Summary
Lenticels can be defined as pores that are the entrance of a continuous aeration system from the atmosphere via the living bark to the secondary xylem in the otherwise protective layers of the periderm. Most work on lenticels has had an anatomical focus but the structure-function relationships of lenticels still remain poorly understood. Gas exchange has been considered the main function of lenticels, analogous to the stomata in leaves. In this perspective review, we introduce novel ideas pertaining to lenticel functions beyond gas exchange. We review studies on lenticel structure, as this knowledge can give information about structure-function relationships. The number of species investigated to-date is low and we provide suggestions for staining techniques for easy categorization of lenticel types. In the follow-up sections we review and bring together new hypotheses on lenticel functioning in the daily “normal operation range”, including regulative mechanisms for gas exchange and crack prevention, the “stress operation range” comprising flooding, drought and recovery from drought and the “emergency operation range”, which includes infestation by insects and pathogens, wounding and bending. We conclude that the significance of dermal tissues and particularly of lenticels for tree survival has so far been overlooked. This review aims to establish a new research discipline called “Phytodermatology”, which will help to fill knowledge gaps regarding tree survival by linking quantitative and qualitative lenticel anatomy to tree hydraulics and biomechanics. A first step into this direction will be to screen more species from a great diversity of biomes for their lenticel structure.
Introduction
Lenticels can be broadly defined as pores embedded in the periderm of trees and shrubs that serve in gas exchange in otherwise impervious outer protective layers (Angyalossy et al. 2016). While most past work on lenticels has had an anatomical focus, the structure-function relationships of lenticels remain poorly understood (Lendzian 2006). The periderm and, in particular, the lenticels are still among the least studied plant parts notwithstanding their protective as well as physiological role in plant functioning (Leite & Pereira 2017; Serra et al. 2022).
Gas exchange is considered the principal function of lenticels (Unger 1836; Haberlandt 1875; Langenfeld-Heyser 1997; Lendzian 2006), analogous to the stomata in leaves. Plants under drought stress close the stomata of their leaves to avoid excessive water loss and cavitation of the water columns in the xylem conduits. While we are still lacking knowledge on whether such safety mechanisms exist in lenticels (Lintunen et al. 2021), mechanisms for drought-induced stomatal closure are nowadays quite well understood (e.g., Martin-StPaul et al. 2017; Buckley 2019; Henry et al. 2019). Some species, especially among angiosperms, have low hydraulic safety margins, closing their stomata quickly (Choat et al. 2012). In this case, the tree must survive on its sugar reserves providing enough water is available for transport to the sinks (Choat et al. 2018; Klein et al. 2018). Cambial activity regarding wood formation, i.e., xylogenesis, under restricted water availability has been intensively investigated (e.g., Plomion et al. 2001; Zhang et al. 2014; Cuny et al. 2015; Friend et al. 2019) and, only recently, attempts were made to link secondary phloem formation to drought events (e.g., Dannoura et al. 2019; Gričar et al. 2019; Prislan et al. 2019; Miller et al. 2020). A strong hint that lenticels play an important physiological role in tree functioning is their annual phellogen activity (e.g., Schönherr & Ziegler 1980; Rosner & Kartusch 2003; Shibui & Sano 2018), which might be related to changing demands in gas exchange between living bark tissues and the environment over the seasons and, when under stress, water availability. Regarding the latter demand, the continuous intercellular network (Klebahn 1884; Rosner & Kartusch 2003) from the atmosphere to the interior stem parts could function in the refilling of embolized conduits via bark moisture (water vapour) uptake (Earles et al. 2016; Cuneo et al. 2018; Liu et al. 2019; Lintunen et al. 2021). In relation to fluctuations in bark water content and subsequent stem diameter changes (e.g., Ilek et al. 2021; Van Stan et al. 2021), we propose that lenticels fulfil important biomechanical functions as the stem axes shrink and swell. Moreover, lenticels also have an influence on the susceptibility of bark and wood to insects and pathogenic fungi (Neger 1922; Rosner & Führer 2002; Nemesio-Gorriz et al. 2019).
Variability in lenticel size and shape. (a) Lime tree (Tilia sp.). (b) Black elder (Sambucus nigra L.). (c) Paper birch (Betula papyrifera Marshall). (d) Tibetan cherry (Prunus serrula Lindl.). (e) Silver poplar (Populus alba L.) with very big rhomboid lenticels. Scale bar = 10 mm.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Little is known about the function of lenticels for tree survival and we still fail to understand the regulatory mechanisms for gas exchange, or moisture release and uptake, which are considered two conflicting functions. To unravel the functional aspects of lenticels, an investigation into the anatomical design of lenticels and the tissues beneath them is required. However, lenticels are extremely complex structures, varying tremendously in size within and between species (Fig. 1). To understand the structure-function relationships of lenticels, a holistic approach is necessary that includes several functional aspects of bark (Rosell 2019) and wood (Baas 1983; Lachenbruch & McCulloh 2014). Here, we review the existing knowledge about lenticels and maintain that lenticels fulfil important functions for woody stems beyond gas exchange.
Lenticel anatomy
Lenticel types and associated anatomy remain unsolved
A lenticel is a structure often visible with the naked eye on the outer surface of the bark of woody species that has continuity through the living bark to the vascular cambium. The actual form of a lenticel can be quite variable (Fig. 1); however, in all cases, lenticels are formed at locations of inhomogeneity in the phellogen (cork cambium) called lenticular phellogen. Lenticular phellogen is more meristematically active than surrounding phellogen (Fig. 2a) and has intercellular spaces in the dormant stage (Fig. 2d). In addition, lenticular phellogen produces phelloderm cells (i.e., living parenchyma) to the interior and filling tissue to the exterior (Trockenbrodt 1990; Angyalossy et al. 2016). Filling tissue (“sc” and “sp” in Fig. 2c–d) can be distinguished from the phellem (Fig. 2e) of the periderm in having intercellular spaces. Comparable to lenticels on the bark surface (Fig. 1), the products of lenticular phellogen can also vary considerably, with filling tissue in lenticels varying in density, extent of inclusions, extractives and waxes, as well as in the volume of intercellular space. We use a sketch (Fig. 2a) of a Picea abies lenticel (Fig. 2b) to show a general anatomical overview and the high structural variability in the filling tissue. Information on lenticel structure is — with some exceptions (Wutz 1955) — available solely for primary lenticels, defined as lenticels in the primary (first-formed) periderm. Secondary lenticels (Wutz 1955) are present in subsequent periderms when the rhytidome has been formed.
Structure of primary Picea abies (L.) Karst. lenticels (b–d) and periderm (e). (a) Sketch of a lenticel in the dormant season. (b) Longitudinal-radial section, early summer. (c) Tangential section, early summer. (d) Tangential section, autumn. (e) Longitudinal-radial section, autumn. Scale bars: (b, c) 250 μm; (d, e) 50 μm. Arrows in (b) point to lenticular phellogen, in (d) at intercellular spaces in the lenticular phellogen and in (e) at crystals. Abbreviations: ch, chlorenchyma; lp, lenticular phellogen; pd, phelloderm; sc, sclereids; se, suberized empty cell of the phellem; sp, suberized filling tissue or phellem cells with polyphenolic content; uf, unsuberized filling tissue. Staining: (b, c) Astra blue/Safranin; (d, e), Sudan Black B.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
DeCandolle (1826) and Unger (1836) described anatomical and potential physiological aspects of lenticels; however, it was not until 1873 that a first attempt to provide an overview on the structural variability of lenticels was published (Stahl 1873). This work was then further developed by Klebahn (1884) and Devaux (1900). Stahl (1873) made a distinction between lenticels that had non-stratified, relatively dense, filling tissues from those with stratified filling tissue having alternating layers of dense and less dense material. Klebahn (1884) categorized three different types of lenticels and provided additional information on the chemical characteristics of the cell walls of the filling tissues in selected species (Table 1). In later works, Wutz (1955) and Langenfeld-Heyser (1997) suggested four lenticel types, whereby the “gymnosperm type”, which was regarded as the most primitive one, represented its own category. Angyalossy et al. (2016) classify three angiosperm lenticel types: (I) where filling tissue is non-stratified and suberized, (II) filling tissue non-stratified and largely unsuberized and a compact layer of suberized cells formed at the end of the growing season, and (III) filling tissue composed of alternating layers of loose unsuberized cells and compact suberized layers (Table 1). Type I is found in, e.g., Liriodendron, Magnolia, Malus, Persea, Populus, Pyrus and Salix, etc., Type II in, e.g., Sambucus, Fraxinus, Quercus and Tilia, etc., and Type III in, e.g., Betula, Fagus, Prunus and Robinia, etc. (Esau 1977; Evert 2006). For most woody species, especially those of tropical regions of the world, we are lacking knowledge on the layering of the filling tissue (Table 1) and remain far from understanding the functional aspects of the different structural varieties.
Gymnosperm lenticel anatomy
Gymnosperm lenticels have been regarded as the simplest type, as the filling tissue is composed of the same type of cells as surrounding phellem except for the presence of intercellular spaces with a sizable production of filling tissue (Stahl 1873; Klebahn 1884; Devaux 1900; Wutz 1955; Esau 1977; Evert 2006). Some species are presumed to have no lenticels per se, although this is probably owing to difficulty in finding them macroscopically. However, tangential periderm sections show evidence of areas with intercellular spaces in the phellem, e.g., in Araucaria heterophylla and Taxus baccata (Klebahn 1884). According to our findings for Taxus baccata (Fig. 3a), we suggest that every woody species produces lenticels of varying abundance, even though they are not easily detected macroscopically. In gymnosperm lenticels, layered (Fig. 3b) as well as non-layered filling tissue (Fig. 3c) can be found (Table 2), which is analogous to the stratification of the periderm (Klebahn 1884). The phellem of Picea abies consists of tannin-filled cells with suberized walls (“phlobaphene cork”), empty cells with suberized walls (“spongy cork”) and mostly unsuberized thick walled sclereids (“stone cork”) (Fig. 2e). However, empty cells with suberized walls are missing in the central region of the lenticels; moreover, thin-walled filling tissue with no suberized walls has no counterpart in the phellem (Parameswaran et al. 1976; Rosner & Kartusch 2003). We would like to add that the term “cork” should be regarded as a non-technical term for the commercial product (Trockenbrodt 1990) and will avoid it hereafter for describing derivatives of the phellogen. Moreover, the terminology “phlobaphene cork” should be avoided because tannins of conifers are merely hydrolysable (proanthocyanidines) rather than condensed tannins (phlobaphenes) (Trockenbrodt 1990). In Larix (Fig. 3d–e) and Cedrus (Fig. 3f) species, densely packed plates of sclereids can be found in the periderm and lenticels (Neger & Kupka 1920). Lenticular phellogen of Chamaecyparis, Juniperus and Thuja produces suberized intercellular free filling tissue with tannins occupying the central part of the lenticel, while at its edges, some unsuberized cell layers with intercellular spaces can be found. Gases can only pass through the latter described parts of the lenticel (Neger & Kupka 1920). Gymnosperm lenticels show a high structural variability and the functional aspects of this variability demands closer observation to properly understand a species’ reaction to environmental stresses. In the following sections, we show that gymnosperm lenticels can be integrated into the angiosperm category system.
Angiosperm list of lenticel categorizations.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Variability in gymnosperm lenticels. (a) Taxus baccata L., dormant season. (b) Picea abies (L.) Karst., dormant season. (c) Developing Ginkgo biloba L. lenticel, dormant season. (d) Larix decidua Mill., dormant season. (e) Larix sibirica Ledeb. lenticel drawing modified from Neger & Kupka (1920). (f) Cedrus libani A.Rich. lenticel drawing modified from Neger & Kupka (1920). Scale bars: (a, c, d) 200 μm; (b) 500 μm. Arrows in (a–c) point to lenticular phellogen and in (d–f) at densely packed plates of filling tissue. Abbreviations: rd, primary resin duct; sc, sclereids; se, suberized empty cell; sp, suberized cell with polyphenolic content.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Gymnosperm list of lenticel categorizations.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Lenticel type I of the IAWA lenticel categorization (Angyalossy et al. 2016). The filling tissue is non-stratified and suberized (Salix-type; Wutz 1955). (a, b) Magnolia × soulangeana Soul.-Bod. (c, d) Carpinus betulus L. (e, f) Populus × canescens (Aiton) SM. (g, h) Salix caprea L. Sampling was done in the dormant season, except for Populus (e, f), in early summer. Scale bars: (a, c, g) 200 μm; (b, d) 50 μm; (e, f, h) 100 μm. Arrows point to lenticular phellogen. Abbreviations: co, collenchyma; se, suberized empty filling tissue; sp, suberized filling tissue with polyphenolic content. Staining: (a, c, e, g) Astra blue/Safranin; (b, d, f, h) Astra blue/Safranin & Sudan Black B.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Angiosperm lenticels: Type I anatomy
In lenticel type I, all cells of the filling tissue have suberized cell walls. Lenticular phellogen of Magnolia produces loose suberized filling tissue and some layers of more densely packed suberized cells filled with tannins that resemble phellem cells, with the exception of the intercellular spaces between them (Fig. 4a). Sudan Black B gives clear histological evidence for suberin and hydrophobic fatty substances such as waxes in the phellem and lenticel filling tissues (Fig. 4b) than the traditionally used Sudan III or IV stains, especially when cells have brown/orange polyphenolic contents (Rosner & Kartusch 2003). Type I can also be found in Carpinus betulus (Fig. 4c–d) and in Populus alba (Fig. 4e–f). Klebahn (1884) had previously observed that lenticels of several Populus species show special characteristics; their lenticular phellogen is not ‘watch-glass-shaped’ but ‘wedge-shaped’ and the loose filling tissue is tangentially elongated (Fig. 4e). Type I was termed the “Salix-type” by Wutz (1955). The filling tissue in Salix caprea is less homogenous than in other species of Type I; dense layers of tannin-filled cells alternate with empty, more loosely packed filling tissue (Fig. 4g). Probably for the aforementioned reason or as a result of the high structural variability in Salicaceae, Klebahn (1884) categorized Salix species as Type II (Table 1). However, all filling tissue produced by Salix caprea lenticular phellogen has suberized cell walls, as indicated by the dark blue staining with Sudan Black B (Fig. 4h).
Overview of key terms used in bark anatomy with a focus on stems.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Angiosperm lenticels: Type II anatomy
The lenticel Type II, where filling tissue is non-stratified, largely unsuberized and a compact layer of suberized cells is formed at the end of the growing season, has also been termed the “Sambucus-type” (Wutz 1955). Lenticels of Sambucus nigra (Fig. 1b) have often been used to describe lenticel anatomy, especially the developmental stages as bark ages; however, as Klebahn (1884) stated, the species proved unsatisfactory for the study of the structure of older lenticels owing to the collapsed and dried out state of the tissue with the majority of cells containing polyphenols. The aforementioned tissue condition obscures histochemical detection of suberin. Staining with Sudan Black B showed that the filling tissue of young Sambucus nigra primary lenticels (Fig. 5a) is mostly suberized (insert in Fig. 5b). However, primary lenticels of Sambucus canadensis have a more distinct layering with a greater quantity of unsuberized filling tissue (Esau 1977; Evert 2006). The lenticels of Quercus petraea (Fig. 5c) are a more typical representative of Type II. The filling tissue of Quercus petraea is mostly unsuberized; however, towards the end of the growing season, several cell rows of suberized filling tissue are produced (Fig. 5d).
Lenticel Type II of the IAWA lenticel categorization (Angyalossy et al. 2016). (a, b) Sambucus nigra L. (c, d) Quercus petraea (Mattuschka) Liebl. (e, f) Picea abies (L.) Karst.). The filling tissue is largely unsuberized and a compact layer of suberized cells is formed at the end of each growing season (Sambucus-type; Wutz 1955). Sampling was carried out in the dormant winter season, except for Picea (e, f), in early summer. Scale bars: (a) 200 μm, (d, f) 50 μm; (b, c, e) 100 μm. Arrows point to lenticular phellogen. Abbreviations: co, collenchyma; se, suberized empty filling tissue; sc, sclereids; sp, suberized filling tissue with polyphenolic content; uf, unsuberized filling tissue. Staining: (a, c, e) Astra blue/Safranin; (b, d) Astra blue/Safranin and Sudan Black B; (f) Sudan Black B.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Consistent with the annual production of filling tissue and the high number of unsuberized cells, primary lenticels of the conifer Picea abies can be categorized as Type II (Rosner & Kartusch 2003). The annually produced filling tissue consists mostly of unsuberized cells, where both densely packed sclereids and thin-walled — often collapsed — loosely arranged cells can be found (Fig. 5e). The cell wall chemistry of the filling tissue of P. abies compares well to the unsuberized filling tissue of angiosperms (Parameswaran et al. 1976). Unsuberized filling tissue in P. abies alternates with layers of tannin-filled cells with suberized walls, where the latter can also contain crystals (Fig. 5f).
Lenticel Type III of the IAWA lenticel categorization (Angyalossy et al. 2016). Each year, lenticular phellogen produces alternating layers of loose unsuberized cells and compact, suberized, layers (Prunus-type; Wutz 1955). (a, b) Prunus dulcis L. (c, d) Betula pendula Roth. (e, f) Fagus sylvatica L. Sampling was carried out in the dormant winter season, except for Fagus sylvatica (e, f), in early summer. Scale bars: (a–c, e) 100 μm; (d, f) 50 μm. Arrows point to the lenticular phellogen. Abbreviations: se, suberized empty filling tissue; sp, suberized filling tissue with polyphenolic content; uf, unsuberized filling tissue. Staining: (a, c, e) Astra blue/Safranin; (b, d, f) Astra blue/Safranin and Sudan Black B.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Angiosperm lenticels: Type III anatomy
In lenticel Type III, which has been termed the Prunus-type by Wutz (1955), the filling tissue is composed of alternating layers of loose unsuberized cells and compact suberized layers (Esau 1977; Evert 2006; Angyalossy et al. 2016). Lenticels of Prunus dulcis (Fig. 6a) show alternating layers of 2–3 rows of suberized filling tissue as well as layers of up to five rows of often collapsed loose filling tissue (Fig. 6b); both types of cells can contain polyphenolic contents. Young two-year-old branches contained four suberized and unsuberized layers produced by lenticular phellogen in the previous growing season. The annual rhythm of lenticular phellogen activity can easily be detected by the polyphenolic content in the suberized cells as only the last formed suberized layer of the season contains polyphenolic contents, while the earlier formed suberized layers appear empty (Fig. 6b). Klebahn (1884) made similar observations in Prunus padus. The lenticels of Betula pendula (Fig. 6c) also show the typical layering of lenticel type III; however, the annual increments are not easily detectable because in all suberized layers, cells with polyphenolic contents can be found (Fig. 6d). We counted six annual wood increments in a sampled branch and found nine layers of suberized filling tissue in the centre of the lenticel. The phellogen of young Betula maximowicziana lenticels in the main stem can produce up to three layers of suberized filling tissue (Shibui & Sano 2018). In our Betula pendula sample (Fig. 6c–d) we found that in most years only one set of suberized and unsuberized filling tissue was formed in the slower growing branches (Wutz 1955) or parts of the filling tissue layers had been shed. In Fagus sylvatica, several layers of suberized and loose unsuberized filling tissues are produced annually (Fig. 6e). The last formed suberized layer is distinguishable from layers previously formed in the same year because it consists of more cell rows and the suberin deposits in the walls of the filling tissue (dark blue staining with Sudan Black B) are much thicker (Fig. 6f). The cells in the terminally formed layer resemble normal phellem cells in Fagus sylvatica f. purpurea, with the exception that they are interspersed by intercellular spaces (Jacob et al. 1989). In addition, cells of the terminal suberized layer in Fagus sylvatica lenticels are less tangentially stretched than the phellem cells (Fig. 6e–f).
Overview of the annual production of filling tissue in the three lenticel types. Sketches represent lenticels in the dormant season. (a) Type I, Salix caprea L. in the dormant season; only suberized filling tissue is produced that may contain polyphenolic contents. (b) Type II, Picea abies L. Karst. sampled in early summer (photo left side); first (lignified) sclereids are produced, thereafter loose unsuberized tissue and finally a denser layer of suberized filling tissue with polyphenolic contents. (c) Type III, Prunus dulcis L. in the dormant season; the annual production comprises four layers of unsuberized and suberized tissues, respectively, whereby the suberized cells of the last formed layer contain polyphenols. — Stars indicate the lenticular phellogen. Abbreviations: ch, chlorenchyma; co, collenchyma; lp, lenticular phellogen; pd, phelloderm; se, suberized empty phellem cells or filling tissue; sp, suberized phellem cells or filling tissue with polyphenolic content; uf, unsuberized filling tissue. For terminology see Table 3. Staining: Astra blue/Safranin and Sudan Black B.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Common lenticel categories for all woody species and suggestions for further investigations
In summary, lenticular phellogen can produce several different anatomical types of filling tissue: (1) empty suberized cells, (2) suberized cells with polyphenolic content, which may also contain crystals (3) unsuberized loose filling tissue with lignified or non-lignified walls and (4) thick-walled and often lignified sclereids, which may contain polyphenolic contents (Fig. 7). We propose to place gymnosperms into the angiosperm categorization, making three lenticel types in total (Esau 1977; Evert 2006; Angyalossy et al. 2016) and keep this categorization until we gain more information about the species-specific variability in the stratification of the filling tissue. When describing the stratification of filling tissues, we avoided the term “closing layer”, used in earlier reviews for a dense suberized layer formed towards the end of the growing season (Trockenbrodt 1990) since this “terminal layer”, and, even the lenticel meristem contains intercellular spaces (Klebahn 1884; Esau 1977; Evert 2006; Angyalossy et al. 2016).
For detection of intercellular spaces and cell wall chemistry, semi-thin (1–2 μm) — or even thinner — sections are required. Embedding lenticels in hydrophilic resins such as 2-hydroxyethyl metacrylate (Rosner & Kartusch 2003) allows both semi-thin sections and histochemical detection of suberin. Combined staining with Astra blue-Safranin (e.g., Gerlach 1984; Angyalossy et al. 2016) and Sudan Black B (Rosner & Kartusch 2003) is a straightforward technique to examine filling tissue for both lignin and suberin (Fig. 7). Through this technique, non-lignified cell walls are stained light blue, while lignified cell walls are stained pink (Figs 5e, 7b). In addition, suberized walls display as dark blue while polyphenolic contents appear bright red (Fig. 7a–c), rather than brown/orange (Fig. 5f). The coloration can notably provide clues to the annual production of filling tissue as only the last formed suberized layer of the season may contain polyphenolic contents (Fig. 7c). The annual activity of lenticular phellogen differs between the three lenticel categories: Type I and II (Fig. 7) produce one dense suberized layer that is ruptured annually, while type III produces several suberized layers that may stay intact for years (Schönherr & Ziegler 1980; Shibui & Sano 2018). It is thus assumed that lenticels and not only wood (Anfodillo et al. 2013; Rosner 2013; Lachenbruch & McCulloh 2014) and living bark (Rosner et al. 2001; Jyske & Hölttä 2015; Rosell 2019; Rosell et al. 2021) are designed in relation to functional demands that differ through seasons, cambial age and environment.
In the following chapters of this perspective review, we outline three operation ranges for lenticel functioning to better understand how lenticels work under normal and stressed conditions. Lenticel functioning in the “normal operation range” includes the daily balance of bark gas exchange (Chan et al. 2018; Wittmann & Pfanz 2018) and water deficits (Geurten 1950) as well as dealing with the physiological demands of the stem that change over the seasons (Rosner & Kartusch 2003; Shibui & Sano 2018). The “stress operation range” comprises the contribution of lenticels to survival from hypoxia (e.g., Bertolde et al. 2012; Pangala et al. 2014; Yang & Li 2016), drought stress and recovery by refilling of embolized conduits (Cuneo et al. 2018) aided by bark photosynthesis (Nilsen 1995; Vick & Young 2009; Vandegehuchte et al. 2015; Rowland et al. 2018). The role of lenticels for recovery and defence in the “emergency operation range” when trees are bent (Rosner 1998), wounded or attacked by insects (Rosner & Führer 2002) and fungal pathogens, has been underestimated or totally overlooked in studies to-date (Serra et al. 2022). During a tree’s life cycle, all three scenarios will play crucial roles in tree health. However, to understand how lenticels function under stress requires a greater understanding of anatomical modifications to lenticel and periderm structure within all three scenarios.
Lenticel functioning in the normal operation range
Gas exchange and possible regulatory mechanisms
Through lenticels, the periderm is permeable to oxygen, carbon dioxide (e.g., Lendzian 2006; Tarvainen et al. 2014; Rowland et al. 2018) and other gaseous agents including some air pollutants such as methane, nitrous oxide (Pangala et al. 2014; Pitz et al. 2018; Welch et al. 2019) and O3 (Matyssek et al. 2002), organic volatile compounds, for instance monoterpenes (Vanhatalo et al. 2015) and also to water vapour (Oren & Pataki 2001; Ellsworth & Sternberg 2015). Water vapour can carry organic substances dissolved in it, such as methanol, acetone and acetaldehyde (Rissanen et al. 2018). For physiological functions, the “degree of opening” rather than the surface area covered by lenticels is thought to be relevant (Langenfeld-Heyser 1997). Structural features that might contribute to physiological functions could be the shape of the filling tissue cells, the volume of intercellular space in lenticular phellogen (Rosner & Kartusch 2003), or the extent of lenticular phelloderm and of tissues with chloroplasts (chlorenchyma) as well as the continuity of intercellular spaces connecting metabolically active sites from plant body to atmosphere (Langenfeld-Heyser 1997).
The permeability of lenticels to gases could be influenced by the seasonal activity of lenticular phellogen (Rosner & Kartusch 2003). Seasonal changes in the hydration status of the bark may have an impact on the permeability of lenticels (Lendzian 2006). Geurten (1950) suggested a mechanism that is analogous to the closing of stomata or “incipient drying”, the latter being defined as a contraction of bark due to turgor loss and the resulting shrinkage of cell walls followed by a decrease in the volume of intercellular volume. The swelling pressure required to increase lenticel permeability could be enhanced by hydration of cell wall pectin or degradation of cell contents (Jacob et al. 1989). The special lenticel type “Klappenventile” (cap valve), found in some conifers (Neger & Kupka 1920) and in aerial roots of Philodendron (Neger 1922), might allow sealing of the periderm under the impact of drought stress and eventual bark shrinkage, since the layer of densely packed sclereids (“sclerophelloids”) is without intercellular spaces (Fig. 3f). Combining cutting-edge techniques such as Raman spectroscopy (e.g., Gierlinger et al. 2012; Bock et al. 2021) and micro-CT (e.g., Earles et al. 2016; Cuneo et al. 2018; Crouvisier-Urion et al. 2019) will help to answer questions regarding the opening and closing mechanisms of lenticels.
Bark gas exchange and chronological adjustment of both lateral cambia
Bark gas exchange under normal conditions includes daily and seasonal management of CO2 recycling and O2 shortages (Chan et al. 2018; Wittmann & Pfanz 2018). Cells of living tissue immediately beneath superficial periderm contain chloroplasts capable of photosynthesis (Langenfeld-Heyser 1989; Langenfeld-Heyser et al. 1996; Aschan et al. 2001; Pfanz et al. 2002; Wittmann et al. 2006; Pfanz 2008; Wittmann & Pfanz 2014; Wittmann & Pfanz 2018). Photosynthesis in the living bark allows young stems to compensate for more than 50% of their respiratory carbon losses, an often overlooked component of carbon balance in woody plants (Wittmann & Pfanz 2008b; De Roo et al. 2020). Refixation of CO2 in the living bark is important for the carbon economy, especially during a leafless state (Aschan et al. 2001; De Roo et al. 2019). In the latter regard, bark photosynthesis is likely to make an important if not critical contribution to the carbon pool prior to leaf-flushing. In temperate regions, lenticular phellogen is annually active (Klebahn 1884; Wutz 1955; Jacob et al. 1989; Rosner & Kartusch 2003) but for a much shorter period than the vascular cambium (Klebahn 1884; Shibui & Sano 2018). During the dormant season an intact layer of densely packed, suberized cells can be found in all types of lenticels (Figs. 7 and 8a). In Picea abies, the production of new filling tissue starts later at higher elevations depending on climatic conditions (Rosner & Kartusch 2003). Lenticular phellogen is highly active and produces at first thick-walled sclereids and later thin-walled filling tissue, especially in early summer (Fig. 8b–c). The production of numerous rows of filling tissue eventually results in rupture of the suberized layer produced during the previous year (Fig. 8d). The new suberized layer is produced towards the end of the growing season (Fig. 8e), when cambial activity has ceased (Rosner 1998). The suberized layer may be ruptured either owing to the production of new filling tissue and/or from stem expansion due to secondary growth, implying that bark permeability is greater during periods of heightened cambial activity in P. abies and other species with lenticel type II anatomy (Fig. 5). In species that have the lenticel type III anatomy (Fig. 6), one or more layers of suberized filling tissue may remain intact during periods of increased wood growth (Shibui & Sano 2018). Information on the annual activity of lenticular phellogen and its timing with the vascular cambium is thus of high relevance regarding seasonal changes in sensitivity to drought stress in woody plants. A basic knowledge of seasonal changes in lenticel tissues culminating in differences in lenticel permeability (Lendzian 2006) will also help to better understand the relationships between stem respiration and stem growth (Pruyn et al. 2003; 2005; Chan et al. 2018). The research following this direction should not be only focused on temperate but also tropical species.
Annual activity of lenticular phellogen of Picea abies (L.) Karst. in Central Europe (Kindberg, Austria, 47°33′N, 15°26′E, altitude 1200 m (a.s.l.)). (a) Dormancy in November (10.11.1996). (b) Lenticular phellogen activity in early summer (07.06.1997). (c) Filling tissue production in summer (06.07.1996). (d) Lenticel activity in late summer (04.08.1997). (e) Production of the terminal layer in autumn. Scale bars: (a–d) 50 μm; (e) 25 μm. Stars indicate the lenticular phellogen. pd, phelloderm; sc, sclereids; sp, suberized filling tissue with polyphenolic content; uf, unsuberized filling tissue. Staining: Toluidine blue.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
Regulation of bark transpiration in the normal operation range
Transpiration through bark is possible (e.g., Catinon et al. 2012; Beikircher & Mayr 2013; Ellsworth & Sternberg 2015; Hölttä et al. 2018; Lintunen et al. 2021) and can account for up to 5% of annual water loss in temperate deciduous forests (Oren & Pataki 2001). During the vegetation period in Central Europe, bark transpiration reaches its highest values in late morning and lowest from 12 pm to 2 pm (Geurten 1950). Daily stem contractions induced by reversible tension-driven or osmotically-driven changes in bark water content (e.g., Chan et al. 2016; Zweifel et al. 2016; Mencuccini et al. 2017; Hölttä et al. 2018) may influence the diffusion resistance of bark. Required is an investigation into the regulatory mechanisms of lenticels and passive water-loss, i.e., a passive decrease in intercellular volume (“incipient drying”) from shrinkage, as well as active osmotically driven processes. Trees may be able to modify, to some degree, their lenticel size and total percentage area according to local climate. Size and percentage area of primary lenticels was found to be significantly higher in Picea abies trees of a similar height growing at higher elevations (1200 m) than non-autochthonous plantations at lower elevations (300 m) in Austria (Rosner 1998). The lower lenticel coverage (area %) in trees from the lowland site (1.9%) compared to the higher elevation site (2.7%) was interpreted as an adaptation to site-specific precipitation and air humidity; however, more research is required to test this hypothesis. The number of primary lenticels per given bark surface area in P. abies decreases with trunk diameter, but their size increases linearly, whereby the latter results in a significant increase in the lenticel area percentage (Rosner 1998). According to Geurten (1955), bark transpiration in P. abies increases with trunk diameter, either within an individual tree or when measured at the same height in trees with different ages and diameters at breast height. The size of primary lenticels may thus be a much better predictor of potential stem transpiration than lenticel quantity. More knowledge about the aforementioned relationships is required for a range of species, and to see if differences in lenticel size and lenticel coverage exist among provenances as well as to what extent they can adapt to local climate.
Functions beyond gas exchange: lenticels and crack prevention
The dead tissues outside the phellogen (i.e., the phellem, the filling tissue of the lenticels and the rhytidome) passively exchange water vapour with the atmosphere. This hygroscopic response can be observable daily and its magnitude is linked to the bulk density and porosity of the bark tissues (Ilek et al. 2021; Van Stan et al. 2021). Daily stem swelling and shrinkage are a challenge for the tree regarding the prevention of crack formation in the periderm. Moreover, heavy and long-lasting rainfall events following severe droughts during the growing season would result in crack formation if bark design was not adapted to such climate scenarios. We therefore postulate that the pattern of lenticel distribution is strongly linked to the prevention of cracks associated with bark hygroscopicity. Moreover, the different types of filling tissue arrangement in lenticels (Figs 4–6) and the spatial distribution of the tissues beneath them, e.g., collenchyma, sclereids and fibres, may be strongly linked to biomechanical functioning associated with dimensional changes. Lenticels can be aligned vertically (e.g., Tilia, Fig. 1a) or perpendicularly to the stem axis (e.g., Betula, Prunus, Fig. 1c–d). The upper and lower parts of vertically-aligned lenticels and the middle part of horizontally-aligned lenticels likely contain reinforced tissues either in the filling tissue or in the tissues beneath them. The hygroscopicity of the lenticels should also be strongly dependent on the quantity of unsuberized tissue, which differs tremendously among species (Figs 4–6). The rhytidome structure is largely dependent on the spatial arrangement of primary lenticels in the first-formed periderm, which varies considerably among taxa (Braun 1955). Little knowledge exists on the structure of secondary lenticels in the rhytidome along with the total area they cover (Wutz 1955). For Robinia pseudoacacia, Salix alba, Quercus robur and Pinus sylvestris, Wutz (1955) reported that the characteristics of the filling tissue of secondary lenticels do not differ greatly from primary lenticels, except that the number of cells is lower and lenticular phellogen is less active. In other words, lenticels of subsequent periderms may not have an annual growth rhythm as observed in primary lenticels of species from temperate zones. According to Rosner (1998), the percentage of primary lenticel area in the superficial periderm does not differ much from the secondary lenticel area in the rhytidome along trunks of Picea abies; both cover about 1–3% of total trunk area. However, when a thick rhytidome (with multiple subsequent periderms) has been formed, the biomechanical function of preventing shrinkage cracks due to the spatial arrangement of secondary lenticels presents a compelling trade-off to gas exchange and hydraulic functions. Novel micro-CT imaging techniques (e.g., Earles et al. 2016; Cuneo et al. 2018; Crouvisier-Urion et al. 2019) will be helpful to investigate buffering from the destructive effects of stem swelling and shrinkage.
Lenticel functioning in the stress operation range
Bark water loss during drought and heat waves
The intercellular spaces of the lenticels in plant stems form a continuous network (Fig. 2d) from the atmosphere to the bark and via the rays to the wood (Klebahn 1884; Rosner & Kartusch 2003). This continuous network of lenticels is advantageous during periods of sufficient water supply but might eventually cause severe stress during drought and heat wave events. While bark transpiration might be relatively small compared to the level of transpiration from the tree when water supply is sufficient and stomata are open, after drought-induced stomatal closure, this deficit may become critical (Wittmann & Pfanz 2008a; Lintunen et al. 2021; Van Stan et al. 2021). Increased bark transpiration has a cooling effect at the cost of water loss; accordingly, not only a lower number but also smaller lenticels tend to be found on sun-exposed sides of tree stems (Geurten 1950) and the percentage area of lenticels (measured in transverse sections) in the cork of Quercus suber L. decreases with stem height (Costa et al. 2021).
It is highly likely that waxes cover the inner surface of the lenticels — in particular at the interface to the atmosphere — to prevent the entrance of liquid water, since accessibility to gas molecules must be maintained even during rainfall (Lendzian 2006). In leaves, structural changes of the cuticular waxes that occur above a species specific phase transition temperature of between 32 and 38°C result in a steep increase in cuticular conductance and thus also in transpiration (Burghardt & Riederer 2006). Temperatures above 44°C result in irreparable damage of cuticular waxes and, coincidently, an increase in water permeability (Schönherr et al. 1979). Complete melting of cuticular waxes can be visually observed at temperatures ranging from 66 to 74°C. At such high temperatures, which are easily reached during forest fire events, increased volume expansion of the cutin polymer might cause additional defects to the wax barrier resulting in an increase in cuticular water permeability (Burghardt & Riederer 2006). The suberized phellem of Quercus suber has a relatively high thermal stability and low thermal conductivity (Pereira 2015). The insulating effect will depend on the thickness of the tissues outside the youngest phellogen and probably ultimately on the quantity of suberized filling tissue in the lenticel. It remains to be investigated if extremely high air temperatures and bark surface temperatures from direct sun exposure have a negative impact on suberin structure in the filling tissue that would eventually result in an increase in bark transpiration. The morphological structure of lenticels might help to prevent the bark beneath from localized heating because the loose filling tissue protruding from the periderm surface has a thermal insulating effect. In some species, lenticels are also not gateways for optimal light penetration into stems; transmittance is higher in the lenticel free periderm than in the lenticel regions (Manetas & Pfanz 2005). Moreover, the chloroplasts in the cortex tissue beneath the lenticels appeared more shade acclimated, with lower PSII efficiency and a reduced potential to dissipate excess light (Manetas & Pfanz 2005).
Carbon balance during drought and heat stress
Wetland trees are sources for greenhouse gases such as carbon dioxide, methane and nitrous oxide emitted from tree stems (Lendzian 2006; Pangala et al. 2014; Pitz et al. 2018; Rowland et al. 2018; Welch et al. 2019). Prolonged drought stress boosts carbon losses in addition to losses from drought-induced tree mortality. The worst case scenario would be a reversal of the global forest carbon sink (Rowland et al. 2018). More knowledge of species and age specific differences in periderm and lenticel structure would help when selecting trees that have better sink/source relationships regarding greenhouse gases. Drought- and heat-stressed trees show a higher rate of stem respiration and thus CO2 efflux if the stress occurs during periods of enhanced growth or from sapwood undergoing hydraulic recovery (Wittmann & Pfanz 2007; Rowland et al. 2018). Light-driven bark photosynthesis was found to level off or decrease at 35–40°C in temperate angiosperm species (Wittmann & Pfanz 2007). When a tree loses its leaves from the impact of drought stress or when stems are severed from their water supporting parts, i.e., the roots (e.g., cuttings used for clonal production), bark photosynthesis helps maintain water and sugar supply and promote organ development (Aschan et al. 2001; Liu et al. 2018). Stem CO2 efflux during bark and corticular photosynthesis is highest in young shoots and reduces with age (Wittmann & Pfanz 2008b; Tarvainen et al. 2014; Rowland et al. 2018). Bark photosynthesis is especially crucial for survival of young trees owing to less carbon storage reserves. Young trees may thus efficiently recycle or take up CO2 given their high quantity of chlorenchyma tissues and thinner periderm (Vandegehuchte et al. 2015). Young trees or branches of older trees show, however, a higher cambial activity, suggesting higher annual meristematic activity in lenticular phellogen. Early summer drought might result in an earlier production of the terminal layer of suberized filling tissue and/or the production of filling tissue with smaller intercellular spaces, thus influencing gas exchange. If early summer drought would result in an earlier production of a terminal layer of suberized filling tissue, lenticular phellogen might be reactivated when water becomes available in mid-to-late-summer.
Recovery from drought stress: evaporative enhancement
Lenticels may play a role in recovery from drought stress due to sapwood refilling by means of direct moisture uptake or bark transpiration. Such an evaporative enhancement due to bark transpiration might fulfil important physiological functions for recovery from embolization after summer drought events or in spring (Oren & Pataki 2001; Ellsworth & Sternberg 2015; Martín-Gómez et al. 2016; Hölttä et al. 2018). For instance, sap-flow in Quercus virginiana can be observed up to a fortnight before bud-break and one of the driving forces could be bark transpiration. The average daily water loss through bark in a temperate deciduous forest stand is about 0.02–0.03 mm (Oren & Pataki 2001). The distance from bark to the sapwood where conductivity loss occurs is short, facilitating embolism reversal and recovery from drought stress. Defoliation and leaf shading experiments using a range of conifer and deciduous species also indicate that isotopic enrichment of stem water can only be explained by transpiration via bark (Ellsworth & Sternberg 2015; Martín-Gómez et al. 2016). This so-called “evaporative enhancement” is lower through the bark of mature trees and it is suggested that the thin bark on younger branches and expanding buds may make a higher contribution to water vapour exchange with the atmosphere (Oren & Pataki 2001). Venturas et al. (2017) state in their review that a more holistic approach to xylem transport might include bark-xylem exchange processes in addition to long-distance axial transport.
Recovery from drought stress: water uptake via bark
Earles et al. (2016) observed that coastal redwood (Sequoia sempervirens) can rehydrate leaves via foliar and bark water uptake during fog/rain events. Moreover, Cuneo et al. (2018) reported that vessels of Vitis roots can be refilled via water uptake in regions where a periderm had been formed. Also, in Pinus halepensis, uptake of moisture via bark was found to occur during the night (Lintunen et al. 2021). Regarding the refilling of the secondary xylem via bark, symplastic transport via plasmodesmata (Earles et al. 2016; Secchi et al. 2017) might be possible, since bark water uptake rates correlate with xylem osmotic potential. In a living tree, intercellular spaces of lenticels are, however, not filled with water, but can be artificially infiltrated (Schönherr & Ziegler 1980). In experiments where solutions were sprayed on branches, the permeability of the periderm exceeded the uptake of water by needles; the main moisture uptake of a twig is therefore supposed to occur via the bark (Katz et al. 1989). Groh et al. (2002) reported that isolated angiosperm lenticels have a 10–40× higher permeance rate for liquid water than surrounding tissues. Extracting waxes from the filling tissue or the phellem did not result in a further increase of permeability. Water permeance of isolated Betula potaninii lenticels was found to be highest in July–August and lowest in September–April (Lendzian 2006). Another hint for xylem refilling from bark water uptake is that larger lenticels can be found on the “shaded-side” of a tree stem (Geurten 1950). Questions can be raised if a trade-off in uptake of liquid water with gas exchange is at all possible (Lendzian 2006). It remains uncertain if uptake of water is a controlled process after a period of severe summer or winter drought stress or if safety barriers exist analogous to the Casparian strip of the endodermis of young primary roots and conifer needles (Mayr et al. 2014); this needs to be investigated.
Hypoxia and hypertrophied lenticels
Photosynthesis in bark and woody tissues does not only play a role in the plant carbon economy (e.g., Vandegehuchte et al. 2015; Bloemen et al. 2016; De Baerdemaeker et al. 2017); it is also important for preventing low oxygen-limitations of respiration (Wittmann & Pfanz 2014; Wittmann & Pfanz 2018). Corticular photosynthesis can actively increase oxygen concentration, preventing living cells in tree stems succumbing to phases of daily oxygen shortage (Wittmann & Pfanz 2018). Severe oxygen shortage due to waterlogging (hypoxia) results in the development of hypertrophied lenticels (Ghouse & Yunus 1974). Warmer winters in northern latitudes in the near future will increase snow-melt frequency and rainfall, in turn increasing the risk of soil waterlogging (Wang et al. 2016). Flood tolerant species or genotypes are especially known to develop hypertrophied lenticels (e.g., Topa & McLoed 1986; Aronen & Häggman 1994; Parelle et al. 2007; Ruas et al. 2011; Bertolde et al. 2012; Le Provost et al. 2016; Wang et al. 2016; Yang & Li 2016). Wetland plants in particular are sources of methane emissions (e.g., Pangala et al. 2014; Pitz et al. 2018). Lenticel density (number/area) increases in plants growing at a high water table (Wang et al. 2016). In addition, the increase in lenticel density is related linearly to methane emissions from bark (Pangala et al. 2014). Bark anatomy beneath lenticels is expected to be modified by the degree of flood tolerance. The latter could indicate that where species can develop hypertrophied lenticels, they could have a lower number of sclerenchyma cells in the living bark beneath lenticels.
Lenticel functioning in the emergency operation range
The contribution of lenticels to the constitutive defence system and trade-offs with other functions
The anatomy of the living bark tissues, i.e., the phelloderm, cortex and secondary phloem, may vary considerably beneath lenticels and lenticel-free periderm (Wetmore 1926) in accordance with roles lenticels must fulfil for plants to function. While quantitative studies on the topic are scarce (Rosner & Führer 2002), some qualitative observations exist (Chan 1986; Langenfeld-Heyser et al. 1996; Graça & Pereira 2004). The loose arrangement of filling tissue may also represent an ‘Achilles heel’ to attacks by pathogenic fungi (Nemesio-Gorriz et al. 2019) and invasion by insects, e.g., bark beetles, due to the scarcity of resin canals beneath lenticels compared to lenticel-free periderm (Rosner & Führer 2002). Thus, a trade-off may exist between constitutive plant defence and gas exchange or moisture uptake. It is likely that bark beetles sense attracting and repelling chemical signals emanating through the lenticels (Franceschi et al. 2005). In boreal forests, high stem-born emission rates (single bursts lasting for 12 hours) of monoterpenes were observed in Pinus sylvestris shortly before the onset of vascular cambium production (Vanhatalo et al. 2015). Such monoterpene bursts could be a consequence of winter recovery and might be related to tracheid-refilling or to phenological changes in the epithelial cells surrounding resin ducts. Constitutive defence mechanisms in lenticels could be the accumulation of polyphenols in the filling tissue of lenticels to a higher extent than in the surrounding periderm. We found examples for higher accumulation of polyphenols in the filling tissue of all three lenticels types, e.g., in Salix caprea (Fig. 4g), Quercus petraea (Fig. 5d), Picea abies (Fig. 2b) and Betula pendula (Fig. 6c). Chan (1986) found crystals in the walls of lenticel tissues in Agathis australis. Inclusions of sclerified cells are mostly restricted to the lenticels, or in the nearby vicinity, in young Quercus suber stems (Graça & Pereira 2004). Lenticels are thus not necessarily open entry regions for insects and pathogens. For instance, root lenticels of P. abies are not more easily penetrable to the pathogenic fungus Heterobasidion annosum than surrounding periderm (Peek & Liese 1972). We still lack information on the role of lenticels in constitutive and inducible defences against insects and fungal/bacterial pathogens.
Emergency lenticels in wound response tissue (a–c) and next to a wound with strong resin flow (d). (a) Magnolia × soulangeana Soul.-Bod. (b) Malus sylvestris (L.) Mill. (c) Ginkgo biloba L. (d) Picea abies (L.) Karst. White arrows point to lenticels.
Citation: IAWA Journal 43, 3 (2022) ; 10.1163/22941932-bja10090
The role of lenticels in wound healing
The role of lenticels in wound healing has not yet been investigated. We postulate that lenticels play an important role in wood, living-bark and periderm repair processes. Accelerated metabolic processes associated with the formation of wound response tissue and the production of defence substances such as traumatic resin canals (Morris et al. 2020) might demand a higher level of aeration (Fig. 9a–c). In addition, extensive resin (conifers) or gum (angiosperms) flow from wounds can seal the periderm locally (Fig. 9d). The formation of new lenticels or increased meristematic activity of lenticular phellogen in such “compartmentalized regions” could be crucial to maintain gas exchange between bark and the atmosphere (Fig. 9d). Refixation of CO2 is also of high importance for carbon budgets during pathogen-induced leafless states in deciduous trees (Aschan et al. 2001). However, when bark is locally infected by pathogenic fungi (Pfanz et al. 2015) or when trees possess a large quantity of decayed wood, bark chlorenchyma has an extremely reduced photosynthetic capacity (Johnstone et al. 2014). Photosynthetic activity in the adjacent, healthy, regions should therefore be of high importance for the formation of reaction zones, i.e., dead parenchyma filled with antimicrobial polyphenolic compounds that form a barrier to restrict the spread of decay (Morris et al. 2020).
The contribution of lenticels to keep stems upright
Another emergency scenario is when trees are passively bent due to mechanical perturbation, e.g., by wind, or when they bend actively during competition for light. Trees form reaction wood to maintain biomechanical stability. In conifers, annual rings widen and the wood becomes dense on the leeward side (opposite direction of the stress) and is termed compression wood (Timell 1986). Extreme wind exposure results in permanent perturbation of the main stem. Picea abies grown on a wind-exposed site had bigger lenticels and a higher lenticel percentage area on the leeward side compared to the windward side of the stem (Rosner 1998). In angiosperms, tension wood is formed with specific anatomical characteristics (gelatinous layer in the secondary cell wall) on the windward side (Bamber 2001). In a recent study, Clair et al. (2019) found that living bark becomes thicker on the tension wood side, while underlining the important role of bark fibres for actively correcting bending and in regaining straight growth. Thicker living bark is also more prone to swelling and shrinking and a higher area of lenticels may fulfil the biomechanical function of crack prevention. The role of lenticels for keeping stems upright needs to be investigated since bark photosynthesis demands greater gas permeability.
Conclusions and outlook
Research on structure-function relationships of the secondary dermal tissues (periderm, rhytidome, lenticels) with the goal of learning how trees cope with extreme climatic and emergency scenarios requires greater attention to understand why some tree species survive and others not. The role of lenticels for tree survival in particular has so far been overlooked. The bark of tree stems, branches and roots covers several — sometimes competing — functions necessary for tree survival. The periderm provides armour against desiccation, heat and mechanical injury. However, photosynthetically active cells in the phelloderm, the cortex and non-conducting secondary phloem demand gas exchange between them and the atmosphere, which is achieved through lenticels. There is only scarce knowledge about the functional trade-offs in bark tissues and, as far as we know, no existing knowledge exists on how lenticels can be modified in accordance with physiological and biomechanical demands during a trees’ life cycle. If these basic relationships are known, we will be in a better position to understand survival strategies of trees to different stressors. Classical light microscopy and histological staining together with cutting-edge high-resolution 3-dimensional imaging techniques and up-to-date imaging technologies such as confocal Raman spectroscopy will offer new opportunities to investigate anatomy and chemistry of lenticels and related tissues. Establishing the new research discipline “Phytodermatology” (ancient Greek: phyton, plant; derma, skin; logia, science or study of) will help to fill knowledge gaps regarding tree survival by linking quantitative and qualitative periderm/rhytidome and lenticel anatomy to tree hydraulics and biomechanics. The new discipline should aim towards relating lenticel morphology, anatomy and ontogeny to tree survival under different stress scenarios, including the normal operation range during a tree’s life cycle, alongside the stress and emergency operation ranges. Regarding the normal operation range, we need information on: (a) how the permeability of the periderm varies among species as well as within an individual tree, (b) if the variability among species in the stratification of filling tissue and cell wall characteristics give clues to the opening mechanisms of lenticels, (c) how the annual activity of lenticular phellogen is timely linked with cambial activity and (d) if the permeability of lenticels for gaseous substances and water changes over seasons and how this permeability influences the hydration status of bark. Research on the stress operation range should unravel (a) how lenticels modify their stratification and structure under the impact of drought stress, (b) if the activity of lenticular phellogen is crucial for water supply after periods of drought stress, especially after leaf loss, and (c) how bark anatomy beneath lenticels is modified by the degree of flood tolerance. Concerning the operation for/under emergency including constitutive defence, wound healing and bending, we lack knowledge about: (a) the differences in the quantity of insect repellents and fungicides in bark tissue beneath lenticels and the surrounding periderm, (b) whether cells of the filling tissue contain more polyphenols than cells of phellem and (c) whether lenticel density is higher in both wound response tissue and reaction wood. The proposed Phytodermatology research will demand interdisciplinary work among plant anatomists, plant taxonomists, plant physiologists, bio-physicists, ecologists, foresters and tree breeders. Future research on the structure-function relationships of lenticels should be done on trees and shrubs from different biomes to gain greater knowledge on how conserved lenticel types are within monophyletic clades.
Corresponding author; email: Sabine.rosner@boku.ac.at
Acknowledgement
We thank Pieter Baas, Marcelo R. Pace and an anonymous reviewer for their valuable input.
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Footnotes
Edited by Marcelo R. Pace