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ANNUAL REVIEWS

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Annu. Rev. Phytopathol. 2014.52:427-451. Downloaded from www.annualreviews.org Access provided by Cornell University - Weill Medical College on 07/01/17. For personal use only.

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Plant Cell Wall–Degrading Enzymes and Their Secretion in Plant-Pathogenic Fungi Christian P. Kubicek,1 Trevor L. Starr,2 and N. Louise Glass2 1 Austrian Center of Industrial Biotechnology, 8010 Graz, Austria; email: [email protected] 2 Department of Plant and Microbial Biology and the Energy Biosciences Institute, University of California, Berkeley, California 94720-3102; email: [email protected], [email protected]

Annu. Rev. Phytopathol. 2014. 52:427–51

Keywords

First published online as a Review in Advance on June 16, 2014

plant cell wall, cellulases, hemicellulases, pectin, plant pathogens, enzyme secretion, transcriptional regulation

The Annual Review of Phytopathology is online at phyto.annualreviews.org This article’s doi: 10.1146/annurev-phyto-102313-045831 c 2014 by Annual Reviews. Copyright  All rights reserved

Abstract Approximately a tenth of all described fungal species can cause diseases in plants. A common feature of this process is the necessity to pass through the plant cell wall, an important barrier against pathogen attack. To this end, fungi possess a diverse array of secreted enzymes to depolymerize the main structural polysaccharide components of the plant cell wall, i.e., cellulose, hemicellulose, and pectin. Recent advances in genomic and systemslevel studies have begun to unravel this diversity and have pinpointed cell wall–degrading enzyme (CWDE) families that are specifically present or enhanced in plant-pathogenic fungi. In this review, we discuss differences between the CWDE arsenal of plant-pathogenic and non-plant-pathogenic fungi, highlight the importance of individual enzyme families for pathogenesis, illustrate the secretory pathway that transports CWDEs out of the fungal cell, and report the transcriptional regulation of expression of CWDE genes in both saprophytic and phytopathogenic fungi.

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INTRODUCTION

Annu. Rev. Phytopathol. 2014.52:427-451. Downloaded from www.annualreviews.org Access provided by Cornell University - Weill Medical College on 07/01/17. For personal use only.

The ability to parasitize and cause damage to living plants has evolved independently in many branches of the tree of life (http://tolweb.org), including in bacteria, fungi, and protista. Approximately 10% of all fungal species identified can cause disease in more than 10,000 different plants (49). This ability to cause disease is reflected in the variability in fungal adaptation to plant hosts and also in their mode of pathogenesis. Some phytopathogens invade and colonize all tissues of a host plant, whereas others attack only particular tissues or organs (e.g., leaves, roots, stems, or seeds). Most plant pathogens, including nearly all biotrophic and many necrotrophic fungi, cause disease in only one or a few plant species, whereas other pathogens, including many root-infecting and soft rot–inducing fungi, have a wide host range (113, 126). When a fungus encounters a potential host, the cell wall is an important barrier that plants use to limit pathogen attack (46, 123). Plant cell walls are heterogeneous structures, composed of polysaccharides, proteins, and aromatic polymers. The composition and structure of the cell wall differ significantly among plant lineages, but nevertheless they share basic principles: all contain cellulose microfibrils embedded in a matrix of pectin, hemicellulose, lignin, and structural proteins (21, 98), although differences occur in the relative amounts of these compounds in different groups of plants. For example, type I cell walls found in noncommelinoid monocots and dicots are generally rich in xyloglucan and pectin. In contrast, type II cell walls of commelinoid monocots contain glucuronoarabinoxylan as the major noncellulosic polysaccharide. In addition to the physical complexity of the plant cell wall, its structure changes as the plant grows and develops. In both monocot and dicot species, during the maturation of the cell wall from a primary to secondary wall, the amount of xyloglucan, pectin, and structural proteins decreases, whereas the amount of xylan and lignin increases (5). To overcome the barrier of the plant cell wall, phytopathogenic fungi produce enzymes, which focus on the deconstruction of cellulose, xylan, and pectin, that are capable of degrading cell wall polymers [cell wall–degrading enzymes (CWDEs)]. These enzymes are particularly important for phytopathogenic fungi that do not have specialized penetration structures. In addition, all phytopathogenic fungi require these enzymes during late stages of invasion (38). For example, many plant-pathogenic fungi actively kill and degrade plant tissue to utilize the liberated monoand oligosaccharides for growth and reproduction. Plants protect themselves against destruction of their cell walls by producing proteins that inhibit microbial CWDEs; e.g., inhibitors of pectindegrading enzymes are common in dicots and noncommelinoid monocots, and inhibitors of xylandegrading enzymes are common in grasses (102). The production of these inhibitors by plants has, in turn, driven the evolution of some CWDE groups of phytopathogenic fungi toward inhibitorresistant enzymes (55). In some phytopathogenic fungi, there is evidence for production of different amounts of specific CWDEs, depending on whether the plant host is a monocot or dicot (24, 61).

ENZYMES INVOLVED IN PLANT CELL WALL DEGRADATION The complexity of the plant cell wall is mirrored by a diverse arsenal of CWDEs produced by fungi (40). Each type of enzyme that can degrade structural polysaccharides is represented in multiple families that share sequence and structural similarities. Enzymes that are capable of hydrolytically cleaving glycosidic bonds in oligo- or polysaccharides (including cellulose and hemicellulose) are generally summarized under the term glycoside hydrolases (GHs). GHs are distributed among various gene families that are genetically and structurally different. A classification concept based on amino acid similarity of GH members [carbohydrate active enzymes (CAZymes)] (47) correlates with enzyme mechanisms and three-dimensional structures of the respective proteins. Murphy 428

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et al. (79) manually curated GH families found in fungal genomes. A total of 453 characterized GHs from 131 different fungal (mostly ascomycete) species were retrieved that represented 44 of the 115 CAZy GH families. The annotated genes and proteins have been compiled in a searchable online database [mycoCLAP (characterized lignocellulose-active proteins of fungal origin); http://mycoCLAP.fungalgenomics.ca/].

Annu. Rev. Phytopathol. 2014.52:427-451. Downloaded from www.annualreviews.org Access provided by Cornell University - Weill Medical College on 07/01/17. For personal use only.

Cellulose-Degrading Enzymes The canonical view of hydrolysis of cellulose involves the action of two types of cellulases in an exo/endosynergy, followed by a β-glucosidase that hydrolyzes the soluble cellodextrin oligomers to glucose. However, it has been recognized for some time now that the differentiation of cellulases into endoglucanases and exoglucanases (cellobiohydrolases) is an oversimplification: Cellulases have evolved overlapping modes of action ranging from totally random endoglucanases through processive endoglucanases to strictly exoacting, highly processive cellobiohydrolases (64, 118). The exact role of individual enzymes with different degrees of processivity and endoactivity in cellulose degradation is not known. Analyses of the genome sequences of more than 40 ascomycete and basidiomycete species show that these enzymes are confined to a relatively low number of GH families (132) (Table 1): strictly processive exocellulases (cellobiohydrolases found in GH families 6 and 7, which are usually present as only one or two isoenzymes) and endocellulases (properly called endo-β-1,4-glucanases), which are distributed throughout a larger number of GH families (GH families 5, 7, 12, and 45). β-Glucosidases are predominantly found in the GH1 and GH3 families, but these families also contain other glycosidases. A feature typical for most, but not all, cellulases, and also found in some other CWDEs, is the presence a polysaccharide-binding domain connected by a loop hinge region, which aids in the binding of cellulases to their insoluble substrate (8). Recent work in Neurospora crassa and in Thermoascus aurantiacus showed that some members of the GH61 family encode a novel class of copper-dependent enzymes, now referred to as lytic polysaccharide monooxygenases (LPMOs) (4, 93). LPMOs catalyze an oxidative cleavage of cellulose in the presence of an external electron donor, thus exhibiting synergy with hydrolytic enzymes in biomass depolymerization. Other proteins (called cellulase-enhancing proteins), for example, the expansin-like protein swollenin identified in Trichoderma reesei, synergistically raise the activity of the cellulases but do not exhibit any enzymatic activity on cellulose themselves (100).

Hemicellulose-Degrading Enzymes Hemicellulose is a term used to describe the noncellulosic polysaccharides of the plant cell wall that comprise xyloglucans, xylans, and galactomannans. Although the linkage and sugars in the core chains are different between these major polysaccharides, the side-chain substituents often comprise the same sugar and the same linkage, and therefore the same enzymes are involved in their cleavage. Table 2 presents an overview on enzymes required for complete depolymerization of these three major classes of hemicellulose. Fungi use both nonspecific and specific groups of endo-β-(1→4)-glucanases for hydrolysis of the backbone chain of xyloglucan. Endo-β-1,4-glucanases with xyloglucanase activity can be found in GH5, GH12, GH16, and GH44 families, and all but the last are present in fungal genomes. The specific xyloglucanases, which completely lack or display only very low activities toward carboxymethyl-cellulose and β-glucan, belong to the GH74 family. Endo-1,4-β-xylanases (1,4-β-D-xylan xylanohydrolase; EC 3.2.1.8) cleave the glycosidic bonds in the xylan backbone. The topology selected for hydrolysis depends on the chain length, the degree www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

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Table 1 Enzymes and proteins involved in cellulose depolymerization in saprotrophic and plant-pathogenic fungia Cellulases Nutritional style

Annu. Rev. Phytopathol. 2014.52:427-451. Downloaded from www.annualreviews.org Access provided by Cornell University - Weill Medical College on 07/01/17. For personal use only.

Saprotrophic

Hemibiotrophic

Fungi

Host

GH6

β-Glycosidases

Accessory enzymes GH61b

GH7

GH5

GH12

GH45

GH1

GH3

Allomyces macrogynus

NA

0

0

4

0

0

0

0

3

CBM 0

Arthrobotrys oligospora

NA

2

6

21

1

4

2

9

26

112

Aspergillus clavatus

NA

2

4

9

3

0

3

12

8

15

Aspergillus flavus

NA

1

3

15

5

0

3

24

8

5

Aspergillus fumigatus

NA

1

4

13

4

1

5

18

8

17

Aspergillus nidulans

NA

2

3

16

1

0

3

20

11

6

Aspergillus niger

NA

1

0

5

1

0

3

14

3

1

Aspergillus oryzae

NA

1

3

12

4

0

3

23

9

3

Aspergillus terreus

NA

2

4

19

6

0

3

21

13

15

Chaetomium globosum

NA

4

6

10

4

2

1

13

45

34

Neosartorya fischeri

NA

2

5

16

5

1

5

19

8

22

Neurospora crassa

NA

3

5

6

2

1

1

11

16

21

Neurospora tetrasperma

NA

3

6

6

1

1

1

11

15

19

Penicillium chrysogenum

NA

2

2

13

3

0

3

17

10

6

Penicillium marneffei

NA

1

2

8

3

2

3

14

2

17

Talaromyces stipitatus

NA

1

2

11

3

1

2

22

1

20

Trichoderma reesei

NA

1

2

8

2

1

2

13

4

14

Cochliobolus sativus

M

3

6

15

3

3

3

15

29

11

Dothistroma septosporum

G

0

1

11

3

1

2

10

4

1

Fusarium graminearum

M

1

2

14

4

1

3

22

14

12

Glomerella graminicola

M

3

6

15

6

2

1

18

32

21

Magnaporthe oryzae

M

3

5

13

4

1

2

18

24

25

Moniliophthora perniciosa

D

2

3

11

4

2

2

11

16

5

Mycosphaerella graminicola

M

0

1

9

0

1

1

16

4

0 (Continued )

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Table 1 (Continued )

Cellulases Nutritional style

Annu. Rev. Phytopathol. 2014.52:427-451. Downloaded from www.annualreviews.org Access provided by Cornell University - Weill Medical College on 07/01/17. For personal use only.

Necrotrophic

Fungi

Host

GH6

Accessory enzymes

β-Glycosidases

GH7

GH5

GH12

GH45

GH1

GH3

GH61b

CBM

Botryotinia fuckeliana

M/D

1

3

16

3

2

4

16

10

18

Dichomitus squalens

G

1

3

20

3

0

4

8

15

17

Fomitiporia mediterranea

M/D

2

2

19

4

0

5

8

13

6

Fusarium oxysporum

M/D/G

1

3

22

4

1

6

32

17

14

Fusarium verticillioides

M

1

3

19

4

1

7

22

14

14

Gaeumannomyces graminis

M

5

6

14

2

4

1

17

28

18

Heterobasidion annosum

G

1

1

17

4

1

2

11

10

16

Magnaporthe poae

M

3

7

10

2

3

2

14

22

11

Nectria haematococca

D

1

3

18

6

1

5

38

13

12

Phaeosphaeria nodorum

M

4

5

18

3

2

2

15

30

10

Pyrenophora teres

M

3

3

13

2

3

3

12

26

12

Pyrenophora tritici

M

3

3

13

2

3

3

12

26

11

Sclerotinia sclerotiorum

D

2

3

14

4

2

3

13

9

19

Verticillium albo-atrum

D

4

6

13

5

1

4

21

22

22

Verticillium dahliae

D

4

6

13

6

2

2

16

29

29

a

Data collected from Reference 132. A lytic polysaccharide monooxygenase. Abbreviations: CBM, cellulose-binding domain; D, dicots; G, gymnosperms; M, monocots; NA, not applicable. b

of branching, and the presence of substituents. Most fungal xylanases have been identified as members of GH10 and GH11. Family GH10 xylanases are abundant in fungi, and a particularly high number are found in plant-pathogenic and saprobic fungi. These enzymes also attack decorated forms of xylan (7) but tolerate only a low number of unsubstituted consecutive β-1,4-xylopyranosyl units in the main chain. Some fungi also possess an additional GH family with xylanase activity, GH30. This family accommodates glucuronoxylan xylanohydrolases. The substrate specificity of one of the two GH30 members in the genome of T. reesei, XYN4, has been studied in detail (120). XYN4 displays greater activity toward unsubstituted xylans or acetylated methylglucuronic acid xylans than the GH10 and GH11 xylanases, and, most importantly, produces xylose as its main product and may thus be termed xylo-β-1,4-xylanase. β-Xylosidases, which are especially needed in fungi that do not possess GH30 xylanases, are distributed in families GH3, GH43, and GH54. www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

431

432

Fungi

Kubicek 2 4 4 3 1 4 4 7 4 4 4 3 1 2 1

NA NA NA NA NA NA NA NA NA NA NA NA NA NA

Aspergillus flavus

Aspergillus fumigatus

Aspergillus nidulans

Aspergillus niger

Aspergillus oryzae

Aspergillus terreus

Chaetomium globosum

Neosartorya fischeri

Neurospora crassa

Neurospora tetrasperma

Penicillium chrysogenum

Penicillium marneffei

Talaromyces stipitatus

Trichoderma reesei

4

0

GH10

NA

NA

NA

Host

· ·

Starr

Glass 3

4

3

1

2

2

4

9

2

4

2

2

3

4

3

3

8

GH11

5

6

5

1

2

2

1

2

1

0

2

0

1

0

2

1

0

GH30

7

6

5

1

6

6

11

9

8

2

4

8

8

3

7

13

7

GH74

6

4

2

1

0

0

5

2

4

3

4

3

5

3

2

2

0

GH27

1

1

0

1

0

0

2

0

3

2

1

3

2

2

2

0

0

GH36

α-Galactosidases

0

0

0

1

1

1

0

1

0

1

1

3

0

1

0

1

13

GH26

β-Mannanase

3

12

7

14

7

7

20

16

20

19

8

18

18

20

13

9

2

GH43

0

2

1

3

1

1

2

2

4

3

3

3

2

4

2

1

0

GH51

2

3

4

1

1

1

1

0

1

1

0

1

1

1

1

1

0

GH54

α-Arabinosidases

1

2

2

1

0

0

2

4

3

2

0

2

2

2

2

1

0

GH62

1

4

4

2

2

2

5

2

4

5

4

3

5

7

2

2

0

GH35

β-Galactosidases

1

2

2

1

1

1

1

1

2

1

1

1

1

1

1

0

0

GH67

1

1

0

0

2

2

2

2

3

7

0

1

2

6

2

1

0

GH115

β-Glucuronidases

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Aspergillus clavatus

Arthrobotrys oligospora

Allomyces macrogynus

Xyloglucanases

ARI

Saprotrophic

Nutritional style

Xylanases

Table 2 Hemicellulolytic enzymes in saprotrophic and plant-pathogenic fungia

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Necrotrophic

4 5 5 8 2 8

M/D M/D/G M M G M

Fomitiporia mediterranea

Fusarium oxysporum

Fusarium verticillioides

Gaeumannomyces graminis

Heterobasidion annosum

Magnaporthe poae

2

M

Mycosphaerella graminicola

5

4

D

Moniliophthora perniciosa

G

6

M

Magnaporthe oryzae

Dichomitus squalens

9

M

Glomerella graminicola

2

5

M

Fusarium graminearum

M

0

G

Dothistroma septosporum

Botryotinia fuckeliana

5

M

www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

433 2

0

4

3

3

0

0

2

1

2

5

6

2

2

5

2

2

2

2

3

2

2

0

1

1

1

2

0

2

2

4

5

7

7

6

11

6

2

4

3

5

7

5

3

4

3

4

2

1

3

4

5

4

1

1

3

3

2

2

3

1

0

1

3

2

0

0

0

1

0

0

1

2

1

1

1

0

0

0

0

0

0

2

0

0

0

1

0

0

1

16

5

16

22

30

6

7

6

10

3

19

17

19

11

16

2

1

2

2

3

1

2

2

2

1

3

2

2

1

2

0

0

1

1

1

0

0

1

1

0

1

1

1

1

1

1

0

1

1

1

0

0

1

1

0

4

2

1

1

2

2

4

2

3

6

2

3

4

2

2

0

5

3

1

4

1

0

1

3

2

0

0

0

0

0

1

1

1

1

1

(Continued )

2

1

2

3

1

6

4

2

1

1

4

3

3

0

3

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Cochliobolus sativus

ARI

Hemibiotrophic

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Kubicek

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Starr

Glass 3

8 4 4 2 4 4

D

M M M D D D

Phaeosphaeria nodorum

Pyrenophora teres

Pyrenophora tritici

Sclerotinia sclerotiorum

Verticillium albo-atrum

Verticillium dahliae

5

4

3

3

4

7

3

GH11

1

1

0

2

3

2

0

GH30

8

8

3

4

5

5

5

GH74

a Data collected from Reference 132. Abbreviations: D, dicots; G, gymnosperms; M, monocots; NA, not applicable.

GH10

Host

Fungi

Nectria haematococca

Xyloglucanases

4

3

3

4

4

2

1

GH27

0

1

0

2

1

1

1

GH36

α-Galactosidases

0

0

0

0

0

0

0

GH26

β-Mannanase

21

22

5

16

16

13

32

GH43

2

2

2

2

2

2

4

GH51

1

1

1

1

1

1

1

GH54

α-Arabinosidases

0

0

0

2

2

3

2

GH62

4

5

4

4

4

4

8

GH35

β-Galactosidases

1

1

0

1

1

1

0

GH67

3

4

1

2

2

2

3

GH115

β-Glucuronidases

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Nutritional style

Xylanases

ARI

Table 2 (Continued )

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Some of these enzymes, e.g., bifunctional β-D-xylopyranosidase/α-L-arabinofuranosidase of T. reesei, contain several different enzyme activities (48). The mannan-degrading enzymes comprise β-mannanase (1,4-β-D-mannan mannohydrolase; EC 3.2.1.78) and β-mannosidase (1,4-β-D-mannopyranoside hydrolase; EC 3.2.1.25) (76). βMannanases are found in GH5, subfamilies 7 and 8, and GH26. Although the latter also contains enzymes with β-1,3-xylanase activity, enzymes belonging to this family characterized from fungi (e.g., from Aspergillus spp., Humicola spp., N. crassa, Penicillium chrysogenum, and Orpinomyces spp.) were exclusively β-mannanases. β-Mannosidases belong to GH1 or GH2 families.

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Pectin-Degrading Enzymes The main chain of pectin is composed of so-called hairy and smooth regions, the former receiving its name from its significantly sized side chains. The structural differences between these two regions have implications for enzymes necessary to degrade these regions, with the hairy regions needing several additional accessory enzymes. Given that the building block of pectin is a sugar acid, the glycosidic bond can be cleaved by hydrolysis [using polygalacturonases (PGs) EC 3.2.1.15 and 3.2.1.67] and by a nonhydrolytic reaction called β-elimination, which uses pectin lyases (EC 4.2.2.10) and pectate lyases (EC 4.2.2.2). Polygalacturonidases comprise endo- and exoacting enzymes, both of which are accommodated in family GH28. Members are present in all fungal genomes but with variable numbers (Table 3). They are best studied in the saprophytic fungi Aspergillus niger and Rhizopus oryzae (because enzymes from these two fungi are used in the food and feed industry) but also in some phytopathogenic fungi (29, 66). GH28 also contains rhamnogalacturonan hydrolases that cleave the α-1,2-glycosidic bonds formed between D-galacturonic acid and L-rhamnose residues in the hairy regions both by an exo- or endomechanism (125). The polysaccharide lyases (PLs) (EC 4.2.2.-) cleave uronic acid–containing polysaccharide chains via a β-elimination mechanism to generate an unsaturated hexenuronic acid residue and a new reducing end. The CAZy database has classified these enzymes into 21 families (68). PL families 1, 3, and 9 contain fungal pectin/pectate lyases, and family 11 includes fungal rhamnogalacturonan lyases. The homogalacturonan backbone of pectin varies in its degree of methylation from a highly methylated and relatively hydrophobic form to a fully demethylated and highly charged form, which are consequently called pectine and pectate, respectively. Enzymes attacking these structures are termed pectin lyase and pectate lyase, respectively, with the latter being calcium ion dependent for activity. All pectin lyases characterized at present are found in PL1, whereas fungal pectate lyases are found in PL1 and PL3.

Side-Chain Cleaving and Other Accessory Enzymes In addition to the enzymes described above, additional enzymes are required to cleave the linkage to side chains, to remove modifications (such as methyl esters and acetylation), or to split linkages to lignin. A detailed review about these enzymes and their properties has recently been published (63).

OCCURRENCE OF CELL WALL–DEGRADING ENZYMES IN FUNGAL GENOMES As mentioned above, the arms race between plants and phytopathogenic fungi has led to a significant multiplicity in CWDEs. Although some fungi infect both dicot and monocot species, most fungi can only infect either dicots or monocots, which are characterized by different cell wall compositions. It is therefore not surprising that a different enzymatic arsenal for cell wall degradation www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

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Table 3 Pectin depolymerizing enzymes in saprophytic and plant-pathogenic fungia Polygalacturonases Lifestyle

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Saprotrophic

Hemibiotrophic

Necrotrophic

Fungi

Host

GH28

Polygalacturonate lyases

GH78

PL1

PL3

PL4

PL9

Allomyces macrogynus

NA

4

0

2

0

0

0

PL11 0

Arthrobotrys oligospora

NA

7

5

8

3

1

1

0

Aspergillus clavatus

NA

3

0

2

1

2

0

0

Aspergillus flavus

NA

22

14

12

3

3

1

0

Aspergillus fumigatus

NA

13

5

6

3

3

1

0

Aspergillus nidulans

NA

10

9

8

5

4

1

1

Aspergillus niger

NA

20

7

4

0

2

0

0

Aspergillus oryzae

NA

21

10

12

3

5

1

0

Aspergillus terreus

NA

8

4

7

3

3

1

0

Chaetomium globosum

NA

1

1

7

4

2

1

0

Neosartorya fischeri

NA

13

7

6

3

3

1

0

Neurospora crassa

NA

2

1

1

1

1

0

0

Neurospora tetrasperma

NA

2

0

1

1

1

0

0

Penicillium chrysogenum

NA

5

6

5

1

3

0

0

Penicillium marneffei

NA

7

2

2

0

0

0

0

Talaromyces stipitatus

NA

8

2

0

0

0

0

0

Trichoderma reesei

NA

4

1

0

0

0

0

0

Cochliobolus sativus

M

4

6

0

5

4

0

5

Dothistroma septosporum

G

4

1

0

0

2

0

1

Fusarium graminearum

M

6

9

0

7

3

1

7

Glomerella graminicola

M

7

7

0

4

3

1

4

Magnaporthe oryzae

M

4

2

0

1

1

0

4

Moniliophthora perniciosa

D

3

9

0

2

0

0

0 3

Mycosphaerella graminicola

M

Botryotinia fuckeliana

M/D

Dichomitus squalens

G

2

2

0

1

0

0

21

7

0

2

0

0

7

7

0

0

0

1

0

4

Fomitiporia mediterranea

M/D

15

1

0

0

0

0

3

Fusarium oxysporum

M/D/G

15

11

1

7

3

2

16

Fusarium verticillioides

M

8

11

0

7

3

4

8

Gaeumannomyces graminis

M

3

3

0

1

1

0

2

Heterobasidion annosum

G

8

2

0

0

1

0

2

Magnaporthe poae

G

2

3

0

1

1

0

4

Nectria haematococca

D

11

14

1

11

6

1

11

Phaeosphaeria nodorum

M

5

4

0

2

4

0

4

Pyrenophora teres

M

5

3

0

3

4

0

1

Pyrenophora tritici

M

6

3

0

3

4

0

4

Sclerotinia sclerotiorum

D

17

4

0

0

0

0

4

Verticillium albo-atrum

D

14

16

1

13

5

3

10

Verticillium dahliae

D

12

17

1

12

5

2

9

a Data collected from Reference 132. Abbreviations: D, dicots; G, gymnosperms; M, monocots; NA, not applicable.

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is present in these fungi. For example, dicot phytopathogens generally have more genes encoding pectin-degrading enzymes (particularly GH28 PGs and GH105 unsaturated rhamnogalacturonyl hydrolases) than fungi that are pathogenic on monocots (Table 3). These data correlate well with the higher amount of pectin in the cell walls of dicots. Some other dicot pathogens, such as Nectria haematococca, Verticillium albo-atrum, and Verticillium dahliae, also have a greater number of pectate lyase genes. However, although dicot and monocot species have different amounts of hemicellulose in their cell walls, this difference is not reflected in the diversity or number of CWDEs related to hemicellulose degradation in genomes of phytopathogenic fungi. Zhao et al. (132) investigated the genomes of 103 fungi to discover their CWDE content and biodiversity. They found that phytopathogenic fungi in general contain a larger number of CAZymes, such as carbohydrate esterases (CE5 cutinases and xylan esterases) and PL1 pectate lyases, which is reflected in most GH families. However, a few subtle differences were identified. For example, the genomes of most biotrophic fungi, which derive nutrients from living tissues, have the least number of CAZymes and are devoid of GH6 cellobiohydrolases, of the copper-containing lytic polysaccharide monooxygenases in GH61, of GH78 β-glucanases, and of pectate lyases of groups PL1 and PL3. However, the biotroph Cladosporium fulvum is a notable exception to this generalization, as the genome contains significantly more CAZymes in the β-glucosidase family GH3, GH31 α-glucosidases, and GH43 α-L-arabinofuranosidases, endo-α-L-arabinanases, and β-D-xylosidases. A phylogenetic analysis of the PL1 pectate lyase genes revealed that most clades contained only one or a few fungal taxa, and none of the taxa contained representatives of all ancestral paralogs. These data suggest that different subsets of ancestral paralogs may have been lost in certain fungal taxa during evolution (132). This analysis also revealed that the PL1 paralogs of particular phytopathogenic fungal taxa, such as Fusarium spp., underwent lineage or species-specific gene duplication events. In Macrophomina phaseolina, one of the most destructive necrotrophic fungal pathogens [which infects more than 500 plant species throughout the world (114)], the highest number of CEs (especially CE9 and CE10) was detected as compared to all other sequenced fungal genomes (51). Phytopathogenic fungi also have fewer GH76 α-1,6-mannanases and enzymes with CBM1 cellulose-binding domains or CBM18 and CBM50 chitin- and chitin/peptidoglycan-binding domains. Given that the latter domains are part of the chitinolytic enzyme system that fungi use to modify their cell walls and to attach to competing fungi during antagonism, it is tempting to speculate that the lifestyle of biotrophic fungi does not require chitinases for defense. Intriguingly, in Botrytis cinerea and Sclerotinia sclerotiorum, several of the CBM18 modules are found in CE4 chitin deacetylases. Amselem et al. (2) speculated that this arrangement contributes to reducing the release of chitooligosaccharides, which are known to lead to defense responses by host plants. However, necrotrophic plant pathogens, which derive nutrients from dead host cells, are characterized by the highest number and diversity of CAZymes in their genomes. Most of these species are members of the phyla Sordariomycetes, Dothideomycetes, and Leotiomycetes (132).

ROLE OF CELL WALL–DEGRADING ENZYMES IN PLANT PATHOGENESIS A number of studies have been conducted in which the transcriptional responses of fungi grown in planta or in media intended to mimic the plant environment have been assessed. These studies revealed an upregulation of genes encoding secreted proteins, including effectors, which modulate the intracellular host environment and genes encoding CWDEs. For example, when the causative agent of corn smut disease, Ustilago maydis, is grown in an Arabidopsis thaliana infection model, www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

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genes for the degradation of cellulose, including an endoglucanase, and for the degradation of hemicellulose, including an arabinofuranosidase and a xylanase, are upregulated (72). In the rice blast fungus Magnaporthe oryzae, genes encoding cellulases, hemicellulases, and pectate lyases increase in expression level during infection, relative to a minimal media control (73). The observation that phytopathogenic fungi have an expanded arsenal of CWDEs and express genes encoding CWDEs during plant infection challenges the hypothesis that they are not virulence factors. However, whether individual CWDEs indeed fulfill this role has yet to be demonstrated for most plant pathogens. Early studies using strains deleted for various CWDEencoding genes failed to establish a clear role for any particular enzyme as a key pathogenicity factor (3, 121). However, owing to the redundancy of function provided by another enzyme or suite of enzymes common in CWDEs, these findings are not necessarily an argument against the action of CWDEs as virulence factors. Recent examples indicating a positive correlation between certain CWDEs and virulence include studies of the necrotroph pathogen B. cinerea, the wilt pathogen V. dahliae, and the blotch fungus Mycosphaerella graminicola, among others (10, 31, 35, 58). The GH28 PGs are CWDEs for which some evidence as virulence factors is available. These are the first enzymes to be secreted by phytopathogens when they encounter plant cell walls (27). Deletion of the PG pecA gene in Aspergillus flavus reduces lesion development in cotton bolls, whereas expression of this gene in an A. flavus strain that lacks PG increases the size of lesions (107). A PG gene is required for full virulence of B. cinerea on different hosts (119) and of Alternaria citri on citrus fruit (52). Importantly, a Claviceps purpurea strain carrying a deletion of two PG genes (cppg1 and cppg2) renders the fungus nearly nonpathogenic on rye without affecting its vegetative properties (83). The role of PGs as virulence determinants appears to be rather specific for the type of disease rather than the taxonomy of the fungus. Isshiki et al. (52) compared the effect of PGs in A. citri, the cause of Alternaria black rot, and Alternaria alternata rough lemon pathotype, the cause of Alternaria brown spot. These two species are morphologically indistinguishable pathogens of citrus, yet A. citri causes rot by macerating tissues, whereas A. alternata causes necrotic spots by producing a host-selective toxin. Although the PGs produced by these two fungi have similar biochemical properties, and the genes are highly similar, an A. citri strain carrying a PG knockout was reduced in its ability to cause black rot symptoms on citrus as well as in the maceration of potato tissue, whereas similar mutants in A. alternata were unchanged in pathogenicity. The importance of PGs in the fungal attack is also reflected in the formation of PG-inhibiting proteins (PGIPs) by plants, which are located in the plant cell wall, thereby counteracting the action of PGs (26); the PGIPs do not inhibit the plants’ own PGs (16). Furthermore, the restriction of fungal PG activity by PGIPs leads to the accumulation of long-chain oligogalacturonides, which are capable of eliciting defense responses in plants (17, 96). Little evidence is available for other GHs as virulence determinants in plant pathogenesis. One rare example is the xylanase Xyn11A of B. cinerea (10): Its disruption caused a 30% decrease in extracellular xylanase activity, but reduced the average lesion size by more than 70%. However, the defect in virulence in this mutant has been shown to be a loss of necrotizing activity, as opposed to loss of xylanase activity of Xyn11A (81). Additional examples come from approaches that use RNAi to knock down entire sets of GH enzymes. In M. oryzae, RNAi was used to knock down gene expression of xylanases in GH10 and GH11 families as well as endo- and exocellulases in the GH6 and GH7 families (80, 127). A decrease in the production of numerous enzymes was associated with a concomitant decrease in virulence. Interestingly, knockdowns of the GH10 and GH11 families yielded greater defects in virulence as compared with knockdowns of GH6 and GH7 families, although it was unclear if this result was due to a greater importance of xylanases during plant infection (127).

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SECRETION OF FUNGAL CELL WALL–DEGRADING ENZYMES

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To attack plant cell walls, CWDEs must be secreted across the fungal plasma membrane. In addition, extracellular enzymes must be protected against adverse conditions that prevail on the exterior of the cell, such as very low protein concentration, adverse pH, and proteolytic attack (28). Most of our detailed information on the fungal secretory pathway comes from studies on the yeast Saccharomyces cerevisiae (103), but knowledge based on selected steps within this process is also available for some filamentous fungi (mostly Trichoderma and Aspergillus). The role of secretion of CWDEs and their importance in phytopathogenic fungi and virulence is currently not well characterized, with the exception of recent work in the rice blast pathogen M. oryzae (see below) (39).

General Overview of Fungal Protein Secretion The secretory pathway of fungi involves several cellular endomembrane systems. This pathway transports proteins destined for other cellular locations, such as the plasma membrane, the vacuole/lysosome, or the golgi complex, as well as extracellular proteins (for reviews, see 19, 50, 99, 108, 110) (Figure 1a). Proteins destined for the secretory pathway often carry signal sequences at their N termini, the so-called signal peptide. There are two pathways that recognize this signal peptide: processes associated with cotranslational translocation and those with posttranslational translocation (133). During cotranslational translocation, the newly translated signal peptide sequence is recognized by the signal recognition particle (SRP), causing a temporary stop in translation. The SRP directs the ribosome, mRNA, peptide ternary complex to the translocation complex on the membrane of the endoplasmic reticulum (ER). The translocation complex is composed of several proteins including SEC61, which forms a channel through the ER membrane (23). Once docking has been achieved, translation continues and the nascent polypeptide chain is translocated into the ER lumen simultaneously with its translation. In the process of posttranslational translocation, fully synthesized precursor polypeptides associate with cytosolic molecular chaperones, which maintain the precursor polypeptides in a soluble state, of the Hsp70 and Hsp40 heat shock protein family. A heterotrimeric complex of three membrane proteins (Sec62p, Sec71p, and Sec72p) recognizes the signal peptide enabling the passage of the polypeptide through the Sec61ptranslocon complex. The importance of the process for secretion of CWDEs is evident based on the observation that transcriptional expression of the translocation components is increased when N. crassa is grown on cellulose (6, 20). Additionally, in M. oryzae, the LHS-1 homolog, which is a nucleotide exchange factor for BiP/Kar2 (112), is important for pathogenesis, and as a deletion mutant was defective for translocation of proteins and showed reduced infection ability as well as significantly decreased xylosidase, arabinosidase, glucanase, and laccase activity (130). Once they arrive in the ER, CWDEs are folded into their final conformation, which is assisted by chaperones that bind to hydrophobic regions of the polypeptide and prevent unfavorable interactions. The major ER chaperone BiP/Kar2 assists in this process as proteins enter the lumen (12, 77). Many secreted proteins, including CWDEs, have disulfide bridges. Disulfide bond formation is catalyzed by the protein disulfide isomerases or the ERV1 family proteins that bear thiol oxidase activity (85, 106). Additionally, the isomerization of peptide bonds preceding a proline residue is catalyzed by the prolyl-peptidyl cis/trans isomerases (9). Calnexin plays a special role in the folding of nascent glycoproteins by mediating correct folding and processing of the N-glycans (discussed below). If this quality control mechanism fails to yield a properly folded protein, the glycoprotein dissociates from calnexin and enters a quality control checkpoint exerted by the BiP/Kar2 chaperone complex (75). This complex preferentially binds to hydrophobic patches of a misfolded protein. Prolonged binding of a misfolded protein to either www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

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calnexin or the BiP complex results in activation of the unfolded protein response (UPR) and targeting of the polypeptides to ER-associated protein degradation (ERAD) (11, 56).

Unfolded Protein Response

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The accumulation of unfolded proteins in the ER activates UPR, resulting in the triggering of a gene expression program that adjusts the protein-folding capacity of the cell according to need (22, 87). The UPR is induced under conditions of secretory stress, which can be brought about by environmental perturbations that inhibit the ability of proteins to fold in the ER, as well as by increased secretory load. The primary transcriptional activator of the UPR is the HAC1/HacA transcription factor, which is both induced in expression and processed into an active form during stress. This transcription factor is induced during growth of N. crassa on plant cell wall material (6) as well as in T. reesei and Aspergillus species during secretion of heterologous proteins (43, 101).

a

Cell wall

Structure Osmotic resistance Secretion barrier Plasma membrane

Septum

Glycoprotein modification Protein sorting

Translocation Folding ER Glycosylation ERAD UPR Nucleus Vesicle biogenesis

Spitzenkörper Golgi

Degradation Storage

) lar

(tu

Vacuole (round)

bu

Tip trafficking

o cu Va

Exocyst

le

PM t-SNAREs

Endocytosis Protein recycling EE Vacuolar pathway

LE

Budding vesicle

Secreted proteins

b Plant cell wall Plant plasma membrane

BIC Plant nucleus

Plant cytosol ER Nucleus Nucleus Golgi

Golgi

ER

Bulbous invasive hypha Plant PM-derived EIHM

Exocyst

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t-SNARE SSO1

Apoplastic effector proteins

Cytosolic effector proteins

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Deletion of hacA in the human pathogen and saprotroph Aspergillus fumigatus caused damage to the fungal cell wall, reduced protease secretion, and reduced virulence in a mouse model (95). The importance of HAC1/HacA has also been demonstrated in phytopathogenic fungi. For example, during infection of sorghum by Bipolaris sorghicola, transcription of hacA was increased in parallel with expression of CWDEs (129). Additionally, Joubert et al. (54) showed that a mutant variant of the necrotrophic fungus Alternaria brassicicola that lacks hacA exhibited decreased virulence, as well as decreased protein secretion, including decreased esterase and protease activities, highlighting the importance of the UPR in CWDE secretion.

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Glycosylation Simultaneously with translocation across the ER and folding, carbohydrates are attached to most of the newly synthesized secreted proteins via either an asparagine (N-glycosylation) or serine/ threonine residue (O-glycosylation); many proteins carry both types of glycosylation. These modifications influence the secretion of proteins and their resulting enzymatic activities. These pathways are discussed briefly here, but the reader is directed to several excellent reviews that specifically treat the subject in greater detail (36, 42, 67, 104). Little work has been done to characterize the role that glycosylation plays in secretion of CWDEs from phytopathogenic fungi, with the exception of analyzing the effects of impaired Olinked glycosylation. In fungi, O-linked glycosylation involves the initial addition of a single mannose residue to serines or threonines of proteins within the ER lumen. This process is catalyzed by protein-O-mannosyltransferases (PMTs), which fall into three different phylogenetic classes (69). These enzymes have been shown to have unique protein substrate specificities, as strains carrying deletion of each PMT result in differential protein glycosylation defects and cellular phenotypes (41, 62, 78). Mutations in the PMTs result in defects in glycosylation of not only soluble secreted proteins but also of cell surface glycoproteins, which can result in major changes to cell morphology and growth as well as virulence. For example, in Aspergillus nidulans, a strain carrying a deletion ←−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−− Figure 1 Secretion in vegetative and invasive hypha. (a) General model of the compartments and pathways of the secretory pathway in vegetative hyphae in filamentous fungi, as influenced by information from yeast and filamentous fungal studies. Tip-directed and lateral secretion are depicted, as is the localization of vesicle docking (exocyst) and fusion (t-SNARE) machineries at the plasma membrane. Arrows show potential routes of vesicle-mediated transport between the compartments. Compartments of the endocytic-vacuolar pathway are included, as is the Spitzenkorper, which is specialized for tip-directed secretion. Budding vesicles ¨ containing secreted proteins are also shown. (b) Depiction of the distinct secretory pathways utilized by Magnaporthe oryzae during in planta invasive growth, as proposed by Giraldo et al. (39). Apoplastic effectors are secreted by a pathway that utilizes the golgi apparatus. These effectors reside in the apoplastic space between the hyphal exterior and the extrainvasive hyphal membrane (EIHM), which is derived from the plant cell plasma membrane (shown in blue). Effectors delivered to the plant cytoplasm are first secreted into the biotrophic interfacial complex (BIC) via a pathway that is presumably golgi-independent. Note that the exocyst is utilized by both pathways, whereas the t-SNARE SSO1 is more important for secretion into the BIC. Spitzenkorper markers are present in the BIC-associated cell. The role of the Spitzenkorper in ¨ ¨ cytoplasmic effector secretion is unclear, and therefore this structure is not shown. Additionally, the Spitzenkorper is not present in the bulbous invasive hypha, and therefore apparently not required for ¨ apoplastic effector secretion, although this structure is present when M. oryzae grows vegetatively. Adapted with permission from Reference 39. Abbreviations: EE, early endosome; ER, endoplasmic reticulum; ERAD, endoplasmic reticulum–associated degradation; LE, late endosome; PM, plasma membrane; t-SNARE, target soluble N-ethylmaleimide-sensitive factor attachment protein receptor; UPR, unfolded protein response. www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

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of pmtA causes hyphal cell walls to display altered carbohydrate content with reduced skeletal polysaccharides (84). In U. maydis, deletion of pmt4 resulted in reduced virulence, both at early and later time points of infection. It was hypothesized that the early defect in virulence was due to a defective cell wall stress response, whereas the defect at the later infection time point was independent of cell wall defects. The authors observed alterations in the glycosylation patterns of secreted effector proteins and suggested this as the cause of reduced virulence at the later time point (37). In B. cinerea, individual deletions of the three genes encoding PMT enzymes yielded strains with reduced virulence, including reduced ability to adhere and penetrate plant surfaces, likely caused by fungal cell wall defects, as the mutants exhibited poor growth, swollen cells, and a loss of extracellular matrix. Additionally, altered glycosylation profiles in CWDEs important for virulence, including glucoamylase, α-L-arabinofuranosidase and endopolygalacturonase 1, were observed (41).

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Vesicle-Mediated Protein Trafficking After being synthesized and folded, secreted proteins exit the ER and are delivered to the golgi apparatus for further modification and sorting to their final destination (i.e., plasma membrane, export, or vacuole). Vesicle formation and transport as well as fusion processes are controlled by a large number of proteins and protein complexes, some of which function in multiple membrane trafficking events, whereas others are specific. Key regulators of the different steps of the pathway are small GTPases. These include the SAR1/ARF1-type GTPases involved in vesicle budding, Rho-type GTPases involved in membrane scaffolding, and the Rab-type GTPases involved in vesicle tethering (50). After budding, vesicles are transported from the donor membrane to the target membrane, where they are tethered and fused. Vesicle tethering events are mediated by Rab GTPases, which bind to specific membrane compartments and recruit particular sets of effector proteins, including oligomeric tethering complexes (50). In S. cerevisiae, the Rab protein Sec4 recruits the exomer complex to the PM and thus determines the site of exocytic vesicle fusion (45). Exocyst localization has been shown to localize to the hyphal tip, the main site of vesicle fusion, in A. nidulans, N. crassa, and M. oryzae. In N. crassa, it has been shown that the tip is the main site of vesicle fusion (39, 97, 117). Sec4 function in A. niger has been shown to be nonessential, whereas Sec4 function is essential in S. cerevisiae (92). However, in Colletotrichum lindemuthianum, expression of the SEC4 homolog (CLPT1) is induced when the fungus is grown on pectin. When a dominant negative version of CLPT1 was overexpressed in C. lindemuthianum, secretion of pectinases was reduced with a concomitant reduction in plant penetration and pathogenesis (109). After tethering, vesicle fusion events are mediated by the v-SNARE- and t-SNARE-type proteins on the vesicle and on the target membrane, respectively. The SNARE proteins recognize and bind each other during the vesicle docking stage and provide specificity for each vesicle fusion event (60, 71). A systematic study of SNAREs in Aspergillus oryzae showed localization to a variety of intracellular compartments, providing insights about possible secretory routes (65). Indeed, the localization of PM SNARE proteins in filamentous fungi has pointed to specific transport routes for lateral secretion. In S. cerevisiae, PM vesicle fusion begins with the formation of binary complexes between two t-SNAREs, Ssop1/2p and Sec9. This complex then binds the vSNARE Snc1p (1). Two genes encoding syntaxin-like t-SNAREs are present in filamentous fungal genomes, including N. crassa, A. oryzae, A. niger, and T. reesei. Interestingly, in T. reesei, SSO1 interacts with the v-SNARE Snc1p in the subapical/basal regions of the hyphae, whereas SSO2 interacts with SNC1 in the apical regions (124). These data suggest that the two SSO proteins are involved in distinct secretion pathways in filamentous fungi. Further evidence for alternate routes is the observation that some proteins do not apparently pass through the Spitzenkorper, a specialized ¨ 442

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collection of vesicles localized just under the growing hyphal tip of filamentous fungi that serves as a vesicle supply center for tip-directed vesicles (97). For example, amino acid permeases and the major plasma membrane proton pump PMA-1 do not localize to the Spitzenkorper, but are ¨ secreted to the lateral wall (32, 44). These results support the concept of exocytosis to subapical hyphal regions (94). In A. oryzae, it has been shown that extracellular α-amylase-GFP rapidly accumulates in the periplasm between the plasma membrane and cell wall of the septa (94). In T. reesei, CBH1 localized along the length of the cell surface, suggesting lateral secretion (82). The existence of multiple secretion pathways in filamentous fungi may provide an explanation for their high secretion capacity.

A Case Study of Protein Secretion in the Phytopathogenic Fungus Magnaporthe oryzae Comprehensive molecular and cell biological studies of protein secretion in phytopathogenic fungi are scarce. However, recent work by Giraldo et al. (39) evaluated the role of protein secretion in pathogenicity of M. oryzae on rice. The authors provided evidence for two distinct routes for secretion of effector proteins during biotrophic invasion of the host (Figure 1b). Apoplastic effectors accumulate extracellularly at the host-pathogen interface and are actively secreted via the conventional secretory process described above. In contrast, cytoplasmic effector proteins destined for delivery inside rice cells are secreted by a different pathway that involves the exocyst complex and biotrophic interfacial structures (BICs). The latter are plant-derived, membrane-rich structures that associate with invasive hyphae (59). Cytoplasmic effector proteins accumulated predominantly inside the BIC-associated cells in exo70 and sec5 mutants rather than at hyphal tips, suggesting that these effector proteins may be secreted predominantly from BIC-associated cells. The t-SNARE Sso1 also localized adjacent to the BIC, and sso1 mutants showed defects in normal BIC development. However, it is currently unclear whether other phytopathogenic fungi have similar pathways or have developed different secretory pathways for effector proteins and possibly CWDEs during plant pathogenesis.

REGULATION OF CELL WALL–DEGRADING ENZYME FORMATION IN RELATION TO PLANT PATHOGENESIS Understanding the genetic pathways that regulate how plant-pathogenic fungi respond to their environment is paramount to developing effective mitigation strategies against disease. In addition, identification of the transcriptional activators that govern the expression of CWDE-encoding genes would be an effective means to reassess the relevance of these enzymes to plant pathogenesis. Regulation of expression of CWDE genes has so far been a subject of major interest in saprophytic fungi, which are forced by the need to manipulate their transcription toward improved enzyme mixes for lignocellulose hydrolysis for the generation of monosaccharides for biofuels and biorefineries (40, 105, 134). The regulation of CWDEs in plant-pathogenic fungi has received considerably less attention. Most of the work has concentrated on wide-domain regulators, such as carbon catabolite repression and pH control, whose involvement in CWDE expression has already been demonstrated in saprophytic fungi.

Carbon Catabolite Repression Carbon catabolite repression (CCR) is a global regulatory mechanism found in a wide range of microorganisms that serves to ensure the preferential utilization of glucose or other easily www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

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metabolizable carbohydrates over less-favorable carbon sources. Its regulatory role in the expression of CWDEs in saprophytic fungi has been well established (90, 115). Despite the importance of this process to fungal physiology, little is known about the proteins mediating CCR in filamentous fungi. The only well-documented component is the C2H2-type transcriptional regulator CreA/CRE1, which functions as the main repressor protein in this process (33). Despite the central role of this protein in saprophytic fungi, there is little evidence for its importance during CWDE expression during plant pathogenesis. For example, in A. citri, overexpression of creA had little effect on endopolygalacturonase formation, although the mutant caused severe black rot symptoms in both the central axis and juice sac areas, indicating a role for CreA in another process relevant to pathogenesis (57). These data are comparable to those by Dean & Timberlake (25), who demonstrated that inoculation of excised plant tissues with conidia of A. nidulans leads to formation of necrotic, water-soaked lesions, within which the organism sporulates but that were independent of CreA. Similarly, CreA did not influence CWDE expression in A. brassicicola (18). However, in Fusarium oxysporum f. sp. lycopersici, CRE1 cooperates with the F-box protein FRP1 in regulating the expression of several CWDEs (53), but unfortunately the effect of this regulation on pathogenicity has not been studied. More recently, Fernandez et al. (34) employed a genome-wide approach to identify three new mediators of CCR in M. oryzae: the sugar sensor Tps1 (a trehalose phosphate synthase), an Nmr1-3 inhibitor protein, and a multidrug and toxin extrusion pump, Mdt1. Fernandez et al. demonstrated that Tps1, in response to glucose-6-phosphate sensing, triggers CCR via the inactivation of Nmr1-3. Mdt1 regulates glucose assimilation downstream of Tps1 and is necessary for sporulation, pathogenicity, and nutrient utilization. A strain carrying a deletion of tps1 and mdt1 resulted in overexpression of a β-glucosidase and an exoglucanase gene and caused defects in pathogenesis. The authors speculated that overexpression of genes encoding CWDEs may disturb the carbon assimilatory balance during the early phase of infection and consequently impair the plant-pathogenic ability. In S. cerevisiae, CCR involves the action of the serine/threonine kinase Snf1p (86), which regulates nuclear entry of Mig1p by phosphorylation (whose DNA binding domain is orthologous to that of CreA/CRE1) (30). A possible role for Snf1 orthologs in CCR in phytopathogenic fungi has been tested, but a clear role during pathogenesis has not emerged. For example, in Cochliobolus carbonum, a strain carrying a deletion of snf1 resulted in the downregulation of several CWDEencoding genes (XYL1, XYL2, XYL3, XYL4, XYP1, ARF1, MLG1, EXG1, PGN1, and PGX1) and also reduced virulence at the stage of penetration of the plant (122). Similarly, knocking out snf1 in F. oxysporum resulted in a reduced expression of several genes encoding CWDEs and delayed the progression of wilt symptoms. However, in M. oryzae, a strain carrying a deletion of snf1 did not affect the expression of the CWDE genes but significantly reduced the prepenetration stage, including conidial germination and appressorium formation (131).

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pH Control and Pathogenesis Control of gene expression by pH is another global regulatory circuit in fungi. Its relevance to plant pathogenesis stems from the fact that pH is one of the major ambient traits that affects the activity of virulence factors secreted by the pathogen; hence, a pH sensing-response system enables a pathogen to tailor its arsenal to best fit its host (91). This regulatory system has been studied in detail in the saprophyte A. nidulans (88, 89). Alkaline pH is signaled from the plasma membrane to the endosomal membrane, where the zinc finger transcription factor PacC undergoes a pHdependent proteolysis-enabling activator function for alkaline-responsive genes and repressor function for acid-responsive genes. The components of this signal transduction pathway include 444

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palA, B, C, F, H, and I, which are also present in the genomes of other fungi (89). CWDEs, particularly xylanases, were recognized to be controlled by environmental pH and later shown to be regulated by the PacC pH regulatory system. The involvement of pH-controlled gene expression in fungal virulence has been documented in several systems. Tissue alkalinization during a Colletotrichum gloeosporioides attack on avocado enhances the expression of the pectate lyase pelB (74). Loss-of-function mutants in the C. gloeosporioides pacC ortholog, pac1, showed an 85% reduction of pelB transcript expression, delayed PL secretion, and dramatically reduced virulence. In contrast, pacC knockout mutants of Fusarium oxysporum were more virulent in root infection assays with tomato plants than was the wild-type strain, whereas pacC overexpression strains were significantly reduced in virulence, indicating that PacC acts as a negative regulator of virulence in this fungus (15).

Specific Regulators of Cell Wall–Degrading Enzyme Gene Expression The Zn(II)Cys(VI) transcriptional activator XlnR/XYR1 is a major positive activator of the expression of CWDEs in a number of filamentous fungi, although the enzymes that are actually regulated by it differ in different fungi (13, 105, 134). In N. crassa and A. nidulans, expression of genes encoding cellulolytic enzymes is regulated by a Zn(II)Cys(VI) transcriptional activator, CLR2/CLRB (20), whereas genes involved in hemicellulose degradation are regulated by a homolog of XlnR (116). Similarly, in F. oxysporum, a strain carrying a deletion of xlnR lacked transcriptional activation of structural xylanase genes in culture and during infection of tomato plants, as well as in dramatically reduced extracellular xylanase activity, but did not affect virulence on tomato plants (14). So far, a specific regulator of pectinase or pectin/pectate lyase gene expression in fungi has not been reported. In S. cerevisiae, expression of the gene encoding the pectinolytic enzyme PGU1 is regulated by the transcription factors TEC1 and STE12 (70). A BLASTP search against several fungal genomes present in the JGI or Broad databases did not reveal the presence of genes encoding a protein similar to TEC1 (C.P. Kubicek, unpublished data). However, homologs to the Ste12 transcription factor are specific to the fungal kingdom and are present in all members of the subphylum Pezizomycotina. In S. cerevisiae, Ste12p regulates mating and invasive/pseudohyphal growth. In phytopathogenic fungi, Ste12 knockout strains are impaired in the development of the highly melanized appressoria and reduced virulence was shown for Cryphonectria parasitica, B. cinerea, A. brassicicola, and F. oxysporum (reviewed in 128). In the latter species, pathogenesis on tomato tissues and the ability to penetrate cellophane membranes were impaired, and amylase and cellulase activities were also reduced. However, Cho et al. (18) did not find a reduction of utilization of polysaccharides in STE12 deletion strains of A. brassicicola. Srivastava et al. (111) identified a zinc-finger-family transcription factor, AbVf19 in A. brassicicola, which was important for lesion development. Its deletion in A. brassicicola led to a decrease in expression of some cellulase and pectinase genes.

CONCLUDING REMARKS Evidence for the need of plant CWDEs for attack by plant-pathogenic fungi on their hosts can be deduced from differences in the enzyme arsenal between saprophytic and plant-pathogenic fungi, as well as between plant-pathogenic fungi that utilize different modes of pathogenesis. However, functional evidence for the essentiality of CWDEs in virulence has been reported for only some enzymes (particularly those involved in pectin/pectate depolymerization). In addition, some of the transcription factors that regulate expression of genes encoding CWDEs have been identified www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

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and have further supported the critical role of these enzymes in plant pathogenesis. The present evidence suggests that there is no general mechanism that can explain the behavior of all plantpathogenic fungi, but significant differences in the importance of specific CWDEs or CWDE families are evident even between species from the same genus. In this regard, we note that there is particularly a general lack of knowledge about the transcription factors governing the expression of CWDE-encoding genes during plant infection and cell wall utilization and, consequently, the plant-derived signals that activate these fungal transcription factors. In addition, the possible importance of the secretory pathway by which these enzymes are transported out of the cell has been almost completely neglected, although recent exciting results with M. grisea have shown its importance for pathogenesis and further demonstrated significant differences to the mechanism used by saprophytic fungi. Further work on the characterization of transcription and secretion of CWDEs in different plant-pathogenic fungi will enable the elucidation of biochemical mechanisms that may identify new specific targets to protect plants and expedite development of new agents for combating plant-pathogenic fungi.

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DISCLOSURE STATEMENT The authors are not aware of any affiliations, memberships, funding, or financial holdings that might be perceived as affecting the objectivity of this review.

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62. Kriangkripipat T, Momany M. 2009. Aspergillus nidulans protein O-mannosyltransferases play roles in cell wall integrity and developmental patterning. Eukaryot. Cell 8:1475–85 63. Kubicek CP. 2012. Fungi and Lignocellulose Biomass. New York: Wiley and Sons 64. Kurasin M, Valjamae P. 2011. Processivity of cellobiohydrolases is limited by the substrate. J. Biol. Chem. 286:169–77 65. Kuratsu M, Taura A, Shoji JY, Kikuchi S, Arioka M, Kitamoto K. 2007. Systematic analysis of SNARE localization in the filamentous fungus Aspergillus oryzae. Fungal Genet. Biol. 44:1310–23 66. Lara-M´arquez A, Zavala-P´aramo MG, Lopez-Romero E, Camacho HC. 2011. Biotechnological poten´ tial of pectinolytic complexes of fungi. Biotechnol. Lett. 33:859–68 67. Lehle L, Strahl S, Tanner W. 2006. Protein glycosylation, conserved from yeast to man: a model organism helps elucidate congenital human diseases. Angew. Chem. Int. Ed. 45:6802–18 68. Lombard V, Bernard T, Rancurel C, Brumer H, Coutinho PM, Henrissat B. 2010. A hierarchical classification of polysaccharide lyases for glycogenomics. Biochem. J. 432:437–44 69. Lommel M, Strahl S. 2009. Protein O-mannosylation: conserved from bacteria to humans. Glycobiology 19:816–28 70. Louw C, Young PR, van Rensburg P, Divol B. 2010. Regulation of endo-polygalacturonase activity in Saccharomyces cerevisiae. FEMS Yeast Res. 10:44–57 71. Malsam J, Kreye S, Sollner TH. 2008. Membrane fusion: SNAREs and regulation. Cell. Mol. Life Sci. 65:2814–32 72. Martinez-Soto D, Robledo-Briones AM, Estrada-Luna AA, Ruiz-Herrera J. 2013. Transcriptomic analysis of Ustilago maydis infecting Arabidopsis reveals important aspects of the fungus pathogenic mechanisms. Plant Signal Behav. 8:pii:e25059 73. Mathioni SM, Belo A, Rizzo CJ, Dean RA, Donofrio NM. 2011. Transcriptome profiling of the rice blast fungus during invasive plant infection and in vitro stresses. BMC Genomics 12:49 74. Miyara I, Shafran H, Kramer Haimovich H, Rollins J, Sherman A, Prusky D. 2008. Multi-factor regulation of pectate lyase secretion by Colletotrichum gloeosporioides pathogenic on avocado fruits. Mol. Plant Pathol. 9:281–91 75. Molinari M, Galli C, Vanoni O, Arnold SM, Kaufman RJ. 2005. Persistent glycoprotein misfolding activates the glucosidase II/UGT1-driven calnexin cycle to delay aggregation and loss of folding competence. Mol. Cell 20:503–12 76. Moreira LR, Filho EX. 2008. An overview of mannan structure and mannan-degrading enzyme systems. Appl. Microbiol. Biotechnol. 79:165–78 77. Morrow MW, Janke MR, Lund K, Morrison EP, Paulson BA. 2011. The Candida albicans Kar2 protein is essential and functions during the translocation of proteins into the endoplasmic reticulum. Curr. Genet. 57:25–37 78. Mouyna I, Kniemeyer O, Jank T, Loussert C, Mellado E, et al. 2010. Members of protein Omannosyltransferase family in Aspergillus fumigatus differentially affect growth, morphogenesis and viability. Mol. Microbiol. 76:1205–21 79. Murphy C, Powlowski J, Wu M, Butler G, Tsang A. 2011. Curation of characterized glycoside hydrolases of fungal origin. Database 2011:bar020 80. Nguyen QB, Itoh K, Van Vu B, Tosa Y, Nakayashiki H. 2011. Simultaneous silencing of endo-β-1,4 xylanase genes reveals their roles in the virulence of Magnaporthe oryzae. Mol. Microbiol. 81:1008–19 81. Noda J, Brito N, Gonzalez C. 2010. The Botrytis cinerea xylanase Xyn11A contributes to virulence with its necrotizing activity, not with its catalytic activity. BMC Plant Biol. 10:38 82. Nykanen M, Saarelainen R, Raudaskoski M, Nevalainen K, Mikkonen A. 1997. Expression and secretion of barley cysteine endopeptidase B and cellobiohydrolase I in Trichoderma reesei. Appl. Environ. Microbiol. 63:4929–37 83. Oeser B, Heidrich PM, Muller U, Tudzynski P, Tenberge KB. 2002. Polygalacturonase is a pathogenicity factor in the Claviceps purpurea/rye interaction. Fungal Genet. Biol. 36:176–86 84. Oka M, Maruyama J-I, Arioka M, Nakajima H, Kitamoto K. 2004. Molecular cloning and functional characterization of avaB, a gene encoding Vam6p/Vps39p-like protein in Aspergillus nidulans. FEMS Microbiol. Lett. 232:113–21 www.annualreviews.org • Plant Cell Wall–Degrading Enzymes

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Annual Review of Phytopathology Volume 52, 2014

How Way Leads on to Way Isaac Barash p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 1 Harnessing Population Genomics to Understand How Bacterial Pathogens Emerge, Adapt to Crop Hosts, and Disseminate Boris A. Vinatzer, Caroline L. Monteil, and Christopher R. Clarke p p p p p p p p p p p p p p p p p p p p p p p19 New Insights into Mycoviruses and Exploration for the Biological Control of Crop Fungal Diseases Jiatao Xie and Daohong Jiang p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p45 Altering the Cell Wall and Its Impact on Plant Disease: From Forage to Bioenergy Qiao Zhao and Richard A. Dixon p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p69 Network Modeling to Understand Plant Immunity Oliver Windram, Christopher A. Penfold, and Katherine J. Denby p p p p p p p p p p p p p p p p p p p p p p p93 The Role of Trees in Agroecology and Sustainable Agriculture in the Tropics Roger R.B. Leakey p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 113 Plant-Parasitic Nematode Infections in Rice: Molecular and Cellular Insights Tina Kyndt, Diana Fernandez, and Godelieve Gheysen p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 135 Mechanisms of Nutrient Acquisition and Utilization During Fungal Infections of Leaves Jessie Fernandez, Margarita Marroquin-Guzman, and Richard A. Wilson p p p p p p p p p p p p 155 Governing Principles Can Guide Fungicide-Resistance Management Tactics Frank van den Bosch, Richard Oliver, Femke van den Berg, and Neil Paveley p p p p p p p p p 175 Virus Infection Cycle Events Coupled to RNA Replication Pooja Saxena and George P. Lomonossoff p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 197

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Novel Insights into Rice Innate Immunity Against Bacterial and Fungal Pathogens Wende Liu, Jinling Liu, Lindsay Triplett, Jan E. Leach, and Guo-Liang Wang p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 213 The Activation and Suppression of Plant Innate Immunity by Parasitic Nematodes Aska Goverse and Geert Smant p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 243

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Protein Kinases in Plant-Pathogenic Fungi: Conserved Regulators of Infection David Turr`a, David Segorbe, and Antonio Di Pietro p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 267 Speciation in Fungal and Oomycete Plant Pathogens Silvia Restrepo, Javier F. Tabima, Maria F. Mideros, Niklaus J. Grunwald, ¨ and Daniel R. Matute p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 289 The ABCs and 123s of Bacterial Secretion Systems in Plant Pathogenesis Jeff H. Chang, Darrell Desveaux, and Allison L. Creason p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 317 Induced Systemic Resistance by Beneficial Microbes Corn´e M.J. Pieterse, Christos Zamioudis, Roeland L. Berendsen, David M. Weller, Saskia C.M. Van Wees, and Peter A.H.M. Bakker p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 347 Fifty Years Since Silent Spring Lynn Epstein p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 377 Localizing Viruses in Their Insect Vectors St´ephane Blanc, Martin Drucker, and Marilyne Uzest p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 403 Plant Cell Wall–Degrading Enzymes and Their Secretion in Plant-Pathogenic Fungi Christian P. Kubicek, Trevor L. Starr, and N. Louise Glass p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 427 Meta-Analysis and Other Approaches for Synthesizing Structured and Unstructured Data in Plant Pathology H. Scherm, C.S. Thomas, K.A. Garrett, and J.M. Olsen p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 453 Networks and Plant Disease Management: Concepts and Applications M.W. Shaw and M. Pautasso p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 477 Small RNAs: A New Paradigm in Plant-Microbe Interactions Arne Weiberg, Ming Wang, Marschal Bellinger, and Hailing Jin p p p p p p p p p p p p p p p p p p p p p p 495 Predisposition in Plant Disease: Exploiting the Nexus in Abiotic and Biotic Stress Perception and Response Richard M. Bostock, Matthew F. Pye, and Tatiana V. Roubtsova p p p p p p p p p p p p p p p p p p p p p p p p 517

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Susceptibility Genes 101: How to Be a Good Host Chris C.N. van Schie and Frank L.W. Takken p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 551 Horizontal Gene Transfer in Eukaryotic Plant Pathogens Darren Soanes and Thomas A. Richards p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 583 Errata

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Plant cell wall-degrading enzymes and their secretion in plant-pathogenic fungi.

Approximately a tenth of all described fungal species can cause diseases in plants. A common feature of this process is the necessity to pass through ...
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