Pathological Anatomy and Histochemistry
of Leucostoma Canker on Stone Fruits and Other Selected Fungal Cankers of Deciduous Fruit
Trees
Alan R. Biggs, West Virginia University
1. Introduction
2.1. Pathological Anatomy of Leucostoma Cankers
2.1.1. Peach
2.1.2. Sweet cherry
2.2. Histochemical Aspects of Leucostoma Infection
2.3. Temporal Changes in the Anatomy of the Infection Court and Pathogen Resistance
3.1. Pathological Anatomy
4. Valsa Canker
4.1. Pathological Anatomy
5.1. Pathological Anatomy
6. Comparative Anatomy of Periderm Formation in Response to Wounds and Fungal Pathogens
7. Canker Expansion and Routes of Re-entry for Pathogens
8. Conclusions
1. Introduction
There are a great many fungal pathogens of fruit trees which cause twig and stem cankers and which also cause leaf spots and/or fruit rots. There are some fungal pathogens, however, which primarily cause stem cankers and which cause major economic losses in some regions. This latter group of pathogens will be the focus of this chapter. Infection of deciduous fruit trees by canker-causing fungi usually involves breaching the host's passive barriers to pathogen ingress via wounding. Wounding in orchard trees can be caused by agents such as insects, humans, lightning, wind, hail, animals, and nutritional and physiological disorders. Alternatively, natural leaf abscission and cold temperature injury also play key roles in creating infection courts for these pathogens. This chapter describes the pathological anatomy and histochemistry of fungal cankers of deciduous fruit trees, focusing, on only four diseases: those caused by Leucostoma spp. and Botryosphaeria spp. on stone fruits, but also found on pome fruits, and Nectria galligena and Valsa ceratosperma, found primarily on pome fruits. Most of the other canker diseases described in the literature are similar to these examples.
Leucostoma canker, also called perennial canker, Cytospora canker, and Valsa canker, is recognized as an important disease of peach in the northern portions of the region suitable for production of temperate fruits, including Canada, and the northeastern United States.1 It is mentioned as part of a complex set of factors associated with peach tree short life syndrome in the southeastern United States.2 It is equally important on other stone fruits including prune and plum in California 3 and Idaho.4 In Europe, the disease is important on apricot, peach, sweet cherry, and is part of the disease complex of stone fruits called 'apoplexy'.5-7 The disease also occurs in South America and Japan.8,9 The host range of the two causal organisms is similar and they may be found on a wide variety of species within, as well as outside, the family Rosaceae, including apple; apricot; Sitka mountain ash; blackthorn; black, Japanese, flowering, pin, sour, and sweet cherries; chokecherry; nectarine; Russian olive; peach; pear; common, Damson, and wild plums; prune; serviceberry; and golden willow. The pathological anatomy of this disease has been the subject of several investigations.
The fungi which cause Leucostoma cankers on stone fruits, Leucostoma cincta (Pers. ex Fr.) Höhn. [= Valsa cincta (Pers. ex Fr.) Fr.] and L. persoonii (Nits.) Höhn. [= Valsa leucostoma (Pers. ex Fr.) Fr.], are ascomycetes in the order Diaporthales, family Valsaceae. The imperfect stages of these fungi, Leucocytospora cincta (Sacc.) Höhn. [= Cytospora cincta Sacc.] and Leucocytospora leucostoma (Pers.) Höhn.) [= Cytospora leucostoma Sacc.], respectively, are encountered most commonly in the field.
The appearance of Leucostoma canker varies depending on the part of the tree infected and the specific pathogen involved. Infections of small twigs appear as sunken, discolored areas, often with alternating zonation lines, usually around winter-killed buds or leaf scars from the previous year's foliage (Fig. 1). Nodal infections, usually but not always associated with L. cincta, are observed 2 to 4 weeks after bud break. The infected tissues become darker with time and an amber colored gum may ooze from the infected tissue unless the twig is killed entirely. Previous year's shoots that develop in the center of the tree are quite susceptible to nodal infections and, if left untreated, the fungus can invade rapidly into scaffold limbs and large branches. Branch cankers that result from this type of infection will have dead twigs or twig stubs at the center of the canker.
Cankers that form on the main trunk, branch crotches, scaffold limbs, and older branches are the most conspicuous expression of fungal infections. Usually the first external symptom of such cankers is the copious quantity of amber-colored gum. Gum production is a natural response of the tree to irritation, but that due to infection by Leucostoma spp. is excessive to the point of being detrimental. As the cankers age, the gum becomes dark brown, the infected bark dries out and cracks open, exposing blackened tissue beneath. Cankers appear elliptical along the length of the stem.
Beginning in the late spring and continuing through the summer months, the tree grows rapidly and is able to resist further penetration of the fungus into healthy tissues. During this time, the tree may form a callus ring around the canker but the fungus usually invades this tissue again in late fall or early spring when the tree is dormant and can not actively resist penetration. The yearly alternation of callus production and canker extension produces a canker with concentric callus rings. Where tree defenses are compromised by environmental stresses, callus production may be inhibited and cankers may appear more diffuse. Branch or twig infections may produce leaf symptoms during the growing season. Leaves on an infected branch often turn yellow, droop down, and may wilt and die. Dead twigs and branches are usually covered with a multitude of pin-head sized black pycnidia erupting through the dead bark.
A. Pathological Anatomy of Leucostoma Cankers
1. Peach
In 1-year-old peach shoots exhibiting tip dieback from natural infection in the field by either L. cincta or L. persoonii 10, fungal mycelium was present within the cankered area in the cortical tissue, phloem, xylem vessels, and pith. The mycelium was intracellular in the xylem vessels and pith. The pathogens appeared to spread via the pits in these tissues. In the phloem and cortex, fungal growth was intercellular. Mats of fungus mycelium separating adjacent rows of cells often were seen in this region (Figs. 2 - 4).
At the transition zone between healthy and diseased tissues, mycelium was detected regularly in the xylem vessels but was observed infrequently in cortical tissue, phloem and pith. Mycelium could be observed in vessels, but not in other tissues, up to 1 cm beyond the visible margin of the canker. The walls of xylem vessels located up to 2 cm beyond the margin of the cankers exhibited a brown discoloration, primarily in the vessels near the pith but extending throughout the wood with increased proximity to the canker margin (Figs. 5 and 6). The lumens of many xylem vessels, both within the cankered region and beyond the canker margin, became plugged with gum which darkened with age.10
In 1-year-old peach bark inoculated in the field with mycelium of L. persoonii, fungal colonization of cortical and phloem tissues was initially diffuse although mycelial aggregations formed within 3 weeks.11 Host responses to inoculation over the initial one-week period were largely confined to the vascular cambium which gave rise to a barrier zone of gum ducts present in the xylem and forming a distinct band (Figs. 8 - 13). Gum ducts continued to enlarge during the 3 to 8 weeks following inoculation and a second row of ducts was formed in some samples. Gum oozed from the inoculated sites during the 11-month period of study, except during periods of low rainfall and during dormancy. Plugging of vessel elements with gum was sporadic and tyloses were observed only rarely. Diffuse mycelial wedges were present within the gum duct barrier zone in the xylem (Fig. 7).
Wound periderm (Figs. 10 and 11) was formed in the bark tissues 2 to 3 weeks after inoculation, however mycelial aggregates accumulated in the necrotic tissue, followed by the eventual penetration of wound periderm tissues by mass action and ingress into living tissue. Formation of a new wound periderm occurred and the proliferation of the fungus was again slowed. Five months after inoculation, the pathogen could be observed penetrating into living tissues by breaching the wound periderm in the bark and along the barrier zone of gum ducts in the xylem. In general, colonization of woody tissue was poor and was confined to the outer xylem (Figs. 14 and 15). Pith tissues remained free from fungal colonization.
Samples taken 11 months after inoculation, at the beginning of the second growing season, showed extensive necrosis of noninvaded bark tissues 2 to 3 mm in advance of fungal colonization.11 The vascular cambium had produced another layer of barrier zone gum ducts. All tissues located outside the previous season's xylem appeared necrotic, suggesting that living tissues had succumbed after the onset of cambial activity. Observations on the presence of necrosis in advance of colonization, which occurred twice in the study, were emphasized. The role of a toxin in pathogenesis has been previously reported or suggested for this disease 12-14 and other diseases caused by Cytospora spp. 15-16
The comparative anatomy of 2-year-old peach stems from relatively resistant and susceptible cultivars inoculated with mycelium of L. cincta and L. persoonii has been described 17, and these observations have helped resolve some of the discrepancies among earlier studies.10,11,18 At 7 days postinoculation, the cultivar Candor had longer cankers than the cultivar Sunhaven following inoculations with either fungus (Fig. 16). Colonization around the stem circumference was more extensive for L. persoonii on both cultivars relative to L. cincta (Fig. 17). Regression analyses of the relationship between postinoculation time and canker length showed that Candor was colonized more aggressively by L. persoonii than any of the other host/pathogen combinations. In addition, Sunhaven inoculated with L. persoonii showed more aggressive colonization than either of the two cultivars inoculated with L. cincta. Interestingly, there was a second period of rapid increase in canker length, width, and colonization rate between days 28 and 35 in both cultivars inoculated with L. persoonii (Figs. 16 and 17).
The extent of fungal invasion, as determined by isolation, varied with time and the particular cultivar/pathogen combination (Table 1). Fourteen days after inoculation, all cultivar/pathogen combinations showed approximately equal invasion of bark and current year's xylem. Fungi were not recovered from 1- or 2-year-old xylem or pith. However, by 28 days postinoculation, the cultivar Candor, inoculated with L. persoonii, (the more susceptible cultivar paired with the more aggressive fungus) yielded this fungus from 1- and 2-year-old xylem located under the point of inoculation and 1 cm proximal and distal to this point. The extent of xylem colonization in current year's xylem was equal to that found in bark, i.e. the margin of visible symptoms. Notwithstanding, by day 56, L. persoonii could be isolated from all tissues, including the pith; and the extent of colonization in 2-year-old xylem and pith exceeded the visible canker margin in the bark by 4 cm proximally (the maximum distance examined) and 1 cm distally.
The extent of colonization in the cultivar Sunhaven inoculated with L. persoonii , and in Candor inoculated with L. cincta, was similar. Pathogens from both cultivars were isolated infrequently from 2-year-old xylem and, when found, were limited to the region under the point of inoculation. In 1-year-old and current year's xylem, L. persoonii was equally advanced relative to bark symptoms in Sunhaven, whereas in Candor, L. cincta was in xylem in advance of visible bark symptoms 3 cm proximally and 1 cm distally. Isolations of L. cincta from Sunhaven (the more resistant cultivar paired with the less aggressive fungus), were limited to bark and current year's xylem. Xylem colonization was ahead of that in bark by 1 cm proximal to the visible canker margin but equal to that in bark distally. No pathogens were detected in wounded, noninoculated control tissues.
In mechanically wounded, control tissues and in tissues colonized by L. persoonii and L. cincta, lignified and suberized cells formed a boundary separating living tissues from necrotic tissues and the environment. The location and chemical qualities of these tissues may influence or impart pathogen resistance properties in bark and xylem. In cankered tissues, several discontinuities in bark and xylem boundaries were observed which occurred in the absence or as a result of infection.17 In noninoculated wounds inflicted with a cork borer, discontinuities in suberin formation occurred at the juncture of the suberized wound periderm with the original periderm and with primary phloem fibers; the centripetal surface of the wound callus as it grew across the exposed xylem; and, in the xylem, the presence of nonsuberized vessels, trachieds, and axial parenchyma. Where suberin was absent in xylem and in bark adjacent to fibers, increased deposition of lignin-like material was observed. The periderm juncture and callus discontinuities appeared time dependent and remained nonsuberized for 2 and 4 weeks, respectively.
In tissues inoculated with L. persoonii or L. cincta, suberization was less continuous than in wounds. Suberization was observed only infrequently in xylem although deposition of lignin-like substances occurred in a visible barrier zone. In infected bark, the pattern of suberization was similar to that of wounds, although the shapes of wound phellem cells were altered in some samples. Altered phellem appeared thin-walled and rounded, compared with the relatively thick, rectangular phellem seen at wounds. The junctions of suberized tissues with the original periderm and\or primary phloem fibers often were colonized by wedges of fungus mycelium. In both cultivars inoculated with L. cincta, wound periderm formation appeared effective in limiting colonization in bark during the 8 week course of study. With the Sunhaven/ L. cincta combination, xylem responses (described below) were associated with limited colonization relative to the other cultivar/fungus spp. combinations.
The nonsuberized, centripetal region of the callus was readily colonized by both pathogens in the early stages of callus formation. As callus tissues progressed in their development, the suberized boundary comprised of current year's xylem parenchyma was often ruptured from the inside by the coalescing of enlarged gum ducts. The rupture then provided access for the hyphae to the gum duct region, thus facilitating longitudinal invasion in a manner similar to that described by Wisniewski et al.11 In addition, the rupture allowed fungal access to parenchymatous tissues of the differentiated callus, resulting in rapid circumferential spread of mycelium in the bark. Direct penetration of well-formed wound periderm (at least three cells thick) was observed once in this study.17 Mycelia were more likely to separate adjacent rows of phellem, parallel to the axis of the tissue, after access to the tissue was gained via lenticels or growth cracks. Primary ligno-suberized boundary tissues and wound periderms with one to two layers of phellem were penetrated readily by hyphal aggregations produced by both fungi.
In bark of both Sunhaven and Candor inoculated with L. persoonii and L. cincta, necrophylactic periderm formed at 21 to 28 days after inoculation and was associated with an effectively delimited lesion in both cultivars inoculated with L. cincta, the less pathogenic fungus. For cultivars inoculated with L. persoonii, the wound periderm was not as thick, had fewer cells, and was associated with only a slight inhibition of lesion expansion followed by a second rapid increase in canker length as the periderm was breached between 28 to 35 days after inoculation 19 (Figs. 16 and 17). In contrast, Hebard et al.20 observed that periderms in American chestnut were produced by the host and then breached by the chestnut blight fungus, Cryphonectria parasitica, at 18 or 28 days after inoculation depending on the virulence of the isolate. In resistant chestnuts, some virulent isolates continued to expand lesions up to 30 days after inoculation.21
Numbers of suberized phellem cells formed following inoculations of peach cultivars Sunhaven and Candor with L. persoonii or L. cincta differed significantly depending upon fungus species, cultivar, and the orientation of the canker margin (longitudinal or transverse) (Fig. 18). Fungus species alone accounted for the greatest amount of variation in cell number (29.3%), followed by the cultivar x orientation interaction (14.1%). For example, trees inoculated with L. cincta had nearly twice the numbers of necrophylactic periderm cells as those inoculated with L. persoonii. At 56 days postinoculation, bark tissues colonized by L. cincta possessed significantly more suberized phellem relative to the other fungus species x time combinations. The numbers of phellem cells were significantly less for Candor in the longitudinal orientation (at the proximal and distal canker margins) relative to the other cultivar x orientation combinations. Xylem tissues also became suberized in response to wounding or fungal infection. Suberin was observed in ray parenchyma cells of 1- or 2-year-old xylem present at the time of wounding or inoculation, and in current year's xylem formed after inoculation and at a distance from the point of inoculation. In the latter case, suberization was limited to xylem parenchyma in the gum duct zone as described by Wisniewski et al.11
It was shown, with quantitative fluorescence microphotometry, that wounded bark of the more resistant cultivar Sunhaven had 73% more suberin relative to the cultivar Candor.17 Furthermore, the frequency of stem wounds exhibiting suberization at 8 days postwounding was 100% for Sunhaven and only 33% for Candor. Interestingly, amounts of suberin in wounded xylem of Candor was slightly higher that of Sunhaven. Tissue deposition patterns for lignin and suberin following wounding were independent of cultivar. For example, Sunhaven formed more suberin in bark and more phloroglucinol-positive material in xylem than Candor. In contrast, Candor formed more lignified material in bark and more intensively suberized xylem ray parenchyma than Sunhaven.
2. Sweet Cherry
Cankers caused by L. persoonii on sweet cherry in Germany 7 are initiated in the bark via wounds in the periderm and through leaf scars injured by autumn frosts during the leaf abscission period. Symptoms of the latter type of infection consisted of cankers with dead buds or fruiting spurs in their center. In winter and early spring, the pathogen grows mainly in the bark and less so in the xylem. In summer and fall, the fungus is found growing only in xylem. Because of this alternating, two types of cankers develop on sweet cherry.
The first type of canker is characterized by bark necrosis and wound periderm formation by the host in late spring. Cankers cease to enlarge until the following winter when the fungus, which has survived in cavities around the wound periderm, circumvents the wound periderm and invades healthy bark tissue. In this type of canker, which always originates from bark infections, the fungus is not able to grow from the older xylem or pith into the bark.
In the second type of canker, the fungus can be seen in young xylem immediately beneath the vascular cambium. Goring maintains that the fungus kills the cambial cells by the action of a toxin, resulting in heavy gum production in the bark external to the dead cambium. The pathogen could not be isolated from the necrotic bark tissues in July. Wound periderm production in bark is suppressed in this canker type, also due to the action of the putative toxin. This second canker type is found only on large trees which exhibited extensive necrosis on the trunk and main branches. It was suggested that in the first canker type the current year's xylem shields the vascular cambium from the effects of the fungus which is growing in the older wood.
B. Histochemical Aspects of Leucostoma Infection
Few histochemical studies have been conducted on tissues infected with Leucostoma spp. 11,17,18 These studies have shown that lignin (and/or lignin-like substances, hereafter referred to collectively as lignin) and suberin are the main constituents of cell walls in the boundary zone that is formed prior to wound periderm formation in bark.18 The lignin in this ligno-suberized tissue can be detected with phloroglucinol/HCl, but not with the Mäule reaction or the chlorine-sulfite test. The first phloroglucinol-positive reactions were observed 72 hr after inoculation or wounding. In inoculated tissues, lignin was detected in crushed phloem sieve elements, sieve element cell walls, and xylem vessel plugs and vessel pits, up to 2 cm in advance of hyphae. As the boundary zone developed in the bark, the red phloroglucinol chromophore was first seen in the corners of the compound middle lamellae of adjacent hypertrophied cells. The cell walls in the boundary zone then became uniformly reactive over the next 4 days for wounded tissues. In inoculated tissues, lignification can be delayed up to 21 to 28 days, or, in certain locations, it may not develop.17,18 Results of tests for tissue aldehydes (Schiff reagent prior to periodic acid hydrolysis, orcinol) gave color reactions in the same positions as described for phloroglucinol/HCl.
Tests with Azure B as a histochemical reagent for lignin showed that this test was not specific for lignin in peach tissues.18 Although some green coloration was imparted to lignified cell walls with this reagent, the reaction was weak. Azure B also showed strong affinity for the walls of cells in a zone that appeared amber-yellow without staining, and which appeared strongly reactive with the periodic acid-Schiff test for polysaccharide substances. Toluidine blue O also yielded poor color differentiation for lignified tissues.
Nonwounded, healthy periderm exhibited a strong reaction with many of the histochemical reagents for suberin in these studies.11,17,18 Observations with phosphine GN (chrysaniline yellow), Sudan IV, Sudan black B, and epifluorescence microscopy were the most reliable, whereas results of tests with crystal violet, Sudan III, and methylene blue were variable.18 Suberized wound periderms appeared pinkish with Sudan IV and examination of sections treated with Sudan IV with blue-violet fluorescence excitation showed bright red fluorescence associated with only the most recently suberized cells. Application of phosphine GN and examination with ultraviolet fluorescence excitation induced a silvery-white fluorescence of all suberized cells. Nonstained tissues mounted in glycerol and examined with ultraviolet fluorescence exhibited violet autofluorescence of suberized tissues, however in boundary zone tissues this was often masked by the bright blue autofluorescence of lignin. As the boundary zone developed, in both control wounds and inoculated tissues, cells which had become lignified formed a thin suberin lining on the inner sides of the cell walls.18,19,22,23 These linings were most easily visualized under ultraviolet fluorescence in tissues reacted with phloroglucinol/HCl or toluidine blue O (fig. 12). These reagents acted to quench any lignin autofluorescence, similarly Sudan black B quenched suberin autofluorescence, helping to confirm further the chemical nature of these cell wall layers.18
Suberized cells were of two basic types. Suberization occurred in tissues that were present in the wound reaction zone or in advance of colonization. The tissue was comprised of dedifferentiated cells with lignified walls lined with lamellar suberin. Cell division was not involved in the formation of this tissue. Suberization of extant tissues occurred following formation of the impervious boundary zone of the bark in the inner periderm, cortex, and primary and secondary phloem. Secondary suberization in the bark occurred after cell division and resulted in the formation of new phellogen and its derivatives. Secondary suberized cells occurred in the necrophylactic periderm which formed in all wounded or colonized bark tissues.17 Callose deposits in infected peach bark and xylem appeared as bluish-green fluorescent flecks when stained with aniline blue and examined under ultraviolet excitation. Callose was regularly associated with the amber-yellow, polysaccharide-rich zone in wounded and infected tissues. It was observed as early as 24 hr after wounding or inoculation and it occurred at the margins of the wound or infection court, about 5 to 10 cells in from the surface. The intensity and frequency of callose deposition was not influenced by fungal colonization except in necrotic sieve elements observed 7 days postinoculation. In callus tissue, callose was seen in some poorly differentiated phloem sieve elements at 35 days postinoculation.
Degradation of pectin and cellulose was a regular feature of tissues colonized by L. persoonii.18 For pectic substances, the ruthenium red reaction and the iron absorbtion test 18
showed degradation of pectic substances adjacent to mycelial aggregations as early as 7 days postinoculation. Similar results were obtained with histochemical tests for cellulose (birefringence, IKI-H2SO4, and zinc-chloriodide).
C. Temporal Changes in the Anatomy of the Infection Court and Pathogen Resistance
Following wounding, the process of establishing boundaries in the infection court between healthy tissues and the external environment confers resistance to infection in many host/pathogen interactions. Wounds generally become increasingly less susceptible to infection with age.21,24-26 Resistance to infection in this manner is probably related to a number of nonspecific plant responses leading up to and including formation of primary ligno-suberized tissues and secondary wound (or necrophylactic) periderm. The major structural components of these tissues are lignin and suberin.27 See the online wound section at this site for a more thorough discussion of wound anatomy and the interaction of pathogens and wounds.
3. Botryosphaeria Canker of Peach
Peach tree fungal gummosis was first observed in central Georgia in 1970 28 and is now reported throughout Alabama, Florida, Georgia, and Louisiana. Three species of Botryosphaeria have been associated with the disease and wound inoculations with B. obtusa (Schw.) Shoem. and B. rhodina (Berk. and Curt.) Arx produced symptoms identical to those produced by B. dothidea (Moug. ex Fr.) Ces. & DeNot.29 The infection court for all three species is pruning wounds, although B. dothidea also can penetrate lenticels.30,31 Penetration of lenticels by the two former species has not been observed, but these species can invade lesions caused by B. dothidea. Isolation of B. obtusa and B. dothidea from naturally infected tissue was more common than B. rhodina. B. dothidea was more active in the summer, whereas B. obtusa predominated in cankers in the winter and spring.32
The disease is characterized by numerous sunken necrotic lesions about 7 to 14 mm in diameter and usually located at lenticels. Gum exudation occurs in excess. Shallow, round to oval, brown, gummy lesions, about 12 to 25 mm in diameter, can be seen upon removal of the outer bark. Symptoms of the disease usually occur first on the trunk during the second or third growing season. As the disease progresses, scaffold limbs and twigs may be affected. On young branches lenticels may be swollen but gumming does not occur. Reproductive stroma of the causal organisms are produced in infected lenticels.
Inoculation of wounded peach bark with spores of B. obtusa and B. dothidea provided an interesting contrast to the mycelial inoculation studies described above for Leucostoma canker.33 Initial colonization by the fungus occurred on the wound surface, in necrotic bark tissue on the wound margin, and in necrotic xylem tissue to a depth of about 200 m. In inoculated trees, few macroscopic symptoms of infection were present during the eight week course of the study, and normal wound healing responses appeared macroscopically and microscopically to be proceeding similar to the noninoculated wounds. Histologically, no differences in lignin and suberin deposition, periderm formation, and callus formation were observed between noninoculated wounds and wounds inoculated with either fungus after one and two weeks incubation. However, by 28 days after inoculation for both fungi, new callus tissue was being colonized by fungal hyphae located on the surface of the xylem beneath the nonsuberized ventral callus surface. The nonsuberized ventral region of callus was primarily parenchymatous with a ligno-suberized outer layer, and was the focal point for fungal pathogenesis. The actions of fungal toxins as irritants to this nonsuberized, highly meristematic region is probably critical to successful pathogenesis. By eight weeks, the breakdown of tissue in this region was directly associated with fungal hyphae in close ventral proximity. Direct fungal penetration of the callus, although only occasionally observed, was not required to induce gum pockets, cambial alterations, and tissue breakdown. These effects are consistent with the production of a fungal toxin or toxins as described recently for B. obtusa.34 Intracellular hyphae of both Botryosphaeria spp. were observed colonizing cortical and callus parenchyma, and xylem ray parenchyma, vessels and tracheids. Intercellular hyphae were observed in phloem fibers, necrotic cortical parenchyma external to wound periderm, and necrotic ligno-suberized parenchyma on the callus surface.
4. Valsa Canker of Apple
Valsa canker, caused by the fungus Valsa ceratosperma (Tode ex Fr.) Maire (anamorph Cytospora sacculus (Schwein.) Gvritischvili) is an important disease of apple in the Pacific Rim countries, including Japan, China, and Korea.35 It is only found occasionally on pear and quince. In northern Japan, the disease is especially severe with more than 35% of orchards affected to some degree. The disease is manifested as cankers around fruit scars, twig stubs, branch crotches, and sites of winter injury or mechanical injury to the bark. The surface of infected bark may appear swollen and water soaked and, when wet, the centers of cankers may appear pinkish. Infections are commonly seen as elongated cankers on the upper sides of the larger limbs of older trees. Most new lesions appear in spring, between March and late April.
Cankers elongate rapidly in the spring and early summer. Canker growth is slow in the remainder of summer and during the winter. The fungus can be isolated from about 5 mm beyond the margin of actively growing cankers. Single hyphae of Valsa ceratosperma rapidly invade the cortical and phloem tissues during the spring and early summer. Colonization proceeds similarly, albeit more slowly, during the dormant season. A transition zone of collapsed cells separates infected from noninfected tissue. As temperatures increase in the summer, the host forms a lignified layer in the transition zone, followed by formation of cork layers with thin-walled phellem cells. During this period, the fungus forms mycelial aggregates which penetrate and destroy wound periderm tissues by mass action.36 However, repeated formation of cork layers eventually results in a decreased rate of lesion development. Wound periderm composed of thick-walled cells is formed by August, however, as temperatures decline in the autumn, healthy tissue is re-invaded by the fungus which grows through defective phellem layers between the periderm and cortex or between the phloem and xylem. The authors concluded that the wound cork layers acted as a temporal barrier to mycelial invasion and that the rate of formation of periderm depended upon host metabolic activity.36
5. Nectria Canker of Apple, Pear and Quince
Nectria canker, caused by the fungus Nectria galligena Bres. (anamorph Cylindrocarpon heteronemum (Berk. & Broome) Wollenweb.) occurs in North America, Chile, Australia, New Zealand, northern continental Europe, the United Kingdom, South Africa, and Japan. In North America, the disease, also known as European, apple or crotch canker, occurs in the northeastern United States and southeastern Canada and westward to the Pacific Coast. The fungus which causes Nectria canker has a broad host range, including species of aspen, beech, birch, elm, and hickory, among others.37
The first visible symptoms of infection are observed at nodes and appear as elliptical, sunken areas. The infected area may appear water-soaked and darker than the adjacent, healthy tissue. Infected twigs and branches may be girdled as the cankers enlarge causing the collapse of portions of the shoot distal to the infection. If shoots are not killed during the first year they are infected, the host may produce an annual layer of callus surrounding the area colonized by the fungus. The following year, the fungus invades and kills the callus tissue and, when this cycle of healing and reinvasion occurs annually, the cankers develop a concentric or zonate appearance. Gelatinous sporodochia of the fungus are common on cankers during wet weather and the bright red to orange perithecial stage often appears on older cankers in late fall or winter.
Nectria galligena is able to enter the host through wounds inflicted by pruning, frost injury, breakage caused by ice and snow 38, and small dead branch stubs.39,40 In apple, N. galligena enters the host through pruning wounds, frost injury, infections caused by Venturia inaequalis and Neofabrea malicorticis, and leaf scars.41,42 For the fungus to become well established, wounds must be to the depth of the vascular cambium. The host is able to confine the pathogen to the cortex when wounds are shallow. The fungus is able to colonize all of the tissues which it penetrates. In the cortex and phloem, hyphae are intercellular in the early stages of infection but intracellular hyphae were observed during the later stages. The fungus grows vigorously in the phloem region and enters the xylem via the medullary rays. Although the pathogen is unable to penetrate suberized cell walls 43, it grows freely in the vessels and tracheids and spreads in the xylem through the pits. Intracellular phenolic compounds, gum in the xylem, and wound lignin at the levels observed in this host/pathogen interaction do not deter fungal colonization.
6. Comparative Anatomy of Periderm Formation in Response to Wounds and Fungal Pathogens
In tree bark, delimitation of fungal cankers ultimately occurs via the formation of ligno-suberized boundary tissue derived from cells present at the time of inoculation and necrophylactic periderm derived from newly differentiated phellogen.18,19,44,45,46 The same statement is accurate for "healing" processes which begin immediately after wounding. For some cankers, barriers may be effective for preventing continued colonization and diseased tissues may be sloughed. For established perennial cankers, bark boundary tissues, which also may contain extensive xylem inclusions,46 are either directly penetrated or circumvented annually resulting in a series of concentric callus ridges.
The direct involvement of the ligno-suberized layer and/or wound periderm, or by-products generated during the differentiation of these tissues, in the resistance of woody plants to fungi has never been demonstrated conclusively. Several lines of evidence indicate that periderms function to prevent desiccation of internal layers, prevent inward movement of pathogen toxins or enzymes, and serve as physical barriers to colonization. The process of generating new tissues also may result in lignification of pathogen hyphae 17 or in the production of fungistatic compounds which act as phytoalexins.44
Several recent papers have described the formation of necrophylactic periderm and associated tissues in response to wounding and pathogen inoculation of peach trees.17,19,33 Some notable differences were observed. For example, pathogens affect the position or location of periderm and related tissues in wounded bark relative to their location in the absence of pathogens (Figs. 19 and 20). In wounds, new tissues are regenerated usually within 1-2 mm of the wound surface.18 When wounds are colonized by aggressive pathogens, the formation of new tissues may be several centimeters from the infection court. While this may seem obvious and, perhaps, insignificant, the location of natural defensive barriers in trees is of paramount importance for the proper management of canker diseases.
Pathogens affect the morphology and differentiation of new tissues and may retard or inhibit the formation of periderm and/or the deposition of suberin in wound periderm and xylem barrier zones (Figs. 21 and 22).7,11,19,33,36 Such alterations could influence the effectiveness of the periderm in limiting further spread of the pathogen within the host.
Few hosts are able to wall out the initial invasion of a pathogen with the formation a single ligno-suberized boundary zone and necrophylactic periderm, as usually occurs at wounds. Instead, the tree often forms a series of periderms, each exhibiting varying degrees of development, within a single season in response to the advancing fungus. The fungus will continue to spread in the host tissue until conditions become more favorable for the host to form a complete periderm and less favorable for the pathogen to penetrate further. This is why many canker lesions on juvenile stems have visible concentric areas of discoloration during the initial establishment phase. For perennating cankers, the tree also produces annual rings of callus tissue which represent the successful formation of necrophylactic periderm and callus tissue for that year but which gives no clue as to the number of attempts during that particular year.
7. Canker Expansion and Routes of Re-entry for Pathogens
Very few studies on hardwood cankers, in general, or fruit tree cankers, in particular, have presented evidence to show conclusively how fungi invade the phellogen layer to reinfect the phloem after the canker has stopped expanding for the season. For many canker-causing fungi, the main mode of reinfection is by direct hyphal penetration of the wound periderm. Penetration may be either direct via mass action of hyphae or indirect via hyphal penetration of fibers or sclereids which occasionally bridge wound periderms and connect diseased areas with healthy tissue. Penetration of periderms by mass action has been described for Leucostoma canker of peach,11 Nectria canker of apple,42 and Valsa canker of apple.36 In contrast, Cryphonectria parasitica, which forms diffuse cankers on American chestnut, grows under wound periderm before it forms48 and thus can rapidly girdle stems. In addition, Hebard et al.20 showed that mycelial fans of this fungus also could penetrate developing wound periderms directly.
Other avenues of reinfection occur in natural and pathogen-induced discontinuities along the wound periderm and new callus tissue produced by the host in response to invasion. For example, L. persoonii can circumvent wound periderm by colonizing the nonsuberized ventral callus region or by growing between the original periderm and wound periderm external to the cortex, or between the wound periderm and phloem fibers.18,19 Similar routes of reinfection were described for Valsa canker of apple.36
The lumens of phloem fibers also may provide a means of entry for fungi into healthy tissues delimited by wound periderm. Apple bark colonized by N. galligena42 and poplar bark colonized by Cytospora chrysosperma15 and may be reinvaded in this manner. Similarly, Eutypella parasitica may reinvade maple bark through sclereids or other discontinuities in the wound periderm.49 Colonization of phloem fibers also has been described for Leucostoma canker of peach18, and illustrated for Botryosphaeria canker of peach.33
Ashcroft50 proposed that fungi such as N. galligena which colonize the xylem may reinvade the bark by growing through the medullary rays and into the wound wood (CODIT wall 4) and ray parenchyma into healthy bark that was not delimited by wound periderm. This has never been demonstrated although French49 mentions it as a possibility for E. parasitica.
It is likely that most fungal canker pathogens can re-invade the host by any or all of the above routes. Reinvasion is accomplished by single hyphae or fungal mass action in combination with toxin production, and may be facilitated by environmental factors, such as cold injury to the new phellogen, and host physiological status.
8. Conclusions
Histological studies have provided a rudimentary understanding of the host/pathogen interaction between woody hosts and canker-causing fungi. We know that resistance to canker pathogens is largely nonspecific and therefore we must pay increasing attention to the host if progress in understanding these diseases is going to continue. Elucidating the factors that trigger periderm formation in woody plants could lead to improved control of these diseases via manipulation of the host reaction following pruning. The relative role of resistance in bark and xylem tissues is not understood and more research is needed to elucidate the relative importance of the many and varied contributions of partial resistance in woody plants. We know little about the importance and extent of pathogen variation for these diseases. We know that environment plays a major role in the interaction between host and pathogen, although only few studies have attempted to quantify environmental influences. Because of this, we lack the information required to predict accurately the occurrence of canker diseases. More importantly, we lack the knowledge required to formulate reliable plant health criteria to avoid canker diseases.
Literature Cited
1. Biggs, A.R., Integrated control of Leucostoma canker of peach in Ontario, Plant Dis., 73, 869, 1989.
2. Ritchie, D.F., and Clayton, C.N., Peach tree short life: A complex of interacting factors, Plant Dis., 65, 642, 1981.
3. DeVay, J.E., Gerdts, M., English, H., and Lukezic, F.L., Controlling Cytospora canker in President plum orchards in California, California Agriculture, 28, 12, 1974.
4. Helton, A.W., and Konicek, D.E., Effects of selected Cytospora isolates from stone fruits on certain stone fruit varieties, Phytopathology, 51, 152, 1961.
5. Babos, K., Rozsnyay, Zs. D., and Klement, Z., Apoplexy of apricots. V. Pathological and histological investigations of the apoplexy of apricots, Acta Phytopathol. Acad. Sci. Hung., 11, 71, 1976.
6. Portilla, M.T., and Tuset, J.J., Chancros en fruitales de hueso: Leucocytospora cincta (Sacc.) Höhnel un coelomycete activo en el melocotonero, Inv. Agrar. Prod. Prot. Veg., 1, 259, 1986.
7. Goring, M.-C., Zur Ätiologie der Valsa-Krankheit an Süsskirsche. Histologische Untersuchungen über Eintrittspforten und Ausbreitung von Leucostoma persoonii (Nits.) Hohnel, Mitt. Biol. Bundesanst, Land. Forstwirtsch, Berlin-Dalhem v. 162, 1975.
8. Biggs, A.R., Leucostoma Canker, in Compendium of Stone Fruit Diseases, Ogawa, J. M. and Zehr, E. I., Eds., APS Press, St. Paul, 1992, 20.
9. Togashi, K., Studies on the pathology of peach canker, Bull. Imp. Coll. Agric. For., Morioka, Japan, 16, 178 pp., 1931.
10. Tekauz, A. and Patrick, Z.A., The role of twig infections on the incidence of perennial canker of peach, Phytopathology, 64, 683, 1974.
11. Wisniewski, M., Bogle, A.L., and Wilson, C.L., Histopathology of canker development on peach tress after inoculation with Cytospora leucostoma, Can. J. Bot., 62, 2804, 1984.
12. Rozsnyay, D. S. and Barna, B., Apoplexy of apricots IV. Studies on the toxin production of Cytospora (Valsa) cincta Sacc., Acta Phytopathologica Academiae Scientiarum Hungaricae, 9, 301, 1974.
13. Tsakadze, T. A., The action of the toxin of Cytospora leucostoma on the plant cell, Bull. cent. bot. Gdn, Moscow (In Russian) 35, 75, 1959.
14. Svircev, A. M., Biggs, A. R., and Miles, N. W., Isolation and partial purification of phytotoxins from liquid cultures of Leucostoma cincta and L. persoonii, Can. J. Bot., 69, (in press), 1991.
15. Biggs, A.R., Davis, D.D., and Merrill, W., Histology of cankers on Populus caused by Cytospora chrysosperma, Can. J. Bot., 61, 563, 1983.
16. Koganezawa, H., and Sakuma, T., Possible role of breakdown products of phloridzin in symptom development by Valsa ceratosperma, Ann. Phytopathol. Soc. Jpn., 48, 521, 1982.
17. Biggs, A.R., Comparative anatomy and host response of two peach cultivars inoculated with Leucostoma cincta and L. persoonii, Phytopathology, 76, 905, 1986.
18. Biggs, A.R., Boundary zone formation in peach bark in response to wounds and Cytospora leucostoma infection, Can. J. Bot., 62, 2814, 1984.
19. Biggs, A.R., Intracellular suberin: occurrence and detection in tree bark. Int. Assoc. Wood Anat. Bull. n.s., 5, 243, 1984.
20. Hebard, F.V., Griffin, G.J., and Elkins, J.R., Developmental histopathology of cankers incited by hypovirulent and virulent isolates of Endothia parasitica on susceptible and resistant chestnut trees. Phytopathology, 74, 140, 1984.
21. Russin, J.S., and Shain, L., Initiation and development of cankers caused by virulent and cytoplasmic hypovirulent isolates of the chestnut blight fungus, Can. J. Bot., 62, 2660, 1984.
22. Biggs, A.R., Detection of impervious tissue in tree bark with selective histochemistry and fluorescence microscopy, Stain Technol., 60, 299, 1985.
23. Biggs, A.R., and Stobbs, L.W., Fine structure of the suberized cell walls in the boundary zone and necrophylactic periderm in wounded peach bark. Can. J. Bot., 64, 1606, 1986.
24. Butin, H., Über den Einfluss des Wassergehaltes der Pappel auf ihre Resistenz gegenüber Cytospora chrysosperma (Pers.)Fr., Phytopathol. Z., 24, 245, 1955.
25. Cline, M.N., and Neely, D., Wound-healing process in geranium cuttings in relationship to basal stem rot caused by Pythium ultimum, Plant Dis., 67, 636, 1983.
26. Bostock, R.M. and Middleton, G.E., Relationship of wound periderm formation in resistance to Ceratocystis fimbriata in almond bark, Phytopathology, 77, 1174, 1987.
27. Kolattukudy, P.E., Biochemistry and function of cutin and suberin, Can. J. Bot., 62, 2819, 1984.
28. Weaver, D.J., A gummosis disease of peach trees caused by Botryosphaeria dothidea, Phytopathology, 64, 1429, 1974.
29. Britton, K.O., and Hendrix, F.F., Three species of Botryosphaeria cause peach gummosis in Georgia, Plant Dis., 66, 1120, 1982.
30. Weaver, D.J., Role of conidia of Botryosphaeria dothidea in the natural spread of peach tree gummosis, Phytopathology, 69, 330, 1979.
31. Pusey, P.L., Influence of water stress on susceptibility of nonwounded peach bark to Botryosphaeria dothidea, Plant Dis., 73, 1000, 1989.
32. Britton, K.O., and Hendrix, F.F., Population dynamics of Botryosphaeria spp. in peach gummosis cankers, Plant Dis., 70, 134, 1986.
33. Biggs, A.R. and Britton, K.O., Presymptom histopathology of peach trees inoculated with Botryosphaeria obtusa and Botryosphaeria dothidea, Phytopathology, 78, 1109, 1988.
34. Venkatasubbaiah, P., Sutton, T. B., and Chilton, W. S., Effect of phytotoxins produced by Botryosphaeria obtusa, the cause of black rot of apple fruit and frogeye leaf spot, Phytopathology, 81, 243, 1991.
35. Sakuma, T., Valsa canker, in Compendium of Apple and Pear Diseases, Jones, A. L. and Aldwinckle, H. S., Eds., APS Press, St. Paul, 1990, 39.
36. Tamura, O. and Saito, I., Histopathological changes of apple bark infected by Valsa ceratosperma (Tode ex Fr.) Maire during dormant and growing periods, Ann. Phytopathol. Soc. Japan, 48, 490, 1982.
37. Grove, G. G., Nectria canker, in Compendium of Apple and Pear Diseases, Jones, A. L. and Aldwinckle, H. S., Eds., APS Press, St. Paul, 1990, 35.
38. Lortie, M., Pathogenesis in cankers caused by Nectria galligena, Phytopathology, 54, 261, 1964.
39. Grant, R.J. and Childs, T.W. Nectria canker of northeastern hardwoods in relation to stand improvement, J. For., 38, 797, 1940.
40. Zalasky, H., Penetration and initial establishment of Nectria galligena in aspen and peachleaf willow, Can. J. Bot., 46, 57, 1968.
41. Wiltshire, S.P., Studies on the apple canker fungus. I. Leaf scar infection, Ann. Appl. Biol., 8, 182, ,1921.
42. Crowdy, S.H., Observations on apple canker. III. The anatomy of the stem canker, Ann. Appl. Biol., 36, 483, 1949.
43. Krähmer, H., Wound reactions of apple trees and their influence on infections with Nectria galligena, J. Plant Dis. Prot., 87, 97, 1980.
44. Mullick, D.B., The non-specific nature of defense in bark and wood during wounding, insect and pathogen attack, Recent Adv. Phytochem., 11, 395, 1977.
45. Biggs, A.R., Merrill, W., and Davis, D.D., Discussion: Response of bark tissues to injury and infection, Can. J. For. Res., 14, 351, 1984.
46. Biggs, A.R., Anatomical and physiological responses of bark tissues to mechanical injury, in Defence Mechanisms of Woody Plants Against Fungi, Blanchette, R.A. and Biggs, A. R., Eds., Springer-Verlag (Berlin), 1992, chap. 2.
47. Biggs, A.R., Responses of angiosperm bark tissues to canker and canker rot fungi, in Defence Mechanisms of Woody Plants Against Fungi, Blanchette, R.A. and Biggs, A. R., Eds., Springer-Verlag (Berlin), 1992, chap. 3.
48. Bramble, W.C., Reaction of chestnut bark to invasion by Endothia parasitica, Am. J. Bot. 23, 89, 1934.
49. French, W.J., Eutypella canker on Acer in New York, State Univ. Coll. For. Tech. Publ. No. 94, Syracuse, NY, 1969.
50. Ashcroft, J.M., European canker of black walnut and other trees, West Virginia Agr. Expt. Sta. Bull. 261, 1934.
