Skin-Set, Wound Healing, and Related Defects

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:471

Trimsize:165×240 MM

Chapter 22

Skin-Set, Wound Healing, and Related Defects Edward C. Lulai USDA-ARS, Northern Crop Science Laboratory, 1307 18 Street N, Fargo, ND, USA

22.1 INTRODUCTION The physiology and biochemistry of the development of resistance to tuber skinning injury (idiom = skin-set), wound healing, and wound-related defects are of global importance because of the magnitude of food and financial losses impacted by these processes. The amount of these losses is difficult to determine because of the large range of infections, bruise defects, water vapor loss, and various quality issues that are affected by inadequate skin-set and slow wound healing. Collectively, minor to serious wounding and bruising can average 40%, resulting in serious food quality and loss problems and the creation of grower–processor contracts with stringent incentives to reduce these losses (Hampson et al., 1980; Brook, 1996). This chapter will discuss important physiological and biochemical research that impacts these costly wound-related problems. This chapter will discuss the formation and maturation of native tuber periderm and its relationship to tuber skin-set development in section 22.2. Skinning wounds are difficult to control during harvest unless the tuber periderm has matured so that the skin is set and resistant to skinning injury. The structure of tuber periderm and the maturational changes that result in resistance to tuber skinning injury are of fundamental importance in developing physiological approaches to enhance skin-set and reduce associated losses. The current status of this relatively young research area is described. The following section, 22.3, discusses tuber wound-healing in detail. The process of wound-induced suberization to heal skinned, cut, and so-called bruised areas covers a vast research plane. Sections 22.3 includes information on the induction and regulation of suberization, and the composition, biosynthetic pathways, assembly, and molecular structure of suberin. These areas of suberin physiology and biochemistry are not fully understood, but they are of great importance in mitigating infection, defect development, and other losses. Suberin is somewhat of an enigma that is often misunderstood and poorly described in conjunction with wound healing and wound periderm development. Consequently, the section covers what is currently considered appropriate terminology and description of suberin. The final section, 22.4, discusses wound-related defects including shatter bruise, blackspot bruise, growth cracks, and skinning. These defects directly affect food losses and market quality. The physiology and biochemistry of these defects are discussed in relation to wounding and suberization. Potato Biology and Biotechnology: Advances and Perspectives D. Vreugdenhil (Editor) 2007 Published by Elsevier B.V.

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

472

28-4-2007

9:37 a.m.

Page:472

Trimsize:165×240 MM

Lulai

22.2 NATIVE PERIDERM AND SKIN-SET 22.2.1 Native periderm formation A well-developed intact periderm with its suberin biopolymer provides the primary barrier against disease, insects, dehydration, and physical intrusions for the potato tuber (Lulai, 2001a). These critical roles of protecting and preserving the tuber are indicative of the importance of the native periderm and wound periderm development. Prior to development of the native periderm, an epidermis exists for a short time on the youngest tubers of approximately 1 cm or less in diameter. Epidermal tissues are created as the underground stem of the potato plant, i.e. the stolon, swells to form the nascent potato tuber (Artschwager, 1918; Peterson and Barker, 1979). Stomata are scattered in the epidermis and permit gas exchange. The native periderm forms from epidermal tissues of the nascent potato tuber. Tuber periderm consists of three distinct types of cells; each cell type is grouped into separate layers: (1) phellem, (2) phellogen, and (3) phelloderm (Fig. 22.1). Periderm formation in the emergent tuber is initiated by periclinal division of the epidermal and subepidermal cells, but the meristematic layer of periderm cells, referred to as the phellogen or cork cambium, is formed from the hypodermis (Peterson and Barker, 1979). The phellogen then divides outwardly forming the phellem of the periderm. The phellem is a corky material consisting of several layers (approximately 4–10, depending on genotype, environment, and stage of growth) of well-organized, rectangular suberized cells located at the very surface of the tuber. The innermost layer of periderm cells, the phelloderm, neighbors the phellogen which utilizes substrate sources and starch collectively found in the phelloderm and neighboring cortical cells as is evidenced by the lack of starch granules in these cells. The phellem and phelloderm cells are derivatives of the phellogen and as such are organized into the same rectangular file as that of the original phellogen cell from which the file originates. During periderm development, the meristematic action of the Phellem tensile component Periderm Phellem (skin)

Phellogen shear component

Phellogen Phelloderm Cortical cells

Fig. 22.1. Outline of the cell walls comprising tuber native periderm (immature) and neighboring cortical cells illustrating the two types of fractures that occur on tuber skinning/excoriation injury. Skinning injury occurs by (1) exceeding the tensile strength of the phellem resulting in a fracturing of the phellem cell walls and tearing of the fabric-like skin (phellem tensile component) and (2) a fracture of the phellogen cell walls (phellogen shear component). (Courtesy: E.C. Lulai, Am J. Potato Res. 2002).

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:473

Trimsize:165×240 MM

473

Skin-set, wound healing, and related defects

Fig. 22.2. A fully suberized tuber lenticel visualized by treatment with Sudan III/IV. The arrow on the left side points to the red coloration created by the partitioning of Sudan III/IV into the suberin poly(aliphatics) domain. Other cell wall material is visible because of autofluorescence. (Courtesy: E.C. Lulai, S. Goecke and T.J. Wirta).

phellogen also forms lenticels below stomata that existed in the previous epidermal tissues (Artschwager, 1918; Peterson and Barker, 1979) (Fig. 22.2). As the tuber grows, the radial and longitudinal dimensions increase, thereby requiring commensurate growth of the periderm tissues to maintain the protective suberized covering on the tuber surface. The meristematic action of the phellogen layer continues to form cells necessary to replace sloughed cells and maintain the native periderm on the expanding surface of the growing tuber. During tuber growth, the surface of the periderm may remain smooth, as it does in red- and white-skinned genotypes, or it may develop a rough surface. These rough surfaces of the potato tuber are referred to as russeted or netted skins. The biology of native periderm and wound-induced suberization to heal wounds and bruises that breach or damage the native periderm are of great importance in minimizing disease, tuber rot during storage, and the development of various defects. Among the most common and yet problematic types of tuber wounds are those created by tuber skinning injury. 22.2.2 Skin-set: a part of native periderm maturation During growth and for a period during potato plant senescence, the periderm covering the tuber is immature and as such is fragile and susceptible to abrasion. These abrasion-type wounds are frequently referred to as scuffing, skinning injuries, or skinning wounds. This type of periderm damage can be extensive during harvest and handling operations, and results in costly disease, dehydration, and defect development that adversely affects all sectors of the potato industry including processing, fresh market, and seed (Hampson et al., 1980; Lulai and Orr, 1993, 1994). The ensuing losses are consistent with the

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

474

28-4-2007

9:37 a.m.

Page:474

Trimsize:165×240 MM

Lulai

250- to 1000-fold increase in the initial rate of water vapor loss from skinned areas compared with non-skinned areas of freshly harvested tubers (Lulai and Orr, 1995). Water vapor loss from freshly harvested tubers that are damage-free is up to 28 times greater than that from mature tubers (Lulai and Orr, 1994). The process of periderm maturation is initiated after growth ceases, and the potato vines die or begin to die in the field (Murphy, 1968). The decades-old approach employing vine/haulm killing or desiccation remains the standard means of promoting periderm maturation and related development of resistance to skinning injury. However, results are varied, and reasonably good skin-set development generally requires about 3 weeks of periderm maturation following vine-killing treatment (Lulai and Orr, 1993). Cultural practices and conditions, such as excessive amounts of fertilizer applied or remaining in the soil because of dry growing conditions followed by rain near the time of harvest, may drastically increase the time required for skinset (Stark and Love, 2003). In addition to cultural conditions, potato genotype is a major factor in skin-set development; this can be broadly illustrated where russeted genotypes generally mature more quickly whereas red-skinned genotypes often develop skin-set slowly (Lulai and Orr, 1993). Some potato genotypes are deemed unacceptable for commercial production because inherent skin-set development is exceedingly slow, leading to excessive skinning wounds, disease, and development of related defects. The term ‘skinning’ has often been loosely used to imply that the entire periderm becomes physically detached from the underlying tuber cells; this is not the case as will be discussed in this chapter. The term excoriation (Latin: ex – out of; corium – skin) has been used to describe this same type of skinning wound but with emphasis that the entire periderm does not become detached (Lulai and Freeman, 2001). The terms excoriation and skinning will be used interchangeably to streamline this discussion. The development of resistance to excoriation, or skin-set, is an important part of periderm maturation that occurs during and after potato vine senescence as well as postharvest (Lulai and Orr, 1993, 1995; Bowen et al., 1996; Pavlista, 2002). Although slow or hindered development of resistance to excoriation during periderm maturation is a serious and costly problem, little is known about the physiology of susceptibility and resistance of the potato tuber to skinning injury. 22.2.3 Skin-set and native periderm physiology Tuber periderm is composed of (1) phellem (suberized cells), (2) phellogen (cork cambium), and (3) phelloderm (parenchyma-like cells derived from the phellogen) tissues (Reeve et al., 1969). Analysis of mature tuber periderm, however, may not produce easily identifiable phellogen or phelloderm (Lyshede, 1977). A study of all three periderm cell types in immature and mature periderm was needed to determine maturational changes. Only recently have these periderm cell structures been clearly illustrated for easier identification and associated morphological description (Lulai and Freeman, 2001; Lulai, 2002). The suberization processes involved in phellem development are only partially characterized (Kolattukudy, 1980, 2001; Lulai and Morgan, 1992; Thomson et al., 1995; Bernards and Lewis, 1998; Lulai and Corsini, 1998; Lulai, 2001a; Bernards, 2002). Considering the long history of potato cropping and the breadth and depth of global potato research, it is surprising that earlier identification was not made of the type of

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:475

Trimsize:165×240 MM

475

Skin-set, wound healing, and related defects

periderm cells and cellular changes involved in susceptibility and resistance to tuber excoriation. The lack of fundamental information, particularly at the cellular level, describing the simplest aspects of susceptibility and resistance to excoriation, has hampered the development of effective, rational approaches to describe periderm maturation and associated skin-set development. In turn, there has been a lack of technological advancements necessary to move toward solving the costly problem of tuber skinning injuries that occur during harvest and that hinder successful long-term storage of tubers. The lack of research led to non-scientific explanations for skinning and skin-set, which resulted in postulates incorrectly ascribing skin thickness, periderm thickness, and suberization as determinants of susceptibility and resistance to tuber skinning in immature and mature tubers. These characterizations of skin-set often incorrectly refer to the skin, i.e. phellem, as the periderm of the potato tuber even though the skin constitutes but one of the three types of cells that make up the periderm (Reeve et al., 1969). Because these postulates and idioms arose without scientific investigation or verification, they have become entrenched as descriptive vernaculars and they have been appropriately found in various reviews (Hiller et al., 1985; Peterson et al., 1985; de Haan, 1987; Hiller and Thornton, 1993). Research advancements have moved toward new information and hypotheses describing periderm maturation and excoriation. The ability to objectively measure the status of skin-set development is an important requisite for this research. A few techniques have been developed to objectively measure the total resistance to skinning during periderm maturation (Ostby et al., 1990; Halderson and Henning, 1993; Lulai and Orr, 1993; Muir and Bowen, 1994; Bowen et al., 1996). All of these techniques rely on measurement of the physical resistance to skinning injury, i.e. the tangential or torsional force required to mechanically shear the phellem from the tuber. Results obtained using the basic principle for these techniques were quantitatively related to observed tuber skinning injury (Pavlista, 2002). The ability to objectively measure the development of resistance to skinning injury is essential for assessing the effectiveness of cultural practices intended to address skin-set development and for uncovering physiological factors associated with susceptibility and resistance to excoriation. However, a uniformly acceptable means of objectively measuring skin-set has not been adopted. Postharvest controlled environment studies, in conjunction with objective measurement of skin-set, have shown that for some genotypes low relative humidity may hasten periderm maturation and the development of resistance to excoriation in freshly harvested tubers (Lulai and Orr, 1993). Periderm maturation was more rapid in tubers from cultivars with characteristically higher water vapor loss, particularly russeted genotypes (Lulai and Orr, 1994). Periderm maturation and skin-set development did not relate to phellem/skin thickness, phellem/skin weight, or phellem histology. Also, the method of haulm destruction did not influence skin morphology (Lulai and Orr, 1993, 1994; Bowen et al., 1996). These results suggested that the first layer of fully hydrated cells within the periderm, i.e. the phellogen, should play an important role in tuber periderm maturation and skin-set development. Until recently, there was no published information available on the changes that occur within the cork cambium/phellogen of potato tuber periderm as growth ceases and as the periderm matures (Lulai and Freeman, 2001). Extensive studies had been conducted on the structure, ultrastructure, cytology, and biochemistry of the vascular cambium of perennial woody plants and taproots as the plants cycle through

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

476

28-4-2007

9:37 a.m.

Page:476

Trimsize:165×240 MM

Lulai

growth and dormancy. These are periods when the vascular cambium correspondingly cycles from being meristematically active to inactive (Catesson, 1994; Catesson et al., 1994; Chaffey et al., 1998; Lachaud et al., 1999). The changes in cell wall architecture of the vascular cambium from perennial plants may be a poor model for the changes in cork cambium/phellogen from periderm tissues of annual plants such as potato tubers. In potato tuber, the cells of the lateral meristem irreversibly change from meristematically active to inactive. However, as noted in section 22.2.4, the overall changes in cell wall morphology occurring in the vascular cambium as it enters dormancy are very similar to those found in tuber phellogen as it becomes meristematically inactive upon periderm maturation. 22.2.4 Periderm architecture and skinning injury Lulai and Freeman (2001) investigated the cellular architecture of immature and mature tuber periderm at the light and electron microscope levels and developed a new paradigm for susceptibility and resistance to tuber skinning injury. They confirmed that the periderm consisted of phellem, phellogen, and phelloderm cells and defined the physiology of skinning by showing that the phellem portion of the periderm comprises what had been loosely referred to as the skin. They further showed that skin thickening and suberization are not the source of development of resistance to skinning injury upon periderm maturation. Instead, the tissue separation responsible for skinning injury occurred solely within the phellogen layer (cork cambium) of immature periderm. As discussed earlier, in section 22.2.1, the phellogen is a lateral meristem from which the periderm is formed (Artschwager, 1918, 1924; Peterson and Barker, 1979). The cell walls of the phellogen mechanically connect and hold the phellem, i.e. skin, to the underlying phelloderm cells, which are tough and rigidly connected to the neighboring cortical tissues. The phellogen cells of immature periderm are meristematic and have thin walls that are easily fractured. This fracturing was found to be synonymous with skinning injury. The walls of these cells were shown to strengthen and thicken considerably and were no longer susceptible to fracture upon development of full resistance to skinning injury and periderm maturation. Although the changes that occur in tuber phellogen cells during periderm maturation and the susceptibility of immature phellogen cell walls to fracture prior to periderm maturation are only beginning to be studied, the biomechanics and fracture of plant cell walls in general have been described (Niklas, 1992a,b). During tuber skinning, two physical fractures or breakages occur within the periderm. The modulus associated with each of these fractures consists of the stress and strain related to (1) tearing of the fabric-like phellem/skin when the tensile strength is exceeded and (2) shearing type of fracture incurred by the radial cell walls in the phellogen. Lulai and Freeman (2001) showed that the phellogen is the specific area of contiguous fracture upon skinning and that phellogen cell wall strengthening is a determining factor for skin-set development. Lulai (2002) quantitatively determined the role of phellem/skin tensilerelated fractures and shear-related fractures in susceptibility and resistance to skinning injury and in skin-set development (Fig. 22.1). The relative strength of the phellem tensile component was nearly constant for the time points tested during periderm maturation for each cultivar and did not measurably increase as the periderm approached maturation. These results indicated that the phellem tensile component did not significantly contribute

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:477

Trimsize:165×240 MM

477

Skin-set, wound healing, and related defects

to skin-set development. However, the force required for fracture of the phellogen shear component did increase upon periderm maturation. Results indicated phellogen shear was the major determinant for development of resistance to skinning injury and confirmed phellogen cell wall strength as the determinant for susceptibility and resistance to tuber excoriation. Collectively, this information showed that the nature of tuber phellogen cell wall strengthening is of major importance to tuber excoriation and skin-set. 22.2.5 Cellular changes associated with skin-set Research has been conducted to determine maturational changes that occurred in phellem, phellogen, and phelloderm cell walls in native periderm in comparison with changes observed in wound periderm (Sabba and Lulai, 2002, 2004, 2005). As the wounded tuber tissue began to heal, it formed a closing layer (Fig. 22.3) where the walls of existing parenchyma cells became suberized. In conjunction with the formation of the closing layer, a wound phellogen formed under the closing layer. The meristematic action of the wound phellogen formed a wound phellem and a wound phelloderm. This newly formed wound periderm was subject to similar susceptibility and later resistance to excoriation as that of the native periderm. Thickening of wound phellogen cell walls, after meristematic activity ceased, was evident at the light microscope level. Both the phellogen and the phelloderm are often difficult to discern. Sabba and Lulai (2004, 2005) resolved this difficulty by developing a technique employing toluidine blue O, which differentially stained periderm cell walls. The technique proved to be useful in immunolocalization

Closing layer

Wound periderm

Phellem

Phellogen Phelloderm

100 μm

Fig. 22.3. Suberized closing layer and wound periderm viewed using epifluorescence microscopy 31 days after wounding. Autofluorescence (AF) reveals only those walls that have accumulated poly(phenolic) material and will not show poly(aliphatic) material without further cytochemical treatment. Note suberized cells of the closing layer, i.e. existing parenchyma cells that were induced to suberize prior to development of a wound periderm. The wound periderm consists of (1) phellem cells, suberized cells derived from phellogen meristematic action, visualized by poly(phenolic) AF; (2) phellogen, cork cambium or meristematic layer, not suberized and poorly visible under these AF conditions; and (3) phelloderm, derived from phellogen, does not suberize and is weakly AF, and therefore not visible. Under the wound periderm are parenchyma cells; in this case, cortical parenchyma cells that weakly AF and are therefore not visible. (Courtesy R.P. Sabba and E.C. Lulai).

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

478

28-4-2007

9:37 a.m.

Page:478

Trimsize:165×240 MM

Lulai

studies where, in separately stained parallel sections, the orthochromatic staining (blue) of phellem walls could be used to identify these cells (Sabba and Lulai, 2004, 2005). Adjoining phellogen and neighboring phelloderm layers, which stained metachromatically (violet) in native and wound periderm, could then be separately identified based on their location within the file of periderm cells and their morphology. Identification of these cells in native and wound periderm is essential in the interpretation of immunolocalization data and other biological investigations that involve wound healing and periderm maturation. Results from histological and immunolabeling studies of immature and mature native periderm indicated that pectin and extensin depositions are associated with the processes involved in the thickening and related strengthening of phellogen walls upon meristematic inactivation of the phellogen and development of resistance to skinning injury (Sabba and Lulai, 2004). Histological and immunolocalization data indicated that native periderm maturation and associated development of resistance to skinning injury are accompanied by an increase in unesterified pectin in the walls of phellogen cells (Sabba and Lulai, 2002, 2004, 2005). Unesterified pectin can impart rigidity to the cell walls through calcium bridges. Similar processes have been shown to occur in the cambium of aspen in association with cessation of meristematic activity and the onset of dormancy and thickening of cambial walls (Micheli et al., 2000). Pectins are methyl-esterified before they are transported into the wall, and as such, immature cells tend to be characterized by highly esterified pectins that prevent calcium pectate cross-linking and associated strengthening of the cell wall. Older, fully differentiated cells have more rigid walls that are characterized by less esterification and more calcium pectate cross-linking (Goldberg et al., 1989). Interestingly, pre- and postharvest skin-set measurements of maturing tubers in some instances actually show a decrease and later an increase in the resistance to skinning injury (Lulai, 2002; Rolf Peters, personal communication, EAPR, 2005). These mechanical measurements indicated that phellogen cell wall strength may at times decrease prior to cell wall thickening and further skin-set development. The biochemical nature of these anomalies is not known, but they are likely related to reversible compositional changes in the thin cell walls of the phellogen as the tuber undergoes some form of stress imparted by pre- and/or postharvest practices or conditions. Immunocytological analysis showed that some changes occurred in the localization of pectin and extensin epitopes in periderm cell walls during native periderm maturation (Sabba and Lulai, 2005). The antibodies JIM5 and JIM7 recognize a range of homogalacturonan (HG) epitopes (50% and less esterified, and up to 90% esterified, respectively); LM5 and LM6 recognize rhamnogalacturonan (RG)-I epitopes (-galactan tetrasaccharide and -l-arabinan pentasaccharide, respectively); and LM1 recognizes an extensin epitope. The walls of the three types of potato periderm cells labeled differentially for these pectin and extensin epitopes. While the phelloderm labeled equally well for all the epitopes tested, most of the phellem only labeled abundantly for the HG epitope recognized by JIM7 and was lacking in the HG epitope recognized by JIM5, as well as the RG-I and extensin epitopes. Most significantly, labeling of the phellogen layer varied between immature and mature periderm. Cell walls of meristematically active phellogen were lacking in (1,4)--galactan and extensin epitopes, as well as those for HG. Upon maturation of the periderm and development of resistance to excoriation, labeling for all

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:479

Trimsize:165×240 MM

479

Skin-set, wound healing, and related defects

these epitopes increased dramatically in phellogen cell walls. These results imply that an increase in the presence of HG, RG-I, and extensin polymers in these walls coincide with meristematic inactivation and periderm maturation. HG would be expected to form calcium pectate and extensin would be expected to become cross-linked in phellogen walls to provide increased strength and reinforcement once meristematic activity is terminated (Sabba and Lulai, 2005). The (1,4)--galactan epitope recognized by LM5 in phellogen walls at the end of periderm maturation may also be involved in the strengthening of these walls. By contrast, the (1,5)--l-arabinan epitope recognized by LM6 is present in phellogen walls before and after periderm maturation, and is apparently not specifically associated with periderm maturation or a discernable role in the thickening of phellogen walls upon meristematic inactivation. The changes in the epitopes associated with these cell wall polymers provide special insight into the maturational processes that occur during periderm maturation and skin-set development. Collectively, these immunolocalization responses for the targeted cell wall polymers indicate pectin deposition and de-esterification, and extensin depositions are associated with phellogen cell wall strengthening upon inactivation of the phellogen layer as a lateral meristem and maturation of the periderm in potato tuber. Interestingly, both histological and immunolabeling analyses, distinctly different chemistries, indicated differences between phellogen cell walls of wound versus native periderm. Results from both immunolocalization and histochemical techniques indicated that in wound periderm there is no increase in phellogen cell wall pectin upon wound periderm maturation (Sabba and Lulai, 2004). The reason for these apparent differences remains to be determined. Although various studies have been conducted to characterize tuber parenchyma cell walls (Li and Showalter, 1996; Bush and McCann, 1999; Oomen et al., 2002; Oomen, 2003; Obro et al., 2004), information on the physiology of periderm maturation remains sparse. Determination of the biochemical nature and regulatory mechanisms for strengthening of radial phellogen cell walls is important in the development of technologies to hasten skin-set and reduce associated market quality problems. The mechanisms regulating development of resistance to excoriation and biological markers for progress have not been determined. In conjunction with these knowledge gaps, the physiological and biological basis for slow/hindered skin-set development is not known, nor are the mechanisms known for the unpredictable reversal of skin-set development. Much more needs to be learned about the biology of potato periderm maturation so that effective technologies may be created to enhance skin-set development.

22.3 WOUND HEALING 22.3.1 The process of tuber wound healing Tubers that are skinned, nicked, or bruised during harvest or cut for seed lack the robust protection provided by the suberized layer of the native periderm, i.e. native phellem. Rapid wound healing is essential to avoid infection, desiccation/shrinkage, and defect

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

480

28-4-2007

9:37 a.m.

Page:480

Trimsize:165×240 MM

Lulai

development after injury. Tuber wound responses and wound healing involve many biological processes, perhaps the most important of which is wound-induced suberization (suberin biosynthesis). Regardless of the type of cells and associated polymeric structures covering the plant surface, wounding induces suberization to protect the damaged area (Dean and Kolattukudy, 1976). Wound-induced suberization of tuber tissue involves two stages during which two types of cells are suberized. The first stage involves formation of a ‘closing layer’ whereby the walls of existing cells at the wound site suberize, i.e. accumulate suberin biopolymers; ‘closing layer formation’ is also often referred to as ‘primary suberization’. Following the formation of the closing layer, the second stage involves development of a ‘wound periderm’ whereby files of new cells are formed and suberized below the closing layer; ‘wound periderm development’ is also often referred to as ‘secondary suberization’ (Section 22.3.6). A fully suberized tuber wound is resistant to infection by both bacteria and fungi (Lulai and Corsini, 1998). Suberin is a complex of suberin poly(phenolics) (SPP) and suberin poly(aliphatics) (SPA) that are cross-linked by glycerol, embedded with soluble waxes, and laminated to the interior side of the plant cell wall (Bernards, 2002). The terms wound healing and suberization are often loosely used interchangeably. Other forms of resistance are required to provide some degree of protection against infection and rot until suberization of the wound has been completed. These resistance processes and associated mechanisms are not fully understood, are generally temporal, and include various hypersensitive responses, the oxidative bursts, phytoalexin production, pathogenesis-related proteins, and other mechanisms (Lyon, 1989; Beckman, 2000; Pérombelon, 2002). Some resistance responses at the wound site appear to be related to the appearance of phenolic compounds (Lyon, 1989; Nolte et al., 1993; Beckman, 2000), some of which may be precursors of SPP (Bostock and Stermer, 1989). This observation is consistent with the correlative relationship found between strong temporal resistance and the rapidity of suberization across diverse genotypes (Lulai and Corsini, 1998, unpublished results). Importantly, a fully suberized surface provides the final, most durable, and broad ranging barrier for protectin against pests, desiccation, and infection (Lulai and Corsini, 1998). Rapid suberization is essential for protecting tubers that are injured during growth, during harvest and handling, and upon seed cutting. Suberization is also responsible for closure of lenticels as orifices for gas exchange and as a portal of entry for pests (Wigginton, 1973; Banks and Kays, 1988; Scott et al., 1996; Tyner et al., 1997) (Fig. 22.2). The soluble waxes embedded in the suberin matrix control water loss and prevent desiccation (Soliday et al., 1979; Lulai and Orr, 1994; Schreiber et al., 2005a). For the sake of conciseness, the discussion in section 22.3.2 will be primarily directed to potato tuber suberin and suberization. 22.3.2 Induction of suberization Upon cellular damage incurred during wounding or stress, a wide range of signals is perceived, which induces various protective or wound-related responses including suberization (de Bruxelles and Roberts, 2001; Kolattukudy, 2001; Leon et al., 2001; Schilmiller and Howe, 2005; Wasternack et al., 2006). A four-fold increase in translational activity is induced within 1 h of wounding, and a range of transcripts appears including those involved in primary suberization (Vayda and Morelli, 1994). Despite their importance,

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:481

Trimsize:165×240 MM

481

Skin-set, wound healing, and related defects

the signal(s) that induce suberization are enigmas that require further study. Several mitosis-related signals were hypothesized to be involved (Rosenstock and Kahl, 1978). Although induction of cell division is an essential part of wound periderm formation, it does not address the critical induction of suberin accumulation on the existing cell walls in the closing layer, i.e. during primary suberization resulting from a wound or from other stresses. Suberization of existing tuber parenchyma cells, i.e. similar to closing layer formation/primary suberization, has been demonstrated without mechanical wounding (Lulai, 2005; Lulai et al., 2006). The hollow heart disorder is a different type of example; it may be described as an internal growth-related wound with no cells exposed to the external environment; yet, cells neighboring the hollow heart area are suberized and are compositionally and ultrastructurally similar to that of wound periderm (Dean and Kolattukudy, 1977; see Chapter 23, Sowokinos, this volume, for more information on hollow heart). The signals involved in wound-, pathogen-, and other stress-induced suberization have not been determined. Rosenstock and Kahl (1978) asked the fundamental question ‘what is the primary event after mechanical disturbance of a tissue?’ and whether a primary event triggers the sequence of reactions in a wound cell. They hypothesized that one of the first primary changes upon wounding is that of a change in osmotic potential of the cell at the wound site. Many wound-related or wound-induced signals have been studied (de Bruxelles and Roberts, 2001; Leon et al., 2001; Schilmiller and Howe, 2005). However, the signal(s) and related mechanism(s) that induce suberization in potato tuber have not been identified. 22.3.3 Regulation of suberization In association with induction, the regulation of suberization is of equal importance, yet poorly understood. The plant hormone ethylene has been shown to be involved with various kinds of plant stress including wound response (Abeles et al., 1992; Bleecker and Kende, 2000; Ciardi and Klee, 2001; de Bruxelles and Roberts, 2001). However, the involvement of ethylene in wound-induced suberization had not been determined until recently. Using various inhibitors of ethylene biosynthesis and action, Lulai and Suttle (2004) determined the involvement of ethylene in wound-induced suberization. Ethylene biosynthesis was found to be stimulated by tuber wounding. Separate analysis for accumulation of SPP and SPA on wound-induced cell walls showed that ethylene is not required for wound-induced suberization of the closing layer or subsequent suberization associated with wound periderm development. Soliday et al. (1978) employed a gravimetric technique, diffusive resistance, to measure water vapor loss in wound-healing tissue. Results from this technique reflected wax deposition as an indirect assessment of wound-healing responses and the effect of various hormone treatments. They found that the development of resistance to water vapor loss during wound healing was inhibited by indole-3-acetic acid (IAA) and cytokinin treatments, but stimulated by abscisic acid (ABA) treatments. They further found that ABA and other ‘suberization induction factors’ could be washed from the tuber tissue, thereby inhibiting suberization. ABA treatment of cultured potato cells also resulted in increased accumulation of suberin components, waxes, and enzymes involved in suberin biosynthesis including peroxidase, phenylalanine ammonia lyase

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

482

28-4-2007

9:37 a.m.

Page:482

Trimsize:165×240 MM

Lulai

(PAL), and -hydroxy-fatty acid dehydrogenase (Cottle and Kolattukudy, 1982b). Lulai and Orr (1995), using a sensitive porometric technique for direct electronic measurement of water vapor loss, found that ABA treatment hastened development of resistance to water vapor loss during the first day of wound healing but made little difference after that time point when SPP and later SPA began to accumulate. Schreiber et al. (2005a) conducted detailed analysis of wax deposition during periderm maturation and obtained results consistent with that of Lulai and Orr (1995). Schreiber et al. (2005a) also found that even though periderm permeability quickly approached a nearly constant value during periderm development and maturation, significant wax formation continued. These results suggest that regulation of wax accumulation was still active even though water vapor loss had been minimized. Collectively, the above results indicate that ABA is involved in the regulation of wax accumulation and that water vapor loss is controlled by wax accumulation during wound healing, but water vapor loss may not be quantitatively used for assessing suberization. Lulai and Suttle (2005) employed other technologies to determine the involvement of ABA in tuber woundinduced suberization. Liquid chromotagraphy-mass spectrometry (LC-MS) was used to determine changes in tuber ABA content during wound healing. A specific xenobiotic inhibitor of carotenoid (ABA precursor) biosynthesis was used to determine the effect of metabolically blocking ABA biosynthesis during wound healing. Specific cytological techniques were used to directly assess accumulation of SPP and SPA on suberizing cell walls of inhibitor-treated tuber tissue during wound healing. The ability to block ABA biosynthesis provided a non-correlative approach to help determine the role of ABA in the regulation of wound responses. Tuber wounding resulted in an increase in ABA concentration. Inhibition of ABA biosynthesis resulted in a diminution of tuber woundhealing responses including reduced PAL activity, a mild delay in SPP accumulation, a more noticeable delay in SPA accumulation on suberizing cell walls, and a significant hampering in the reduction of water vapor loss. These results clearly indicate that ABA has a role in wound-induced suberization but that other signaling factors could be involved. Jasmonic acid (JA) is included in the wide range of signaling/regulatory compounds induced upon wounding and is involved in a range of wound responses such as insect and disease resistance (Choi et al., 1994; Koda and Kikuta, 1994; Negrel et al., 1995; de Bruxelles and Roberts, 2001; Schilmiller and Howe, 2005; Wasternack et al., 2006). Yet, information on the involvement of JA in wound-induced suberization is sparse. Two hydroxycinnamoyl transferases involved in suberin biosynthesis were shown to be influenced by treatment with ABA; however, JA treatment did not significantly modify the time course or intensity of the induction of these enzymes during wound healing (Negrel et al., 1995). Wound-induced PAL activity and the associated accumulation of phenolics in purple-flesh tubers with varying anthocyanin concentrations did not show a response to methyl jasmonate treatment unless anthocyanin concentrations were low (Reyes and Cisneros-Zevallos, 2003). Although under current investigation, there is little other information available concerning the involvement or regulatory roles of JA in tuber wound healing and suberization. Many signaling compounds, in addition to those discussed in this section, are induced upon wounding, yet their involvement in the regulation of suberization has not been determined.

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:483

Trimsize:165×240 MM

483

Skin-set, wound healing, and related defects

22.3.4 Environmental effects on suberization After induction, the rate of suberization is influenced by genotype (Lulai and Corsini, 1998), type or severity of wound or bruise (Lulai and Orr, 1995; Thomson et al., 1995), physiological age (Thomson et al., 1995; Kumar and Knowles, 2003), a range of possible chemical treatments (Nolte et al., 1987; Gronwald, 1991; Oosterhaven et al., 1995), and environmental conditions (e.g. relative humidity, temperature, and aeration including oxygen/carbon dioxide concentrations) (Artschwager, 1927; Wigginton, 1974; Dean, 1989; Morris et al., 1989; Schaper et al., 1993). Relative humidity above 80%, preferably 90–95%, is needed to ensure that the cells at the wound site do not desiccate and die. Low relative humidity after wounding may lead to tissue dehydration and result in a layer of dead non-suberized cells over the wound. A layer of dead desiccated cells will not resist penetration by pathogens or prevent water vapor loss and should not be confused with suberized cells. Excessive humidity may result in a film of water over the wounds, thereby restricting oxygen supply and causing cell proliferation both of which inhibit suberization. If relative humidity and oxygen supply are favorable, the most important environmental factor affecting suberization is temperature. Suberization increases from 2.5 to 25  C. The rate of suberization increases approximately three-fold between 5 and 10  C, increases another three-fold between 10 and 20  C, and is greatest near 25  C (Artschwager, 1927; Wigginton, 1974; Dean, 1989; Morris et al., 1989). Suberization is prevented at temperatures of 35  C and above. Excessively warm storage temperatures directly after harvest can hamper suberization and result in infection and rapid decay. Ample oxygen (air is approximately 21% oxygen) is required for suberization. Low oxygen concentrations and carbon dioxide concentrations above ambient (air is approximately 0.03% carbon dioxide) inhibit suberization and wound periderm formation. Tuber wounding from harvest and seed-cutting operations can quickly result in a three- to fivefold increase in respiration, which within 24 h can increase another three- to five-fold (Laties, 1978). Wound respiration may be 25 times that of the intact tuber by 24 h after wounding. Fresh wound respiration is derived primarily from lipid. Within 24 h after wounding, carbohydrate becomes the respiratory substrate. Carbon dioxide concentrations increase while oxygen concentrations decrease within the potato storage; both conditions inhibit suberization. Proper ventilation of freshly harvested tubers or cut seed is essential in maintaining oxygen and carbon dioxide concentrations that promote suberization. Unless properly ventilated, the carbon dioxide concentration in a storage bin of freshly harvested skinned and bruised tubers can increase to well over 100 times that of ambient air (Schaper et al., 1993). 22.3.5 Characteristics of the biopolymers that form suberin The identification and description of suberin has been confusing in part because of its complex and poorly understood composition and ultrastructure, and its similarities to the plant polymers lignin and cutin (Kolattukudy, 1980; Cottle and Kolattukudy, 1982a; Kolattukudy, 2001; Bernards, 2002). Lignin is composed of a dense polymeric matrix of aromatic/phenolic monomers, and cutin is composed of hydroxy and epoxy aliphatic monomers (Kolattukudy, 1980; Lewis et al., 1999). The biopolymer suberin is composed

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

484

28-4-2007

9:37 a.m.

Page:484

Trimsize:165×240 MM

Lulai

of both phenolic/aromatic and aliphatic/hydrophobic polymeric domains (Kolattukudy, 1980, 2001), which are spatially separate (Lulai and Morgan, 1992) and cross-linked by glycerol (Moire et al., 1999; Graca and Pereira, 2000; Bernards, 2002). Because of the polymeric and compositional nature of the phenolic/aromatic and aliphatic/hydrophobic domains, they are now referred to as the suberin poly(phenolic) domain (SPPD) and suberin poly (aliphatic) domain (SPAD) (Bernards, 2002). The SPPD had been referred to as lignin-like (Kolattukudy, 1980; Cottle and Kolattukudy, 1982a). The complex biosynthesis and macromolecular assembly of the SPP biopolymer on the cell wall may have similarities to that of lignin and involve dirigent proteins and sites (Lewis et al., 1999; Davin and Lewis, 2000). However, the SPPD lacks the dense matrix of guaiacyl and syringyl phenylpropane units characteristic of lignin (Lapierre et al., 1996). Detailed labeling and spectroscopic studies clearly showed that the phenolic domain of suberin has a very low monolignol content, about one-tenth that of wood or straw, and instead is largely composed of hydroxycinnamic acids and associated derivatives (Bernards et al., 1995; Lapierre et al., 1996; Bernards, 2002). Also, confusion arises because lignin and the SPPD react similarly to histological analyses, such as phloroglucinol and other treatments, and both autofluoresce under commonly used epifluorescent microscopy. Results of Lulai (2005) may be interpreted to indicate that, in some cases, pathogen-induced accumulation of SPP on the cell wall may have been mistaken for lignification in other published reports. SPAs, like cutin, are composed of fatty acid monomers. However, unlike cutin, suberin aliphatics are made up of -hydroxy and  -dicarboxylic acids, which can be of equal or longer chain length than cuticular aliphatics and lack the characteristic cuticular epoxy and internal chain hydroxy groups (Kolattukudy and Dean, 1974; Dean and Kolattukudy, 1977; Kolattukudy, 1980). Bernards (2002) indicated a possible role for epoxy fatty acids in hydroxylation reactions involved in SPA biosynthesis. Although SPP and SPA accumulations are tightly coupled, if conditions are not favorable, SPP accumulation on the cell wall may terminate before completion and SPA accumulation will not occur as has been demonstrated in tuber pink-eye tissues (Lulai et al., 2006). There have been no reports of wound-induced SPA accumulation on the primary cell wall without first accumulating SPP about the entire cell wall nor have there been demonstrations of SPA accumulation on a polyphenolic matrix that had been fully characterized as lignin. These features help define the two biopolymers, SPP and SPA, that form the phenolic/hydrophilic and aliphatic/hydrophobic domains of suberin. Molecular genetic similarities and differences between cutin and suberin continue to be uncovered (Yephremov and Schreiber, 2005). 22.3.6 Suberization: closing layer and wound periderm formation The hierarchy for wound-induced accumulation of SPP and SPA on the suberizing cell walls also differs. The process of closing layer formation, i.e. primary suberization, is cell specific, occurring one cell layer at a time beginning with the first layer of viable cells neighboring the wound site. SPP accumulates on the cell walls in a segmented fashion, first on the outer tangential walls of the cells in the first layer followed by accumulation on the radial cell walls and then the inner tangential cell walls (Lulai and Corsini, 1998). After SPP accumulation on the cell wall is complete, SPA begins to accumulate

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:485

Trimsize:165×240 MM

485

Skin-set, wound healing, and related defects

and does so relatively uniformly about the inner perimeter of the cell. SPA does not accumulate in a pronounced segmented fashion like that of SPP. As SPA accumulates, presumably in conjunction with glycerol, over the SPP in the cells of the first layer, SPP accumulates in a segmented fashion on the walls of the cells in the second cell layer. SPA accumulation follows completion of SPP deposition on the second layer and so on. Generally, under favorable conditions, primary suberization will continue until there are about two or three layers of suberized parenchyma cells forming the closing layer. At this time, the wound phellogen will have sufficiently developed and will form additional suberized cells, i.e. wound phellem, directly beneath the closing layer. In addition, a layer of wound phelloderm cells is formed beneath the wound phellogen (Lulai and Corsini, 1998; Lulai, 2001a). These three cell types, i.e. phellem, phellogen, and phelloderm, comprise the wound periderm (Reeve et al., 1969), but it is the suberized cells of the closing layer and wound phellem that provide the durable protective barrier (Lulai and Corsini, 1998). It is important to note that primary suberization, i.e. formation of the closing layer, involves suberization of existing cell walls. Therefore, these suberized cells do not derive from progenitor cells in meristems, i.e. not from a distinct cambium/meristematic layer. Arguably, these suberized cells may play a more important role than those of the wound phellem because they provide the first suberized form of durable and broad-ranging resistance to infection over the wound (Lulai and Corsini, 1998). Interestingly, the biological differences involved in suberization of closing layer cells and that of wound phellem cells derived from the wound periderm, i.e. secondary suberization, are not known. Development of the closing layer provides a unique perspective compared with that of wound phellem development because SPP accumulation must occur in conjunction with the existing primary cell wall. Whether there are fine biosynthetic and structural differences between suberized cell walls of the closing layer and wound phellem is not certain. 22.3.7 Suberin biosynthesis and structure Suberization requires biosynthesis of phenolic, aliphatic, and glycerol monomers and assembly of these monomers into the two separate biopolymeric domains, the SPPD and the SPAD. Together, the SPPD and the SPAD comprise the suberin barrier. The composition, biosynthesis, and associated macromolecular assembly of these phenolic and aliphatic biopolymers into their respective domains on the cell wall continue to be described through hypothetical models and schemes because the pathways and mechanisms of assembly are only partially elucidated. Suberin biosynthesis is an important area of research that is complicated by the distinctly different biochemical pathways required for SPP and SPA syntheses, assembly, glycerol cross-linking, and placement on/in the cell wall. Detailed issues associated with SPP and SPA biosyntheses have been reviewed and will not be repeated in this chapter (Bernards and Lewis, 1998; Bernards, 2002). The SPPD and SPAD have been shown to be spatially separate and their time courses for wound-induced accumulation on the cell wall initiated at distinctly different times (Lulai and Morgan, 1992; Lulai and Corsini, 1998). SPP biosynthesis occurs prior to that of SPA (Kolattukudy, 1980). Furthermore, nuclear magnetic resonance (NMR) data (Stark and Garbow, 1992) and ultrastructural analysis also show that the SPAD is distinct

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

486

28-4-2007

9:37 a.m.

Page:486

Trimsize:165×240 MM

Lulai

Fig. 22.4. Transmission electron micrograph of a suberized cell wall neighboring an active phellogen layer. The upper primary cell wall is suberized and includes suberin poly(phenolic) material and darkly stained suberin poly(aliphatic) material. The suberin poly(aliphatics) domain (SPAD) is spatially separate from the primary cell wall. The delamination of the SPAD from the primary cell wall (arrow) illustrates the spatial separation of the two suberin polymeric domains. The thin and fragile radial cell wall ∗  of the active phellogen has fractured. (Courtesy: E.C. Lulai and T.P. Freeman).

and spatially separate from the SPPD and primary cell wall (Fig. 22.4). Although spatial features and architecture of the SPPD accumulated on/into the primary cell wall are not clear, spectroscopic data indicate that the aromatic carbons of the SPPD and glycosidic carbons of cell wall polysaccharides are in proximity, perhaps 0.05 nm (Yan and Stark, 2000). Wound signals induce enzymes of the phenylpropanoid pathway to provide the phenolic precursors for the biosynthesis of SPP (Kolattukudy, 1980) (Fig. 22.5A). The first enzyme of this pathway, PAL, which produces cinnamic acid for further biosynthesis, is essential and its inhibition prevents formation of the SPP barrier (Cottle and Kolattukudy, 1982a; Hammerschmidt, 1984; Bostock and Stermer, 1989; Bernards and Lewis, 1998). Hydroxycinnamic acid derivatives, primarily ferulic acid and N -feruloyltyramine in wound-induced suberin, are channeled to comprise a major part of the SPPD (Bernards and Lewis, 1992; Bernards et al., 1995, 2000; Negrel et al., 1996). Ferulate esters of long-chain fatty alcohols, hypothesized to act as bioplasticizers, are found in the wound periderm (Bernards and Lewis, 1992; Bernards, 2002); however, little has been indicated about the presence of these alkyl ferulates in native periderm. Information on the composition of the SPPD has largely been obtained by analysis of end products from harsh chemical degradation treatments, which inherently alter the structure of the final monomers to be identified. NMR analysis of wound periderm confirmed that sinapyl and guaiacyl structures are part of the SPPD (Yan and Stark, 2000), and LC-MS techniques have been developed, which quantitatively show that there are differences in the biosynthetic flux of phenylpropanoids, i.e. formation and conversion of specific monomers into SPP during tuber wound healing (Matsuda et al., 2003).

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Starch

VLCFA Glycerol-P

Gutter:4.6 pc

Ferulate fatty esters

Poly(Aliphatic) domain, Domain, waxes Waxes

9,10-epoxyα,ω-dioic acids (predominantly 18:0)

9:37 a.m. Page:487

Text Width:30 pc

Trimsize:165×240 MM

Depth:45 Lines

Fig. 22.5A. (Caption on page 489)

28-4-2007

Font Size:10/12

O2 NADP+ SAM SAHC NADPH p-coumaroylCoASH 71 FeruloylCaffeoyl- 72 78? Poly(Phenolic) tyramine Phenylalanine Tyramine tyramine tyramine domain Benzoates * Acetyl 78? CoASH CoASH CoA NH4+ 58 75 CoASH 70 CoASH 70 O2 NADH †Hydroxycinnoyl74 Cinnamate NADPH O2 NAD+ 73 CoA’s O H O + 2 2 NADPH NADP+ H2O 59 NADP 78 NADPH NADH SAM SAHC SAM SAHC 61 62 p - Coumaric Caffeic Ferulic 63 62? 5-OH-ferulic Sinapic H2O2 acid acid ATP acid acid acid ATP ATP 77 CoASH ATP CoASH CoASH .CoASH 60 O 2 60 60 ADP, Pi 60 O2 ADP, Pi O2 ADP, Pi ADP, Pi NADP+ + NADPH SAM SAHC 76 NADPH NADP Phenylpropanoid SAM SAHC NADP+ 64 metabolism p-Coumaroyl 66 Caffeoyl- 65 Feruloyl5-OH-feruloyl- 65? SinapoylNADPH O2 CoA CoA CoA † CoA CoA† NADPH Hydrogen peroxide 67 67 67 generation SAM SAHC + + O NADP+ O2 NADP 2 NADP SAM SAHC NADPH p-Coumaryl- NADPH68 Sinap5-OH-conifer- 62 Conifer62 68 CaffeylAldehyde aldehyde aldehyde aldehyde aldehyde 69 69 69 Monolignols

Job code: PBBA

Shikimic acid pathway

9,10-diOH-α,ω-dioic acids (predominantly 18:0)

Ch22-N51018

Margins:Top:5 pc

Carbohydrate metabolism

α,ω-dioicacids

Elsevier AMS

Basal Fonts:Times

1oAlcohols Sucrose

35

37

DHAP

20:0 22:0 24:0 26:0 28:0 30:0 32:0

NADPH

Gutter:4.6 pc

29 NADP+ β-HydroxyacylCoA

Chain elongation

32

?

NADPH

18:1-ω-OH-FA

NADP+

NADP+ ω-oxo-FA

O2 NADPH

34

1° Alcohols

NADP+

α,ω-dioic acids

O2 NADPH

39 NADP+

9,10-diOH-α,ω-dioic acids (predominantly 18:0)

9,10-epoxy-ω-OH-FA

NADPH

41 NADP+ 9,10-epoxy-ω-oxo-FA O2 NADPH

42 NADP+

9,10-epoxy-α,ω-dioic acids (predominantly 18:0)

28-4-2007

44 NAD+

Puly(Aliphatic) domain, waxes

VLCFA Glycerol-P

Page:488

Text Width:30 pc

9:37 a.m.

Font Size:10/12

NAD(P)H

ω-oxo-9,10-diOH-FA

33

NAD+ NAD(P)+

38

NADPH

NADH

31 NADH

40

ω-OH-FA

43

trans-Δ2enoyl-CoA

NADP+

9,10,ω-triOH-FA

NADP+

30 H2O

NADP+

36

9,ω-diOH-FA O2 NADPH

37

Lipid modification

O2 NADPH

Job code: PBBA

β-Ketoacyl-CoA

O2 NADPH

NADP+

18:1

28

9,10-diOH-FA

Ch22-N51018

Margins:Top:5 pc

co2

36

9-OH-FA

27 NADH

Carbohydrate metabolism

NADP+

O2 NADPH

NAD+

Malonyl-CoA

O2 NADPH

NADP+

Elsevier AMS

Basal Fonts:Times

16:0 18:0

Fatty acid biosynthesis

Pyruvate

O2 NADPH

Ferulatefatty esters

45 CoASH

Fig. 22.5B. (Caption on page 489)

†HydroxycinnoylCoA’s

Trimsize:165×240 MM

Depth:45 Lines

Phenylpropanoid metabolism

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:489

Trimsize:165×240 MM

489

Skin-set, wound healing, and related defects

Fig. 22.5. Scheme for the biosynthesis of the suberin poly(phenolic) (SPPD) and poly(aliphatic) (SPAD) domains, respectively. Figure 5A and B is part of a total scheme developed by Bernards (2002) and represents a composite of knowledge derived from a number of different plant species, but principally derived from potato and maize. Subcellular localization of the processes is not depicted. The enzymes involved are identified by number as in the original scheme; a few enzymes within the scheme are hypothetical ∗ : 1–26, 46–57, and 84–85 involve carbohydrate metabolism, fatty acid biosynthesis, shikimic acid pathway, and ammonia recovery and may be found in Bernards (2002); 27, stearoyl-ACP 9 -desaturase; 28, -ketoacyl-CoA synthase III; 29, -ketoacylCoA reductase; 30, -hydroxyacyl-CoA dehydratase; 31, enoyl-CoA reductase; 32, fatty acyl--hydroxylase; 33, -hydroxyacid dehydrogenase; 34, -oxoacid dehydrogenase; 35, fatty acyl-9-hydroxylase∗ ; 36, 9( hydroxy fatty acyl-10-hydroxylase∗ ; 37, 9(10)-hydroxy fatty acyl--hydroxylase∗ ; 38, 9,10,-trihydroxyacid dehydrogenase∗ ; 39, 9,10-dihydroxy--oxoacid dehydrogenase∗ ; 40, -hydroxyacid-9,10-epoxide synthase∗ ; 41, 9,10-epoxy--hydroxyacid dehydrogenase∗ ; 42, 9,10-epoxy--oxoacid dehydrogenase∗ ; 43, reductases∗ ; 44, glycerol-3-phosphate dehydrogenase; 45, hydroxycinnamoyl-CoA; 1-alkanol hydroxycinnamoyl transferase; 58, phenylalanine ammonia-lyase; 59, cinnamate-4-hydroxylase; 60, 4-coumaroyl-CoA ligase; 61, p-coumaric acid 3-hydroxylase; 62, caffeic acid 3-Î-methyltransferase; 63, ferulic acid 5-hydroxylase; 64, p-coumaroylCoA-3-hydroxylase; 65, caffeoyl-CoA-3-O-methyltransferase; 66, hydroxycinnamoyl-CoA-5-hydroxylase∗ ; 67, cinnamoyl-CoA oxidoreductase; 68, hydroxycinnamaldehyde hydroxylase∗ ; 69, coniferyl alcohol dehydrogenase; 70, hydroxycinnamoyl-CoA : tyramine hydroxycinnamoyltransferase; 71, p-coumaroyltyramine-3hydroxylase∗ ; 72, caffeoyltyramine-O-methyltransferase∗ ; 73, hydroxycinnamoyl-CoA-7-hydroxylase∗ ; 74, (7hydroxy)-hydroxycinnamoyl-CoA reductase∗ ; thiolase∗ ; 76, NAD(P)H-dependent oxidase; 77, superoxide dismutase or spontaneous; 78, peroxidase. Reactions denoted by solid lines are known while those denoted by broken lines are hypothetical or assumed. Shaded boxes denote known precursors incorporated into the suberin poly(aliphatic) domain (SPAD) and suberin poly(phenolic) domain (SPPD). (Courtesy: M.A. Bernards and the Can. J. Bot., 2002, revised).

Peroxidases and H2 O2 formation are induced upon wounding and are required for suberization, possibly in a peroxidase-mediated coupling process cross-linking phenolics within the SPPD and attachment to the cell wall (Espelie et al., 1986; Bernards et al., 1999, 2004; Bernards and Razem, 2001; Razem and Bernards, 2003). A model has been constructed whereby phenolics are synthesized in the cytoplasm, transported to the cell wall (by an unknown mechanism), and polymerized by a wall-associated peroxidase into the SPPD (Bernards and Razem, 2001). The polymeric attachments to the cell wall are at discrete sites (Stark and Garbow, 1992). Specific peroxidases have been determined to preferentially cross-link suberin phenolics but not monolignols (Bernards et al., 1999). In vitro studies have shown that there are several specific combinations of crosscoupling products formed by peroxidase-catalyzed polymerization of hydroxycinnamic acids (Arrieta-Baez and Stark, 2006). The identity of these coupling products provides important insight into the molecular architecture of suberin and the structural requisites that impart durable protective properties to the barrier. The construction of the SPPD is particularly perplexing when the ambiguities of SPP biosynthesis, phenolic transport, and polymeric assembly are combined with polymeric attachment at discrete sites on the cell wall in conjunction with the segmented hierarchy for SPP accumulation on the cell wall. Biosynthesis of the SPAD involves fatty acid synthesis and the creation of aliphatic monomers from palmitic (16:0), stearic (18:0), and oleic (18:1) acids (Fig. 22.5B). The suberin aliphatics are composed primarily of -hydroxy acids and the corresponding dicarboxylic acids and smaller amounts of long > C20  chain acids and corresponding alcohols (Kolattukudy, 1978). Fatty acid monomers are substrate to a chain elongation system (Kolattukudy, 1978), which now appears to be microsomal and

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Basal Fonts:Times

OH OCH3

Ch22-N51018

Margins:Top:5 pc

P O H2C

O O

O

Gutter:4.6 pc

CH2OH HN C O O

O OCH3

O

O

O

OCH3 O

C

O

O

HO O

O

HOCH2

O

O OCH3 O

C

O

O O

S

O O HO

O

O

HO

O

O

O

OH

O O HOH2C

C OCH3 HOH2C

O C

O

O C

O O

O O

OCH3

P

O O

HO O

O

O

O

O

O

O OCH3

H3CO

OCH3

O O

OCH3

OH ( S)

S

C

HO ( S)

OH

S

Page:490

Text Width:30 pc

O

O O

OCH3

S

O

O (H3CO)

S

9:37 a.m.

Font Size:10/12

O O OCH3

S

28-4-2007

CH2OH (COO )

O O

O

HO ( S)

OCH3

O C O

O

HN C O

S

O O

O

3

O

O

OCH3 OH

O

OCH3 OCH

(H3CO)

O

O

O

C

S

O C

O

O

O

S

O

Job code: PBBA

HO

HO

S

O O O

O OCH3

CH2OH

O

O O

P

Primary cell wall

Fig. 22.6. (Caption on page 491)

Suberin poly(aliphatic) domain

Trimsize:165×240 MM

Depth:45 Lines

Suberin poly(phenolic) domain

Suberin lamellae

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:491

Trimsize:165×240 MM

491

Skin-set, wound healing, and related defects

Fig. 22.6. Tentative model for the structure of potato suberin. The suberin poly(phenolic) domain (SPPD) is covalently attached to carbohydrate units of the cell wall. Glycerol units are used to cross-link suberin poly(aliphatic) monomers and account for the lamellae found within the suberin poly(aliphatic) domain SPAD. The SPAD is illustrated using two lamellae and is covalently attached to the SPPD through easily hydrolyzed ester linkages. Very long-chain fatty acids and waxes are not illustrated within this diagram but are thought to be embedded in the aliphatic domain. Ferulates esterified to long-chain fatty alcohols are integrated into the aliphatic domain and are thought to act as plasticizers. The constituents are connected to C, carbohydrate; P, phenolic; S, suberin (phenolic or aliphatic). (Courtesy: M.A. Bernards and the Can. J. Bot., 2002, revised).

elongate through malonyl-CoA as substrate (Schreiber et al., 2000, 2005b). The fatty acids are presumed to undergo -hydroxylation, similar to cutin, and then further conversion to dicarboxylic acids in reactions that may have some uniqueness to SPA biosynthesis (Kolattukudy, 1980). The -hydroxy fatty acids are converted to -oxo fatty acids through an NADP-dependent oxidoreductase (i.e. -hydroxy fatty acid dehydrogenase) that is wound inducible. A constitutive -oxoacid dehydrogenase catalyzes the conversion of the -oxoacid to the dicarboxylic acid. These enzymes have been separated and the dehydrogenase enzyme purified and characterized. The catalytic properties of the -hydroxy fatty acid dehydrogenase appear to be distinct from alcohol dehydrogenase (Agrawal and Kolattukudy, 1977, 1978a,b; Kolattukudy, 2001). Kinetic studies involving chain-length substrate specificity suggest that the pocket of the active site of the -fatty acid dehydrogenase enzyme limits binding of longer chain >C18  substrate, which may explain the presence of dicarboxylic acids in the range of C18 and -hydroxy fatty acids of longer chain length (Kolattukudy, 2001). Tracking the flux of aliphatic monomers of the SPAD during wound healing further demonstrated that the newly formed acids had two possible metabolic fates: (1) desaturation and oxidation to form suberin C18 -hydroxy and dioic acids, and (2) elongation to form very long-chain fatty acids C20 –C28  associated with reduction to 1-alkanols, decarboxylation to n-alkanes, and minor amounts of hydroxylation (Yang and Bernards, 2006). In some plant tissues, fatty 9,10-epoxides may be hydroxylated during suberization, thereby creating a hypothetical role for these compounds in SPA biosynthesis (Bernards, 2002). Glycerol is brought into the SPA biosynthetic scheme and incorporated into the polymer at about the same rate as that of the major/diagnostic suberin monomers, -alkanedoic acids (dicarboxylic acids), thus suggesting cross-linking of aliphatic and aromatic domains (Moire et al., 1999). The cross-linking hypothesis is supported by ultrastructural data, which show that the alternating electron-transparent regions found within the electron-dense areas of the suberin aliphatic domain have dimensions similar to that created by a C-22 hydrocarbon located between two glycerol molecules (Schmutz et al., 1996; Moire et al., 1999). The lamellae dimensions match with the tentative model for the structure of potato suberin (Fig. 22.6). The source for glycerol in the suberin model is hypothesized to be dihydroxyacetone phosphate (Fig. 22.5B) (Bernards, 2002). However, the complete pathway for these processes is not fully known including how these specialized fatty acids and glycerol are assembled to form the SPAD. Nor is it clear how these monomers or oligomers are transported on to the cell wall after some apparent resident time elsewhere while construction of the SPPD is being completed. As with the SPPD, the construction of the SPAD becomes more complicated and perplexing when attempting to integrate the biosynthesis

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

492

28-4-2007

9:37 a.m.

Page:492

Trimsize:165×240 MM

Lulai

and coordinated macromolecular assembly of the SPA biopolymer as it accumulates in a non-segmented fashion over the SPPD on the cell wall. 22.3.8 Suberization and resistance to infection Suberization and loosely defined wound periderm development have been suggested to play various roles in resistance to tuber infection (Nnodu et al., 1982; O’Brien and Leach, 1983; Nolte et al., 1987; Bostock and Stermer, 1989; Lyon, 1989; Vaughn and Lulai, 1991; Pérombelon, 2002). However, resistance to bacterial and fungal infections was often poorly related to the various definitions for suberin and wound periderm; consequently, the roles of suberin in resisting bacterial and fungal infections had not been clearly resolved. Lulai and Corsini (1998) determined that the hierarchy for SPP and SPA accumulations during wound healing is responsible for the differential development of resistance to bacterial and then fungal penetration. Total resistance to bacterial infection occurred after the SPP accumulation was complete along the contiguous outer tangential walls of the first layer of cells (2–3 days) in the closing layer. However, the SPP accumulation offered no protection against fungal infection even after SPP accumulation advanced to the adjoining radial and inner tangential cell walls. Fungi could breach the SPP barrier and thereby provide an entry for bacterial infections such as Erwinia carotovora ssp. carotovora. Lulai and Corsini (1998) indicated that this process is responsible for the confusing development of resistance to bacteria, followed by fungal infection and subsequent susceptibility to bacterial infection. Resistance to infection by the fungal agent that causes dry rot, Fusarium sambucinum, began to develop after accumulation of SPA was initiated. The final form of full resistance to fungal infection occurred upon completion of SPA accumulation on the first layer of cells. The specificity of the SPA domain as a barrier to fungi has been further demonstrated at the cellular level by blockage of advancement of fungal hyphae within pink-eye-afflicted tubers (Lulai et al., 2006). The knowledge gaps and ambiguities associated with wound healing and woundinduced suberization are of great importance because of the critical roles that rapid suberization plays in minimizing food-quality deterioration, shrinkage, defects, and perhaps the most easily and widely recognized problem of infection by a wide range of bacterial and fungal pathogens.

22.4 RELATED DEFECTS 22.4.1 Wound-related tuber defects Various forms of wound-related defects, i.e. those caused by different types of bruising and growth anomalies, are responsible for a range of tuber quality problems and losses. Reviews indicate that bruising and associated defects may range from 9 to 40% and that they are influenced by genetics, cultural conditions/practices, and handling procedures (Storey and Davies, 1992; Brook, 1996; McGarry et al., 1996). The types of defects and associated disorders that may fit this general category are variable. Many of these defects and disorders are poorly described because of the dearth of published information and

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:493

Trimsize:165×240 MM

493

Skin-set, wound healing, and related defects

the frequent use of untested hypotheses as fact. The discussion herein will primarily deal with the wound-related issues of these defects and will not delve deeply into other details. 22.4.2 Shatter bruising and tuber cracking Shatter bruising and cracking results from mechanical impact stresses incurred during potato harvest and handling operations (Thornton, 2001). The internal shattering or cracking of tuber tissue to the surface is a serious problem because of the impaired suberization inherent with this type of wound. This type of bruise damage results in a combination of problems including disease, severe defects created by discoloration, and the development of scars from the wound. The impairment of suberization in these cracks is inherent and is a significant cause for the exacerbation of the defect. The impaired suberization of these bruise wounds exemplifies the differences in wound healing created by irregular versus smooth wound surfaces, and altered oxygen and carbon dioxide concentrations at the wound site (Section 22.3.4). Wound healing in the microenvironment of a shatter bruise or tuber crack can be especially hindered because of the irregular wound surface and the confinement of the narrow wound crevasse compared with that of a smooth cut. The amount of cell damage and respiration increases proportionally with the larger wound surface area created by the irregular fracture of the shatter bruise. The carbon dioxide concentrations would be expected to increase and the oxygen concentrations decrease in the confined area formed by the wound crevice of a shatter bruise. Ventilation into the confined area of a tuber crack is difficult. Roughened or irregular wound surface areas severely hamper development of contiguous SPP accumulation in the formation of a closing layer and consequently also impair formation of a wound periderm. The irregular surface area and likelihood of a hypoxic environment predispose shatter bruises and cracks to hindered suberization and increased susceptibility to infection (Vayda et al., 1992; Lulai and Corsini, 1998; Lulai, 2001a,b). Higher tuber turgor appears to increase susceptibility to dynamic failure, i.e. shattering and cracking, whereas higher temperature decreases failure rates (Smittle et al., 1974; Bajema et al., 1998). Tuber size also influences failure properties; tubers weighing more than 340 g are significantly more likely to undergo dynamic failure in the form of shatter bruise/cracking whereas tubers of 170–340 g showed little difference in failure rates throughout their range (Baritelle and Hyde, 1999). Managing tuber turgor, especially on rain-fed land, and managing harvest temperature can create unworkable approaches to minimize shattering. Further improvement of mechanical harvesting procedures and biologically improving the rate of suberization and cell–cell adhesion are feasible approaches to develop technologies to reduce shatter bruise/cracking defects. For example, polygalacutronic acid of low ester content has been shown to play a role in cell–cell adhesion at ‘edge-of-face regions’ of tuber parenchyma cells (Parker et al., 2001). 22.4.3 Blackspot and pressure/crush bruising Blackspot is an internal discoloration of the tuber cortical parenchyma found beneath an impact or pressure bruise. A significant amount of research has been conducted on impact-induced blackspot bruise development, while there has been less research

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

494

28-4-2007

9:37 a.m.

Page:494

Trimsize:165×240 MM

Lulai

on pressure bruising and pressure bruise-induced blackspot development. In general, tubers of higher specific gravity have a lower bruise threshold, a lower bruise resistance, or both (Baritelle and Hyde, 2003). Depending on the genotype, less severe impact bruising and impact bruising sustained by tubers at warmer temperatures or by tubers with lower turgor tend not to result in tuber shattering or cracking (Smittle et al., 1974; Brook, 1996; McGarry et al., 1996; Bajema et al., 1998; Baritelle and Hyde, 1999, 2001). Instead, these impact bruises result in cellular damage or physiological disruption and the development of blue-black or gray-black discoloration in tissues beneath the tuber surface. These dark bruise discolorations are referred to as blackspot. Although other secondary mechanisms have been implicated, further evidence indicates that the discolorations are primarily a result of oxidation of tyrosine by polyphenol oxidase and subsequent reactions to form melanin, which in combination with cysteine influences coloration of the pigment (Dean et al., 1992, 1993; Stevens and Davelaar, 1996, 1997; Edgell et al., 1998; Stevens et al., 1998; Partington et al., 1999; Laerke et al., 2002). Antioxidants normally found within the tuber, such as ascorbic acid, may be involved in limiting the enzymatic formation of melanin (Delgado et al., 2001b; Pawelzik et al., 2005). Intracellular compartmentation is an important factor in determining susceptibility to blackspot development (Laerke et al., 2002). McGarry et al. (1996) pointed out that these wound-related discolorations involve compromising the integrity of intracellular membranes and bringing the polyphenol oxidase enzyme, located in the plastid membrane, i.e. amyloplast, into contact with phenolic substrates that are putatively in the vacuole. There is evidence that certain field applications of soluble calcium during plant growth will enhance tuber calcium concentrations and reduce blackspot bruising presumably through improved cell membrane stability and stronger cell walls (Karlsson et al., 2006). Although blackspot development is indicative of cellular disruption within the tuber, indications of wounding on the surface of the tuber are often difficult to detect. Blackspot detection is accomplished by peeling to expose the darkened areas within the cortical parenchyma. Surface detection of wounds, including less obvious bruised areas that involve damaged periderm, which may develop into blackspot or a surface blemish, may be detected in freshly harvested tubers by testing with catechol or other peroxidizable substrate that produces a detectable color difference. Freshly bruised tubers are submerged in a solution of catechol and then visually inspected for oxidative color change of the catechol on the tuber surface (Gould, 1989). Methods have been developed to assess the biochemical potential of potato tubers to discolor and darken in vitro. However, it appears that the conditions for in vitro analysis, possibly including oxygen concentration, are not always fully consistent with the biochemical potential of tubers to discolor and develop blackspot (Delgado et al., 2001a; Laerke et al., 2002). Although the mechanisms associated with the development of blackspot discoloration have been partially characterized, little is known about other wound responses that may or may not be induced by bruising including suberization of cells in the proximity of this type of internal wound. Pressure bruising or crushing occurs at the tuber–tuber contact points or tuber–storage structure contact points during storage and is caused by excessive pressures created by potato pile heights of 3.5–4.0 m or greater (Schaper and Yaeger, 1982). The constant pressure created by the weight of the pile on tubers located at lower depths in a potato storage bin may cause the tuber surfaces at the points of contact to become flattened or

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:495

Trimsize:165×240 MM

495

Skin-set, wound healing, and related defects

indented creating a surface blemish. Cellular damage in the cortical parenchyma under the pressure bruise area may result in the development of blackspot discolorations (Lulai et al., 1996). Although this periderm damage may not be visible to the eye, the pressure often creates a layer of crushed cells and debris, presumably from the phellogen and possibly phelloderm. This layer is found under the tough phellem cells that are also often damaged. High water vapor conductance resulting from wounding normally rapidly declines during the first 24–36 h after the tuber is cut (Lulai and Orr, 1995). However, water vapor conductance through the pressure bruised areas was approximately three times higher than that of control areas and remained high, without decline, for up to 4 days after removal from the pressures created by the storage environment. These results indicate that there was no induction of wax biosynthesis/accumulation as these cells were damaged by storage pressures. Nor was there induction of wax biosynthesis/accumulation at these bruise points after the tubers were removed from the pressures created by the storage environment and after atmospheric oxygen was readily available. Most, but not all, of the pressure bruise areas showed no induction of suberization. Wound healing responses, including suberization and reduced water vapor conductance, as an indication of wax deposition, were inhibited in tuber cells damaged by pressure bruising and in neighboring cells (Lulai et al., 1996). The continued pressure and tight tuber contact at the pressure bruise site would restrict the supply of oxygen from the ventilation stream to the damaged cells. However, oxygen would be available to the pressure bruise areas after removal from the bin. The changes in oxygen availability to cells compromised by pressure bruise could influence blackspot development during and after removal from storage. Proper humidification to reduce loss of turgor is important in minimizing cellular damage and defect development associated with pressure bruising. 22.4.4 Growth cracks Tuber growth cracks are fully healed wounds that develop during tuber growth. These cracks occur after rain or irrigation prompts rapid water uptake resulting in increased tuber turgor pressure and growth (Storey and Davies, 1992; Hiller and Thornton, 1993; Thornton, 2001). Internal pressure exceeds the tensile strength of the surface tissues during tuber enlargement; the resulting tissue failure results in a physical cracking of the tuber surface. The cracks frequently occur on the bud end of the tuber, usually extend lengthwise in the tubers, and vary in length and depth. There is little published information describing the physiology of growth cracks and associated wound healing. The apparent lack of disease associated with growth cracks suggests that these wounds heal readily as the tubers grow in the soil. Growth cracks may also result from various infections and chemical treatments. 22.4.5 Skinning Tuber skinning injury is a superficial wound generally inflicted by mechanical forces during harvest and handling, which results in the fracture of fragile radial phellogen cell walls of the native periderm and loss of the protective layer of phellem cells (Section 22.2). Radial phellogen cell walls connect and physically hold the tuber phellem (skin) in place.

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

496

28-4-2007

9:37 a.m.

Page:496

Trimsize:165×240 MM

Lulai

These cell walls are thin and fragile while the phellogen of the native periderm is meristematically active during growth; tubers are susceptible to skinning/excoriation during this time. As the potato plant senesces and growth terminates, phellogen meristematic activity ceases, the phellogen cell walls strengthen/thicken, and the tuber becomes resistant to skinning, i.e. skin-set develops. Skinning results in discoloration of the wounded area, release of water sequestered within tuber cells, loss of turgor and increased susceptibility to associated pressure bruising and blackspot development, and creation of areas that are open to infection. Red-skinned genotypes are generally most susceptible to skinning injury and require more time for periderm maturation and proper skin-set development. The reason for red-skinned genotypes being predisposed to hindered skin-set development is not known. The red-skin pigment is not synthesized and replaced during wound healing, thus leaving a light off-color blemish amid the red skin. Depending on the maturity of the potato plant at the end of the growing season, no less than 10 days and generally 21 days of periderm maturation are routinely recommended for proper skin-set before harvest (Lulai and Orr, 1993; Struik and Wiersema, 1999; Olsen et al., 2003; Stark and Love, 2003). The wounding caused by skinning injury is frequently categorized as a bruise and can contribute to significant increases in carbon dioxide concentrations in storage bins directly after harvest (Mazza and Siemens, 1990; Schaper et al., 1993). The increased carbon dioxide concentration adversely affects suberization and can increase the reducing sugar concentrations and darkening of processed food products (Mazza and Siemens, 1990). Skinning injuring is among the most common yet troublesome and costly problems of the potato industry.

22.5 SUMMARY A healthy intact periderm is essential for protecting the potato tuber from desiccation, infection, pests, and other intrusions that cause food quality and supply problems and financial losses. A comprehensive understanding of the physiological and biochemical processes associated with skin-set, wound healing, and wound-related responses to bruising is essential for the development of technologies to minimize these problems. Advances have been made; however, the understanding of these processes is far from complete. Research has shown that skin-set development is not caused by skin thickening, suberization, or increased tensile strength of the skin. Instead, phellogen cell wall strengthening and cell wall thickening within the periderm have been shown to be responsible for the development of resistance to tuber skinning injury. Immunolocalization data indicate that accumulation and modification of specific cell wall polymers are among those processes involved in skin-set development. Further information is needed on the biological processes that are involved in tuber phellogen cell wall strengthening and skin-set development. The biological signals that initiate and regulate skin-set development have not been elucidated. This lack of biological information hinders development of technologies to predict, initiate, and/or enhance the development of resistance to skinning injury. Wound-induced suberization is perhaps the most important facet of wound healing. Tubers cut for seed, wounded by various means including harvest/handling processes, and tubers challenged by various biotic and abiotic stresses utilize suberization as the

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:497

Trimsize:165×240 MM

497

Skin-set, wound healing, and related defects

most durable form of protection. A competent suberized barrier protects the tuber from infection, dehydration, and various intrusions. The SPPD has been shown to provide resistance to bacterial but not fungal infection. The SPAD is architecturally distinct and spatially separate from the phenolic domain and is required to provide the final barrier to fungal infection. Associated waxes are part of the protective mechanism that controls water vapor loss. The induction and regulation of suberin biosynthesis, the biosynthetic pathways as well as the assembly and molecular architecture of the suberin biopolymer are of fundamental importance in enhancing wound healing and solving wound-related problems. A better understanding of these processes and the structure of suberin is on the horizon. However, it is obvious that further research is required in all these areas to provide the basis for the development of new technologies that will enhance suberization and mitigate associated food quality and loss problems. Lastly, wound-related responses are crucial elements of the various bruise-related problems and losses described in section 22.4.

ACKNOWLEDGEMENTS The author attempted to include references pertinent to the topics and apologies to those authors who published papers that were cited indirectly through review articles or for other reasons were not directly included in this chapter.

REFERENCES Abeles F.B., P.W. Morgan and M.E. Saltveit (eds), 1992, In: Ethylene in Plant Biology, p. 26. Academic Press, N.Y. Agrawal V.P. and P.E. Kolattukudy, 1977, Plant Physiol. 59, 667. Agrawal V.P. and P.E. Kolattukudy, 1978a, Arch. Biochem. Biophys. 191, 452. Agrawal V.P. and P.E. Kolattukudy, 1978b, Arch. Biochem. Biophys. 191, 466. Arrieta-Baez D. and R.E. Stark, 2006, Phytochemistry, 67, 743. Artschwager E.F., 1918, J. Agric. Res. 14, 221. Artschwager E., 1924, J. Agric. Res. 27, 809. Artschwager E., 1927, J. Agric. Res. 27, 995. Bajema R.W., G.M. Hyde and A.L. Baritelle, 1998, Trans. ASAE 41, 741. Banks N.H. and S.J. Kays, 1988, J. Am. Soc. Hortic. Sci. 113, 577. Baritelle A.L. and G.M. Hyde, 1999, Trans. ASAE 42, 159. Baritelle A.L. and G.M. Hyde, 2001, Postharvest Biol. Technol. 21, 331. Baritelle A.L. and G.M. Hyde, 2003, Postharvest Biol. Technol. 29, 279. Beckman C.H., 2000, Physiol. Mol. Plant Pathol. 57, 101. Bernards M.A., 2002, Can. J. Bot. 80, 227. Bernards M.A. and N.G. Lewis, 1992, Phytochemistry 31, 3409. Bernards M.A. and N.G. Lewis, 1998, Phytochemistry 47, 915. Bernards M.A. and F.A. Razem, 2001, Phytochemistry 57, 1115. Bernards M.A., W.D. Fleming, D.B. Llewellyn, R. Priefer, X. Yang, A. Sabatino and G.L. Pluourde, 1999, Plant Physiol. 121, 135. Bernards M.A., M.L. Lopez, J. Zajicek and N.G. Lewis, 1995, J. Biol. Chem. 270, 7382. Bernards M.A., D.K. Summerhurst and F.A. Razem, 2004, Phytochem. Rev. 3, 113. Bernards M.A., L.M. Susag, D.L. Bedgar, A.M. Anterola and N.G. Lewis, 2000, J. Plant Physiol. 157, 601. Bleecker R.C. and H. Kende, 2000, Annu. Rev. Cell Dev. Biol. 16, 1.

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

498

28-4-2007

9:37 a.m.

Page:498

Trimsize:165×240 MM

Lulai

Bostock R.M. and B.A. Stermer, 1989, Annu. Rev. Phytopathol. 27, 343. Bowen S.A., A.Y. Muir and C.T. Dewar, 1996, Potato Res. 39, 313. Brook R.C. (ed.), 1996, In: Potato Bruising, How and Why, Emphasizing Blackspot Bruise, p. 1. Running Water Publishing, Haslett, MI. Bush M.S. and M.C. McCann, 1999, Physiol. Plant. 107, p. 201. Catesson A.M., 1994, Int. J. Plant Sci. 155, 251. Catesson A.M., R. Funada, D. Robert-Baby, M. Quinet-Szel, J. Cu-Ba and R. Goldberg, 1994, IAWA J. 15, 91. Chaffey N.J., P.W. Barlow and J.R. Barnett, 1998, New Phytol. 139, 623. Choi D., R.M. Bostock, S. Avdiushko and D.F. Hildebrand, 1994, Proc. Natl. Acad. Sci. U.S.A. 91, 2329. Ciardi J. and H. Klee, 2001. Ann. Bot. 88, 813. Cottle W. and P.E. Kolattukudy, 1982a, Plant Physiol. 69, 393. Cottle W. and P.E. Kolattukudy, 1982b, Plant Physiol. 70, 775. Davin L.B. and N.G. Lewis, 2000, Plant Physiol. 123, 453. de Bruxelles G.L. and M.R. Roberts, 2001, Crit. Rev. Plant Sci. 20, 487. de Haan P.H., 1987, In: Rastovski, A., A. van Es. et al. (eds), Storage of Potatoes: Post-harvest Behavior, Store Design, Storage Practice, Handling, p. 371. Pudoc, Wageningen, The Netherlands. Dean B.B., 1989, Plant Physiol. 89, 1021. Dean B.B., N. Jackowiack and S. Munck, 1992, Potato Res. 35, 49. Dean B.B., N. Jackowiak, M. Nagle, J. Pavek and D. Corsini, 1993, Am. Potato J. 70, 201. Dean B.B. and P.E. Kolattukudy, 1976, Plant Physiol. 58, 411. Dean B.B. and P.E. Kolattukudy, 1977, Plant Physiol. 59, 48. Dean B.B., P.E. Kolattukudy and R.W. Davis, 1977, Plant Physiol. 59, 1008. Delgado E., J. Poberezny, E. Pawelzik and I. Rogozinska, 2001a, Am. J. Potato Res. 78, 389. Delgado E., M.I. Sulaiman and E. Pawelzik, 2001b, Potato Res. 44, 207. Edgell T., E.R. Brierley and A.H. Cobb, 1998, Ann. Appl. Biol. 132, 143. Espelie K.E., V. Franceschi and P.E. Kolattukudy, 1986, Plant Physiol. 81, 487. Graca J. and H. Pereira, 2000, J. Agric. Food Chem. 48, 5476. Goldberg R., P. Devillers, R. Prat, C. Morvan, V. Michon and C. Hervé du Penhoat, 1989, In: N.G. Lewis and M.G. Paice (eds), Plant Cell Wall Polymers – Biogenesis and Biodegradation, p. 312. American Chemical Society, Washington D.C. Gould W.A. 1989, In: W.A. Gould, J.R. Sowokinos, E. Banttari, P.H. Orr and D.A. Preston (eds), Chipping Potato Handbook, p. 15. The Snack Food Association, Alexandria VA. Gronwald J.W., 1991, Weed Sci. 39, 435. Hammerschmidt R., 1984, Physiol. Plant Pathol. 24, 33. Hampson C.P., T.J. Dent and M.W. Ginger, 1980, Proceedings Conference Potato Tuber Damage, Ann. Appl. Biol. 96, 366. Halderson J.L. and R.C. Henning, 1993, Am. Potato J. 70, 131. Hiller L.K., D.C. Koller and R.E. Thornton, 1985, In: P.H. Li (ed.), Potato Physiology, p. 389. Academic Press Inc., New York. Hiller L.K. and R.E. Thornton, 1993, In: R.C. Rowe (ed.), Potato Health Management, p. 87. APS Press, The American Phytopathological Society, St. Paul, MN, U.S.A. Karlsson B.H., J.P. Palta and P.M. Crump, 2006, HortScience, 41, 1213. Koda Y. and Y. Kikuta, 1994, Plant Cell Physiol. 35, 751. Kolattukudy P.E., 1978, In: G. Kahl (ed.), Biochemistry of Wounded Plant Tissues, pp. 43. Walter de Gruyter, Berlin, New York. Kolattukudy P.E., 1980, Science 208, 990. Kolattukudy P.E., 2001, Biopolyesters. In: T. Scheper (ed.), Advances in Biochemical Engineering Biotechnology, Vol. 71, p. 1. Springer-Verlag, Berlin. Kolattukudy P.E. and B.B. Dean, 1974, Plant Physiol. 54, 116. Kumar G.N.M. and N.R. Knowles, 2003, Physiol. Plant. 117, 108. Lachaud S., A.M. Catesson and J.L. Bonnemain, 1999, C.R. Acad. Sci. III-Vie 322, 633. Laerke P.E., J. Christiansen and B. Veierskov, 2002, Postharvest Biol. Technol. 26, 99. Lapierre C., B. Pollet and J. Negrel, 1996, Phytochemistry 42, 949. Laties G.G., 1978, In: G. Kahl (ed.), Biochemistry of Wounded Plant Tissues, p. 421. W. de Gruyter, Berlin.

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

28-4-2007

9:37 a.m.

Page:499

Trimsize:165×240 MM

499

Skin-set, wound healing, and related defects

Leon J., E. Rojo and J.J. Sanchez-Serrano, 2001, J. Exp. Bot., 52, 1. Lewis N.G., L.B. Davin and S. Sarkanen, 1999, Carbohydrates and their derivatives including tannins, cellulose and related lignins. In: Sir D.H.R. Barton and K. Nakanishi (eds-in-chief), Comprehensive Natural Products Chemistry, Vol. 3, p. 617. Elsevier Science Ltd, Oxford, U.K. Li S.-X. and A.M. Showalter, 1996, Physiol. Plant. 97, 718. Lulai E.C., 2001a. In: W.R. Stevenson, R. Loria, G.D. Franc and D.P. Weingartner (eds), Compendium of Potato Diseases, p. 3. APS Press, St. Paul, MN, U.S.A. Lulai E.C., 2001b, In: W.R. Stevenson, R. Loria, G.D. Franc and D.P. Weingartner (eds), Compendium of Potato Diseases, p. 6. APS Press, St. Paul, MN, U.S.A. Lulai E.C., 2002, Am. J. Potato Res. 79, 241. Lulai E.C., 2005, Am. J. Potato Res. 82, 433. Lulai E.C. and D.L. Corsini, 1998, Physiol. Mol. Plant Pathol. 53, 209. Lulai E.C and T.P. Freeman, 2001, Ann. Bot. 88, 555. Lulai E.C. and W.C. Morgan, 1992, Biotech. Histochem. 67, 185. Lulai E.C. and P.H. Orr, 1993, Am. Potato J. 70, 599. Lulai E.C. and P.H. Orr, 1994, Am. Potato J. 71, 489. Lulai E.C. and P.H. Orr, 1995, Am. Potato J. 72, 225. Lulai E.C. and J.C. Suttle, 2004, Postharvest Biol. Technol. 34, 105. Lulai E.C. and J.C. Suttle, 2005, 16th Triennial Conference of the EAPR, Abstracts of papers and posters, II poster presentations, p. 775. Lulai E.C., M.T. Glynn and P.H. Orr, 1996, Am. Potato J. 73, 197. Lulai E.C., J.J. Weiland, J.C. Suttle, R.P. Sabba and A.J. Bussan, 2006, Am. J. Potato Res., 83, 409. Lyon G.D., 1989, Plant Pathol. 38, 313. Lyshede O.B., 1977, In: Yearbook of the Royal Veterinary and Agricultural University, p. 68. Copenhagen, Denmark. Matsuda F., K. Morino, M. Miyashita and H. Miyagawa, 2003, Plant Cell Physiol. 44, 510. Mazza G. and A.J. Siemens, 1990, Am. Potato J. 76, 121. McGarry A., C.C. Hole, R.L.K. Drew and N. Parsons, 1996, Postharvest Biol. Technol. 8, 239. Micheli F., B. Sundberg, R. Goldberg and L. Richard, 2000, Plant Physiol. 124, 191. Moire L., A. Schmutz, A. Buchala, B. Yan, R.E. Stark and U. Ryser, 1999, Plant Physiol. 119, 1137. Morris S.C., M.R. Forbes-Smith and F.M. Scriven, 1989, Physiol. Mol. Plant Pathol. 35, 177. Muir A.Y. and S.A. Bowen, 1994, Int. Agrophysics 8, 531. Murphy H.J., 1968, Am. Potato J. 45, 472. Negrel J., S. Lotfy and F. Javelle, 1995, J. Plant Physiol. 146, 318. Negrel J., B. Pollet and C. Lapierre, 1996, Phytochemistry 43, 1195. Niklas K.J., 1992a, In: K.J. Niklas (ed.), Plant Biomechanics. An Engineering Approach to Plant Form and Function, p. 48. The University of Chicago Press, Chicago. Niklas K.J., 1992b, In: K.J. Niklas (ed.), Plant Biomechanics. An Engineering Approach to Plant Form and Function, p. 234. The University of Chicago Press, Chicago. Nnodu E.C., M.D. Harrison and R.V. Park, 1982, Am. Potato J. 59, 297. Nolte P., G.A. Secor and N.C. Gudmestad, 1987, Am. Potato J. 64, 1. Nolte P., G.A. Secor, N.C. Gudmestad and P.J. Henningson, 1993, Am. Potato J. 70, 649. O’Brien V.J. and S.S. Leach, 1983, Am. Potato J. 60. 227. Obro J., J. Harholt, H.V. Scheller and C. Orfila, 2004, Phytochemistry 65, 1429. Olsen N., G.E. Kleinkopf and J.C. Stark, 2003, In: J.C. Stark and S.L. Love (eds), Potato Production Systems, p. 309. The University of Idaho Extension/The University of Idaho Agricultural Communications, Moscow, ID, U.S.A. Oosterhaven K., K.J. Hartmans, J.J.C. Scheffer and L.H.W. van der Plas, 1995, Physiol. Plant. 93, 225. Ostby P.B., A.Y. Muir and F.N. Zender, 1990, Abstracts 11th Triennial Conference, European Association for Potato Research, Edinburgh, Scotland, p. 176. Oomen R.J.F.J. 2003, In planta modification of the potato tuber cell wall. Thesis, Wageningen University, The Netherlands. Oomen R., C.H.L. Doeswijk-Voragen, M.S. Bush, J.-P. Vincken, B. Borkhardt, L.A.M. van den, Broek, J. Cosar, P. Ulvskov, A.G.J. Voragen, M.C. McCann and R.G.F. Visser, 2002, Plant J. 30, 403.

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines

Elsevier AMS

Ch22-N51018

Job code: PBBA

500

28-4-2007

9:37 a.m.

Page:500

Trimsize:165×240 MM

Lulai

Parker C.C., M.L Parker, A.C. Smith and K.W. Waldron, 2001, J. Agric. Food Chem. 49, 4364. Partington J.C., C. Smith and G.P. Bolwell, 1999, Planta 207, 449. Pavlista A.D., 2002, Am. J. Potato Res. 79, 301. Pawelzik E., N.U. Haase, A. Schulze and R. Peters, 2005, Abstracts 16th Triennial Conference, European Association for Potato Research, Bilbao, Spain, p. 375. Pérombelon M.C.M., 2002, Plant Pathol. 51, 1. Peterson R.L. and W.G. Barker, 1979, Bot. Gaz. 140, 398. Peterson R.L., W.G. Barker and M.J. Howarth, 1985, In: P.H. Li (ed.), Potato Physiology, p. 123. Academic Press Inc., New York. Razem F.A. and M.A. Bernards, 2003, J. Exp. Bot. 54, 935. Reeve R.M., E. Hautala and M.L. Weaver, 1969, Am. Potato J. 46, 361. Reyes L.F. and L. Cisneros-Zevallos, 2003, J. Agric. Food Chem. 51, 5296. Rosenstock G. and G. Kahl, 1978, In: G. Kahl (ed.), Biochemistry of Wounded Plant Tissues, p. 623. Walter de Gruyter & Co., Berlin, New York. Sabba R.P. and E.C. Lulai, 2002, Ann. Bot. 90, 1. Sabba R.P. and E.C. Lulai, 2004, Am. J. Potato Res. 81, 119. Sabba R.P. and E.C. Lulai, 2005, J. Am. Soc. Hort. Sci. 130, 936. Schaper L.A., M.T. Glynn, and J.L. Varns, 1993, Appl. Eng. Agric. 10, 89. Schaper L.A. and E.C. Yaeger, 1982, Trans. ASAE 25, 719. Schilmiller A.L. and G.A. Howe, 2005, Curr. Opin. Plant Biol. 8, 369. Schmutz A., A.J. Buchala and U. Ryser, 1996, Plant Physiol. 110, 403. Schreiber L., M. Skrabs, K. Hartmann, D. Becker, C. Corsagne and R. Lessire, 2000, Biochem. Soc. Trans. 28, 647. Schreiber L., R. Franke and K. Hartmann, 2005a, Planta 220, 520. Schreiber L., R. Franke and R. Lessire, 2005b, Phytochemistry 66, 131. Scott R.I., J.M. Chard, M.J. Hocar, J.H. Lennard and D.C. Graham, 1996. Potato Res. 39, 333. Smittle D.A., R.E. Thornton, C.L. Peterson and B.B. Dean, 1974, Am. Potato J. 51, 152. Soliday C.L., B.B. Dean and P.E. Kolattukudy, 1978, Plant Physiol. 61, 170. Soliday C.L., P.E. Kolattukudy and R.W. Davis, 1979, Planta 146, 607. Stark R.E. and J.R. Garbow, 1992, Macromolecules 25, 149. Stark J.C. and S.L Love, 2003, In: J.C. Stark and S.L. Love (eds), Potato Production Systems, p. 329. The University of Idaho Extension/The University of Idaho Agricultural Communications, Moscow, ID, U.S.A. Stevens L.H. and E. Davelaar, 1996, Phytochemistry 42, 941. Stevens L.H. and E. Davelaar, 1997, J Agric. Food Chem. 45, 4221. Stevens L.H., E. Davelaar, R.M. Kolb, E.J.M. Pennings and N.P.M. Smit, 1998, Phytochemistry, 49, 703. Storey R.M.J. and H.V. Davies, 1992, In: P.M Harris (ed.), The Potato Crop, The scientific basis for improvement, p. 507. Chapman and Hall, London. Struik P.C. and S.G. Wiersema (eds), 1999, In: Seed Potato Technology, p. 219. Wageningen Pers, Wageningen, The Netherlands. Thomson N., R.F. Evert and A. Kelman, 1995, Can. J. Bot. 73, 1436. Thornton R.E., 2001, In: W.R. Stevenson, R. Loria, G.D. Franc and D.P. Weingartner (eds), Compendium of Potato Diseases, p. 84. APS Press, St. Paul, MN, U.S.A. Tyner D.N., M.J. Hocart, J.H. Lennard and D.C. Graham, 1997, Potato Res. 40, 181. Vaughn S.F. and E.C. Lulai, 1991, Physiol. Mol. Plant Pathol. 38, 455. Vayda M.E., L.S. Antonov, Z. Yang, W.O. Butler and G.H. Lacy, 1992, Am. Potato J. 69, 239. Vayda M.E. and J.K. Morelli, 1994, In: W.R. Belknap, M.E. Vayda and W.D. Park (eds), The Molecular and Cellular Biology of the Potato, p. 188. CAB International, Wallingford, U.K. Wasternack C., I. Stenzel, B. Hause, G. Hause, C. Kutter, H. Maucher, J. Neumerkel, I. Feussner and O. Miersch, 2006, J. Plant Physiol., 163, 297. Wigginton M.J., 1973, Potato Res. 16, 85. Wigginton M.J., 1974, Potato Res. 17, 200. Yan B. and R.E. Stark, 2000, J. Agric. Food Chem. 48, 3298. Yang W.L. and M.A. Bernards, 2006, Plant Signal. Behav. 1, el. Yephremov A. and L. Schreiber, 2005, Plant Biosyst. 139, 74.

Basal Fonts:Times

Margins:Top:5 pc

Gutter:4.6 pc

Font Size:10/12

Text Width:30 pc

Depth:45 Lines