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4.3 Degradation of Walls and Membranes by Microbial Enzymes

D.F. BATEMAN and H.G. BASHAM

1. Introduction

One characteristic feature of many phytopathogenic organisms is their ability to produce an array of enzymes capable of degrading the complex of the plant (BATEMAN and MILLAR, 1966; , 1967; ALBERSHEIM et aI., 1969; WOOD, 1973) and membrane constituents (PORTER, 1966; TSENG and BATEMAN, 1968). These enzymes usually are produced inductively. Generally, they are extracellular, highly stable and present in infected host tissues. In most plant caused by microbial agents, cell walls are penetrated, tissues are colonized, and permeability of host cells is altered. A brief summary of our understanding of cell wall and membrane structure, coupled with knowledge of the enzymes capable of degrading the components of these structures, and an analysis of the association of these enzymes with diseased , should enable us to make an appraisal of their involvement in pathogenesis and point the way to an objective consideration of this area of physiology.

2. Structural Components

2.1 Cell Wall Composition and Structure

The wall is that structure surrounding the exterior to the plasmalemma. This structure may be viewed as a two-phase system--- a dispersed phase of microfibrils and a complex continuous matrix. During develop• ment cell walls undergo conspicious changes in structure, form and function, and they exhibit dynamic, rapid changes in their various constituents (NEVINS et aI., 1968; BERLYN, 1970; NORTHCOTE, 1972). Cell walls in young tissues are composed primarily of polysaccharides and a structural rich in (LAM• PORT, 1970). In older tissues, walls may also contain . Traditionally, the cell wall has been divided into three functional-structural regions: middle , primary wall and secondary wall. Middle lamella is that region where the walls of two cells join in a tissue system. The primary wall is the first wall region formed with definite organization, and the most dynamic of the wall regions. Secondary wall is that wall portion added after cell elongation is complete. Differences between wall regions relate to chemical composition and the degree of organization; transitions from region to region are not abrupt but intergrade gradually into one another (Fig. 1).

R. Heitefuss et al. (eds.), Physiological Plant © Springer-Verlag Berlin · Heidelberg 1976 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 317

Microfibrils

Continuous matrix

Middle Primary Secondary Lumen lamella wall wall

Hemicellulose

Fig. I. Distribution in the plant cell wall of the major wall consti• Protein tuents

The wall polysaccharides have historically been grouped into the pectic substan• ces, , and cellulose (NORTHCOTE, 1963). This grouping is based upon solubilities of the constituents rather than upon their chemical compositions. The pectic substances are composed primarily of rhamnogalacturo• nans, galactans and arabans (ASPINALL, 1970); they are extracted with cold and hot water and solutions of chelating agents. The hemicellulosic fraction includes , , and heteropolymers of , galactose and man• nose. These substances are extracted in alkaline solutions. Cellulose is the residual wall polysaccharide remaining after the above extractions. This empirical means of wall fractionation leads to the isolation of heterogeneous mixtures of wall polysaccharides and polysaccharide fragments. The pectic substances are the primary constituents of the middle lamella and are structural elements in the primary wall (MCCLENDON, 1964; TALMADGE et a!., 1973). The major component of the pectic fraction is a high molecular weight consisting of a backbone of IX-I,4-linked D-galacturonopyranose intersper• sed with 1,2-linked rhamnopyranose. The uronic carboxyls may be methylated and the uronide moieties may be acetylated at positions 2 and 3. The neutral , consisting of lX-l,3- and lX-l,5-linked L-arabinofuranose and linear polymers of f3-1,4-linked D-galactopyranose, may serve as a bridge between the rhamnogalacturonan and the hemicellulosic wall components (TALMADGE et a!., 1973). Hemicelluloses are major constituents of the primary and secondary wall re• gions. Functionally they link the pectic and cellulosic fractions (BAUER et a!., 318 D.F. BATEMAN and H.G. BASHAM:

1973; NORTHCOTE, 1972). , a chain of fJ-1,4-linked D-glucopyranose residues with terminal branches of 1X-1,6-linked xylopyranose, is a constituent of primary cell wall. This molecule is linked covalently to the pectic fraction and by hydrogen bonds to the surface of cellulose fibrils (BAUER et aI., 1973). The xylans, consisting of fJ-l,4-1inked xylopyranose chains, are widely distributed throughout higher . Xylans commonly have side branches of fJ-I,3-linked arabinofuranose and 1X-1,2-linked D-glucuronopyranose (or its 4-methyl ether). The numbers and kinds of branch residues are characteristic for different plant groups. Some xylans are acetylated at carbons 2 and 3 of the xylopyranose residues (ASPINALL, 1970). Other hemicellulosic polymers include , a heteropolysaccharide of glucopyranose and mannopyranose linked fJ-I,4, and galactoglucomannan, a glucomannan with 1,6-linked galactopyranose side branches. The glucose to man• nose ratios in these polymers vary depending upon the source, but ratios of 1 : 3 and 1: 2 are common in coniferous and deciduous , respectively (ASPINALL, 1970). The mannans and galactomannans are fJ-l,4-linked mannopyranose chains, with the latter having 1,6-linked galactopyranose side branches. Both of these polymers occur in higher plants, but they do not appear to be significant as structural cell ~all constituents. Cellulose is the most abundant substance found in the plant kingdom. It exists as elementary and microfibrils and constitutes the major structural component of cell walls. In primary wall, cellulose fibrils have a more or less random orienta• tion, but in secondary wall they occur in parallel lamellae, the different layers being oriented in different directions (MUHLETHALER, 1967). Chemically, cellulose is made up of long chains of fJ-l,4-linked D-glucopyranose. The individual chains are flat, ribbon-like molecules stabilized by hydrogen bonds between adjacent glucose residues. The chains are associated with each other in fibrils through interchain hydrogen bonds between hydroxyl groups at carbon 6 and the glycosidic oxygens of adjacent chains (NORTHCOTE, 1972). Cellulose fibrils contain both cry• stalline and amorphous regions. There is some disagreement about the diameter of these fibrils and the arrangement of the crystalline and amorphous regions within them (FREY-WYSSLING, 1969; PRESTON, 1971). One view holds that fibrils contain a crystalline core surrounded by amorphous cellulose, another holds that amorphous regions are randomly distributed within fibrils. The diameters of cellu• lose fibrils most often quoted range from 3.5-35 nm. The occurence and function of a structural protein in cell walls has been a controversial subject (LAMPORT, 1970). Most investigators now accept the cell wall as the location of a hydroxyproline-rich (KEEGSTRA et aI., 1973; NORTHCOTE, 1972). This structural protein may be deposited during maturation of the primary wall (SADAVA and CHRISPEELS, 1973; SADAVA et aI., 1973). The components of this protein are primarily galactose and arabinose. Arabinose is glycosidically linked to the hydroxyl of the hydroxyproline moiety, and galactose is believed to be covalently linked to serine (LAMPORT et aI., 1973). A recently proposed model of primary cell walls (Fig. 2) indicates that structural protein is covalently linked to the pectic substances through the fraction (KEEGSTRA et aI., 1973). Another major cell wall constituent, characteristic of woody species, is lignin (FERGUS et aI., 1969). Deposition of lignin occurs after cell wall maturation. The 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 319

Cellulose fibril & ~ \ III III ~; ~ III III III

Rhamnogalacturonan: -,..1---- .....1 L

Xyloglucan : III III III I

L. - Linked (Arabino) Galactan: ~

3,6 -Linked (Arabino) Galactan:

Fig. 2. Proposed model of primary plant cell wall. (KEEGSTRA et aI., 1973)

monomeric subunits of lignin are the oxidation products of sinapyl, coniferyl, and p-hydroxycinnamyl alcohols which condense by free radical mechanisms to form an amorphous polymer. The major intermonomer bonds between the substi• tuted monomers in lignin include arylglycerol-p-arylether bonds, phenylcoumaran structures, and biphenyl linkages, along with several other less prevalent linkages, all in a random 3-dimensional array (FREUDENBERG, 1968). Lignin may also be covalently linked to the other polymeric wall constituents (COWLING and BROWN, 1969). The polysaccharide constituents in cell walls are quite resistant to enzymatic decomposition after the walls become lignified, since they are complexed with and masked by the lignin component (DEHORITY et al., 1962). This highly cross• linked polymer reinforces the strength of cell walls in woody tissues. 320 D.F. BATEMAN and H.G. BASHAM:

2.2 Concepts of Membrane Composition and Structure

Membranes are a major structural element of all cells. They provide a semiperme• able barrier between the cell and its external environment and between intracellular compartments. Through specific transport and osmoregulatory functions, mem• branes are responsible for maintaining the basic biochemical integrity necessary for the vital functions in cells. The major constituents of membranes are lipids and . The lipid fraction consists of phospho glycerides such as diacylglycerolphosphorylethanolamine, dia• cylglycerophosphorylcholine, diacylglycerolphosphorylserine, and a variety or relat• ed compounds, consisting of complex alcohols esterified with phosphatidic acid. In addition, sphingolipids, glycolipids, sterols, and several other complex lipids may be found in the lipid fraction of certain membranes (THOMPSON, 1965; KORN, 1966; KORN, 1969). A variety of proteins and can be isolated from membrane preparations. Although little has been achieved in the assignment of functional or structural roles to specific membrane proteins, some of these proteins must be involved in membrane transport and regulatory functions (CARRAWAY et a!., 1971; OSEROFF et a!., 1973; SINGER, 1974). A number of structural models of biological membranes have been proposed to explain known membrane compositions and functions (THOMPSON, 1965; BRAN• TON, 1969; COOK, 1971; SINGER, 1971). The model most consistent with the available data relating to composition, structure and function is the" fluid mosaic" model (SINGER, 1971; SINGER and NICOLSON, 1972; BRETSCHER, 1973; SINGER, 1974). In this model (Fig. 3) the lipids are arranged in a bilayer with their polar head• groups oriented toward the aqueous exterior faces of the bilayer and the hydro• phobic hydrocarbon chains forming the interior of the membrane. The protein components of the membrane are in the globular form with a high proportion of the molecules in the configuration of an a-helix. Polar regions of the protein molecules are located on the exterior faces of the membrane, and nonpolar regions extend into or through the hydrophobic interior. The proteins are thought to migrate laterally within the fluid . Although membrane models which have been developed are based on data from bacterial and systems, the available evidence suggests that these models also apply to plant cell membranes. Biological membranes have similar functions in all living systems, serving as a matrix for a variety of enzyme systems and mediating numerous molecular, , and electron transport functions. Plant mem• branes contain lipid fractions similar to those found in animal membranes (BENSON, 1964; THOMPSON, 1965; OHAD et a!., 1967; KORN, 1969). Purified plasmalemma from higher plants has a sterol to phospholipid ratio comparable to that of animal membranes (1 : 2-1: 3) (HODGES et a!., 1972) and also contains an adenosine triphos• phatase (LAI and THOMPSON, 1971; HODGES et a!., 1972). Electron microscopy of freeze-fractured plant cell membranes (KORN, 1966; BRANTON, 1969) reveals a structure comparable to that observed in similar preparations of animal mem• branes (WEINSTEIN and BULLIVANT, 1967). Many of the proteins and lipids in animal and bacterial cell membranes are complexed with oligo- or polysaccharides. The glycoproteins and glycolipids are thought to exist at the membrane faces and may be involved with transporting structural components through the membrane, providing a variety of specific recog- 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 321

Fig. 3. The lipid-globular protein mosaic model of membrane structure. Phospho• lipids are arranged as a discontinuous bilayer with their polar heads in contact with water. The integral proteins, with the heavy lines representing folded poly• chains, are shown as globular proteins partially embedded in and par• tially protruding from the membrane. Polar amino are at the surface and nonpolar residues are in the embedded regions. (SINGER and NICOLSON, 1972) llltlOn sites, and/or protecting the outer surface of the membrane (SINGER and NICOLSON, 1972; BRETSCHER, 1973; OSEROFF et aI., 1973). Although little work has been done in this area with plant membranes, there are a few reports of visualization of membrane-associated molecules resolved by polysaccharide specific stains in electron micrographs (ALBERSHEIM and KILLIAS, 1963; ROLAND, 1969; ROLAND and VIAN, 1971). The developing technology for isolation and identification of specific plant cell membranes through the use of differential centrifugation (OHAD et aI., 1967; MORRE et aI., 1969; Ltd and THOMPSON, 1971; HODGES et aI., 1972) and stains for plasmalemma (ROLAND, 1969; ROLAND and VIAN, 1971; HODGES et aI., 1972), and the rapid advances in our understanding of the structure and function of similar membranes in and , will encourage research which should increase our knowledge of membrane systems in higher plants.

3. Enzymes Which Degrade Plant Structural Elements

3.1 Cell Wall Degradation

The plant cell wall is a complex, ordered structure, and it constitutes one barrier to microbial invasion of cells. Some appear to have evolved purely mechanical means of transversing this structure, but a vast majority of pathogens studied have evolved enzymes capable of dealing with the various wall constituents. In many instances cell invasion appears to be accomplished by a combination of physical and enzymatic means. When a confronts a plant cell wall it faces a complex barrier composed of polymers with different chemical linkages that require specific enzymes for their degradation. Pathogens have evolved the means to recognize the chemical structures in plant ceIl walls and elaborate the appropriate enzymes to dismantle the various cell wall constituents.

3.1.1 Degradation of the Pectic Fraction A key constituent of the pectic fraction in the plant cell wall is rhamnogalacturonan (TALMADGE et aI., 1973). Rhamnogalacturonans with up to 75% of the uronic acid carboxyls methyl esterified are known as pectinic acids, those with a higher 322 D.F. BATEMAN and H.G. BASHAM:

degree of esterification are referred to as , while the unesterified molecules are known as pectic acids (BATEMAN and MILLAR, 1966). Pectinmethylesterases hydrolyze the methyl esters of IX-I ,4-linked galacturonosyl residues at the carboxyl of the uronide moiety to yield a uronic acid carboxyl and methanol. The activity of this enzyme can be measured by determining either the amount of acid or of methanol produced. Continuous titration of the acid produced in reaction mixtures is the common method of measuring methyl• esterase activity (KERTESZ, 1951). This enzyme is found in higher plants and is produced by many but not all phytopathogenic . Pectinmethylesterases from higher plants are readily adsorbed by plant cell walls in an acid environment. The amount of enzyme free to react with its substrate appears to be regulated by the ionic strength of the environment and pH. The proposed site of binding of the enzyme to the cell wall is the free uronic carboxyl groups of the pectic fraction (JANSEN et aI., 1960). Pectinmethylesterases from higher plants lose activity upon dialysis, but this can be restored by adding a salt to the dialyzed solution. In contrast, pectinmethylesterases produced by certain fungi are not readily adsorbed by plant cell walls and they do not respond to dialysis or salt activation (MCCOLLOCH and KERTESZ, 1947; BATEMAN, 1963a). Mixtures of pectinmethylesterases of host and pathogen origin are probably com• mon in many diseases. The lX-l,4 bonds linking the galacturonosyl moieties in the pectic molecule are split by two enzymatic mechanisms. The enzymes that degrade this bond hydrolytically are known as or pectinmethylgalacturonases. Po• lygalacturonic acid trans-eliminases and pectinmethyl trans-eliminases split this bond by a Iyitic mechanism and are commonly referred to as trans-eliminases or lyases. Lyitic degradation of the glycosidic linkage results in an unsaturated bond between carbons 4 and 5 of a uronide moiety in the reaction product (ALBERS• HElM et a!., 1960b). The pectic enzymes have been grouped according to the following criteria: (1) the mechanism by which the lX-l,4 glycosidic bond is split (i.e. hydrolytic or lyitic), (2) enzyme specificity for a substrate (i.e. pectin or pectic acid), and (3) position in the pectic chain at which cleavage occurs (i.e. random or terminal point of attack) (BATEMAN and MILLAR, 1966; ROMBOUTS and PILNIK, 1972). Hydro• lases which exhibit a distinct specificity for pectin over pectic acid as a substrate are referred to as pectinmethylgalacturonases, whereas those that exhibit preference for pectic acid are called polygalacturonases. If the enzyme attacks its substrate in a terminal manner, i.e. releases only monomeric products, the prefix "exo" is used, e.g. a hydrolase that releases only galacturonic acid from pectic acid would be designated as an exopolygalacturonase, whereas if the attack is random and oligomers are released the enzyme would be called an endopolygalacturonase. It should be recognized that the endo enzymes may not degrade their substrates in a completely random fashion. Endo and exo enzymes are readily distinguished by separation and identification of reaction products and/or by determination of the percentage of lX-l,4 bonds split at a given percent viscosity loss of high molecular weight substrates. Endo enzymes will normally induce a 50% viscosity loss of a high molecular weight substrate with only 0.5-3% hydrolysis whereas 20% hydrolysis or more may be required for an exo enzyme to induce a similar viscosity loss (ROMBOUTS, 1972). 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 323

Endo- and exolyases are produced by a broad range of phytopathogens. Within this group there are enzymes which specifically degrade either methyl esterified or unesterified substrates (DEMAIN and PHAFF, 1957; BATEMAN and MILLAR, 1966). Unlike the exohydrolases which release monomeric reaction products, exolyases normally release unsaturated dimers (OKAMOTO et aI., 1964). A variety of procedures can be used to measure the activities of enzymes that split the a-l,4 bond in pectic polymers. The "cup plate" assay is a convenient procedure for screening large numbers of samples for pectolytic activity (DINGLE et aI., 1953): enzyme is placed in a well cut into an slab containing a pectic substrate, and after incubation a turbid zone develops, upon the addition of acid, if the substrate has been degraded. Loss in viscosity of high molecular weight pectins or pectic acids is a common, sensitive means of estimating activities of endo enzymes that attack these substrates (BELL et aI., 1955; BATEMAN, 1963a). Cleavage of glycosidic bonds is readily quantified by determining the increase in reducing groups in reaction mixtures (NELSON, 1944). The viscosity loss and reducing group assays can be used in combination to help distinguish between endo and exo enzymes, but these techniques do not permit one to determine whether the enzyme is a hydrolase or a lyase. The unsaturated products released from pectic acids and pectins by lyases absorb ultraviolet light maximally at 230 and 235 nm respectively (ALBERSHEIM et aI., 1960b; STARR and MORAN, 1962; NAGEL and ANDERSON, 1965), whereas reaction products released hydrolytically do not absorb light at these wave lengths. The hydrolytic and trans-eliminative reaction products can also be distinguished by their reaction with thiobarbituric acid (TBA) (NEUKOM, 1960; AYERS et aI., 1966). When reaction products of pectic hydrolases are treated with periodate, they are destroyed and do not react with TBA, but when reaction products of the trans-eliminases are treated with periodate, formylpyruvate is produced, which can react with TBA to form a characteristic red chromagen (WEISSBACH and HURWITZ, 1959). Thus, when one combines the chromatography of reaction products (SMITH, 1.,1960; PAGE, 1961) with a combina• tion of the above procedures, the enzymes which degrade the a-l,4 bonds in pectic acids or pectins can be readily distinguished. The activity of hydrolases that degrade pectic acids is often reduced by Ca + + (BATEMAN 1964a; ROMBOUTS and PILNIK, 1972). This presumably is the result of the formation of insoluble pectate. The trans-eliminases are generally activated by Ca + +, and their activity is lost in the presence of chelating agents (BATEMAN, 1966; ROMBOUTS and PILNIK, 1972). The pH optima for the pectic hydrolases are generally acidic, whereas the trans-eliminases, except for a few that degrade pectin, are most active under alkaline conditions (ALBERSHEIM et aI., 1960b; BATEMAN, 1966; HALL and WOOD, 1970; ROMBOUTS and PILNIK, 1972). The neutral sugar polymers most closely associated with the rhamnogalacturo• nans are the galactans and arabans (NORTHCOTE, 1972 ; TALMADGE et aI., 1973). Enzymes that hydrolytically degrade both constituents are produced by a variety of phytopathogenic microorganisms (FUCHS et aI., 1965; KAJI et aI., 1965; KNEE and FRIEND, 1968; COLE and BATEMAN, 1969; VAN ETTEN and BATEMAN, 1969; MULLEN, 1974). Degradation of /3-1,4 galactan is accomplished by exo- and endoga• lactanases, whereas degradation of the a-l,3- and ex-l,5-linked araban is effected by enzymes of the exo type. The activities of galactanases and arabanases are generally estimated by reducing group analysis and chromatographic procedures 324 D.F. BATEMAN and H.G. BASHAM:

(FUCHS et aI., 1965; KNEE and FRIEND, 1968; VAN ETTEN and BATEMAN, 1969; MULLEN, 1974).

3.1.2 Degradation of Hemicelluloses The hemicellulosic fraction of the plant cell wall is readily degraded to monomeric constituents by microbial enzymes; however, few phytopathogenic microorganisms have yet been shown to possess the enzymes needed to degrade the different glycosidic bonds found in the various hemicelluloses. This probably represents a lack of research in this area rather than deficiencies in the ability to produce these enzymes by pathogens. Studies with Sclerotium rolfsii (VAN ETTEN and BATE• MAN, 1969), solani (BATEMAN et aI., 1969), Sclerotinia sclerotiorum (HANCOCK, 1967), roseum 'Avenaceum' (MULLEN, 1974) and Diplodia viticola (STROBEL, 1963) indicate that the hemicellulolytic ability of some plant pathogens is substantial. The [3-1,4 linkages in the hemicellulosic xyloglucan of the primary cell wall are split by (endoglucanase) (BAUER et aI., 1973). The [3-1,4-linked xylans are hydrolyzed by endoxylanases to xylose oligomers including dimers and trimers (STROBEL, 1963; WALKER, 1967). A separate exoxylanase or [3-xylosidase is required to convert the xylobiose to xylose (KING and FULLER, 1968). The complete enzyma• tic decomposition of [3-1,4 mannan chains also requires two enzymes, an endoman• nanase and a [3-mannosidase (REESE and SHIBATA, 1965). Both the xylans and mannans exist as branched polymers, and various glycosidases are involved in removing terminal branch residues from such polymers. The 1,3-linked arabinose residues of xylans can be released by an a-L-arabinofuranosidase (KAJI and YOSHI• HARA, 1970), and various glactosidases are involved in the cleavage of terminal galactose residues. Three enzymes are needed for the complete hydrolysis of galacto• mannan: a [3-1,4 endomannanase, [3-mannosidase and an a-galactosidase (REESE and SHIBATA, 1965). S. rolfsii is known to produce these three enzymes (VAN ETTEN and BATEMAN, 1969). Enzymes that specifically split [3-1,3 and [3-1,6 glucan linkages are also produced by certain fungi (KAJI et aI., 1971; SHIBATA and FUKIM• BARA, 1972). Hemicellulases are generally assayed by determining increase in reducing groups in reaction mixtures and identification of reaction products by chromatography (STROBEL, 1963; REESE and SHIBATA, 1965; VAN ETTEN and BATEMAN, 1969). and mannan polymers are normally insoluble; this has created problems in the use of these polymers as substrates for enzyme assays. Some investigators have rendered these materials soluble by preparing carboxymethyl or hydroxyethyl deriv• atives of the polymers, which facilitates their use as enzyme substrates (HRAZDINA and NEUKOM, 1966; HASHIMOTO et aI., 1971). Most commercial substrates available for assay of these enzymes are contaminated with other polysaccharides, and care must be exercised in the utilization of these preparations for enzyme assays.

3.1.3 Degradation of Cellulose Cellulose is an insoluble, crystalline substance in its native form. Early work indicat• ed that the enzymes produced by certain microorganisms could hydrolyze soluble, 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 325

substituted forms of cellulose such as carboxylmethyl cellulose, yet these enzymes were unable to hydrolyze native cellulose. Other microorganisms, however, utilize native cellulose as a carbon source. Those microorganisms capable of utilizing native cellulose have been termed cellulolytic and were believed to produce an enzyme designated as C 1 . The C 1 enzyme was postulated to act on native cellulose by destroying its crystalline structure and exposing the glucan chains to 13-1,4 endoglucanases, termed Cx enzymes, which degrade the glucan chains to cellobiose (REESE, 1956). Conversion of cellulose to glucose also requires a cellobiase or f3-g1ucosidase. The multienzyme hypothesis of enzymatic decomposition of cellulose has been formulated around the C 1 , Cx and f3-g1ucosidase concept (REESE, 1963; GOULD, 1969). Recent studies clearly indicate the need for enzymes of the C 1 , Cx and cellobiase types for the complete degradation of native cellulose (KING and VESSAL, 1969; HALLIWELL and MOHAMMED, 1971). Cellulolytic microorganisms secrete an enzyme that fragments cellulose fibers into short insoluble particles termed "short fibers" (MARSH and SIMPSON, 1964; HALLIWELL, 1966). The cellulase complex produced by Trichoderma koningii can convert native cellulose to glucose (HALLIWELL, 1966). This enzyme complex has been resolved into four pure components: (a) cellobiase, (b) a CM-cellulase (Cx),

(c) a short fiber-forming component, designated C 2 , and (d) a C j component

(HALLIWELL and MOHAMMED, 1971). Only C j and C 2 are capable of attacking native cellulose when acting alone. When cotton fibers are used as substrate, C 1 exhibits only weak solubilizing ability; C 2 degrades cellulose weakly to yield short fibers, but acts synergistically with the Cx component to promote short fiber production. The four components act together in converting native cellulose to glucose. The C 1 and C 2 enzymes of T. koningii can be equated with the hypothet• ical C 1 postulated by REESE (1956). A number of procedures have been used to evaluate the cellulolytic ability of microorganisms and cellulase preparations. The loss in birefringence of plant cell walls in tissues invaded by pathogens is an indication of enzymatic destruction of crystalline cellulose by pathogenic microorganisms (RAWLINGS and TAKAHASHI, 1952; BATEMAN, I 964b). Cellulase preparations induce a loosening or swelling of cotton fibers. Gravimetric determination of the ability of cotton fibers to absorb more NaOH solution than controls after enzyme treatment is used to quantify this effect of cellulase action and is expressed as the "S-factor" (MARSH, 1953; KELMAN and COWLING, 1965). The mechanism of cellulose loosening remains un• known. The enzymatic fragmentation or short fiber formation can be determined turbidometrically (HALLIWELL, 1966). The quantity of short fibers released from native cellulose may not give an accurate measure of the amount of the C 2 enzyme component present, since other components of the cellulase complex are known to act synergistically with the C 2 component in short fiber formation (HALLIWELL and MOHAMMED, 1971). A common method for measuring cellulase activity on native cellulose is the determination of weight loss of substrate after exposure to cellulolytic microorganisms or cellulase preparations (HALLIWELL, 1963). Chro• matography of reaction products, release of reducing groups, and release of glucose as determined with glucose oxidase are used to measure degradation of soluble cellulose derivatives (BATEMAN, 1964b; KELMAN and COWLING, 1965; HALLIWELL and MOHAMMED, 1971). 326 D.F. BATEMAN and H.G. BASHAM:

Modified are generally more susceptible to enzymatic degradation than are native celluloses, and they are widely used for cellulase assays (HALLIWELL, 1963). Reduction in turbidity of acid-swollen cellulose sols by cellulase has been used to measure cellulolytic enzyme activity (KELMAN and COWLING, 1965). The cellulolytic ability of fungi may be estimated by incorporating a partially crystalline form of regenerated cellulose into a of defined medium on which the organism is grown. Cellulolytic activity is measured on a continuous, cumulative basis by observing the depth of clearing of the opaque medium (RAUTELA and COWLING, 1966). The most common and perhaps the least meaningful assay for evaluating the cellulolytic capacity of an organism involves the use of high molecular weight carboxymethyl substituted cellulose to determine enzyme activity by follow• ing decrease in substrate viscosity or increase in reducing groups (HALLIWELL, 1963). Conversion of cellobiose to glucose is readily measured by determining the production of glucose with glucose oxidase or by chromatographic procedures (HALLIWELL and MOHAMMED, 1971). Each of the assays mentioned can serve a useful purpose in studying cellulolysis, but should be selected and used in appro• priate combinations to answer specific questions. Our current knowledge of the cellulase complex and the role each component plays in degradation of cellulose should serve as a guide for selecting appropriate assay procedures to assess ade• quately the cellulolytic ability of phytopathogenic organisms.

3.1.4 Sequential Events in Cell Wall Degradation Plant cell walls exhibit differential susceptibility to enzymatic decomposition, de• pending upon the combination of enzymes or sequence of treatment with specific enzymes. Abilities of enzymes to degrade model substrates are not necessarily indicative of their ability actually to degrade plant cell walls (KAJI, 1958; GARIBALDI and BATEMAN, 1971). The presence ofa "wall-modifying enzyme" had been consid• ered to be a prerequisite for wall degradation by certain po1ysaccharidases (KARR and ALBERSHEIM, 1970). Such a factor was first demonstrated in Pectinol R-lO, a commercial pectinase preparation, but the wall-modifying enzyme was not resol• ved from an endopolygalacturonase. When the purified wall-modifying enzyme was incubated with cell walls, water-soluble - 70% ethanol-insoluble polymers, rich in uronic acid, were released. It is now reasonably clear that the pectic enzymes serve as wall-modifying enzymes and, by their action, the nonpectic polymers, including the structural protein in cell walls, are rendered more susceptible to enzymatic decomposition (KARR and ALBERSHEIM, 1970; ENGLISH et aI., 1972; TALMADGE et a!., 1973). Based on the concept of primary cell wall structure (KEEG• STRA et aI., 1973), it seems quite reasonable that the endopectic enzymes would have a key function in cell wall decomposition. The pectic enzymes, particularly endopolygalacturonases and lyases, are able to attack unaltered plant cell walls, loosen their structure and render other wall polymers susceptible to enzymatic hydrolysis. 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 327

3.2 Membrane Degradation

3.2.1 Degradation of Phospholipids Plant pathogens are known to produce enzymes that attack the major membrane constituents. A number of specific phospholipases hydrolyze the acylester and phosphate ester bonds of phospholipids (McMuRRAY and MAGEE, 1972; GATT and BARENHOLZ, 1973). Phospholipase A hydrolyzes one of the acylester bonds of diacylglycerophosphoryl compounds, yielding a and a lysophospholi• pid. Where careful investigations have been made, the A-type phospholipases have been shown to exhibit specificity for the acylester at the 1 or 2 position of the glycerol moiety; these enzymes are designated phospholipase Ai or A z respectively, depending upon their specificity. Lysophospholipase hydrolyzes the acylester of monoacylglycerophosphoryl compounds, yielding a fatty acid and the glycerophos• phoryl moiety. These enzymes are specific for the ester at the 1 or 2 position and are designated lysophospholipase Li or L2 respectively. If positional specificity has not been determined, the enzymes are designated phospholipase A or lysophos• pholipase. Phospholipase B clea ves both of the fatty acylesters from a diacylglycero• phosphoryl compound, yielding two fatty acids and the glycerophosphoryl moiety. There is some controversy as to whether phospholipase B is a single enzyme or a closely associated phospholipase A-lysophospholipase complex (SAITO and SATO.1968; McMURRAY and MAGEE, 1972). Phospholipase C hydrolyzes diacylgly• cerophosphoryl compounds to diacylglycerol and the phosphoryl moiety. Phospho• lipase D hydrolyzes diacylphosphoryl compounds to phosphatidic acid and the complex alcohol moiety. This type of enzyme is found only in higher plants. The specific phosphoryl or fatty acid substituents associated with a given phos• pholipid mayor may not affect the rate of enzyme activity, so substrate require• ments for an uncharacterized enzyme must be determined experimentally (DOERY et a1.. 1965; DE HAAS et aI., 1968; McMURRAY and MAGEE, 1972). Detergents, such as Triton XlOO or deoxycholate, are often required for dispersal of substrates for phospholipase assay; a particular detergent may be required as a specific enzyme activator in some cases or may, with other enzymes, act as an inhibitor. Often, Ca + + or other divalent cations are also required for enzyme activity (DE HAAS et aI., 1968; SAITO and SA TO, 1968; McM URRA Y and MAGEE, 1972). Natural lecithins from egg yolk or soybean are most commonly used as substrates for the assay of phospholipases, but a number of pure substrates of known composition can be prepared or purchased to determine enzyme specificity (DE HAAS et aI., 1968; LOWENSTEIN, 1969; GALLIARD, 1971). The site of bond cleavage is determined by separation and identification of the reaction products. Reaction products are separated by their differential solubilities in solvents and/or chromatography; they are identified by thin-layer chromatography, gas-liquid chro• matography, or chemical assays for specific reaction products (DOERY et aI., 1965; DE HAAS et aI., 1968; LUMSDEN and BATEMAN, 1968; LOWENSTEIN, 1969; TSENG and BATEMAN, 1969; GALLIARD, 1971). For routine assay of phospholipases a variety of procedures may be used. The simplest is the cup plate assay, in which an enzyme preparation is placed in a well cut into an agar slab containing lecithin and any required cofactors. Production of a clear or turbid zone around the well containing the enzyme is indicative of phospholipase activity (DOERY et aI., 328 D.F. BATEMAN and H.G. BASHAM:

1965; HABERMANN and HARDT, 1972). All phosphoIipases cleave ester bonds and produce acids as reaction products, thus enzyme activities may be followed potentio• metrically (DE HAAS et aI., 1968). Phospholipase B, phospholipase C and lysophos• phoIipase release acid-soluble phosphorus which can be measured by colorimetric procedures (LUMSDEN and BATEMAN, 1968; LOWENSTEIN, 1969). Activities of enzy• mes that hydrolyse fatty acylester bonds can be determined by following the de• crease of these bonds in reaction mixtures by the hydroxyamate procedure (SNYDER and STEPHENS, 1959; BULOS and SACKTOR, 1971).

3.2.2 Degradation of Proteins

Proteolytic enzymes hydrolyze peptide bonds in proteins. Many of these enzymes also catalyze the formation of peptide bonds and aminotransfer reactions, and hydrolyze ester linkages involving amino acids (SMITH, E.L., 1960). Primary classifi• cation of proteolytic enzymes is based on the catalytic mechanism of the . The serine proteinases contain serine at their active site and are inactivated by diisopropylphosphofluoridate. The thiol proteinases contain cysteine at their active site and are inactivated by thiol reagents such as mercuric chloride or iodoacetic acid; thiol proteinases are seldom produced by microorganisms. Metal proteinases require bound metal such as Mg+ +, Mn + +, Zn + +, or Fe + + for activity, and are inactivated by chelating agents. Acid proteinases contain carboxyl groups at their active site and are characterized by a low pH optimum and activity in the presence of inhibitors that inhibit the other classes of proteinases. Specificity for peptide bonds adjacent to particular amino acids, random or terminal cleavage, and ability to cleave small are specific characteristics of given enzymes; bacterial and fungal proteinases are normally nonspecific with respect to these properties (SMITH, E. L., 1960; MATSUBARA and FEDER, 1970). Most assays for proteinases are based on the increased solubility of reaction products. Large peptides or intact protein molecules in reaction mixtures are precip• itated with trichloroacetic acid and removed by filtration or centrifugation; proteinase activity is determined by measuring, spectrophotometrically or with Folin reagent, the concentration of aromatic amino acids in the trichloroacetic acid soluble fraction (KUNITZ, 1947 ; VAN ETTEN, 1966; PERLMAN and LORAND, 1970). Proteinase activity may also be determined with a variety of insoluble protein• dye complexes by following the solubilization of dye molecules as the protein is hydrolyzed (NELSON et aI., 1961). Assays based on substrate solubilization do not reflect the actual number of peptide bonds broken, and make kinetic interpreta• tion of data difficult; assays utilizing the esterase activity of serine proteinases reveal the number of enzymatic events and facilitate kinetic interpretations. The substrates for the esterase assays are L-et amino acids blocked at the N terminal with an acyl, benzyl, or other moiety and ester linked to a methyl, ethyl, or p-nitrophenyl group. Cleavage of the ester linkage can be followed potentiometri• cally or spectrophotometrically (WALSH and WILCOX, 1970). A number of other assays such as milk or blood clotting and ninhydrin reactions with amino groups may be used for estimating enzymatic hydrolysis of proteins and peptides (PERLMAN and LORAND, 1970). 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 329

3.2.3 Effects of Enzymes on Membranes Membrane degradation by phospholipases and proteinases depends on the condi• tion of the membrane and on the specific type of enzyme involved. Phospholipase A produces lysophospholipids, which have a detergent action and destroy intact membranes (CONDREA and DE VRIES, 1965). This enzyme has been shown to damage membranes of Phaseolus mitochondria (EARNSHAW and TRUELOVE, 1970). Phospho• lipase A is not known to be produced by plant pathogens, but little work has been done on the characterization of phospholipases from these organisms. Phos• pholipase B is produced by Thielaviopsis basicola (LUMSDEN and BATEMAN, 1968), S. rolfsii (TSENG and BATEMAN, 1969), and Botrytis cinerea (TSENG and CHANG, 1970). The effect of this class of enzymes on intact membranes has not been investigated. Phospholipase C appears to have little direct effect on intact mem• branes, but there is a report that this enzyme from carotovora lyses intact cucumber (TSENG and MOUNT, 1974). Phospholipase C does not lyse intact red blood cells or Avena protoplasts (RUESINK, 1971; ZWAAL et aI., 1971). Red blood cell ghosts are extensively degraded by phospholipase C, thus this enzyme does degrade membrane fragments (ZWAAL et aI., 1971). Phospholipase C and phospholipase D apparently cause no damage to tobacco tissue (HUANG and GOODMAN, 1970), and proteinases and/or phospholipase C from E. carotovora cause little or no damage to intact cucumber or potato tissue (MOUNT et aI., 1970; TSENG and MOUNT, 1974). Proteinases cleave certain exposed proteins on the surface of animal and micro• bial membranes, but do little to alter basic membrane structure (ITO and SATO, 1969; BRETSCHER, 1973). Enzymatic degradation of membrane protein is much more extensive with membrane fragments than with intact membranes (ITO and SATO, 1969). Proteinases apparently do not affect isolated Avena protoplasts (RUE• SINK, 1971) but have been reported to lyse cucumber protoplasts (TSENG and MOUNT, 1974). More information is needed before any conclusions can be drawn about the importance of phospholipases and proteinases in the destruction of intact plant membranes. It is apparent that degradation of intact membrane systems cannot be presupposed merely because a pathogen has the capacity to produce proteinases and/or phospolipases.

4. Relationships of Enzymatic Alterations of Structural Constituents to Pathogenesis

The ability of plant pathogens to produce enzymes in culture that degrade model substrates is merely a first step in relating a given enzyme to pathological processes. Too often studies of degradative enzyme systems have focused only on culture work and test-tube . This approach is of course often helpful in examin• ing the types of enzymes a pathogen is capable of producing and in guiding studies relating to infection and disease, but beyond this, the utility of in vitro studies in helping to understand the significance of enzymes of pathogen origin 330 D.F. BATEMAN and H.G. BASHAM:

in pathogenesis is limited. This is because many of the extracellular enzymes in• volved in degradation of structural components of the host are inductive and/or subject to catabolic repression. Also, substrates for the specific enzymes that de• grade wall polymers or membrane constituents do not necessarily exist in a suscepti• ble state in living tissue, but rather in association with other molecules in the structural elements that may render them resistant to enzymatic alteration. The mere demonstration of a degradative enzyme of pathogen origin in infected or diseased tissue does not in itself indicate involvement of that enzyme in pathogenesis or destruction of invaded host tissues (HANCOCK, 1967). The association of a degradative enzyme with diseased tissue should be coupled with a demonstration of depletion of the enzyme's substrate or a modification of the functional capacity of the structural element containing the specific substrate prior to invoking a causal function of the enzyme in tissue breakdown or alteration. Enzymes that degrade the plant cell wall constituents represent one battery of "attacking mechanisms" in the arsenal of many phytopathogenic organisms. Pectic enzymes are produced by of some pathogens upon (BA• RASH, 1968; VERHOEFF and WARREN, 1972). One characteristic feature of disease in plants is the alteration or degradation of structural components of host cell walls during pathogenesis. This is particularly true for diseases caused by facultative parasites (BATEMAN and MILLAR, 1966; WOOD, 1967). Obligate parasites also have the ability to produce cell wall-degrading enzymes (VAN SUMERE et aI., 1957), but the significance of these enzymes in diseases caused by obligate parasites is not yet clear. It would appear, however, that cell wall-degrading enzymes produ• ced by obligate parasites could be important in aiding ingress and colonization of host tissues. Certain mycorrhizal fungi which establish compatible relationships with their hosts are known to produce pectic enzymes (PEROMBELON and HADLEY, 1965; WILLIAMSON and HADLEY, 1970). It is apparent that production of enzymes by mycorrhizal fungi and obligate parasites must be under adequate control lest the invaded tissues become macerated and the host cells killed.

4.1 Alterations of Cell Wall Constituents 4.1.1 Factors Influencing Production and Activity of Cell Wall-degrading Enzymes in Infected Tissues The environment in which a pathogen grows dictates to a large degree the types and quantities of cell wall-degrading enzymes produced. Catabolic repression by of the synthesis of cell wall-degrading enzymes is a common phenomenon in many phytopathogenic organisms (KEEN and HORTON, 1966; PATIL and DIMOND, 1968; MORAN and STARR, 1969; BRATHWAITE and DICKEY, 1970; BIEHN and DI• MOND, 1971 a; BIEHN and DIMOND, 1971 b; SPALDING et aI., 1973; COOPER and WOOD, 1975), and there is evidence to indicate that this factor may be of importance in pathogenesis (HORSFALL and DIMOND, 1957; HORTON and KEEN, 1966; ZUCKER and HANKIN, 1971). A small number of pathogens are known to produce certain cell wall-degrading enzymes in a constitutive manner, for example, the polygalactu• ronases produced by albo-atrum (MUSSELL and STROUSE, 1972) and Aphanomyces euteiches (AYERS et aI., 1969), the polygalcturonase and systems of Helm in thosporum maydis T (BATEMAN et aI., 1973), and the cellulase system of solanacearum (KELMAN and COWLING, 1965). 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 331

Plant cell wall constituents may serve as effective inducers of cell wall-degrading enzymes in plant pathogens. The limited studies that have been made with patho• gens in culture, where isolated cell walls have been used as a carbon source, indicate that cell wall-degrading enzymes are produced in a temporal sequence in which the pectic enzymes appear first and those that degrade xylan and cellulose appear last. This pattern of production of cell wall-degrading enzymes has been reported for Colletotrichum lindemuthianum (ENGLISH et al., 1971), Fusarium oxyspo• rium f. sp. lycopersici (JONES et al., 1972) and F. roseum 'Avenaceum' (MULLEN, 1974). A low molecular weight water-soluble fraction, rich in uronic acid, from tomato cell walls, and a water-soluble, ethanol-insoluble component from , are effective inducers ofpolygalacturonases in F. oxysporum f. sp. lycopersici and Ceratocystis ulmi respectively (BIEHN and DIMOND, 1971 b; JONES et al., 1972). Since the pectic enzymes appear to be the key factors in wall degradation, the occurrence of soluble inducers of these enzymes in plant tissues deserves further study. The pH of the environment in which pathogens grow has a marked influence upon the stability of the enzymes produced and appears to influence production of pectic Iyases (BATEMAN, 1966; HANCOCK, 1966b; BATEMAN, 1967; PERLEY and PAGE, 1971). Many pathogens that produce pectic Iyases do so best in an alkaline environment. Infection of tissues by a number of Fusaria, for example, results in an alkaline shift in the pH of the infected host tissues, and pectic lyases are known to be the predominant pectic enzymes associated with these diseases (BATE• MAN, 1966; HANCOCK, 1968a; MULLEN and BATEMAN, 1971). The dominant pectic enzymes found in tissues infected by S. rolfsii and S. sclerotiorum, which create acid environments by production of organic acids, are the po1ygalacturonases, which are more active under acid conditions (BATEMAN and BEER, 1965; HANCOCK, 1966a). In many instances it appears that the pectic enzymes associated with a given disease are those best adapted to the environment created by the host• parasite interaction. Host environment and/or response limits the growth of saprophytes in living plant tissue, but modification of these limiting factors by a pathogen can often result in the establishment of saprophytes in diseased tissue. It is quite likely that in some instances tissue degradation is accomplished in part by enzymes of saprophytes once disease has been initiated by a primary causal agent. Pectolytic and cellulolytic enzymes are produced by a saprophytic sp. often associated with the noncellulolytic, nonpectolytic causal agent of a carnation wilt, Pseudomonas caryophylli. The enzymes produced by this saprophyte contribute to the rapid destruction of host tissue associated with this disease (BRATHWAITE and DICKEY, 1970). Pectic enzymes produced by a pathogen in culture may be quite different from those enzymes associated with pathogenesis, and studies with the former can be misleading in attempts to understand the involvement of pectic enzymes in a given disease. Much of the work on the involvement of pectic enzymes in of tomato has focused on the polygalacturonases produced by F. oxysporum f. sp. lycopersici in culture; pectic Iyases are produced by this pathogen when it is grown on tomato tissue (BATEMAN, 1967). The occurrence of a polygalacturonate lyase in F. oxysporum f. sp. lycopersici-infected tomato tissue has been recently demonstrated (FERRARIS et al., 1974). Perhaps more attention to the role of the 332 D.P. BATEMAN and H.G. BASHAM:

pectic lyases in Fusarium wilt of tomato would help to resolve the significan~e of pectic enzymes in this disease (METLITSKII and OZERETSKOVSKAYA, 1968). The pectic enzymes produced by R. solani and Penicillium expansum in culture and those that predominate in tissues infected with these pathogens are known to differ considerably (BATEMAN, 1963a; SWINBURNE and CORDEN, 1969). The types of pectic enzymes present in culture filtrates of R. solani differ with culture age and pH; although young cultures with an acid pH contain primarily polygalacturo• nase, older cultures can shift to an alkaline pH and may contain only polygalacturo• nate lyase. The pectic enzyme predominating in the critical phases of pathogenesis by R. solani in bean hypocotyls is , and the pH of lesion extracts remains acidic (BATEMAN, 1967). Certain phenolic compounds, and particularly their oxidation products may function as nonspecific enzyme inactivators or inhibitors, since their free radical intermediates react readily with biochemical constituents capable of electron inter• action, such as -NH2 and -OH groups (PATIL and DIMOND, 1967). Such inhibitors could affect all enzymes. Certain plants also contain tannins and related materials that can function as nonspecific enzyme inhibitors (BATEMAN and MILLAR, 1966). The occurrence of enzyme inhibitors in plants has been correlated with resistance to pathogenic organisms (PRIDHAM, 1959; MAHADEVAN et aI., 1965; LOVREKOVICH et aI., 1967). The importance of naturally occurring enzyme inhibitors, particularly phenolics, in relation to celJ wall degradion in infected tissue needs to be resolved. It is clear that certain cell wall-degrading enzymes can be inactivated in vitro by oxidized phenolics (DEVERALL and WOOD, 1961; BATEMAN, 1967; PATIL and DIMOND, 1967) and that oxidation of phenols occurs in tissues undergoing necrosis (FARKAS and KIRALY, 1962). Apple infected with Sclerotiniafructigena contains oxidized phenols that inactivate polygalacturonase in vitro. These compounds accu• mulate in lesions (COLE, 1958), but active fungal polygalacturonase can be obtained from the lesions as well as from tissue outside the zone of accumulated phenolics (BYRDE et aI., 1973). Also, cell walls in suberized or lignified tissues are difficult to degrade enzymatically (DEHORITY et aI., 1962; Fox et aI., 1971); it is known that lesion development and maceration of tissue in the Rhizoctonia disease of bean occurs prior to development of necrosis or significant increase in phenoloxi• dase or peroxidase levels (MAXWELL and BATEMAN, 1967). It is not clear that cell wall-degrading enzymes are readily inactivated in tissues undergoing pathogenesis. The failure to demonstrate active cell wall-degrading enzymes in water extracts of a diseased tissue has little relevance, since precautions need to be taken in some cases to prevent inactivation of these enzymes by oxidized phenolics during the extraction process. Active preparations of pectic lyase, 13-1,4 galactanase, xylanase, and arabanase are readily obtained from potato tubers infec• ted with F. roseum 'Avenaceum' if precautions are taken to inactivate phenol oxidase in potato tissue prior to enzyme extraction, or if extraction is carried out in the presence of an antioxidant such as 2-mercaptoethanol; the activities of cell wall-degrading enzymes in water extracts of diseased tissues are slight or nil (MULLEN and BATEMAN, 1971; MULLEN, 1974). Active polygalacturonases are readily obtained in water extracts of some necrotic tissues even though it can be demonstrated in vitro that the phenol-oxidizing systems in these tissues are capable of generating products that inactivate these enzymes (DEVERALL and WOOD, 1961; BATEMAN, 1967; BALASUBRAMANI et aI., 1971). A number of factors must 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 333

be considered when examining the influence of phenolic inhibitors on cell wall degradation during pathogenesis: (1) the temporal and spatial relationships among enzyme production, lesion development, and necrosis following infection, (2) possi• ble "compartmentalization" of the oxidative activities and cell wall-degrading enzymes, and (3) the effect of oxidized phenolics plus and lignin-like compo• nents which may complex with wall polymers and render them resistant to enzyma• tic degradation. Plant cell walls also contain proteins, with properties similar to phytoagglutins, capable of binding with and serving as potent inhibitors of certain polygalacturona• ses. A proteinaceous polygalacturonase inhibitor has been demonstrated in water extracts of sweet potato tissue (URITANI and STAHMANN, 1961). Recent studies have focused attention on the possible significance of this group of inhibitors on the enzymatic decomposition of plant cell walls (ALBERSHEIM and ANDERSON, 1971; FISHER et a!., 1973). The protein-inhibitors obtained from bean and tomato cell walls do not inhibit cellulase, xylanase, galactosidase, a-arabinofuranosidase or pectic lyases. Since polygalacturonase is a key enzyme in cell wall degradation by certain pathogens, the significance of these protein-inhibitors in cell walls could be substantial, but their role in vivo has not been determined. It has been demonstra• ted in vitro that the polygalacturonase inhibitor prepared from tomato cell walls protects these walls from degradation by the complex cell wall-degrading enzyme mixture produced in culture by F. oxysporum f. sp. lycopersici (JONES et aI., 1972). Attempts to demonstrate differential inhibition of the polygalacturonases produced by the Ct., [3, and y races of C. lindemuthianum by protein-inhibitors from their respective suscept and nonsuscept bean varieties failed, i.e. the polygalacturonases and the protein-inhibitors from the respective pathogen races and bean varieties appeared to be similar (ANDERSON and ALBERSHEIM, 1972). Aside from the possible effects of catabolic repressors and inducers on enzyme synthesis by pathogens, the pH of the host environment on enzyme stability, and the influence of nonspecific and specific enzyme inhibitors, it is known that the association of cell wall components with each other and the changes that occur during wall maturation can also greatly influence the susceptibility of specific wall components to a given enzyme. Regulation of production of cell wall-degrading enzymes and their effectiveness in infected plant tissues is thus directed by a balance of forces at play in the environment created by the host-pathogen interac• tion. In compatible interactions the production of enzymes by pathogens that degrade plant cell walls appears to be the rule (BATEMAN and MILLAR, 1966; ALBERSHEIM et a!., 1969).

4.1.2 Tissue Maceration

In many plant diseases, particularly the soft rots, tissue maceration is a characteristic symptom. This process involves the separation of cells from each other within a tissue system. The enzymatic basis of tissue maceration was clearly established in early studies (DE BARY, 1886; JONES, 1909; BROWN, 1915), and the phenomenon was attributed to the digestion of the "inter-cellular cement" or middle lamella of the cell walls. Pectic enzymes have long been associated with tissue maceration, but imprecise information on the chemical make-up of the middle lamella (JOSLYN, 334 D.F. BATEMAN and H.G. BASHAM:

1962), and the lack of highly purified enzymes for experimental work prevented resolution of the precise mechanism of tissue maceration prior to the early 1960s (DEMAIN and PHAFF, 1957; BATEMAN, 1963b; SPALDING, 1963; MCCLENDON, 1964; ZAITLIN and COLTRIN, 1964). It is now clear that the endopectic enzymes that split the 0(-1,4 bonds between the galacturonosyl moieties of the pectic fraction, apart from enzymes that degrade other cell wall polymers, can account for the process of tissue maceration (ZAITLIN and COLTRIN, 1964; DEAN and WOOD, 1967; BATEMAN, 1968; BUSH and CODNER, 1968; BYRDE and FIELDING, 1968; HALL and WOOD, 1970; MOUNT et aI., 1970; GARIBALDI and BATEMAN, 1971; BATEMAN, 1972). There is doubt whether enzymes that degrade cellulose, protein, xylan or man• nan-based polymers aid the maceration process (MCCLENDON, 1964; BATEMAN and MILLAR, 1966; MOUNT et aI., 1970). Reports that O(-L-arabinofuranosidase (arabanase) macerates plant tissues have been refuted (BUSH and CODNER, 1968; BYRDE and FIELDING, 1968; COLE and BATEMAN, 1969; COLE and WOOD, 1970). There is a recent unconfirmed report that endo-fJ-1,4 glactanases from Fusarium caeruleum and erythroseptica are capable of causing tissue maceration and host cell death which resembles that induced by endopectic enzymes (COLE and STURDY, 1973). Maceration of plant tissue by Dothidea ribesia has been attrib• uted to a factor, designated" phytolysin ", which is supposedly distinct from pectic enzymes (NAEF-RoTH et aI., 1961; KERN and NAEF-RoTH, 1971). However, it is not clear that pectic enzymes have been completely resolved from phytolysin. At the present time, the pectic enzymes that split the 0(-1,4 bonds between the galacturonosyl moieties in the pectic fraction of the cell wall remain the only enzymes confirmed to cause plant tissue maceration; enzymes that attack these bonds in an endo or random fashion are by far the most efficient in causing maceration. Not all pectic enzymes that degrade isolated polygalacturonides in a random manner have the ability to macerate plant tissue readily. A polygalacturonase has been reported from Clostridium felsineum which failed to macerate tissue (KAJI, 1958). Also, an endopolygalacturonase which failed to macerate tissue was resolved from the macerating enzyme in a culture filtrate of S. fructigena (BYRDE and FIEL• DING, 1962); the macerating enzyme was later shown to be a pectin methyl trans• eliminase (BYRDE and FIELDING, 1968). The polygalacturonate trans-eliminase com• plex produced by Erwinia chrysanthemi can be separated into four isozymes (GARI• BALDI and BATEMAN, 1971). All the isozymes except one with a pI of 4.6 macerate plant tissue; the isozyme with a pI of 4.6 readily attacks pectates isolated from a number of plants and releases soluble uronides from isolated bean hypocotyl cell walls but fails to macerate readily or release soluble uronides from intact tissue. These examples of endopectic enzymes which fail to macerate tissue readily are exceptions; their failure to effect tissue maceration is not understood. A number of procedures have been devised to assess the ability of enzyme preparations to effect tissue maceration. Most procedures employ disks or slices of a homogeneous tissue as substrate. The medullary tissue of potato tubers or the mesocarp tissue of cucumber fruit are the most commonly used substrates for tissue maceration assays, but almost any herbaceous tissue, particularly those containing cells, can be used. One of the earliest procedures for measur• ing tissue maceration involves the estimation of loss in coherence of tissue pieces 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 335 after exposure to enzyme (BROWN, 1915; BATEMAN, 1963 b), tissue coherence being evaluated by determining the ease with which this tissue can be teased apart with the fingers or with forceps. Although this technique is SUbjective, it can be, with practice, a useful rapid method for assessing the macerating ability of enzyme preparations. More objective procedures include assessment of the time required for uniform strips of tissue to break when subjected to a constant stress in the presence of enzyme (DAVISON and WILLAMAN, 1927) or measurement of the force necessary to "burst" or cause a blunt object to penetrate a tissue slice after exposure to enzyme (MCCLENDON and SOMERS, 1960; SHERWOOD, 1966; TA• GAWA and KAJI, 1966). These procedures permit determination of the end-point of maceration, i.e. the time when tissue coherence is completely lost, or the relative amount of maceration which has occurred to a given tissue piece after a given time of enzyme exposure. Procedures based on the release of cells from tissue have been devised which permit continuous measurement of maceration of a given tissue sample over the full course of the process (BATEMAN, 1968; SATO, 1968). Maceration of leaf tissue is commonly measured by suspending leaf strips in enzyme preparations and agitating the mixture; cells freed from the tissue are removed periodically and their numbers assessed by measuring the amount of present in the cells released (ZAITLIN and COLTRIN, 1964; BRATHWAITE and DICKEY, 1971 ). The rate of cell release from tissue slices or disks by macerating enzymes is easily followed by measuring the turbidity of the suspending medium which can be related to cell number per unit volume of medium (BATEMAN, 1968; SA TO, 1968). Procedures in which maceration is determined by cell release offer a number of advantages over other procedures used to estimate tissue maceration. These advantages include: (1) measurements are more objective, (2) actual cell separation, rather than loss in tissue coherence or tissue strength, is measured, (3) measurements are made using the same tissue samples over the entire course of the experiment, and (4) variation between replicates is greatly reduced, since a larger number of tissue pieces may be employed in a given replicate (BATEMAN, 1968). Another method of measuring tissue maceration has been employed to deter• mine the activity of enzymes that cleave the polygalacturonides in plant tissues in a random manner (MUSSELL and MORRE, 1969; CRONSHAW and WOOD, 1973). This procedure involves washing, blotting, and weighing tissue samples after differ• ent periods of exposure to the enzyme preparation. Enzyme activity or maceration is then estimated by the loss in fresh weight of the tissue samples due to cell sloughing. This procedure has one inherent weakness: the assay is based on weight loss of the tissue sample; many factors other than maceration can effect weight loss through changes in cell permeability and loss of water and solutes. Thus, with this assay, one cannot be entirely certain whether the sole factor being meas• ured is maceration or pectic enzyme activity, or whether some of the effect is due to other toxic factors if crude enzyme preparations are being assayed. It has been demonstrated that tissue maceration can be related to the rate of removal of the pectic substances from a tissue sample (TAGA WA and KAJI, 1966), and that the rate of maceration is directly related to the log of the macerating enzyme concentration (BATEMAN, 1968). Chelating agents which remove Ca + + from calcium pectates increase maceration rates by polygalacturonases (ZAITLIN 336 D.F. BATEMAN and H.G. BASHAM: and COLTRIN, 1964; BATEMAN and BEER, 1965), but such agents would be expected to have the reverse effect on tissue maceration by pectic lyases (HEATH and WOOD, 1971 ).

4.1.3 Toxic Effects of Pectic Enzymes Pectic enzymes that cause maceration of plant tissues also cause injury and death of un plasmolyzed plant cells (BROWN, 1915; TRIBE, 1955; SPALDING, 1969; MOUNT et al., 1970; WOOD, 1972). In some infections, degradation of host cell walls and cellular injury are associated with permeability increases detectable in early stages of pathogenesis (FRIEDMAN and JAFFE, 1960; Fox et al., 1972; BYRDE et al., 1973). Polygalacturonate lyase from E. chrysanthemi (BASHAM, 1974) purified to homoge• neity or polygalacturonase from V. albo-atrum (MUSSELL, 1973) causes simultaneous maceration and death of cells in disks of potato tissue. Injury of tissue treated with pectic enzymes is characterized by a rapid, irreversible increase in permeability of affected cells and appears to result from damage to the plasmalemma (TRIBE, 1955; WOOD, 1972; HALL and WOOD, 1973). The procedures used to measure permeability changes in plant cells are based on the ability of cells to plasmolyze or deplasmolyze (BROWN, 1915; FUSHTEY, 1957), on the ability of cells to take up and retain Neutral Red, a vital stain (TRIBE, 1955), on uptake (HANCOCK, 1968 b) or loss (MOUNT et al., 1970) of radioiso• topes, and on loss of solutes or water (WOOD, 1972; BASHAM, 1974) from cells. Leakage of electrolytes from infected or enzyme-treated tissue into the bathing medium can be followed with a conductivity bridge (FRIEDMAN and JAFFE, 1960; LAI et al., 1968; HALL and WOOD, 1973). If the electrolyte loss from tissue is plotted as the log of the percentage of electrolytes remaining in the tissue at the time of measurement, the reaction course is approximately linear with time. This means of expressing cell leakage facilitates the determining of the rate of electro lyte loss (KECK and HODGES, 1973; BASHAM and BATEMAN, 197 5 b). Since no one method is completely adequate for the description of permeability alteration in a given system, results obtained by one method should be checked against those obtained with other methods (COLLANDER, 1959). Injury to potato disks treated with purified polygalacturonate lyase from E. chrysanthemi can be detected as electrolyte loss within 3 min after exposure of the tissue to the enzyme; wall degradation, detected by solubilization of unsaturated uronides, parallels electrolyte loss. Tissue maceration, measured by estimating the ease with which treated tissue can be pulled apart, is first evident 10-15 min after enzyme treatment (Fig. 4) (BASHAM and BATEMAN, 1975a). The toxic effect of pectolytic enzymes on plant cells is inhibited by . Cells plasmolyzed during enzyme treatment retain their ability to accumulate Neutral Red, but mace• ration is not markedly inhibited (TRIBE, 1955; FUSHTEY, 1957). Photosynthetic and transport processes are essentially the same in intact tissue and in plasmolyzed tobacco leaf cells isolated from tissue macerated by pectolytic enzymes (FRANCKI et al., 1971; JENSEN et al., 1971; REHFELD and JENSEN, 1973). Cellular protection in tissue exposed to pectic enzymes occurs at osmotic concentrations at and above the point of incipient plasmolysis (FUSHTEY, 1957; BASHAM and BATEMAN, 1975a). The mechanism of wall degradation and tissue maceration is understood; how• ever, the mechanism by which plasma membranes are damaged by pectic enzymes 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 337

3 60r------,

~ U :::J U 0 0.2 40 Ul ~ I Ul -0'" Ul E .3 ';l. c >..'" i: 0 e u t) .:.:

o Time-min

Fig. 4. Wall breakdown and cell mjury in potato disks treated with endopectic acid lyase. A Wall breakdown (---) as lyase activity in Ilmoles of unsaturated uronides released from tissue, and cell injury (--) as % of electrolytes lost from tissue. B Wall breakdown (---) as maceration estimated by loss of cohesion in tissue disks, and cell injury (-) as "kill" estimated by loss of ability of cells to retain Neutral Red [rated on a 0 (no loss) to 5 (complete loss) scale]. (BASHAM and BATEMAN, 1975a) is unknown. Several hypotheses explaining cell injury by pectic enzymes have been proposed and evaluated. One hypothesis suggests that some specific feature of the pectic enzyme mole• cule, apart from its catalytic ability, is responsible for the toxic interaction. Experi• ments with plant tissues treated with pectic enzymes that differ with respect to net ionic charge, pI, pH optima, and mode of substrate cleavage indicate that none of these specific characteristics of pectic enzymes is necessary for cellular injury. All pectic enzymes that cause tissue maceration simultaneously cause cell death (BASHAM and BATEMAN, 1975a). Another hypothesis suggests that soluble reaction products from tissue macerat• ed by pectolytic enzymes are responsible for cellular injury. However, if pectolytic enzyme activity is eliminated from supernatant fluids of pectolytic digests of plant tissue by heat (FUSHTEY, 1957), dialysis, ultrafiltration, or column chromatography, the resulting preparations are not injurious to plant tissue (BASHAM and BATEMAN, 1975a). Beet tissue is relatively resistant to maceration and injury by pectic enzymes. Beet tissue is not damaged when incubated with purified pectic lyase and a suscepti• ble plant tissue which is readily macerated and killed. Thus, no soluble reaction product is detected in the maceration of susceptible tissue which alters the per• meability of beet cell membranes (BASHAM and BATEMAN, 1975a). Hydrogen perox• ide, known to increase the permeability of plant membranes (SIEGEL and HALPERN, 1965), is a by-product of pectolytic enzyme activity in cauliflower floret tissue 338 D.F. BATEMAN and H.G. BASHAM:

(LUND, 1973) and cotton leaf tissue (MUSSELL, 1973). Studies with disks of potato tissue (BASHAM and BATEMAN, 1975a) indicate that is not gener• ated on maceration, yet cells are killed. Hydrogen peroxide at 1,000 times the concentration found in cotton treated with pectic enzymes (MUSSELL, 1973) did not cause any detectable injury to potato cells. Based on all available evidence, it appears that death of tissue treated with pectic enzymes is not related to soluble products generated during wall breakdown. Since there is evidence that polysaccharides may be directly associated with the plasmalemma (ALBERSHEIM and KILLIAS, 1963; ROLAND, 1969; ROLAND and VIAN, 1971), it has been suggested that pectic enzymes might cause cellular injury by direct interaction with a substrate in or on the plasmalemma (MOUNT et aI., 1970). In contrast, it has been reported that isolated plant protoplasts are not injured by pectic enzyme (PILET, 1973; TSENG and MOUNT, 1974). This problem is difficult to resolve since plasmolysis of cells has been shown to inhibit the toxic effects of pectic enzymes, and isolated protoplasts must be plasmolyzed in order to prevent them from bursting. Any potential sites of enzyme interaction with the plasmalemma could be so altered by plasmolysis of protoplasts that the interaction would not occur. In an attempt to circumvent this difficulty, tobacco protoplasts have been subjected to osmotic stretching in the presence of a purified pectic lyase. Such experiments provided no evidence that bursting of protoplast was related to the presence of pectic enzyme (BASHAM and BATEMAN, 1975a). Although these results do not indicate a direct attack on the plasmalemma by pectic enzymes, it is still possible that such an interaction occurs, but the damage caused by the interaction is not expressed until the membrane is under stress from . Investigation of the ability of labelled enzyme to bind to intact protoplasts or isolated membrane fragments would be consistent with this hypothesis, but such results would not be conclusive. Another hypothesis suggests that membrane damage during wall degradation is caused by failure of the enzyme-damaged wall to provide sufficient mechanical support to the plasmalemma (HALL and WOOD, 1973). Several lines of evidence support this hypothesis: (1) Purified pectic lyase from E. chrysanthemi can solubilize over 50% of the total wall sugars in isolated tobacco and potato cell walls, removing not only galacturonic acid, but also rhamnose, arabinose and galactose. This sol• ubilization, the rapid maceration of enzyme-treated tissue, and the release of unsatu• rated uronides from enzyme-treated potato disks, all confirm that purified lyase causes extensive damage to cell walls (BASHAM and BATEMAN, 1975b). (2) There is a close relationship between wall degradation and membrane damage. If the temperature of the reaction mixture or the concentration of enzyme or substrate (potato disks) is varied, the change in the rate of electrolyte loss is directly propor• tional to change in the rate of release of unsaturated uronides from the cell walls. When enzyme-treated potato disks are rinsed during intermediate stages of treat• ment, the rates of uronide release and electrolyte loss are both markedly reduced (BASHAM and BATEMAN, 1975b). (3) Under conditions of plasmolysis, purified lyase degrades walls but does not cause membrane damage; membranes are damaged only when cells are deplasmolyzed (Fig. 5). Curve A shows the rate of electrolyte leakage from nonplasmolyzed potato tissue treated with purified lyase. Curve B shows the rate of electrolyte leakage from tissue treated with active and heat• inactivated enzyme in the presence of plasmolyzing concentrations of mannitol 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 339

1.8

01 C :~ 1.6 E ~ Ul ~1.4'" g u Qj'" ~1.2 o -'

1.0

o 30 60 Time-min Fig. 5. Effect of plasmolysis and deplasmolysis on electrolyte loss from potato disks treated with endopectic acid lyase. Electrolyte loss is expressed as log of the percentage of the total electrolytes remaining in the tissue. A Lyase plus buffer (20 mM Tris-HCl, pH 8.5). B Autocla• ved lyase plus buffer, lyase plus osmoticum (0.4 M mannitol and I % dextran sulfate in buffer), and autoclaved lyase in osmoticum. After 60 min, disks in osmoticum were rinsed for 30 min in osmoticum without enzyme. C Rinsed disks pre-treated with autoclaved lyase, deplasmolyzed in phosphate buffer (5 mM, pH 7.5). D Rinsed disks pre-treated with lyase deplasmolyzed in phosphate buffer. (BASHAM and BATEMAN, 1975b)

(0.4 M). After 30 min the plasmolyzed tissues were rinsed in five changes of manni• tol solution to remove enzyme and then transferred to distilled water. The rate of electrolyte loss from deplasmolyzing tissue treated with heat-inactivated enzyme and then washed (Fig. 5, curve C) was less than that from tissue treated with active enzyme (Fig. 5, curve A) but was greater than that of control rates (Fig. 5, curve B), indicating that physical damage occurred to cells during rapid deplasmoly• SIS. In deplasmolyzing tissue treated with active enzyme and then washed (Fig. 5, curve D) all electrolytes were lost at a rate corresponding to that of free-space equilibration, suggesting that immediately upon deplasmolysis the membranes lost their ability to retain electrolytes. These data were confirmed under identical experi• mental conditions using the Neutral Red assay for cell viability; plasmolyzed tissue was viable in the presence of active enzyme but died immediately upon rinsing and deplasmolysis. Maceration of tissue occurred at about the same rate in the presence and absence of the plasmolyzing agent. Plasmolyzed potato tissue treated with lyase, and observed by phase contrast microscopy revealed that cells which retain Neutral Red often have the majority of their plasmalemma surfaces in contact with the cell wall; thus mere contact between the damaged wall and 340 D.F. BATEMAN and H.G. BASHAM: the plasmalemma does not bring about membrane (BASHAM and BATEMAN, 1975b). These results are consistent with the hypothesis that injury of the plasma• lemma results from enzymatic damage to the cell wall. With plasmolyzed tissue, enzymatic degradation of the wall occurs, but the plasmalemma is not tightly pressed to the inner wall surface. Rinsing the tissue before deplasmolysis should remove the majority of the pectolytic enzyme and any soluble breakdown products from wall degradation, but the altered wall structure remains. On deplasmolysis the plasmalemma expands, interacts with or is not supported by the damaged wall, and lyses. The experiments described do not rule out the hypothesis that enzymes interact directly with membranes, but the close correlation between wall alteration and cell death under various experimental conditions would not be expected if direct membrane damage by the enzyme were involved. Recently published studies by STEPHENS and WOODS (1975) are in agreement with our work (BASHAM and BATEMAN, 1975a and b). Both investigations support the hypothesis that cell injury results directly from damage to the plant cell wall by pectic enzymes.

4.1.4 Depletion of Cell Wall Polysaccharides during Pathogenesis Demonstration of depletion or alteration of cell wall constituents during pathogen• esis is the most direct way of showing the involvement of cell wall-degrading enzymes in plant disease. Analysis of the pectic substances in apple fruit infected with S. jructigena, which causes a firm rot, revealed a decrease in pectic substances of ca. 15%, while tissues infected with soft rot pathogens, B. cinerea and P. expan• sum, lose 50-70% of this wall fraction (COLE and WOOD, 1961); these pathogens are known to secrete pectic enzymes in host tissues. S. sclerotiorum produces pectic, hemicellulolytic and cellulolytic enzymes during pathogenesis. Sunflower and toma• to tissues infected with this pathogen undergo a reduction in the methoxyl content of 74-93%, and a decrease in pectic acid content of 64-82% within 2 to 4 days after appearance of symptoms (HANCOCK, 1966a). The xylan content of infected sunflower tissue did not decrease even though xylanase was present, but 65-75% and 72-97% of the arabinose and galactose content, respectively, was lost (HAN• COCK, 1967). S. sclerotiorum-infected bean tissues show a decrease in IX-cellulose content within two days after inoculation; nine days after inoculation the loss of IX-cellulose from host cell walls is substantial (LUMSDEN, 1969). Loss in bire• fringence of bean cell walls has been associated with cellulase production by R. so/ani in diseased tissue (BATEMAN, 1964 b). Degradation of cellulose in tobacco and tomato stem tissues infected with P. solanacearum is also known to occur (KELMAN and COWLING, 1965). Recent development of improved procedures for analysis of plant cell wall polysaccharides have greatly facilitated studies of cell wall decomposition during pathogenesis. The neutral sugars in cell wall polymers are readily determined by hydrolysis of cell walls with trifluoroacetic acid followed by enzymolysis and gas chromatography (ALBERSHEIM et aI., 1967; JONES and ALBERSHEIM, 1972). The uronic acids in cell wall hydrolysates can also be determined by gas chromatography or with uronic acid dehydrogenase (BATEMAN et aI., 1970; JONES and ALBERSHEIM, 1972). These new procedures are effective for assay of all cell wall polysaccharides except cellulose. Cell walls isolated from R. solani lesions on bean hypocotyls 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 341

show about 50% reduction in trifluoroacetic acid-hydrolyzable sugar within 24- 30 hours after tissues are inoculated (BATEMAN, et aI., 1969); depletion of galactur• onic acid in cell walls of infected tissues also occurs (BATEMAN, 1970). Infected tissues contain polygalacturonase, /3-1,4 galactanase, arabanase, xylanase and cellu• lase. The removal of galacturonic acid, rhamnose, galactose, and xylose occurs rapidly following infection. Amounts of cell wall constituents recovered in the above studies are expressed as milligram percent of the total cell wall, and loss of a given sugar is reflected by the differences in its recovery from equal weights of wall prepared from healthy and diseased tissues. Since lesions contain less cell wall per unit volume of tissue than healthy tissue, comparisons based on sugars released from equal weights of wall from healthy and diseased tissues give a conservative estimate of wall degradation during pathogenesis. Analysis of bean hypocotyl tissues infected by S. rolfsii, a pathogen that secretes a potent mixture of cell wall-degrading enzymes, revealed that about 60% of the noncellulosic cell wall polysaccharides are removed within 3 days after inoculation if the results are expressed as the recovery of cell wall sugars per unit length of hypocotyl (Fig. 6) (BATEMAN, unpublished). The degradation of isolated host cell walls by pathogen enzymes produced in culture or diseased tissue has been used to estimate the potential of these pathogens to cause wall breakdown in vivo (BATEMAN et aI., 1969; JONES et aI., 1972; MULLEN, 1974). Although this procedure may not reflect the degree of wall degradation that actually occurs in diseased tissue, it is more accurate than the use of model substrates for this purpose. Cell walls in some diseased tissues become melanized and thus extremely difficult to analyze, since such walls are

4W

400

350

320

>. 280 ou ~ 240 >• I E 200 ~ (]l "'- 150

120

80

40

OL...J'-:R~h-aL---l'-:::F:-'u-c"---LA:-'ra Xyl Man Gal Glc Gal A Glc A Fig. 6. Sugar content per cm of hypocotyl of healthy (D) and Sclerotium rolsii-infected (_) bean cell walls from lO-day-old plants 3 days after inoculation. Rha (rhamnose), Fue (fucose), Ara (arabinose), Xyl (xylose), Man (), Gal (galactose), Gle (noncellulosie glucose), Gal A (galacturonic acid), and Glc A (glucuronic acid). (BATEMAN, unpublished) 342 D.F. BATEMAN and H.G. BASHAM:

resistant to hydrolysis by trifluoroacetic acid and enzymes; the use of isolated cell walls from healthy host tissues in such situations may be a preferable approach in evaluating the cell wall-degrading ability of pathogens. Cell walls isolated from juvenile tissues normally are more susceptible to enzymatic degradation than walls from mature tissues (BATEMAN et aI., 1969). In diseases characterized by extensive degradation of infected tissue, the direct analysis of walls from such tissue clearly shows the involvement of wall-degrading enzymes in disease. Cell wall-degrading enzymes may be important in pathogenesis or in diseases not character• ized by extensive tissue degradation. In such cases a histological or cytochemical approach, coupled with the demonstration of specific enzymes in infected tissues, may have merit. Although Ruthenium Red and ferric hydroxylamine have been used to detect uronic acids in cell walls (ALBERSHEIM et al., 1960; RIEDEL and MAl, 1971), and loss in birefringence of cell walls reflects alteration in cellulose, there is a general lack of reliable cytological procedures for detection of localized changes in specific cell wall polymers. The use of fluorescent (GOLDMAN, 1968; ULRICH, 1975) for localizing specific wall-degrading enzymes in infected tissues may aid our understanding of their involvement in host-parasite interactions, but there are numerous technical difficulties with this approach. The enzymatic degradation of lignin is characteristic of the saprophytic break• down of woody cell walls by a variety of wood-rotting basidiomycetes, but the of this degradation is, at best, incompletely understood. The basis of the difficulty in unravelling the mechanism of the enzymatic degradation of lignin lies in the structural complexity of the lignin polymer. A variety of heterogeneous enzyme preparations have been shown to degrade isolated lignin to various degrees and to cleave specific bonds in low molecular weight model compounds, but in no case are the nature of the degradation products from natural lignin, or the enzyme systems involved, well characterized (KIRK, 1971). The reader is referred to several reviews for summaries of these systems and the techniques involved in approaching this complex problem (SCHUBERT, 1965; PEARL, 1967; KIRK, 1971).

4.2 Enzymatic Degradation of Membranes

The increase in membrane permeability characteristic of most host-pathogen inter• actions (THATCHER, 1942; TRIBE, 1955; WHEELER and HANCHEY, 1968; HANCOCK, 1972), cytological observations that membranes are degraded in many plant diseases (CALONGE et aI., 1969; Fox et aI., 1972), and knowledge that plant pathogens can produce phospholipases and proteinases which degrade membrane constituents (PORTER, 1966; VAN ETTEN and BATEMAN, 1965; TSENG and BATEMAN, 1968), can easily lead to the hypothesis that these enzymes are involved in membrane break• down in diseased tissue (TRIBE, 1955; LUMSDEN and BATEMAN, 1968). There has been, however, almost no critical work that indicates that these enzymes are involved in the permeability changes associated with attacking mechanisms of plant pathogens or that these enzymes have the ability to degrade plant membranes. Production of phospholipases in vitro is quite common with pathogens that cause tissue decay (SATOMURA et aI., 1960; TSENG and BATEMAN, 1968; LUMSDEN, 1970; TSENG and CHANG, 1970). Phospholipase activity is also known to increase in diseased tissues, and in a few cases these enzymes have been characterized. 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 343

Phospholipase B is found in bean tissues infected with T. basicola (LUMSDEN and BATEMAN, 1968) and S. rolfsii (TSENG and BATEMAN, 1969), and phospholipase C is produced in culture filtrates and in potato tissue infected by E. carotovora (MOUNT et aI., 1970). Activity of an uncharacterized phospholipase increased in bean leaves infected with Uromyces phaseoli, and there were concomitant changes in lipids and fatty acids in the region of the rust pustules. Increased permeability of infected tissue, however, occurred 2--4 days before the increase in phospholipase activity was observed (HOPPE and HEITEFUSS, 1974). Our knowledge of proteinase production by pathogens is also sparse; many microorganisms produce proteolytic enzymes (MATSUBARA and FEDER, 1970), and these enzymes are often found in diseased tissue (Kut and WILLIAMS, 1962; PORTER, 1966; VAN ETTEN, 1966; MOUNT et aI., 1970; REDDY et aI., 1971). Proteinase activity has been reported in extracts of cucumber leaves infected with Pseudomonas lachrymans, and levels of free amino acids increase in diseased tissue. The apparent proteinase produced by this pathogen in culture was stable to autoclaving (KEEN et aI., 1967a and b). Subsequent work with P. lachrymans cultures revealed that the" proteolytic" factor was not an enzyme, but a lipomucopolysaccharide that complexes with some proteins, making them soluble in trichloroacetic acid (KEEN et aI., 1969) which resulted in a false positive assay for proteinase activity. Thus, the nature and origin of the proteinase activity in extracts of P. lachrymans-infected leaves is not clear. In no case has a careful identification and analysis of the spectrum of potential membrane-degrading enzymes produced by a pathogen in vivo and in vitro been carried out in a critical manner. Attempts to show correlations between of pathogens and their ability to produce proteinases and phospholipases have generally been based on in vitro studies (FRIEDMAN, 1962; KEEN et aI., 1967b; BERAHA et aI., 1972), yet it has been shown that production of these enzymes in vivo and in vitro is not necessarily correlated (HANCOCK and MILLAR, 1965). Cytological investigations show that host membranes in potato tubers infected with E. carotovora (Fox et aI., 1972) are degraded, and it is known from independent studies that this organism produces enzymes that degrade membrane constituents (MOUNT et aI., 1970). Culture filtrates of Phytophthora palmivora were shown to contain a heat-labile factor which destroyed purified cucumber membrane frag• ments (CALONGE et aI., 1969), but no effort was made to identify this factor. Since many non-enzymatic membrane-perturbing factors are known to be produced by plant pathogens (PAGE, 1972), cytological observations of infected tissue or work with crude enzyme preparations tell nothing about the ability of phospholi• pases or proteinases to degrade membranes. Highly purified phospholipase C and proteinase produced by E. carotovora lyse isolated cucumber protoplast (TSENG and MOUNT, 1974). These enzymes, how• ever, do not kill cells of intact potato tissue. Commercial preparations of phospholi• pase C or proteinase do not lyse isolated Avena protoplasts (RUESINK, 1971). Though no firm conclusions can be drawn from these studies, the approach shows promise of providing definitive information. Until it is shown that specific, purified enzymes of pathogen origin can degrade plant membranes in vitro, that such enzymes and the pathogen cause similar mem• brane degradation in host tissue, and that these specific enzymes of pathogen origin are present in diseased tissue, there is no experimental basis for concluding 344 D.F. BATEMAN and H.G. BASHAM:

that membrane degradation in tissue decay is caused by pathogen enzymes. At our current stage of knowledge, the assumption that these enzymes serve as attack• ing mechanisms necessary for the initiation of pathogenesis is even more speculative. Techniques are available for determining the involvement of phospholipases and proteinases in plant diseases, but they must be applied systematically with appropri• ate systems before we can know the significance of these enzymes in pathological processes.

5. Summary and Additional Considerations

When considering the significance of the association and involvement of enzymes produced by pathogens capable of altering or degrading plant structural com• ponents in pathological processes, it is important to view these degradative enzymes as but one of a complex of attacking mechanisms possessed by most pathogens. Many pathogens capable of producing enzymes that degrade the structural com• ponents of plants also possess the ability to produce (WOOD et aI., 1972), organic acids (BATEMAN and BEER, 1965; MIROCHA, 1972; MAXWELL, 1973) and/or growth regulators (SEQUEIRA, 1973) that may function in concert with or in lieu of degradative enzymes during pathogenesis. Also, enzymes of host origin are of probable importance in cell wall and/or membrane damage following infection (TANI, 1967; WILSON, 1973). Cell wall-degrading enzymes, particularly pectic enzymes, play a key role in pathogenesis by soft rot pathogens. They may be just as important in other diseases, but at present there is insufficient experimental evidence to support this point of view. Cell wall-degrading enzymes appear to enable pathogens to invade host tissues and convert the host cell wall polymers into utilizable nutrients. In diseases caused by facultative pathogens, cell wall-degrading enzymes and perhaps mem• brane-degrading enzymes function in host destruction during both pathogenesis and saprogenesis. The pectic enzymes, those that split the IX-l,4 glycosidic bonds in the galacturo• nide fraction of cell walls in a random or endo manner, are responsible for tissue maceration in diseased tissues; the action of these enzymes renders the nonuronide polymers in plant cell walls more susceptible to other wall-degrading enzymes. It is now clear that pectic enzymes are toxic to plant cells and are directly or indirectly responsible for injury to cell membranes and cell death. Because of the multiple effects pectic enzymes have on living plant tissues, and the almost universal ability of plant pathogens to produce them, this group of enzymes is of paramount importance in understanding pathogenesis by many phytopathogenic orgamsms. Our detailed knowledge of the significance and involvement of the hemicellu• lases, , proteinases and phospholipases in pathogenesis is poorly developed at present, but the broad association of these enzymes with plant diseases and the availability of adequate biochemical technology to work with these enzyme systems in diseased plants render these areas of study fertile for exploration. There are two other areas of work impinging upon our understanding of tissue degradation 4.3 Degradation of Plant Cell Walls and Membranes by Microbial Enzymes 345 in pathogenesis that are in need of development: the involvement of enzymes of pathogen origin in the digestion of cutin, and the relationship of synthesis and breakdown of suberin and lignin-like substances in diseased tissues. Some phytopathogenic fungi have been shown to digest cutin (LINSKENS and HAAGE, 1963; VAN DEN ENDE and LINSKENS, 1974), but this problem has not received the attention it deserves. The chemistry of cutin, suberin, and lignin-like substances and their associations with specific cell wall constituents is poorly understood. The resolution of the problems attendant on this area, however, may contribute greatly to our understanding of the significance of cell wall degradation in pathogen• esis. It is evidently necessary to merge more fully our biochemical knowledge of tissue breakdown with cytological and histochemical studies of pathogenesis. Such a combined approach seems essential if the role of degradative enzymes in pathogenesis by obligate parasites and pathogens which do not generate massive destruction of plant tissue is to be understood. Also, attention needs to be focused on those degradative events associated with the initiation of infection and coloniza• tion of plant tissues by pathogens. Too much of our current information is based on analysis of diseased tissues after the critical events in pathogenesis have occurred.

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