table 5). The molecular mass of the EGs from
white rot fungi ranges from 18 to 78 kDa, and they have acidic pI
values of 4.1 to 5.7. As an exception, EG of the straw-decomposing
fungus V. volvacea has a neutral isoelectric point of 7.7 and also a
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Aspergillus species are widely studied due to their relevance to
human health and economic importance. These species include
the industrial workhorses A. niger and A. oryzae as well as the
opportunistic human pathogen A. fumigatus (83). Therefore,
their genomes were also among the first sequenced fungal genomes. The genomes of aspergilli revealed that these species contain unexpectedly abundant sets of plant-biomass-degrading
genes compared to the previously identified genes and enzymes
(19, 84). This demonstrated that without genome sequence data,
predominantly only the genes and enzymes that are highly expressed and produced under laboratory conditions have been
characterized.
A study of six Aspergillus species (A. clavatus, A. flavus, A. fumigatus, A. niger, A. oryzae, and Neosartorya fischeri [teleomorph
of A. fischerianus]) demonstrated that the genome content and
organization of closely related species are very similar (85). However, differences in the contents of plant-polysaccharide-degrading genes of A. nidulans, A. niger, and A. oryzae have been detected.
A. oryzae has a significantly higher number of xylan- and pectinrelated genes than the other two species. Instead, A. nidulans harbors more galactomannan-related genes than A. niger and A.
oryzae, whereas the number of inulin-related genes is highest in A.
niger (49). While the aspergilli do not provide representative
numbers of genes in CAZyme families for all ascomycete fungi,
their generalistic life-style and ability to degrade every plant polysaccharide (49) make them suitable baselines to use for comparisons with other fungi.
Genes related to cellulose degradation. The overall CAZyme
contents of the genomes of basidiomycetes and Aspergillus species
are similar. They both possess several genes encoding GH5 and -12
EGs. Aspergilli and basidiomycetes have similar numbers of genes
encoding CBHs (GH6 and -7). However, aspergilli have notably
more BGL genes in GH3 than do the basidiomycetes (Table 4).
Interestingly, basidiomycetes harbor genes from GH9, while
aspergilli lack the GH9-encoding genes (Table 4). LPMO-encoding genes are present in most of the basidiomycete and aspergillus
genomes (86), but basidiomycetes have more LPMO gene models
(up to 33) than do the Aspergillus species (7 to 9) (Table 4). Some
ascomycetes have similarly high numbers of LPMOs, e.g., 33 in
Podospora anserina (50).
Genes related to hemicellulose degradation. Generally, aspergilli have more genes in the CAZyme families encoding putative
GH11, GH62, and CE5 enzymes than do wood-decaying white rot
and brown rot basidiomycetes (Table 4). GH11 xylanases are absent from the genomes of brown rot fungi (Table 4). GH11 endoxylanases require a different number of nonsubstituted xylose
residues to be able to cleave xylan than GH10 xylanases (87),
which are present in all basidiomycetes (Table 4). This indicates
that the xylan oligosaccharide profile originating from the action
of brown rot xylanases will be different from that originating from
white rot fungi and Aspergillus, which will affect the overall process of xylan degradation by these fungi. Basidiomycete genomes
almost universally lack the genes encoding GH62 enzymes (Table
4). There are representatives of GH67 and GH93 genes in the
genomes of Aspergillus species, while they are almost missing from
basidiomycete genomes. In contrast, genes encoding GH30 enzymes, e.g., -1,4-exoxylanases, are widely present in basidiomycetes and absent from Aspergillus species. Basidiomycetes have
more genes in CE15 and -16 than do aspergilli.
Genes related to pectin degradation. While pectin is a minor
component of wood, both basidiomycetes and Aspergillus species
possess wide and variable sets of genes encoding pectin-degrading
enzymes. Basidiomycetes and aspergilli have up to 20 and 22
genes, respectively, encoding putative GH28 polygalacturonases
and rhamnogalacturonases (Table 4). All the brown rot fungi and
the white rot species C. subvermispora, P. chrysosporium, and Trametes versicolor lack CE12 genes, which encode putative rhamnogalacturonan acetyl esterases.
Rytioja et al.
TABLE 5 Characterized basidiomycete -1,4-endoglucanases and their biochemical properties
Species
White rot
Cerrena unicolor
Dichomitus squalens
D. squalens
D. squalens
Ganoderma lucidum
G. lucidum
Irpex lacteus
I. lacteus
I. lacteus
I. lacteus
I. lacteus
Phanerochaete chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
Polyporus arcularius
P. arcularius
P. arcularius
Sporotrichum pulverulentumc
S. pulverulentumc
S. pulverulentumc
S. pulverulentumc
S. pulverulentumc
Trametes hirsuta
T. hirsuta
Trametes versicolor
Brown rot
Coniophora cerebella
C. cerebella
Fomitopsis palustris
F. palustris
F. palustris
F. palustris
F. palustris
Gloeophyllum sepiarium
(Lenzites sepiaria)
G. sepiarium
Gloeophyllum trabeum
G. trabeum
G. trabeum
G. trabeum (Lenzites trabea)
Serpula incrassata
S. incrassata
S. incrassata
Piptoporus betulinus
Postia placenta
Straw decomposing Volvariella volvacea
Plant pathogen
Yeast
Polyporus schweinitzii
Sclerotium rolfsii
S. rolfsii
S. rolfsii
Ustilago maydis
Rhodotorula glutinis
Enzymea
NCBI protein
database
Molecular
accession no.b mass (kDa) pI
44
42
56
47
55
43
56
16
En I
En II
En III
GH5
GH5
GH5
GH5
GH12
GH45
GH3
GH5
GH5
En-1
E2-A
E2-B
En-1*
cel5A EG36
cel5A EG38
cel5B EG44
cel12A Cel12A
PcCel45A
CMCase I
CMCase II
cel3A CMCase IIIa
T1
T2a
T2b
T3a
T3b
ThEG
rEG*
AAU12275
AAU12275
AAU12276
BAD98315
A
B
GH5
GH12
GH12
EG47
EG35
cel12
eg2
EGII
BAF49602
EGS
EGT
Cel5A
Cel12A
GH5
GH12
GH45
eg1
EG1
egl1
Endo A
Endo B
Endo C
Egl1
AAG59832
Topt
(°C) Reference(s)
4.0
4.8
4.8
4.8
55
55
55
4.0
4.0
50
50
5.6–5.7
4.9
4.3
5.2
4.4–4.6 68
4.4–4.6 68
4.9
52
5.3
4.7
4.4
4.7
4.2
5.0
50
212
213
213
213
16
16
214
215
216
216
217
58, 90
58, 90
58, 90
58, 91
92
218
218
218, 219
220
220
220
220
220
221
221
222, 223
42
39
40
47
35
4.7
4.2
24
85
3.5
55
4.1
4.2
59
62
4.4
⬍3.6
⬍3.6
⬍3.6
2.6–2.8 3.5
70
50
42
7.7
7.5
55
89, 230
45
52
27
78
4.6
4.2
4.5
4.0
4.0
2.3–3.0
4.0
60
74
50
50
231, 232
233
233
233
93
40
8.6
4.5
50
234
45
41
42
28
29
25
49
57
62
35–40
Cel 25
Cel 49
Cel 57
EG1
GH5
38
36
38
44
28
18
39
36
24
32
37
28
38
37
44
50
30
4.8
4.3
4.1
4.7
4.7
pHopt
3.8
3.1
4.9
70
AAB36147
224
224
225
105
105
106
226
227
108
108
103
103
145
228
228
228
109
229
a
Asterisks indicate a heterologously produced enzyme.
b
See http://www.ncbi.nlm.nih.gov/protein.
c
Anamorph of P. chrysosporium.
626
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Gene
Plant Polysaccharide Degradation by Basidiomycetes
neutral pH optimum (7.5) for the hydrolysis of carboxymethyl
cellulose (CMC) (89). The most comprehensive view of characterized enzymes is from the model white rot fungus P. chrysosporium. Three GH5, one GH12, and one GH45 EG of P. chrysosporium have been biochemically characterized (58, 90–92). GH5
EGs of P. chrysosporium hydrolyze CMC more efficiently than
Avicel (90). GH45 EGs have been characterized only for P. chrysosporium and the plant pathogen U. maydis (92, 93). P. chrysosporium GH45 EG hydrolyzes various glycan substrates, preferring
substrates consisting mainly of -1,3/1,4-glucan (92). These endoglucanases show the common synergistic action with CBHs
from GH6 and -7 (90, 92).
P. chrysosporium has seven CBH-encoding genes, and three of
them have been characterized at the protein level (Table 5). These
isoenzymes work synergistically to cleave cellulose at the reducing
and nonreducing ends (94). Multiple plant-polymer-degrading
isoenzymes produced by one species are hypothesized to have
different biochemical properties, such as the substrate specificity
to enhance the degradation of plant biomass. The three-dimensional crystal structure of P. chrysosporium Cel7D (PDB accession
number 1GPI) (95) shows that the catalytic domain is composed
of a -sandwich structure similar to that of ascomycete GH7
CBHI, first solved for the ascomycetous fungus H. jecorina Cel7A
(PDB accession number 1CEL) (96). The crystal structures reveal
that the cellulose binding tunnels of the CBHIs differ significantly,
thus affecting the accessibility of the substrate to the active site. In
P. chrysosporium Cel7D, the cellulose binding tunnel is more open
December 2014 Volume 78 Number 4
than in H. irregulare Cel7A (HirCel7A) (PDB accession numbers
2YG1 and 2XSP) (97), while H. jecorina Cel7A has the most enclosed structure.
Differences in the three-dimensional structures of the six different P. chrysosporium CBH proteins were revealed by homology
modeling, thus supporting the presence of multiple isoenzymes
with different specificities and catalytic mechanisms (95). A function for the multiplicity of cellulolytic-enzyme-encoding genes is
also supported by their expression at different phases of fungal
growth and degradation of plant biomass. For example, V. volvacea has three GH7 CBHI-encoding genes that are expressed during
different stages of mushroom development (98).
GH1 and GH3 -glucosidases of white rot and straw-decomposing fungi have widely variable molecular masses (from 45 to
640 kDa) and isoelectric points (from 3.3 to 8.5) (Table 7). This
diversity is due to the intra- and extracellular localizations of
-glucosidases. The exceptions of -glucosidases with neutral pI
values are those from Fomes fomentarius, P. ostreatus, P. chrysosporium, and V. volvacea (99–102).
The strategy used by brown rot basidiomycetes to degrade cellulose differs from that used by white rot fungi. Instead of using
cellulolytic enzymes, the brown rot fungi rely on highly reactive
oxidants for initial depolymerization of plant polysaccharides (18,
56, 71). Most brown rots are unable to degrade crystalline cellulose, with the majority preferring amorphous cellulose (88). However, some brown rot species, e.g., G. trabeum and Fomitopsis
palustris, have been shown to degrade crystalline cellulose
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FIG 3 Average molecular masses (kDa for monomers) (A), isoelectric points (pI) (B), pH optima (C), and temperature optima (D) of selected CAZymes from
basidiomycetous (first columns, in dark colors) and Aspergillus species (second columns, in light colors). EG, endoglucanase; CBH, cellobiohydrolase; BGL,
-glucosidase; XLN, endoxylanase; MAN, endomannanase; MND, -mannosidase; AGL, ␣-galactosidase; AGU, ␣-glucuronidase; CDH, cellobiose dehydrogenase. The number of characterized enzymes used for calculation of mean values is marked at the root of each column. Error bars show the minimum and
maximum values reported for each biochemical characteristic. ⫺, no mean value was available.
Rytioja et al.
TABLE 6 Characterized basidiomycete cellobiohydrolases and their biochemical properties
Species
White rot
Dichomitus squalens
D. squalens
D. squalens
Flammulina velutipes
F. velutipes
Ganoderma lucidum
G. lucidum
G. lucidum
Irpex lacteus
I. lacteus
I. lacteus
I. lacteus
I. lacteus
Lentinula edodes
L. edodes
Phanerochaete
chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
Polyporus arcularius
P. arcularius
Brown rot
Gene
Enzyme
NCBI protein
database
accession no.a
Ex 1
Ex 2
GH7
GH7
GH7
GH7
GH7
GH7
GH6
cel7b
cel7a
cel7b
cel1
cel2
cel3
cex3
FvCel7A
FvCel7B
Ex-1
Ex-2
CDJ79665
BAJ07534
BAJ07535
BAA76363
BAA76364
BAA76365
BAG48183
Molecular
mass (kDa)
pI
pHopt
Topt
(°C)
39
36
4.6
4.5
5.0
5.0
60
60
50
60
56
49
50
53
56
5.2
5.0
5.6
4.5
4.8
5.0
5.0
50
50
5.0
50
50
60
65
GH7
GH6
GH7
cel7A
cel6B
cbh1-1
CEL7A
CEL6B
Cel7A
AAK95563
AAK95564
CAA38274
GH7
GH7
cbh1-2
cbh1
Cel7B
Cel7C
CAA38275
AAB46373
62
4.9
GH7
GH7
GH7
GH6
GH6
GH7
GH6
cbh1-4
cbh1-5
cbh1-6/7
Cel7D
Cel7E
Cel7F/G
CBH50
CBHII
AAA19802
58
3.8
50
4.9
52
50
3.6
3.6
cbhII
cel1
cel2
5.0
5.0
4.5
70
4.0
65
107
107
104
Agaricus arvensis
Agaricus bisporus
A. bisporus
GH7
GH6
GH7
cel3AC
cel2
CEL3
HM004552b
AAA50607
CAA90422
Straw decomposing
Volvariella volvacea
V. volvacea
GH7
GH6
cbhI
cbhII
CBHI
CBHII
AAD41096
AAD41097
98
98
Coprophilic
Coprinopsis cinerea
C. cinerea
C. cinerea
C. cinerea
C. cinerea
GH6
GH6
GH6
GH6
GH6
CcCel6A
CcCel6B
CcCel6C
CcCel6D
CcCel6E
CcCel6A
CcCel6B
CcCel6C
BAH08702
BAH08703
BAH08704
BAH08705
BAH08706
248, 249
248, 249
248, 249
249
249
White rot plant
pathogen
Heterobasidion
irregulare
GH7
Plant pathogen
Sclerotium rolfsii
Insect symbiont
Termitomyces sp.
Litter decomposing
HirCel7A
130c
52
235
235
61
236
236
16
16
16
189, 237
189
238
239
240
241
241
95, 192, 242
95, 192, 242
94, 95, 192,
242, 243
94, 95, 244
95
6, 95, 242
94
245
246
246
AAB32942
BAF80326
BAF80327
CBHI
CBHII
Coniophora puteana
C. puteana
Fomitopsis palustris
Reference(s)
50
41.5–42.0
GH6
Cellulase IF
52
4.3
247
186
133
4.0
45
97
4.2
37
250
4.4
251
a
See http://www.ncbi.nlm.nih.gov/protein.
b
Gene identifier.
c
Dimer.
(103–106). The genomes of brown rot fungi harbor EG- but rarely
any CBH-encoding genes, with the exception of the species belonging to the order Boletales, which can also degrade crystalline
cellulose (Table 4). GH5 and GH12 EGs from G. trabeum and F.
palustris have also been characterized (Table 5). In microcrystal-
628
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line cellulose (Avicel) cultures, G. trabeum produces GH5 and
GH12 EGs, of which GH5 EG was shown to hydrolyze Avicel into
cellobiose (103). This processive EG has been suggested to compensate for the lack of CBHs in cellulose degradation of G. trabeum. Only C. puteana and S. lacrymans from the order Boletales
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Plant Polysaccharide Degradation by Basidiomycetes
TABLE 7 Characterized basidiomycete -glucosidases and their biochemical properties
Species
White rot
Ceriporiopsis (Gelatoporia)
subvermispora
C. subvermispora
Fomes fomentarius
Phanerochaete chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
Pleurotus ostreatus
P. ostreatus
P. ostreatus
Trametes gibbosa
Trametes versicolor
Sporotrichum pulverulentumc
S. pulverulentumc
S. pulverulentumc
Stereum hirsutum
White rot-like
Brown rot
110
53
58
90
410
45
114
GH3
GH3
GH3
GH3
GH1
GH1
GH3
Straw decomposing Volvariella volvacea
V. volvacea
V. volvacea
Ectomycorrhiza
Insect symbiont
Yeast
wtBGL
rBGL*
bgl1A BG1A*
bgl1B BG1B*
F1
F2
F3
Sclerotium rolfsii
S. rolfsii
S. rolfsii
S. rolfsii
Ustilago esculenta
AAC26490
AAC26489
BAB85988
BAB85988
BAE87008
BAE87009
A1
A2
B1-3
BGL
116
133
53
60
35
50
66
640
300
165
172
165–182
98
I
II
97
102
96
Cel3A Cel3A*
NR
BG1
Litter decomposing Agaricus bisporus
Plant pathogen
cbgl-1
cbgl-2
Schizophyllum commune
S. commune
S. commune
Fomitopsis palustris
Gloeophyllum trabeum
(Lenzites trabea)
Piptoporus betulinus
Poria vaillantii
Enzymea
NCBI protein
database
Molecular
accession no.b mass (kDa) pI
3.5
60
252
3.5
4.5–5.0
5.5
7.0
5.0
4.0–5.2
60
60
45
45
60
252
100
101
101
253
254
255
255
256, 257
257
113
113
99
99
99
258
185, 259
260
260
260
261
7.5
7.3
8.5
4.0
4.0
5.0
3.5
4.3
4.8
4.0–4.5
4.5
4.0–4.5
4.6–5.2 4.0–4.5
3.4
3.3
70
320
36
BGL-I
BGL-II
2.6
40
50
50
40–50
45
5.3
5.8
5.1
262
263
263
5.0
4.5
60
75
146
145, 264
4.0
4.2
60
50
109
265
—d
5.6
7.0
5.0–5.2 6.2
55–60
55–60
AAG59831
158
256
95
102
102
266
BG-1
BG-2
BG-3
BG-4
UeBgl3A* BAK61808
90
90
107
92
110
4.1
4.6
5.1
5.6
4.2
4.2
4.2
4.2
5.0
68
68
68
68
40
187
187
187
187
267
bgl
GH3
4.7
Topt (°C) Reference(s)
CAC03462
bg1
GH3
6.7
pHopt
Tricholoma matsutake
Pisolithus tinctorius
160
450e
3.8
5.0
4.0
60
65
268
269
Termitomyces clypeatus
T. clypeatus
116
110
4.5
5.0
5.0
45
65
270
271
4.7–5.2 70
3.5
45
188
112
Rhodotorula minuta
Sporobolomyces singularis
GH1
bglA
BglA
BAD95570
144
74
4.8
a
Asterisks indicate a heterologously produced enzyme. wtBGL, wild-type BGL.
See http://www.ncbi.nlm.nih.gov/protein. NR, not reported.
c
Anamorph of P. chrysosporium.
d
Available in the GenBank database (http://www.ncbi.nlm.nih.gov/GenBank/).
e
Trimer.
b
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630 mmbr.asm.org
Biochemical studies of basidiomycete LPMOs are at the early
stage, and most analyses have been conducted only at the gene
level (Tables 4 and 8). In fact, enzymes from the white rot fungi P.
chrysosporium and H. irregulare are the only isolated or characterized basidiomycete LPMOs. The structure of P. chrysosporium
LPMO (GH61D) (PDB accession number 4B5Q) (123) together
with five structures of ascomycete LPMOs, i.e., H. jecorina
(Cel61B) (PDB accession number 2VTC), Thielavia terrestris
(GH61E) (PDB accession numbers 3EII and 3EJA), Thermoascus
aurantiacus (GH61A) (PDB accession number 2YET), and N.
crassa (PMO-2 and PMO-3) (PDB accession numbers 4EIR and
4EIS) LPMOs (27, 28, 33, 124), have opened the path to describing
the biochemical properties of various putative LPMOs harbored
in fungal genomes.
Currently, cellulose-cleaving activities of LPMOs have been biochemically confirmed only for GH61D of the white rot fungus P.
chrysosporium (PcGH61D) (125); the above-mentioned ascomycete LPMOs from H. jecorina, T. terrestris, T. aurantiacus, and N.
crassa; and GH61A and GH61B of the ascomycete fungus P.
anserina (27, 28, 31, 32, 126, 127). PcGH61D is not active on
soluble cellooligosaccharides (125), but it is able to oxidize phosphoric acid-swollen cellulose in the presence of ascorbic acid and
to release lactone, which is spontaneously converted to aldonic
acid (125). The copper-bound active site that is common to LPMOs is present in PcGH61D (123). Nevertheless, it has significant
differences in the loop structures near the binding face compared
to the other characterized LPMO structures, which illustrates the
diversity of the LPMOs.
Hemicellulose-Degrading Enzymes
Xylan degradation. Xylanases break down the most common
hemicellulose found in high quantities in hardwoods and cereals.
Of the isolated and characterized basidiomycete xylan-degrading
enzymes, 75% are from white rot fungi (Table 9). Moreover, basidiomycetes produce several xylanase isoenzymes, which is also
reflected in the high copy number of the corresponding genes
present in their genomes (Table 4). The average molecular mass of
basidiomycete endoxylanases is 45 kDa (Fig. 3A and Table 9).
Their isoelectric points vary from 2.8 to 8.0, and the pH optimum
is between 3.0 and 8.0 (Fig. 3B and C and Table 9). Temperature
optima for endoxylanase reactions range from 50°C to 80°C (Fig.
3D and Table 9).
GH10 and -11 endoxylanases from some white rot species, the
litter-decomposing fungus A. bisporus, and the brown rot fungus
G. trabeum have been characterized (Table 9). Optimum pH values of white rot fungal endoxylanases are most often between 4.0
and 6.0, but an alkaline optimum pH of 8.0 has been reported for
C. subvermispora endoxylanase (128). G. trabeum endoxylanase
(GtXyn10A) also exhibits activity for xyloglucan (129). -Xylosidases have been isolated from only a few white rot fungal species,
including C. subvermispora, G. lucidum, P. chrysosporium, and
Phlebia radiata (Table 9). The GH43 -xylosidase of P. chrysosporium has a molecular mass of 83 kDa (130), whereas the molecular
mass of P. radiata -xylosidase is only 27 kDa (131).
The isoelectric points of xylanases, endoxylanases, and -xylosidases from aspergilli vary substantially (19). Variation in pI values is also seen in basidiomycetous xylanases, even though the
number of characterized enzymes is lower than for ascomycetes
(Table 5) (19). Ascomycete xylanases have different specificities
toward the xylan polymer, some being strongly dependent on the
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possess either one or both GH6 and GH7 CBH gene models (Table
4). The GH6 and GH7 CBHs of C. puteana have also been isolated
and characterized (Table 6) (107).
Both basidiomycetes and aspergilli have a complete set of hydrolytic cellulases, including EGs, CBHIs, CBHIIs, and BGLs.
Only some of the brown rot, plant-pathogenic, and ECM fungi
and basidiomycetous yeasts lack CBHs. Similar to the EGs and
BGLs from Aspergillus species (19), several basidiomycete cellulases are able to degrade the backbone of hemicelluloses (88). EGs
from P. chrysosporium, G. trabeum, Piptoporus betulinus, and Sclerotium rolfsii are able to hydrolyze xylan, galactoglucomannan, or
mannan (91, 108–110). P. chrysosporium EG28 has activity toward
xylan, mannan, and CMC (91), and EG of Aspergillus aculeatus
does not hydrolyze cellulose and releases only xyloglucan oligosaccharides from plant cell walls (111). Basidiomycete BGLs
from both GH1 and GH3 have been characterized (112, 113).
Both basidiomycete and aspergillus BGLs show wide substrate
specificity, and they are able to hydrolyze glucose, mannose,
xylose, or galactose units from the corresponding oligosaccharides (19, 88).
CDHs are widely present in both basidiomycetes and ascomycetes. In contrast to the other plant cell wall polysaccharide-degrading enzymes, CDHs have been more commonly studied in
basidiomycetes than in ascomycetes (114), probably because this
enzyme was first found in the white rot fungus P. chrysosporium
(115). So far, CDHs from 12 different white rot species, the brown
rot fungus C. puteana, the coprophilic fungus C. cinerea, and the
plant pathogen S. rolfsii have been characterized (Table 8). The
average molecular mass of basidiomycete CDHs is 96 kDa (Table
8). The CDHs show acidic isoelectric points (from 3.0 to 6.4) and
pH optima (pH 3.5 to 5). The optimum temperature for CDHcatalyzed reactions varies from 50°C to 75°C (Table 8).
The substrate array of the characterized white rot fungal CDHs
is variable. P. chrysosporium CDH is able to oxidize cellobiose and
higher cellodextrins, lactose, mannobiose, and galactosylmannose
(29). C. subvermispora and Trametes hirsuta CDHs are also able to
oxidize maltose (116, 117), while CDH of Irpex lacteus oxidizes
only cellobiose or higher cellodextrins efficiently (118). In addition, the CDHs of Trametes pubescens and Trametes villosa can
oxidize xylobiose (119). The characterized basidiomycete CDHs
have pH and temperature optima of 3.5 to 5.5 and 50°C to 75°C,
respectively (Table 8). The only characterized brown rot fungal
CDH is from C. puteana. It oxidizes both cellobiose and cellooligosaccharides but not glucose, which supports the typical catalytic
properties of CDH (120, 121).
Basidiomycete and ascomycete CDHs are classified into two
subgroups according to their primary amino acid sequences
(122). Class I includes basidiomycete CDHs, which are shorter
polypeptides than the more complex class II ascomycete CDHs,
which have a C-terminal cellulose binding module. In addition,
the linker regions in basidiomycete CDHs are more conserved
than those in ascomycete CDHs (30). To our knowledge, CDHs
from Aspergillus species have not been characterized at the protein
level to date. The ascomycete CDHs isolated from N. crassa have a
broader substrate spectrum and less glucose discrimination than
basidiomycete CDHs (30). While basidiomycete CDHs catalyze
the reactions at acidic pH, ascomycete CDHs are active at neutral
or alkaline pH (41). Whether the differences in the biochemical
characteristics of CDHs between basidiomycetes and ascomycetes
are also valid for Aspergillus species remains to be clarified.
Plant Polysaccharide Degradation by Basidiomycetes
TABLE 8 Characterized basidiomycete cellobiose dehydrogenases and lytic polysaccharide monooxygenases, their biochemical properties, and their
corresponding genes
Life-style
Species
CAZyme
family(ies) Gene
Enzyme
cdh
pHopt
ACF60617
87–98
3.0
AAC49277
97
89
4.2
CDHII
cdh
cdh
cdh
CDH*
CDHI
CDHII
CDH*
cdh
cdh
cdh
TpCDH
CDH 4.2*
CDH 6.4
TvCDH
GfrCDH
CAA61359
AAC32197
ADX41688
AAC50004
BAC20641
AGE45679
60
116
4.0d
5.0
50d
118
272, 273
4.2
4.2
4.2
89
6.4
4.4
4.5–5.0e
104
113
4.0
4.2
5.0e
4.5
102
Brown rot
Coniophora puteana
111
Coprophilic
Coprinopsis cinerea
AA3_1,
AA8
Plant pathogen
Sclerotium rolfsii
AA3_1,
AA8
Heterobasidion irregulare
H. irregulare
H. irregulare
H. irregulare
H. irregulare
H. irregulare
H. irregulare
H. irregulare
H. irregulare
H. irregulare
Phanerochaete
chrysosporium
AA9
AA9
AA9
AA9
AA9
AA9
AA9
AA9
AA9
AA9
AA9
HiGH61A GH61A
HiGH61B GH61B
HiGH61C
HiGH61D GH61D
HiGH61E
HiGH61F
HiGH61G
HiGH61H
HiGH61I
HiGH61J
PcGH61D*
4.5
3.5–4d
4.5
5.5
4.5
4.5
5.0e
4.5–5.0e
4.5
Schizophyllum commune
Lytic polysaccharide
monooxygenases
White rot
Reference(s)
90
92
81
101
110
92
90
97
White rot-like
CDHcc*
Toptc
(°C)
c
4.2
3.8
5.9
3.8
274–276
277, 278
279
279
70
280
60–70 117
119
55
281–283
75
55
60e
70
281
119
284
285
286
287
3.9
120, 121
⬃80
5.0
101
4.2–5.0 3.2–4.8 55
288, 289
4.8
290
290
290
290
290
290
290
290
290
290
125
AFO72232
AFO72233
AFO72234
AFO72235
AFO72236
AFO72237
AFO72238
AFO72239
25
60
114
a
Asterisks indicate a heterologously produced enzyme.
See http://www.ncbi.nlm.nih.gov/protein.
Activity measured with 2,6-dichlorophenolindophenol (DCIP) and cellobiose.
d
Activity measured with cytochrome c.
e
Activity measured with DCIP and lactose.
b
c
substituents of the xylose residues neighboring the attacked residue and others cutting randomly between the unsubstituted xylose residues (19).
Mannan degradation. Only a few basidiomycete mannanases
have been characterized compared to cellulases and xylanases (Table 10). However, basidiomycetes provide an avenue for the iso-
December 2014 Volume 78 Number 4
lation of unique mannan-degrading enzymes according to their
genomes (Table 4). -Endomannanases have been isolated only
from the white rot fungi P. chrysosporium, T. versicolor, and C.
subvermispora; the litter-decomposing fungus A. bisporus; and the
plant pathogen Sclerotium rolfsii (128, 132–138). However, mannanase activities have also been measured in other white rot fungi,
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Cellobiose dehydrogenase
White rot
Ceriporiopsis
AA3_1,
subvermispora
AA8
Irpex lacteus
Phanerochaete
chrysosporium
P. chrysosporium
Pycnoporus cinnabarinus
P. cinnabarinus
P. cinnabarinus
P. cinnabarinus
Trametes hirsuta
Trametes pubescens
Trametes versicolor
AA3_1,
AA8
T. versicolor
AA3_1
Trametes villosa
Grifola frondosa
Phlebia lindtneri
Pycnoporus sanguineus
AA3_1,
AA8
a
NCBI protein
database
Molecular
accession no.b mass (kDa) pI
Rytioja et al.
TABLE 9 Characterized basidiomycete -1,4-endoxylanases and -xylosidases and their biochemical properties
Life-style
Endoxylanase
White rot
Brown rot
Ceriporiopsis (Gelatoporia)
subvermispora
C. subvermispora
Cerrena unicolor
Ganoderma lucidum
Irpex lacteus
I. lacteus
I. lacteus
Lentinula edodes
L. edodes
L. edodes
Phanerochaete
chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
P. chrysosporium
Phlebia radiata
P. radiata
Pycnoporus cinnabarinus
Schizophyllum commune
S. commune
S. commune
Gloeophyllum trabeum
G. trabeum
G. trabeum
G. trabeum
G. trabeum
Laetiporus sulfureus
Postia placenta
Xylanase I
Xylanase B
Xylanase I
Xylanase III
GH11
GH10
GH10
xyn11A XYN11A*
xynA
XynA*
ABZ88797
GH11
GH10
GH10
GH10
GH10
GH11
xynB
xynC
XynA
XynC
XynA
XynB
XynB*
XynC*
XynA*
XynC*
XYNA*
XYNB*
XA-1
XA-2
ABZ88798
ABZ88799
AEK97220
AEK97221
AAG44992
AAG44995
GH11
GH10
Xylanase A
XynB
XynC
GH10
GH10
GH10
GH10
Xyn10A
Xyn10A*
Xyn10B*
GH10
Coprophilic
Coprinopsis cinerea
GH11
Insect symbiont
Termitomyces clypeatus
Termitomyces sp.
Plant pathogen
Phanerochaete
chrysosporium
Phlebia radiata
Xyl10g
pHopt
Topt
(°C)
Reference
79
8.0
50
128
29
44
31
38
38
62
41
5.0
4.0
60
291
212
16
292
293
293
294
295
296
297
AAL04152
35
52
30
50
50
55
48
37
19
16
50
GH43
xynA
PcXyl
CAB05665
Xyn11C*
AFR33049
XD-1
Sclerotium rolfsii
AFW16059
60
60
70
60
50
50
70
60
70
70
70
6.7
4.1
4.0
5.0
60
297
297
298
299
300
300
131
131
277
33
31
30
2.8
3.6
5.0
5.5
5.5
55
50
50
301
302
302
39–42
39
5.0
4.8
4.0
80
3.4
4.5
50
70
3.0
80
303
103
129
129
—c
304
305
AFR33046
AFR33047
AEJ35165
XLNA*
rPcXyl*
5.4
7.6–8.0 4.6–5.2
6.0
6.0
3.6
4.5–5.0
4.5
4.0
4.5
4.5
4.5
5.0
5.0
69
43
Litter decomposing Agaricus bisporus
-Xylosidase
White rot
Enzymea
NCBI protein
database
Molecular
accession no.b mass (kDa) pI
3.8
34
190
6.5
50–60 129
90
87
5.5
5.6
55
306
65–70 307
83
5.0
45
27
170
5.9
130
131
4.5
50
308
a
Asterisks indicate a heterologously produced enzyme.
b
See http://www.ncbi.nlm.nih.gov/protein.
c
Available in the GenBank database (http://www.ncbi.nlm.nih.gov/GenBank/).
such as P. ostreatus (endomannanase [139]) and P. radiata (140).
The molecular mass of -endomannanases (42 to 65 kDa) is lower
than that of -mannosidases (49 to 105 kDa), with the exception
of the C. subvermispora -endomannanase, with an atypically
high molecular mass (150 kDa) (128) (Fig. 3A and Table 10). The
isoelectric points have been determined only for the -endo-
632 mmbr.asm.org
mannanases from S. rolfsii, and their pI values are very acidic,
from 2.8 to 3.5 (135, 136, 138). Similarly, the pH optima of S.
rolfsii -endomannanases are also acidic (pH 2.9 to 3.5), while the
optimum pH range for P. chrysosporium -endomannanase is 4.0
to 6.0 (Table 10). -Mannosidases from the white rot species G.
lucidum and P. radiata, the brown rot fungus Laetiporus (Polypo-
Microbiology and Molecular Biology Reviews
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White rot-like
Species
CAZyme
family
Gene
Plant Polysaccharide Degradation by Basidiomycetes
TABLE 10 Characterized basidiomycete -1,4-endomannanases and mannosidases and their biochemical properties
Life-style
-Endomannanase
White rot
Species
Ceriporiopsis (Gelatoporia)
subvermispora
Heterobasidion irregulare
(Fomes annosus)
Phanerochaete chrysosporium GH5
P. chrysosporium
GH5
P. chrysosporium
(Chrysosporium lignorum)
Litter decomposing Agaricus bisporus
-Mannosidase
White rot
GH5
Enzymea
NCBI protein
database
Molecular
accession no.b mass (kDa) pI
Mannanase I
150
pHopt
Topt
(°C) Reference(s)
4.5
60
3.9–4.2
man5D Man5D*
man5C
ABG79370
ABG79371
65
128
309
4.0–6.0 60
4.11
132
—c
309
133, 134
Cel4b
Sclerotium rolfsii
61
3.5
2.9
S. rolfsii
S. rolfsii
Stereum sanguinolentum
42
47
3.2
2.8
3.58
3.3
72
3.0–3.5 75
Ganoderma lucidum
G. lucidum
Phlebia radiata
P. radiata
P. radiata
49
49
105
90
100
4.2
4.8
4.8
3.8
4.7
GM-1
GM-2
OT-1
Brown rot
Laetiporus (Polyporus)
sulfureus
64
Plant pathogen
Sclerotium rolfsii
58
5.5
5.5
5.5
74
50
50
50
2.4–3.4
4.5
2.5
135, 136,
310, 311
135, 310
138
309
16
16
141
141
141
142
55
135
a
Asterisks indicate a heterologously produced enzyme.
b
See (http://www.ncbi.nlm.nih.gov/protein).
c
Available in the GenBank database (http://www.ncbi.nlm.nih.gov/GenBank/).
rus) sulfureus, and the plant pathogen S. rolfsii have been characterized (16, 135, 141, 142); their pI values are close to 4.5, and the
optimum pH varies from 2.4 to 5.5 (Fig. 3B and C and Table 10).
The average temperature optimum for the -endomannanases
(70°C) is higher than that for -mannosidases (53°C) (Fig. 3D and
Table 10).
Brown rot fungi are specialized in degrading conifers (143),
which contain a higher percentage of mannan than hardwoods
(20). The genomes of basidiomycetes possess several copies of
genes encoding mannan-degrading enzymes assigned to GH2, -5,
and -26 (Table 4). Correspondingly, the brown rot fungus Piptoporus betulinus has been observed to produce -mannanase and
-mannosidase activities (109, 144). However, no mannanolytic
enzymes from the brown rot basidiomycetes have been isolated or
characterized (Table 5). Nevertheless, cellulolytic enzymes of
brown rot fungi have been shown to hydrolyze substrates other
than cellulose. Gloeophyllum sepiarium and G. trabeum produce
-1,4-endoglucanases that cleave galactoglucomannan and xylan,
respectively (108, 145). F. palustris possesses a GH3 -glucosidase
that is active against p-nitrophenylxyloside, p-nitrophenylgalactoside, p-nitrophenylcellobioside, and p-nitrophenylmannoside
(146). Similarly, P. betulinus -glucosidase is able to release galactose, mannose, and xylose from xylan and galactomannan (109).
Aspergillus species produce both -endomannanases and
December 2014 Volume 78 Number 4
-mannosidases, and characterized enzymes have been classified
into GH5 and -2 (19). Altogether, at the level of characterized
enzymes, both -mannanases and -mannosidases of basidiomycetes and aspergilli have gained less attention than several other
CAZymes (Table 10).
Pectin-Degrading Enzymes
Basidiomycete genomes show a high level of variation with respect to
pectin degradation (Table 4), but only a few pectinolytic enzymes
from basidiomycetes have been characterized (Table 11). Instead, the
main focus of studies of pectinases has been on A. niger and other
Aspergillus species. Aspergilli produce variable hydrolases (endoand exopolygalacturonases, endorhamnogalacturonan hydrolases, rhamnogalacturonan rhamnohydrolase, ␣-rhamnosidase,
and rhamnogalacturonan galacturonohydrolases) and lyases
(pectin, pectate, and rhamnogalacturonan lyases), with several
isoenzymes that differ in their specific activity (19).
The pectinolytic enzymes of basidiomycetes isolated or characterized so far are endo/exorhamno- or polygalacturonases from
the white rot fungus P. chrysosporium (147), the plant-pathogenic
species Chondrostereum purpureum (148–150) and S. rolfsii (151,
152), and the basidiomycete yeast Cystofilobasidium capitatum
(153, 154) as well as a rhamnogalacturonan lyase from the white
rot fungus I. lacteus (155, 156) (Table 11). However, there is a great
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Plant pathogen
CAZyme
family
Gene
Rytioja et al.
TABLE 11 Characterized basidiomycete pectin-degrading enzymes and their biochemical properties
Life-style
Species
NCBI protein
CAZyme
database
Molecular
family
Gene Enzymea accession no.b mass (kDa) pI(s)
Endo/exopolygalacturonases
and
rhamnogalacturonases
White rot
Phanerochaete
chrysosporium
Plant pathogen
Rhamnogalacturonan
hydrolase
White rot
4.3, 4.6, 4.7 4.7
Topt (°C) Reference(s)
66
147
GH28
epgA
AAF68401
150
GH28
GH28
GH28
GH28
GH28
GH28
epgB1
epgB2
epgC
epgD
cppg1 PGA
Pg1
PGA
PGA
AAF68402
AAK29433
AAF68403
AAF68404
BAA96351
BAA08102
2.5
150
150
150
150
149
148
151
4.0
4.0
152
152
39
46–48
28–31
Cystofilobasidium
capitatum
PGA
Irpex lacteus
RGH*
8.8
5.2
5.2
44
ACI26689
55
45
7.2
4.5–5.0 40–50
153, 154
155, 156
a
The asterisk indicates a heterologously produced enzyme.
b
See http://www.ncbi.nlm.nih.gov/protein.
potential to find novel pectinases from basidiomycetes with unique
properties because of the diverse ecological niches that basidiomycetes inhabit and the variety of putative pectinase-encoding genes in
their genomes (Table 4). For example, several basidiomycetes, including the white rot fungi Lentinus sp., P. chrysosporium, Pycnoporus sanguineus, and S. commune, have been shown to produce
higher polygalacturonase activities than A. niger on solid wheat
bran cultures in a screening study of 75 basidiomycetes (63).
The plant pathogens S. rolfsii and C. purpureum produce an
array of GH28 pectinolytic enzymes, which suggests that pectin degradation is important to their pathogenicity (148–150, 157, 158). C.
purpureum, which causes a silver leaf disease in apple trees, produces five GH28 PGA isoenzymes corresponding to the five
cloned PGA-encoding genes (150). A phylogenetic analysis has
shown that the PGA-encoding genes of C. purpureum have undergone duplication after the divergence of ascomycetes and basidiomycetes, suggesting an adaptation to a pectin-rich environment
(150). The pectinases of S. rolfsii act under extreme conditions.
Pectin methyl esterase (CE8) of S. rolfsii has a very acidic pH optimum (pH 2.5) and also retains most of its activity at pH 1.1 and
at 10°C (159). In addition, -galactosidase of S. rolfsii tolerates
acidic conditions, and its optimum pH is between 2 and 2.5 (151).
Very few pectin- and other plant cell wall polysaccharide-degrading enzymes have been isolated from basidiomycetous yeasts
(Tables 5, 7, and 11). This corresponds to the limited number of
CAZyme-encoding gene models found in the genome of R. glutinis (Table 4). However, the enzymes of basidiomycete yeasts may
have interesting biochemical properties. For example, C. capita-
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tum uses pectin as a sole carbon source and produces polygalacturonase (GH28), which is active even at 0°C and therefore has
potential to be used in food processing applications (153, 154).
Hemicellulose- and Pectin-Debranching Enzymes
Debranching enzymes that cleave the side chains of hemicelluloses
and pectin work synergistically with the enzymes that cleave the
backbone and main branches of plant polysaccharides (19). Various debranching enzymes from ascomycetes have also been isolated from basidiomycetes, including ␣- and -galactosidases,
␣-arabinofuranosidases, ␣-glucuronidases, acetyl xylan esterases,
a pectin methyl esterase, and feruloyl esterases (Table 12). Despite
the putative gene models present in basidiomycete genomes (Table 4), many debranching enzymes from basidiomycetes still remain
uncharacterized, such as ␣-fucosidase, p-coumaroyl esterase, arabinoxylan arabinofuranohydrolase, endoarabinase, exoarabinase, and
endogalactanase. Due to their importance for the complete degradation of plant biomass, future efforts should be directed at isolating
debranching enzymes from basidiomycetes.
Only a few white rot fungal debranching enzymes that catalyze the
cleavage of the smaller side branches of hemicellulose and pectin
main chains have been characterized (Table 12). P. chrysosporium
produced ␣-glucuronidase at high activity levels in a screening
study of xylan-degrading fungi, where several ascomycetes, such
as Aspergillus awamori and A. niger strains, and basidiomycetes,
such as P. chrysosporium and S. commune, were included (160).
The molecular mass of P. chrysosporium ␣-glucuronidase is 112
kDa, and its isoelectric point is 4.6. It has an acidic pH optimum
Microbiology and Molecular Biology Reviews
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Yeast
Chondrostereum
purpureum
C. purpureum
C. purpureum
C. purpureum
C. purpureum
C. purpureum
C. purpureum
Sclerotium (Corticium)
rolfsii
S. rolfsii
S. rolfsii
42
pHopt
Plant Polysaccharide Degradation by Basidiomycetes
TABLE 12 Characterized basidiomycete-debranching enzymes and their biochemical properties
pHopt
Topt
(°C)
Reference(s)
60
5.1
3.5
60
312
7.0
45
313
6.0
4.5
3.75
70
60–80
16
166
167
165, 314
5.0
5.0
5.0
5.0
5.0
5.0
4.6–5.0
60
60
60
60
60
60
55
169
169
169
169
169
169
168
Calvatia cyathiformis
3.0–5.0
50
315
Sclerotium (Corticium) rolfsii
2.0–2.5
␣-Arabinofuranosidase
White rot
Dichomitus squalens
␣-Galactosidase
White rot
Coprinopsis cinerea
Ganoderma lucidum
G. lucidum
Lenzites elegans
Phanerochaete chrysosporium
GH62
Gene
CcAbf62A
-Galactosidase
Plant pathogen
Yeast
-1,3-Endo/
exogalactanase
White rot
White rot-like
Litter decomposing
Acetyl xylan esterase
White rot
CcAbf62A*
BAK14423
AAG24510,
AAG24511
AgaS-b1
AgaS-b2
AgaS-b3
AgaS-m1
AgaS-m2
AgaS-m3
Flammulina velutipes
Irpex lacteus
49
249c
158d
250c
5.5
4.5
50
40
318
319
30
45
GH43
1,3Gal43A
1,3Gal43A*
BAD98241
55
112
110
4.6
4.4
3.5
3.8
125
3.6
4.5–5.5
CE1
CE1
axe1
PcAxe2
PcAxe2*
AEX99751
321
63
7.0
30–35
58
170
31
7.7
30–45
322
8.0
60
323
Volvariella volvacea
Vvaxe1
VvAXE1*
ABI63599
45
Coprophilic
Coprinopsis cinereae
CcEst1
CcEst1*
BAJ10857
45
Pectin methyl esterase
Plant pathogen
Sclerotium (Corticium) rolfsii
163
162
52
Straw decomposing
EstBC
FaeA
Est1
160
161
3.3
Schizophyllum commune
Auricularia auricula-judae
Pleurotus eryngii
Pleurotus sapidus
60
ADV52250
White rot-like
Feruloyl esterase
White rot
320
Agaricus bisporus
Phanerochaete chrysosporium
P. chrysosporium
316
317
BAK48741
BAH29957
Agu1*
7.2
5.7
3.5
6.7
5.7
5.0
50
FvEn3GAL*
rIl1,3Gal*
Agu1
4.0–4.2
5.0
FvEn3GAL
Il1,3Gal
GH115
4.6
53
GH16
GH43
Phanerochaete chrysosporium
Phlebia radiata
Schizophyllum commune
S. commune
48
60
59
60
55
58
64
99
Sporobolomyces (Bullera)
singularis
Galactan
-1,3-galactosidase
White rot
Phanerochaete chrysosporium
␣-Glucuronidase
White rot
Enzyme
GH36
GH27
Phlebia radiata
P. radiata
P. radiata
P. radiata
P. radiata
P. radiata
Pleurotus florida
Saprobic
a
36
67
55
37
324
3.2
5.2
6.5
5.0
6.0
61–66
50
50
175
173
174
2.5–4.5
45
159
a
Asterisks indicate a heterologously produced enzyme.
b
See http://www.ncbi.nlm.nih.gov/protein.
c
Tetramer.
d
Dimer.
e
Acetic acid- and ferulic acid-releasing activities.
December 2014 Volume 78 Number 4
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pI
Species
Coprophilic
NCBI protein
database
accession no.b
Molecular
mass (kDa)
Life-style
CAZyme
family
Rytioja et al.
636
mmbr.asm.org
kDa) (176) and A. niger FaeA (36 kDa) (177, 178) but lower than
that of A. niger FaeB (74 kDa) (178). P. eryngii FaeA (PeFaeA) and
P. sapidus Est1 are 93% similar at the amino acid sequence level
(173). However, they do not show significant homology to A. niger
FaeA or FaeB, which is also reflected in their biochemical differences. P. sapidus Est1 hydrolyzes arabinosyl esters of ferulic acid
more efficiently than methyl ferulate, which is the commonly used
substrate for feruloyl esterases (174). PeFaeA had higher activity
toward natural substrates, such as feruloylated mono-, di-, and
trisaccharides (F-A, F-AX, and F-AXG, respectively), than toward
the typical synthetic feruloyl ester substrates methyl ferulate,
methyl coumarate, and methyl sinapate (173). In addition, it is
not able to hydrolyze methyl caffeate (173). Both PeFaeA and P.
sapidus Est1 prefer F-AX over F-A and F-AXG of the natural substrates, whereas A. niger FaeA prefers F-A over F-AX (173, 179).
Recently, a novel type of FAE (EstBC), which hydrolyzes both
benzoates and cinnamates, has been described for the jelly fungus
A. auricula-judae (175).
Only two ␣-xylosidases from Aspergillus species (19) and, so far,
none from basidiomycetes have been characterized. The ␣-xylosidases from aspergilli are specific for ␣-linked xylose residues but
show differences in the type of glycosides that they are able to
hydrolyze (19, 180, 181).
Several cellulases, xylanases, mannanases, and pectinases have
been isolated from phytopathogenic basidiomycetes (Table 5).
Most of the characterized enzymes are from S. rolfsii, which has a
dual life-style as a soil saprotroph and a necrotrophic crop pathogen with a wide host range including ⬎500 plant species (157,
182). S. rolfsii causes large economic losses by infecting several
crop and ornamental plants, especially in tropical and subtropical
regions. During infection, S. rolfsii produces oxalic acid as well as
cellulolytic and pectinolytic enzymes (157, 182). Although the genome of another necrotrophic basidiomycete, the white rot fungus H. irregulare, is already available, the genome of S. rolfsii or a
related facultative pathogenic basidiomycete has yet to be sequenced to reveal the full genetic potential of phytopathogens for
carbohydrate degradation.
REGULATION OF PLANT POLYSACCHARIDE DEGRADATION
IN BASIDIOMYCETES AND ASPERGILLUS
Considering the highly varied composition of plant biomass, efficient degradation of these components by fungi depends on the
production of the right combination of enzymes. Therefore, most
genes encoding plant-biomass-degrading enzymes are under the
control of transcriptional regulators. In aspergilli, several transcriptional regulators (all of the Zn2Cys6 type) that activate the
expression of genes involved in plant biomass degradation have
been identified, such as XlnR, AraR, GalR, GalX, and RhaR (183).
None of these regulators have orthologs in basidiomycetes, but
several are found across the phylum Ascomycota (184). This suggests that regulation of plant biomass degradation has developed
after ascomycetes and basidiomycetes diverged during fungal evolution. No specific regulators involved in plant biomass degradation in basidiomycetes have been described, but indications for
such systems can be derived from transcriptome studies with basidiomycetes, although these indications were not explicitly stated
in those reports (17, 18, 70).
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and hydrolyzes short-chain xylooligosaccharides but shows low
activity toward glucuronoxylan polysaccharides and xylans of
birch, oat spelt, and wheat straw (160). ␣-Glucuronidase of the
white rot fungus P. radiata has been shown to act together with an
endoxylanase in the degradation of oat xylan (161). A. niger and A.
tubingensis ␣-glucuronidases are also active mainly on small xylooligomers, and therefore, they are expected to be dependent on
the action of endoxylanases (19). Interestingly, S. commune produced an ␣-glucuronidase that is active against polymeric glucuronoxylan (162) and for which the gene was recently cloned and
demonstrated to be a member of GH115 (163). In contrast to this
enzyme, a GH115 ␣-glucuronidase from the ascomycete Pichia
stipitis was active only on oligomeric substrates (164).
␣-Galactosidases have been isolated from some white rot species and have diverse properties. Both P. chrysosporium and G.
lucidum glucomannan-debranching ␣-galactosidases are produced as tetramers, while the molecular masses of the monomers
are 50 and 56 kDa, respectively (165, 166). P. chrysosporium ␣-galactosidase has an acidic pH optimum of 3.75, and the enzyme is
stable from 0 to 80°C (165). The pH optimum of G. lucidum ␣-galactosidase is 6.0, and its optimum temperature is 70°C (165, 166)
The white rot fungus Lenzites elegans secretes a homodimeric
␣-galactosidase with a molecular mass of 158 kDa (61 kDa for one
subunit) (167). ␣-Galactosidase of L. elegans has an acidic pI value
ranging from 4.0 to 4.2 and a pH optimum of 4.5. This enzyme
shows activity against several ␣-galactosidases and is very thermostable, with an optimal temperature from 60°C to 80°C (167).
In contrast, ␣-galactosidase from the white rot fungus Pleurotus
florida is a monomeric protein with a molecular mass of 99 kDa,
and its temperature optimum is 55°C (168). The white rot fungus
P. radiata produces several isoforms of ␣-galactosidase when
grown in wheat bran- and locus bean gum-containing liquid media (169). Molecular masses of the ␣-galactosidase isoforms of P.
radiata are between 55 and 64 kDa, and their isoelectric points
vary from 3.5 to 7.15. P. radiata ␣-galactosidase isoforms have an
optimum pH of 5.0 and show the highest activity at 60°C (169).
Several different ␣-galactosidases have been purified from aspergilli. In addition, a few endo- and exogalactanases from aspergilli
have been characterized (19), but these enzymes have not yet been
characterized for basidiomycetes.
Acetyl xylan esterases (AXEs), which cleave ester linkages between
acetic acid and xylan or mannan, and feruloyl esterases (FAEs), which
cleave ester linkages between phenolic acid and the arabinose or galactose side chain of xylan or pectin, have been characterized for the
white rot fungi P. chrysosporium, Pleurotus eryngii, Pleurotus sapidus, and S. commune; the jelly fungus Auricularia auricula-judae;
and the coprophilic species C. cinerea (Table 12). The genome of
P. chrysosporium harbors three AXEs, one of which, PcAxe2, has
been biochemically characterized (170). PcAxe2 and AXEs from
Aspergillus ficcum, A. awamori, and A. niger show a similar pH
optimum (7.0). AXEs of A. ficcum, A. oryzae, and A. niger have
slightly higher temperature optima (35°C to 50°C) than that of
PcAxe2 (30°C to 35°C) (170–172). PcAxe2 displays low specific
activity against birchwood xylan. However, synergistic action between PcAxe2 and the P. chrysosporium endoxylanase PcXynC in
xylan degradation has been reported (170).
Only three basidomycete FAEs have been biochemically characterized (173–175). The molecular masses of FAEs of the white
rot fungi P. eryngii and P. sapidus are 67 kDa and 55 kDa, respectively, which are higher than those of the FAEs of A. awamori (35
Plant Polysaccharide Degradation by Basidiomycetes
Repression of Gene Expression in Basidiomycetes
Induction of Gene Expression in Basidiomycetes
CAZyme-encoding genes of basidiomycetes, from the brown rot
fungi F. palustris and G. trabeum to the litter decomposer A. bisporus, are induced when exposed to long polymers of cellulose and
hemicellulose (105, 186, 195). The CCAAT binding complex,
December 2014 Volume 78 Number 4
CONCLUSIONS AND FUTURE PROSPECTS
The increasing number of genome sequences covering the wide
range of diversity of biomass-decomposing fungi has widened
our understanding of the enzymatic machinery that they possess for plant cell wall polysaccharide degradation. Since the
first whole-genome sequencing of a basidiomycete, P. chrysosporium, next-generation sequencing has facilitated a growing
number of genomes and transcriptomes of plant cell wall-decomposing basidiomycetes (197). These complementary “omics”
studies have accelerated the process of discovering novel enzyme
activities involved in plant cell wall decomposition. In line with
the aspergilli, genome sequencing of basidiomycetes has revealed
an unexpectedly large repertoire of GHs. For example, ⬍20% of
the putative GHs of P. chrysosporium were characterized before
the genome sequence was published (59).
At the same time, the genomes of plant-biomass-degrading
basidiomycete fungi have revealed putative novel protein-encoding genes, especially among those without known homology, providing enzymes related to plant polysaccharide degradation for
further characterization. For instance, several new families, such
as the second family of ␣-glucuronidases (GH115) (163) and a
family of glucuronoyl esterases (CE15) (198, 199), have recently
been added to the CAZy database. Furthermore, the novel concept
of oxidoreductive polysaccharide degradation will provide challenges, as its significance in both the basidiomycete and ascomycete fungi is yet to be fully clarified. The abundance of the LPMOencoding genes and the diversity of LPMO sequences and
described activities for basidiomycetes and other plant-biomassdegrading microorganisms suggest that LPMO-catalyzed oxidation has a major role in plant cell wall polysaccharide conversion
(40). The discovery of LPMO changed the concept of cellulose
degradation (200, 201), and a recent study demonstrated the ability of LPMO to cleave not only cellulose but also hemicelluloses
(40). In the near future, LPMO-catalyzed depolymerization of
other plant cell wall polysaccharides besides cellulose and hemicelluloses will most likely be shown. In addition, a new family of
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CreA-mediated repression is induced by monomeric products
such as glucose, fructose, and xylose in ascomycetes (19), but the
mechanism has not been studied for basidiomycetes. However,
CreA homologs have been detected in all basidiomycete genomes
sequenced so far (184). In line with the ascomycetes, various basidiomycetes, such as the white rot fungus Trametes (Coriolus)
versicolor, the litter decomposers A. bisporus and V. volvacea, the
phytopathogen S. rolfsii, and the basidiomycete yeast Rhodotorula
minuta, have CAZyme-encoding genes that are repressed by glucose and other monosaccharides (89, 185–188). Some CAZymeencoding genes are also repressed by monosaccharides that seem
unrelated to the enzymes that these genes encode. For example,
certain cellulolytic genes are inhibited by lactose, xylose, mannose,
and fructose (89, 186, 189). Like ascomycetes, the xynA endoxylanase gene of A. bisporus is repressed by glucose (190).
Copies of a CreA-related binding site, SYGGRT (191), have
been detected in the genomes of basidiomycetes. In the white rot
fungus I. lacteus, a CreA binding site upstream of the cel2 gene
(GH7 and CBHI) has been found between two CAAT boxes, similarly to what was observed for the cbhI gene of A. aculeatus (189).
CreA binding elements were also identified in the glycosyl hydrolase promoters of the white rot fungi C. subvermispora and S. commune (12, 163) and the brown rot fungus P. placenta (18, 70).
However, these elements were not present upstream of the bglA
gene encoding the -galactosidase of the basidiomycete yeast
Sporobolomyces singularis (112). Information from the genomes of
basidiomycetes and studies on their genes encoding CAZymes
suggests that CreA homologs mediate the repression of some
CAZyme-related genes, while other genes are possibly repressed
by other mechanisms or constitutively expressed, such as the
cbh1-1 and cbh1-2 genes from P. chrysosporium (192).
To relieve glucose repression at low sugar levels, the Snf1 protein kinase, found in Saccharomyces cerevisiae and other ascomycetes (193), targets Mig1, which is a functional homolog of CreA
(194). An snf1 gene has also been identified in the phytopathogen
U. maydis and was shown to mediate gene expression of at least
one EG and one PGA (193). In mutants that did not have snf1, EGand PGA-encoding genes were expressed at lower levels in high
concentrations of glucose. However, xylanase gene expression levels were higher in mutants lacking snf1. This suggests that Snf1
negatively regulates xylanase expression and is required for the
induction of EG- and PGA-encoding genes (193). An ortholog of
Snf1, SnfA, has been identified in the brown rot fungus P. placenta
(18), and it may also exist in other basidiomycetes. However, a
direct interaction between Snf1 and CreA has yet to be studied in
basidiomycetes.
All these data suggest that CreA homologs in basidiomycetes likely
affect the expression of a range of CAZymes and respond to the presence of a variety of monosaccharides, similar to what has been described for Aspergillus species (19). The presence of CreA homologs across the fungal kingdom supports a central role for this
regulator in fungal physiology in natural habitats (184).
which enhances the expression of genes located downstream of it,
is found in the promoter regions of the genes involved in cellulose
and hemicellulose degradation in many Aspergillus species (19).
The white rot fungus I. lacteus possesses a CCAAT motif in the
promoter region of cel2 (GH7 and CBHI) (189), which suggests
that at least some basidiomycete CAZyme-encoding genes may be
upregulated by mechanisms analogous to that of ascomycetes. In
the plant pathogen C. purpureum, a CCAAT motif was found before the start codon in all five PGA-encoding genes (150). However, not all basidiomycetous CAZyme-encoding genes have a
CCAAT binding complex. For example, -galactosidase of the
basidiomycete yeast S. singularis does not possess a CCAAT sequence (112).
The expression of multiple genes encoding plant-polysaccharidedegrading enzymes during growth on plant biomass, as evidenced for
several basidiomycete species (9, 12, 17, 70, 196), indicates a regulatory system similar to that described for ascomycetes. The absence of
homologs of the ascomycete regulators suggests that this may be due
to parallel evolution, during which both fungal phyla developed different regulators that perform the same function. This poses an intriguing question regarding how the expression of plant-biomassdegrading enzymes was organized in the ancestral fungi before the
divergence of basidiomycetes and ascomycetes.
Rytioja et al.
8.
9.
10.
11.
ACKNOWLEDGMENTS
J.R. acknowledges funding from the Doctoral Programme in Microbiology and Biotechnology. M.R.M. was supported by European Union grant
(Optibiocat) EU FP7 201401.
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Johanna Rytioja obtained her B.Sc. in Biotechnology at the University of Helsinki, Finland, in
2008. She received her M.Sc. in Biotechnology
from the University of Helsinki in 2010 under
the supervision of Dr. T. Lundell and Dr. M.
Mäkelä, studying the production and reactions
of laccases of the white rot fungus Physisporinus
rivulosus. She is currently finishing her Ph.D. on
biotechnology in the Microbiology and Biotechnology Doctoral Programme at the University of Helsinki, under the supervision of Prof.
A. Hatakka, Dr. M. Mäkelä, and Dr. K. Hildén. Her Ph.D. focuses on carbohydrate-active enzymes of basidiomycete white rot fungi, in particular cellobiohydrolases, cellobiose dehydrogenases, and the set of enzymes produced by the white rot fungus Dichomitus squalens, during growth on diverse
plant biomass substrates.
Kristiina Hildén is principal investigator of the
FungalGeneticsandBiotechnologygroupoftheDepartment of Food and Environmental Sciences of
the University of Helsinki, Finland. She received her
M.Sc. in Genetics from the University of Helsinki in
1995 and a Ph.D. in Genetics from the same University under the supervision of Prof. O. Ritvos in 2003.
She joined the Department of Food and EnvironmentalSciencesoftheUniversityofHelsinkiin2000,
and since then, her research has included various aspects of fungal molecular biology and enzymology.
From 2009 to 2011, she interrupted her research at the University of Helsinki for a
2-year Marie Curie Intra-European Fellowship project at the University of Nottingham,UnitedKingdom.UponreturningtoHelsinki,shedevelopedherownresearch
line at the group, focusing on the functional genomics of plant-biomass-degrading
fungi from various biotopes. In addition, several more applied projects are ongoing
in the group.
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.doi.org/10.1074/jbc.M501024200.
Puls J, Schmidt O, Granzow C. 1987. ␣-Glucuronidase in two microbial
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Biochem. J. 298:751–755.
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.doi.org/10.1111/j.1574-6968.2007.00844.x.
Hashimoto K, Kaneko S, Yoshida M. 2010. Extracellular carbohydrate
esterase from the basidiomycete Coprinopsis cinerea released ferulic and
acetic acids from xylan. Biosci. Biotechnol. Biochem. 74:1722–1724.
http://dx.doi.org/10.1271/bbb.100299.
Jennifer Yuzon obtained her B.Sc. in Microbiology at the University of California, San Diego, in
2011. She received her M.Sc. in Environmental Biology from Utrecht University under the mentorship of Prof. R. de Vries. In the Fungal Physiology
Laboratory of Prof. de Vries, she assisted with culturing of various strains of Agaricus bisporus and
studied their extracellular carbohydrate-active
enzymes. She also investigated the molecular
phylogeny and taxonomy of the lichenized fungus Bagliettoa under the mentorship of Dr. C.
Gueidan at the Natural History Museum, London, United Kingdom. She is
currently in the doctoral program in Plant Pathology at the University of
California, Davis, under the mentorship of Dr. T. Kasuga. Since she joined
Dr. Kasuga’s Phytophthora Genomics Laboratory in 2014, her research interests include the epigenetic mechanisms contributing to the invasive biology and pathogenicity of Phytophthora ramorum.
Annele Hatakka is a Professor in Environmental Biotechnology at the Division of Microbiology and Biotechnology, Department of Food
and Environmental Sciences, University of Helsinki, Helsinki, Finland. Since the 1990s, she has
been the leader of the research group on biotechnology of renewable natural resources. She
obtained her Ph.D. in Microbiology at the University of Helsinki in 1986. The topic was degradation and conversion of lignin, lignin-related aromatic compounds. and lignocellulose
by white rot fungi. She has worked for 1.5 years at the Swedish Forest Products Laboratory (STFI), Stockholm, Sweden, and for shorter periods at the
INRA, France, and the United Kingdom. After working for 24 years in various research positions at the Academy of Finland, she became Professor in
Environmental Biotechnology in 1997, which was converted to a permanent
professorship in 2008. Her main research areas are lignocellulose degradation by white rot and other fungi, lignocellulose-degrading enzymes, applications in the pulp and paper industry, and bioremediation of polluted soils.
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Plant Polysaccharide Degradation by Basidiomycetes
December 2014 Volume 78 Number 4
Miia R. Mäkelä is principal investigator of the
Fungal Genetics and Biotechnology group of
the Department of Food and Environmental
Sciences of the University of Helsinki, Finland.
She received her M.Sc. in Microbiology from
the University of Helsinki in 2000 and a Ph.D. in
Microbiology from the same University under
the supervision of Prof. A. Hatakka and Dr. T.
Lundell in 2009. Her Ph.D. work focused on the
role of oxalate-converting enzymes related to
lignin modification, which has remained a topic
of interest in her research. She has performed postdoc projects related to
various aspects of lignocellulose degradation by fungi and more recently also
on the applications of various enzyme classes in industrial processes. She has
recently expanded her work on oxalate metabolism to a more global look at
carbon metabolism of wood-decaying fungi that also includes the conversion of plant-based monomeric carbon sources by basidiomycete fungi.
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Ronald P. de Vries is head of the Fungal Physiology group of the CBS-KNAW Fungal Biodiversity Center in Utrecht, The Netherlands. He
received his M.Sc. in Molecular Sciences in 1992
from Wageningen University, The Netherlands,
and a Ph.D. in Fungal Molecular Biology from
the same University under the supervision of
Dr. J. Visser in 1999. Afterwards, he performed
postdoc work at Wageningen University, the
Institut Pasteur, and Utrecht University. In
2009, he was hired to build a group on Fungal
Physiology at the CBS-KNAW Fungal Biodiversity Center, which mainly
addresses plant biomass utilization by fungi. His current research focuses
mainly on understanding fungal diversity with respect to plant biomass utilization, using a multidisciplinary approach combining (post)genomics with
molecular biology, physiology, biochemistry, and microscopy. Special emphasis is placed on regulatory systems that control this aspect of fungal life.
As of May 2014, he is also Professor in Fungal Molecular Physiology at
Utrecht University, The Netherlands.