TY - JOUR
AU - Flexas,, Jaume
AB - Abstract In vascular plants, more rigid leaves have been linked to lower photosynthetic capacity, associated with low CO2 diffusion across the mesophyll, indirectly resulting in a trade-off between photosynthetic capacity (An) and bulk modulus of elasticity (ε). However, we evaluated mosses, liverworts, and Chara sp., plus some lycophytes and ferns, and found that they behaved as clear outliers of the An–ε relationship. Despite this finding, when vascular and non-vascular plants were plotted together, ε still linearly determined the cessation of net photosynthesis during desiccation both in species with stomata (either actively or hydro-passively regulated) and in species lacking stomata, and regardless of their leaf structure. The latter result challenges our current view of photosynthetic responses to desiccation and/or water stress. Structural features and hydric strategy are discussed as possible explanations for the deviation of these species from the An–ε trade-off, as well as for the general linear dependency between ε and the full cessation of An during desiccation. Capacitance, cell wall, CO2 assimilation, desiccation, hydric strategy, modulus of elasticity, non-vascular plants, photosynthetic capacity, stomatal regulation Introduction Metabolic features of photosynthetic organs have been linked with structural and nutritional traits along a wide range of plants, defining the so-called ‘leaf economic spectrum’ (LES) (Reich et al., 1992; Wright et al., 2004, 2005; Tosens et al., 2015). Additionally, structural parameters at the subcellular level, such as mesophyll cell wall thickness (Tcw), chloroplast distribution, and other mesophyll traits, have been shown to explain at least part of the LES relationships (Onoda et al., 2017). Furthermore, a novel trade-off between light-saturated net CO2 assimilation (An) and the bulk modulus of elasticity (ε) has been described recently for vascular plants (Nadal et al., 2018), showing that high An is incompatible with rigid leaves. In addition, He et al. (2019) have shown a negative relationship between photosynthetic capacity and leaf toughness, assessed as the force required to tear leaves. However, the mechanistic bases of the An–ε trade-off have not been fully elucidated yet. Since Tcw largely constrains photosynthesis through mesophyll conductance (gm; Evans et al., 2009; Terashima et al., 2011; Onoda et al., 2017), cell wall traits have been suggested to be involved in the An–ε trade-off (Nadal et al., 2018), but other factors may interfere; these include the structure of the whole leaf, cuticle thickness, distribution of fibers, and the arrangement of the often sclerified leaf veins (Onoda et al., 2015; He et al. 2019). Recent studies have found that bryophytes show analogous correlations for the same LES traits described for tracheophytes, although with largely different interception points (Waite and Sack, 2010; Wang et al., 2014; Carriquí et al., 2019; Gago et al., 2019). However, the possible occurrence of a relationship between An and ε in bryophytes has not been tested. Bryophytes usually have low ε (Proctor et al., 1998; Proctor, 1999) and low An compared with other plant groups (Martin and Adamson, 2001; Brodribb et al., 2007; Waite and Sack, 2010; Wang et al., 2014; Carriquí et al., 2019). Therefore, we expect bryophytes to be outliers of the An–ε trade-off. Additionally, the role of ε in the regulation of leaf water relations has not been fully clarified. High ε is usually associated with sclerophylly (Salleo and Lo Gullo, 1990) and has been suggested to improve water uptake from drying soils (Bowman and Roberts, 1985), although ε does not always increase in the dry season, that is, it does not always acclimate (Davis and Mooney 1986; Scholz et al., 2012; but see also Salleo and Lo Gullo, 1990). However, in a comprehensive meta-analysis, Bartlett et al. (2012) proposed a role of high ε in preventing water loss upon reaching the turgor loss point [associated with high relative water content at turgor loss point (RWCTLP)]. On the other hand, low ε allows for a high degree of leaf shrinkage (Scoffoni et al., 2014), possibly related to the ability to support fluctuations in water content and the maintenance of rehydration capacity (John et al., 2018). In bryophytes, a possible relationship between desiccation tolerance and ε has also been suggested, since high elasticity would avoid mechanical stress on cells during dehydration/rehydration cycles (Proctor et al., 1998, 2007; Proctor, 2001). However, the components of ε in both groups of plants must be quite different. Features such as thickened cuticles, lignified epidermal cells, fiber bundles, and the presence of sclereids, which seem to partially determine rigidity in the leaf blades of vascular plants (Salleo and Lo Gullo, 1990), are absent in the gametophyte of bryophytes, although proto-cuticle and lignin-like components have been reported in some specific groups (Cook and Graham, 1998; Vitt and Wieder, 2009). This paper provides evidence for the divergence of bryophytes, charophytes, and some lycophytes and ferns from the An–ε trade-off reported for most tracheophytes, and discusses possible explanations in terms of their differences in anatomy and hydric strategy. Furthermore, the effect of ε in water relations and maintenance of photosynthesis upon dehydration spanning the whole range of phylogeny of land plants is also reported, in order to disentangle the relationship between tissue elasticity and dehydration dynamics. To achieve these goals, measurements of gas exchange and pressure–volume curves were performed for species of angiosperms, bryophytes, charophytes, lycophytes, and filmy ferns (Hymenophyllum spp.), and compared with the data previously reported by Nadal et al. (2018). Besides bryophytes and other groups behaving as outliers of the previously reported trade-off, a surprising and challenging finding is that ε apparently links linearly with the relative water content (RWC) at full cessation of photosynthesis when all species are pooled together, both those that have and do not have stomata, and regardless of their leaf structure, vascularity, hydric strategy, and desiccation tolerance. Materials and methods Plant materials and growing conditions Twenty-six species were analyzed in this work (Table 1): two angiosperms, four filmy ferns, nine mosses, six liverworts, one hornwort, three clubmosses, and one charophyte. All species except Nicotiana tabacum, Craterostigma plantagineum, Huperzia squarrosa, and Isoetes velata were collected in the field. Bryophytes, Chara sp., and Selaginella denticulata were collected in Mallorca (Balearic Islands) or Tenerife (Canary Islands). Hymenophyllum spp. were collected also in the field in Parque Katalapi (Chile). Nicotiana tabacum, C. plantagineum, H. squarrosa, and I. velata were obtained from commercial suppliers. Nicotiana tabacum (cv. Petit Havana) and C. plantagineum were planted in pots (2 litres) in a growth chamber at 23 °C and with a 12 h /12 h photoperiod. Isoetes velata was maintained in the same growth chamber with the roots immersed in a wet sponge. Aquatic specimens of Chara sp. and Fontinalis antipyretica were stored in an aquarium and measured within 1 week. The rest of the species were kept in a wet (watered daily with deionized water) and shaded greenhouse [photosynthetic photon flux density (PPFD) reaching a maximum of 200 µmol m–2 s–1] for at least 1 week before measurements were made. The mosses, liverworts, hornwort, and S. denticulata were stored in perforated trays with native substrate. Filmy ferns and H. squarrosa were stored in a suspended nest with a wide layer of mosses (a mixture of Thuidium tamariscinum, Pseudoscleropodium purum, and Polytrichastrum formosum) as substrate. The nine angiosperm species and three ferns grown as reported in Nadal et al. (2018) were also used for desiccation assays (see below). Table 1. Phylogeny and habitat of the studied species, and measurements performed (indicated with asterisks) Species . Phylum . Family . Habitat a . Gas exchange and P–V curves b . Desiccation assay . Anthoceros agrestis Paton Anthocerotophyta Anthocerotaceae Wet slopes by streams or springs * Ctenidium molluscum (Hedw.) Mitt. Bryophyta Hypnaceae On soils and calcareous rocks * Fissidens serrulatus Bird. Bryophyta Fissidentaceae Stream margins, soils and rocks * * Fontinalis antipyretica Hedw. Bryophyta Fontinalaceae Submerged in streams * Plagiomnium undulatum (Hedw.) T.J. Kop. Bryophyta Plagiomniaceae Damp or waterlogged soils * * Polytrichastrum formosum (Hedw.) G.L. Sm. Bryophyta Polytrichaceae Slopes and damp, shady soils * Polytrichum juniperinum Hedw. Bryophyta Polytrichaceae Exposed, acidic soils and slopes * Pseudoscleropodium purum (Hedw.) M. Fleisch. Bryophyta Brachytheciaceae Forest soils, in the lowlands * * Sphagnum sp. L. Bryophyta Sphagnaceae Peatlands * * Thuidium tamariscinum (Hedw.) Schimp. Bryophyta Thuidiaceae Wet, shaded rocks, soils and wood * * Chara sp. L. Charophyta Characeae Hard water or alkaline ponds * Huperzia squarrosa (G. Forst) Trevis Lycopodiophyta Lycopodiaceae Epiphyte, lithophyte, on infertile soils * * Isoetes velata A. Braun Lycopodiophyta Isoetaceae Wetland habitats * * Selaginella denticulata (L.) Spring Lycopodiophyta Selaginellaceae Wet slopes by streams or springs * * Barbacenia purpurea Hook. Magnoliophyta Velloziaceae Rocky, tropical areas * Craterostigma plantagineum Hochst. Magnoliophyta Scrophulariaceae Shallow rocky soils * * Helianthus annuus L. Magnoliophyta Asteraceae Subtropical, arable land * Nicotiana tabacum L. (cv. Petit Havanan) Magnoliophyta Solanaceae Crop * * Olea europaea var sylvestris L. Magnoliophyta Oleaceae Subtropical, Mediterranean shrublands * Populus nigra L. Magnoliophyta Salicaceae Subtropical/temperate forests * Quercus ilex L. Magnoliophyta Fagaceae Subtropical, Mediterranean forests * Syringa vulgaris L. Magnoliophyta Oleaceae Temperate, rocky shrubland * Triticum aestivum L. Magnoliophyta Poaceae Subtropical, arable land * Vitis vinifera cv. Tempranillo L. Magnoliophyta Vitaceae Subtropical/temperate arable land * Xerophyta viscosa Baker. Magnoliophyta Velloziaceae Rocky, subtropical/arid areas * Conocephalum conicum (L.) Dumort Marchantiophyta Conocephalaceae Rocks and wet soils * Dumortiera hirsuta (Sw.) Nees Marchantiophyta Marchantiaceae Soils or wet rocks near water * Lunularia cruciata (L.) Lindb Marchantiophyta Lunulariaceae Soil, wet rocks and slopes * * Pellia endiviifolia (Dicks.) Dumort. Marchantiophyta Pelliaceae Streams, wet slopes or submerged * * Porella canariensis (F.Weber) Underw. Marchantiophyta Porellaceae Rocks and bark * Saccogyna viticulosa (L.) Dumort. Marchantiophyta Geocalycaceae Shaded, humus-rich soils, calcifuge * Davallia canariensis (L.) Sm. Pteridophyta Davalliaceae Subtropical/tropical forest * Hymenophyllum caudiculatum Mart. Pteridophyta Hymenophyllaceae Epiphyte on wood * * Hymenophyllum dentatum Cav. Pteridophyta Hymenophyllaceae Rocks and decayed wood * * Hymenophyllum dicranotrichum (C. Presl) Sadeb. Pteridophyta Hymenophyllaceae Epiphyte on wood * * Hymenophyllum pectinatum Cav. Pteridophyta Hymenophyllaceae Rocks, organic soils and wood * * Nephrolepis exaltata (L.) Schott Pteridophyta Lomariopsidaceae Subtropical/temperate forest * Phlebodium aureum (L.) J. Sm. Pteridophyta Polypodiaceae Subtropical/tropical forest * Species . Phylum . Family . Habitat a . Gas exchange and P–V curves b . Desiccation assay . Anthoceros agrestis Paton Anthocerotophyta Anthocerotaceae Wet slopes by streams or springs * Ctenidium molluscum (Hedw.) Mitt. Bryophyta Hypnaceae On soils and calcareous rocks * Fissidens serrulatus Bird. Bryophyta Fissidentaceae Stream margins, soils and rocks * * Fontinalis antipyretica Hedw. Bryophyta Fontinalaceae Submerged in streams * Plagiomnium undulatum (Hedw.) T.J. Kop. Bryophyta Plagiomniaceae Damp or waterlogged soils * * Polytrichastrum formosum (Hedw.) G.L. Sm. Bryophyta Polytrichaceae Slopes and damp, shady soils * Polytrichum juniperinum Hedw. Bryophyta Polytrichaceae Exposed, acidic soils and slopes * Pseudoscleropodium purum (Hedw.) M. Fleisch. Bryophyta Brachytheciaceae Forest soils, in the lowlands * * Sphagnum sp. L. Bryophyta Sphagnaceae Peatlands * * Thuidium tamariscinum (Hedw.) Schimp. Bryophyta Thuidiaceae Wet, shaded rocks, soils and wood * * Chara sp. L. Charophyta Characeae Hard water or alkaline ponds * Huperzia squarrosa (G. Forst) Trevis Lycopodiophyta Lycopodiaceae Epiphyte, lithophyte, on infertile soils * * Isoetes velata A. Braun Lycopodiophyta Isoetaceae Wetland habitats * * Selaginella denticulata (L.) Spring Lycopodiophyta Selaginellaceae Wet slopes by streams or springs * * Barbacenia purpurea Hook. Magnoliophyta Velloziaceae Rocky, tropical areas * Craterostigma plantagineum Hochst. Magnoliophyta Scrophulariaceae Shallow rocky soils * * Helianthus annuus L. Magnoliophyta Asteraceae Subtropical, arable land * Nicotiana tabacum L. (cv. Petit Havanan) Magnoliophyta Solanaceae Crop * * Olea europaea var sylvestris L. Magnoliophyta Oleaceae Subtropical, Mediterranean shrublands * Populus nigra L. Magnoliophyta Salicaceae Subtropical/temperate forests * Quercus ilex L. Magnoliophyta Fagaceae Subtropical, Mediterranean forests * Syringa vulgaris L. Magnoliophyta Oleaceae Temperate, rocky shrubland * Triticum aestivum L. Magnoliophyta Poaceae Subtropical, arable land * Vitis vinifera cv. Tempranillo L. Magnoliophyta Vitaceae Subtropical/temperate arable land * Xerophyta viscosa Baker. Magnoliophyta Velloziaceae Rocky, subtropical/arid areas * Conocephalum conicum (L.) Dumort Marchantiophyta Conocephalaceae Rocks and wet soils * Dumortiera hirsuta (Sw.) Nees Marchantiophyta Marchantiaceae Soils or wet rocks near water * Lunularia cruciata (L.) Lindb Marchantiophyta Lunulariaceae Soil, wet rocks and slopes * * Pellia endiviifolia (Dicks.) Dumort. Marchantiophyta Pelliaceae Streams, wet slopes or submerged * * Porella canariensis (F.Weber) Underw. Marchantiophyta Porellaceae Rocks and bark * Saccogyna viticulosa (L.) Dumort. Marchantiophyta Geocalycaceae Shaded, humus-rich soils, calcifuge * Davallia canariensis (L.) Sm. Pteridophyta Davalliaceae Subtropical/tropical forest * Hymenophyllum caudiculatum Mart. Pteridophyta Hymenophyllaceae Epiphyte on wood * * Hymenophyllum dentatum Cav. Pteridophyta Hymenophyllaceae Rocks and decayed wood * * Hymenophyllum dicranotrichum (C. Presl) Sadeb. Pteridophyta Hymenophyllaceae Epiphyte on wood * * Hymenophyllum pectinatum Cav. Pteridophyta Hymenophyllaceae Rocks, organic soils and wood * * Nephrolepis exaltata (L.) Schott Pteridophyta Lomariopsidaceae Subtropical/temperate forest * Phlebodium aureum (L.) J. Sm. Pteridophyta Polypodiaceae Subtropical/tropical forest * a Based on: Casas et al. (2006, 2009) for bryophytes; Yumkham and Singh (2011), Rhazi et al. (2009), and Tryon and Tryon (1982) for Lycopodiaceae; Larsen et al. (2013) for Hymenophyllaceae; Sheath and Wehr (2015) for Characeae; Heath and Heath (2009) for Craterostigma; and Nadal et al. (2018) for the rest of the angiosperms and ferns. b Asterisks indicate newly measured species. Gas exchange and pressure–volume (P–V) curve data for the other species were obtained from Nadal et al. (2018). Open in new tab Table 1. Phylogeny and habitat of the studied species, and measurements performed (indicated with asterisks) Species . Phylum . Family . Habitat a . Gas exchange and P–V curves b . Desiccation assay . Anthoceros agrestis Paton Anthocerotophyta Anthocerotaceae Wet slopes by streams or springs * Ctenidium molluscum (Hedw.) Mitt. Bryophyta Hypnaceae On soils and calcareous rocks * Fissidens serrulatus Bird. Bryophyta Fissidentaceae Stream margins, soils and rocks * * Fontinalis antipyretica Hedw. Bryophyta Fontinalaceae Submerged in streams * Plagiomnium undulatum (Hedw.) T.J. Kop. Bryophyta Plagiomniaceae Damp or waterlogged soils * * Polytrichastrum formosum (Hedw.) G.L. Sm. Bryophyta Polytrichaceae Slopes and damp, shady soils * Polytrichum juniperinum Hedw. Bryophyta Polytrichaceae Exposed, acidic soils and slopes * Pseudoscleropodium purum (Hedw.) M. Fleisch. Bryophyta Brachytheciaceae Forest soils, in the lowlands * * Sphagnum sp. L. Bryophyta Sphagnaceae Peatlands * * Thuidium tamariscinum (Hedw.) Schimp. Bryophyta Thuidiaceae Wet, shaded rocks, soils and wood * * Chara sp. L. Charophyta Characeae Hard water or alkaline ponds * Huperzia squarrosa (G. Forst) Trevis Lycopodiophyta Lycopodiaceae Epiphyte, lithophyte, on infertile soils * * Isoetes velata A. Braun Lycopodiophyta Isoetaceae Wetland habitats * * Selaginella denticulata (L.) Spring Lycopodiophyta Selaginellaceae Wet slopes by streams or springs * * Barbacenia purpurea Hook. Magnoliophyta Velloziaceae Rocky, tropical areas * Craterostigma plantagineum Hochst. Magnoliophyta Scrophulariaceae Shallow rocky soils * * Helianthus annuus L. Magnoliophyta Asteraceae Subtropical, arable land * Nicotiana tabacum L. (cv. Petit Havanan) Magnoliophyta Solanaceae Crop * * Olea europaea var sylvestris L. Magnoliophyta Oleaceae Subtropical, Mediterranean shrublands * Populus nigra L. Magnoliophyta Salicaceae Subtropical/temperate forests * Quercus ilex L. Magnoliophyta Fagaceae Subtropical, Mediterranean forests * Syringa vulgaris L. Magnoliophyta Oleaceae Temperate, rocky shrubland * Triticum aestivum L. Magnoliophyta Poaceae Subtropical, arable land * Vitis vinifera cv. Tempranillo L. Magnoliophyta Vitaceae Subtropical/temperate arable land * Xerophyta viscosa Baker. Magnoliophyta Velloziaceae Rocky, subtropical/arid areas * Conocephalum conicum (L.) Dumort Marchantiophyta Conocephalaceae Rocks and wet soils * Dumortiera hirsuta (Sw.) Nees Marchantiophyta Marchantiaceae Soils or wet rocks near water * Lunularia cruciata (L.) Lindb Marchantiophyta Lunulariaceae Soil, wet rocks and slopes * * Pellia endiviifolia (Dicks.) Dumort. Marchantiophyta Pelliaceae Streams, wet slopes or submerged * * Porella canariensis (F.Weber) Underw. Marchantiophyta Porellaceae Rocks and bark * Saccogyna viticulosa (L.) Dumort. Marchantiophyta Geocalycaceae Shaded, humus-rich soils, calcifuge * Davallia canariensis (L.) Sm. Pteridophyta Davalliaceae Subtropical/tropical forest * Hymenophyllum caudiculatum Mart. Pteridophyta Hymenophyllaceae Epiphyte on wood * * Hymenophyllum dentatum Cav. Pteridophyta Hymenophyllaceae Rocks and decayed wood * * Hymenophyllum dicranotrichum (C. Presl) Sadeb. Pteridophyta Hymenophyllaceae Epiphyte on wood * * Hymenophyllum pectinatum Cav. Pteridophyta Hymenophyllaceae Rocks, organic soils and wood * * Nephrolepis exaltata (L.) Schott Pteridophyta Lomariopsidaceae Subtropical/temperate forest * Phlebodium aureum (L.) J. Sm. Pteridophyta Polypodiaceae Subtropical/tropical forest * Species . Phylum . Family . Habitat a . Gas exchange and P–V curves b . Desiccation assay . Anthoceros agrestis Paton Anthocerotophyta Anthocerotaceae Wet slopes by streams or springs * Ctenidium molluscum (Hedw.) Mitt. Bryophyta Hypnaceae On soils and calcareous rocks * Fissidens serrulatus Bird. Bryophyta Fissidentaceae Stream margins, soils and rocks * * Fontinalis antipyretica Hedw. Bryophyta Fontinalaceae Submerged in streams * Plagiomnium undulatum (Hedw.) T.J. Kop. Bryophyta Plagiomniaceae Damp or waterlogged soils * * Polytrichastrum formosum (Hedw.) G.L. Sm. Bryophyta Polytrichaceae Slopes and damp, shady soils * Polytrichum juniperinum Hedw. Bryophyta Polytrichaceae Exposed, acidic soils and slopes * Pseudoscleropodium purum (Hedw.) M. Fleisch. Bryophyta Brachytheciaceae Forest soils, in the lowlands * * Sphagnum sp. L. Bryophyta Sphagnaceae Peatlands * * Thuidium tamariscinum (Hedw.) Schimp. Bryophyta Thuidiaceae Wet, shaded rocks, soils and wood * * Chara sp. L. Charophyta Characeae Hard water or alkaline ponds * Huperzia squarrosa (G. Forst) Trevis Lycopodiophyta Lycopodiaceae Epiphyte, lithophyte, on infertile soils * * Isoetes velata A. Braun Lycopodiophyta Isoetaceae Wetland habitats * * Selaginella denticulata (L.) Spring Lycopodiophyta Selaginellaceae Wet slopes by streams or springs * * Barbacenia purpurea Hook. Magnoliophyta Velloziaceae Rocky, tropical areas * Craterostigma plantagineum Hochst. Magnoliophyta Scrophulariaceae Shallow rocky soils * * Helianthus annuus L. Magnoliophyta Asteraceae Subtropical, arable land * Nicotiana tabacum L. (cv. Petit Havanan) Magnoliophyta Solanaceae Crop * * Olea europaea var sylvestris L. Magnoliophyta Oleaceae Subtropical, Mediterranean shrublands * Populus nigra L. Magnoliophyta Salicaceae Subtropical/temperate forests * Quercus ilex L. Magnoliophyta Fagaceae Subtropical, Mediterranean forests * Syringa vulgaris L. Magnoliophyta Oleaceae Temperate, rocky shrubland * Triticum aestivum L. Magnoliophyta Poaceae Subtropical, arable land * Vitis vinifera cv. Tempranillo L. Magnoliophyta Vitaceae Subtropical/temperate arable land * Xerophyta viscosa Baker. Magnoliophyta Velloziaceae Rocky, subtropical/arid areas * Conocephalum conicum (L.) Dumort Marchantiophyta Conocephalaceae Rocks and wet soils * Dumortiera hirsuta (Sw.) Nees Marchantiophyta Marchantiaceae Soils or wet rocks near water * Lunularia cruciata (L.) Lindb Marchantiophyta Lunulariaceae Soil, wet rocks and slopes * * Pellia endiviifolia (Dicks.) Dumort. Marchantiophyta Pelliaceae Streams, wet slopes or submerged * * Porella canariensis (F.Weber) Underw. Marchantiophyta Porellaceae Rocks and bark * Saccogyna viticulosa (L.) Dumort. Marchantiophyta Geocalycaceae Shaded, humus-rich soils, calcifuge * Davallia canariensis (L.) Sm. Pteridophyta Davalliaceae Subtropical/tropical forest * Hymenophyllum caudiculatum Mart. Pteridophyta Hymenophyllaceae Epiphyte on wood * * Hymenophyllum dentatum Cav. Pteridophyta Hymenophyllaceae Rocks and decayed wood * * Hymenophyllum dicranotrichum (C. Presl) Sadeb. Pteridophyta Hymenophyllaceae Epiphyte on wood * * Hymenophyllum pectinatum Cav. Pteridophyta Hymenophyllaceae Rocks, organic soils and wood * * Nephrolepis exaltata (L.) Schott Pteridophyta Lomariopsidaceae Subtropical/temperate forest * Phlebodium aureum (L.) J. Sm. Pteridophyta Polypodiaceae Subtropical/tropical forest * a Based on: Casas et al. (2006, 2009) for bryophytes; Yumkham and Singh (2011), Rhazi et al. (2009), and Tryon and Tryon (1982) for Lycopodiaceae; Larsen et al. (2013) for Hymenophyllaceae; Sheath and Wehr (2015) for Characeae; Heath and Heath (2009) for Craterostigma; and Nadal et al. (2018) for the rest of the angiosperms and ferns. b Asterisks indicate newly measured species. Gas exchange and pressure–volume (P–V) curve data for the other species were obtained from Nadal et al. (2018). Open in new tab Photosynthesis measurements An was determined by gas exchange measurements using a GFS-3000 system coupled with an IMAGING-PAM fluorometer (Heinz Walz, Effeltrich, Germany), except for C. plantagineum, which was measured with Li-6400 gas exchange equipment (Li-Cor Inc., Lincoln, NE, USA) as described in Nadal et al. (2018). Fully hydrated non-tracheophytes (n=5–8) were measured with a custom-made moss cuvette following the method of Perera-Castro et al. (2020), avoiding overlapping of shoots (see Supplementary Fig. S1 at JXB online). The cuvette consisted of a gasket stuck to a thin piece of polyester cloth. The measured CO2 leakage of an empty chamber was not significantly different with or without the custom-made moss cuvette being in place. The CO2 concentration was kept at 400 μmol CO2 mol–1 air, relative humidity at 75–85%, blue light PPFD at saturation level (100–1000 µmol m–2 s–1, depending on the species, tested previously with An–PPFD curves), and temperature at 25 °C. The flow rate within the chamber was 750 µmol s–1. An was recorded at steady-state conditions, when diffusional limitations due to external water were null and biochemistry was fully light-adapted (5–20 min; Supplementary Fig. S2), and was normalized to the measured leaf/shoot/thallus area. Pressure–volume curves Pressure–volume curves were performed by alternately weighing and measuring the leaf/shoot/thallus for water potential using a psychrometer (model WP4C, Decagon Device Inc., Pullman, WA, USA), except for H. squarrosa and C. plantagineum, which were measured with a pressure chamber (Model 1000; PMS Instrument Company, Albany, OR, USA). Fully hydrated samples were slowly air-dried in order to define the turgor loss point accurately. ε was calculated as the slope of pressure potential (Ψ P) versus total RWC as described by Sack and Pasquet-Kok (2011). RWC was estimated as RWC=100(FW–DW)/(TW–DW), where FW is the fresh weight of the sample at any time on the curve, DW is the dry constant weight obtained after keeping the samples at 70 °C for 2–3 days, and TW is the turgid weight. TW was estimated as the x-intercept of leaf water potential (Ψ leaf) versus the mass of water (FW–DW) (see Supplementary Fig. S3 for cross-validation between this method to estimate TW and the method for the desiccation assay, described below). Absolute leaf capacitance at full turgor (Cleaf) was determined from the initial slope of the relationship between water potential (Ψ leaf) and RWC, and was normalized by leaf/shoot/thallus area: Cleaf=(TW–DW)ΔRWC/(ΔΨ leaf×area), as described by Sack and Pasquet-Kok (2011). Desiccation assay To test the response of photosynthesis to water status, the RWC at which An reaches 0 μmol CO2 m–2 s–1 (RWC0) was measured in some of the studied species (Table 1). The response of An to RWC was monitored in excised leaves (for vascular species) or detached shoots/thalli (for bryophytes, lycophytes, and filmy ferns) by alternately measuring gas exchange and FW until An reached 0 μmol CO2 m–2 s–1. Examples of An versus RWC curves during desiccation assays are shown in Supplementary Fig. S4. In order to avoid overestimation of TW in bryophytes and filmy ferns, external water was removed by gently shaking the sample and then absorbing water on the surface by pressing it against a dry paper napkin. This method of TW estimation showed a high correlation (r=0.984, P<0.000) that did not differ significantly from a 1:1 relationship with TW estimated as the x-intercept of Ψ leaf versus the mass of non-external water (FW–DW) (Supplementary Fig. S3). Statistical analysis The relationship between An and ε was tested by Pearson’s correlation coefficient. Linear or logarithmic fitting was applied to the relationships between RWC0, ε, and RWCTLP. Mean ε values of angiosperms and ferns, except N. tabacum, C. plantagineum, and filmy ferns, were taken from Nadal et al. (2018). All analyses were performed using R statistical software (R Core Team, 2016). Results and discussion Bryophytes, charophytes, and some lycophytes and ferns behave as clear outliers of the previously reported An–ε relationship No significant relationship between An and ε was found within either the mosses (r=0.069, P=0.859) or liverworts (r=0.586, P=0.221) (Fig. 1A). Furthermore, all the studied bryophytes, as well as the species representing lycophytes, charophytes, and filmy ferns, were positioned as outliers in the An versus ε plot reported by Nadal et al. (2018), except the lycophyte I. velata (Fig. 1B). Fig. 1. Open in new tabDownload slide (A) Relationship between net CO2 assimilation (An) and modulus of elasticity (ε) of mosses, liverworts, hornwort, clubmosses (except Isoetes velata), and charophyte green algae. (B) Contextualization of (A) in the relationship between both parameters described for angiosperms and non-filmy ferns by Nadal et al. (2018) (translucent points), with the addition of Nicotiana tabacum, Craterostigma plantagineum, and I. velata. Arrows indicate the ‘resurrection plants’ C. plantagineum, Barbacenia purpurea, and Xerophyta viscosa. Data for I. velata are included (red point close to the general relationship for vascular species). Dashed line: y= –0.54 + 22.68x, r= –0.641, P<0.0001. Points represent the mean ±SE for each species (n=5–7). Fig. 1. Open in new tabDownload slide (A) Relationship between net CO2 assimilation (An) and modulus of elasticity (ε) of mosses, liverworts, hornwort, clubmosses (except Isoetes velata), and charophyte green algae. (B) Contextualization of (A) in the relationship between both parameters described for angiosperms and non-filmy ferns by Nadal et al. (2018) (translucent points), with the addition of Nicotiana tabacum, Craterostigma plantagineum, and I. velata. Arrows indicate the ‘resurrection plants’ C. plantagineum, Barbacenia purpurea, and Xerophyta viscosa. Data for I. velata are included (red point close to the general relationship for vascular species). Dashed line: y= –0.54 + 22.68x, r= –0.641, P<0.0001. Points represent the mean ±SE for each species (n=5–7). Based on these results, it might be proposed that structural limitations to An in tracheophytes must be driven by one or more traits that might be absent in the outlier groups and/or might be modulated by additional constraints. The reported structural and physiological differences among the phylogenetic groups are summarized in Table 2. At first sight, none of the presented traits can be unequivocally assigned to be responsible for the departure from the previously established An–ε trade-off. Some traits—for example, Tcw, which was suggested to be involved in the aforementioned trade-off (Nadal et al., 2018), the presence of lignin, the frequency of desiccation tolerance, and the thickness of the cuticle—change gradually along phylogeny. The studied ‘resurrection plant’ angiosperms, C. plantagineum, Barbacenia purpurea, and Xerophyta viscosa, combine elastic tissues with high An, like the rest of the angiosperms. The presence of stomata and a vascular system is not essential to link An and ε, as is evidenced by the lycophytes H. squarrosa and S. denticulata, which behave like moss gametophytes in the An–ε relationship (combining low photosynthetic capacity and elastic tissues) despite possessing stomata and a vascular system. Given the absence of a simple answer, we instead aim to provide a more complex case-by-case explanation for the lack of an An–ε trade-off when pooling species for all the studied phyla. Table 2. Divergences in the main structural, chemical, anatomical, and physiological traits of vascular plants, bryophytes, and charophytes Traits . Charophytes . Bryophytes (liverworts, mosses, and hornworts) . Lycophytes . Ferns . Angiosperms . Thickness of cell walls 1 µm Zygnema sp., (Herburger and Holzinger, 2015) 0.67–3.4 µm (Carriquí et al., 2019) 0.2–0.5 µm (Veromann-Jürgenson et al., 2017; Carriquí et al., 2019) 0.17–0.81 µm (Tosens et al., 2015) 0.11–0.54 µm (Onoda et al. 2017) Composition of cell walls (Matsunaga et al., 2004; Sørensen et al., 2011; Roberts et al., 2012) Lack of lignins. Lignin–like polymers Lack of lignins. Only lignans and other phenolic compounds (Ligrone et al., 2008; Popper, 2008; Popper et al., 2011; Ligrone et al., 2012), recently linked to mechanical reinforcement (Brodribb et al., 2020) Lignins present in some groups (e.g. Selaginellales; Weng et al., 2010) Lignins abundant in the secondary cell wall Lignins abundant in the secondary cell wall High amount of mannans and uronic acids (Popper, 2003; Sarkar et al., 2009; Popper et al., 2011) Higher proportion of mannose in the pectin than in ferns and equal percentage of uronic acids (Silva et al., 2011) Mannans in lower proportion in the primary cell wall. Smaller amount of uronic acids. Arabinose-rich polymers in ‘resurrection plants’ (Moore et al., 2013) Poor in cellulose, with callose instead, which explains its outstanding flexibility upon desiccation (Holzinger et al., 2011; Karsten and Holzinger, 2012) Callose present in sieve elements (Burr and Evert, 1973; Warmbrodt and Evert, 1974) Callose generally sparsely produced (0.3–5% of the total cell wall content; Falter et al., 2015), mainly in plasmodesmata formation (Cui and Lee, 2016) In gametophyte generation, small amounts of RG-II-like polysaccharide Same amount of RG-II as in ferns and some angiosperms High amount of RG-II-like polysaccharides Cuticle (Bargel et al., 2004) Lack of cuticle Cuticle as a thin extracellular membrane Thin cuticle (Hübers et al., 2011) Cuticle present. Absent in filmy ferns Cuticle very variable (40–10 000 nm) (Jeffree, 2008) Stomata (Duckett and Pressel, 2017) Lack of stomata Lack of stomata in the gametophyte Stomata present. Passive control Stomata present. Passive control. Stomata absent in filmy ferns (Bravo et al., 2016) Stomata present. Controlled by abscisic acid Leaf structure (Duckett and Pressel, 2017) Unistratose, high transmittance Photosynthetic tissue one/two cell-layered (high transmittance) or with photosynthetic filaments (e.g. Polytrichaceae and Lunulariaceae). Hornworts and thalloid liverworts have multi-layered liquid/mucilage-filled intercellular spaces Pluristratose mesophyll, low transmittance Pluristratose mesophyll, low transmittance (except filmy ferns, which are unistratose with high transmittance; Ebihara et al., 2006) Pluristratose mesophyll, low transmittance Water control (Oliver et al., 2000) Poikilohydric Poikilohydric Homoiohydric Homoiohydric (except filmy ferns and other ‘resurrection ferns’) (Kessler and Siorak, 2007) Homoiohydric (except ‘resurrection plants’) Desiccation tolerance (Farrant et al., 2017) Rare, reported for Zygnema and Klebsormidium spp. (Herburger and Holzinger, 2015) Frequent, especially in mosses and leafy liverworts Uncommon (except Selaginella spp.) More frequent than in angiosperms Uncommon (Gaff and Oliver, 2013) Water-conducting cells (Ligrone et al., 2012) Absent Absent, perforate water conducting cells (hydroids). Protoplast lost at maturity (Kenrick and Crane, 1997). Surrounding stands of hydroids are leptoids, analogous to phoem Xylem (tracheids) present (Friedman and Cook, 2000) Xylem (mostly tracheids) present, highly reduced in filmy ferns Xylem (vessels) present (Carlquist and Schneider, 1997a, b) Traits . Charophytes . Bryophytes (liverworts, mosses, and hornworts) . Lycophytes . Ferns . Angiosperms . Thickness of cell walls 1 µm Zygnema sp., (Herburger and Holzinger, 2015) 0.67–3.4 µm (Carriquí et al., 2019) 0.2–0.5 µm (Veromann-Jürgenson et al., 2017; Carriquí et al., 2019) 0.17–0.81 µm (Tosens et al., 2015) 0.11–0.54 µm (Onoda et al. 2017) Composition of cell walls (Matsunaga et al., 2004; Sørensen et al., 2011; Roberts et al., 2012) Lack of lignins. Lignin–like polymers Lack of lignins. Only lignans and other phenolic compounds (Ligrone et al., 2008; Popper, 2008; Popper et al., 2011; Ligrone et al., 2012), recently linked to mechanical reinforcement (Brodribb et al., 2020) Lignins present in some groups (e.g. Selaginellales; Weng et al., 2010) Lignins abundant in the secondary cell wall Lignins abundant in the secondary cell wall High amount of mannans and uronic acids (Popper, 2003; Sarkar et al., 2009; Popper et al., 2011) Higher proportion of mannose in the pectin than in ferns and equal percentage of uronic acids (Silva et al., 2011) Mannans in lower proportion in the primary cell wall. Smaller amount of uronic acids. Arabinose-rich polymers in ‘resurrection plants’ (Moore et al., 2013) Poor in cellulose, with callose instead, which explains its outstanding flexibility upon desiccation (Holzinger et al., 2011; Karsten and Holzinger, 2012) Callose present in sieve elements (Burr and Evert, 1973; Warmbrodt and Evert, 1974) Callose generally sparsely produced (0.3–5% of the total cell wall content; Falter et al., 2015), mainly in plasmodesmata formation (Cui and Lee, 2016) In gametophyte generation, small amounts of RG-II-like polysaccharide Same amount of RG-II as in ferns and some angiosperms High amount of RG-II-like polysaccharides Cuticle (Bargel et al., 2004) Lack of cuticle Cuticle as a thin extracellular membrane Thin cuticle (Hübers et al., 2011) Cuticle present. Absent in filmy ferns Cuticle very variable (40–10 000 nm) (Jeffree, 2008) Stomata (Duckett and Pressel, 2017) Lack of stomata Lack of stomata in the gametophyte Stomata present. Passive control Stomata present. Passive control. Stomata absent in filmy ferns (Bravo et al., 2016) Stomata present. Controlled by abscisic acid Leaf structure (Duckett and Pressel, 2017) Unistratose, high transmittance Photosynthetic tissue one/two cell-layered (high transmittance) or with photosynthetic filaments (e.g. Polytrichaceae and Lunulariaceae). Hornworts and thalloid liverworts have multi-layered liquid/mucilage-filled intercellular spaces Pluristratose mesophyll, low transmittance Pluristratose mesophyll, low transmittance (except filmy ferns, which are unistratose with high transmittance; Ebihara et al., 2006) Pluristratose mesophyll, low transmittance Water control (Oliver et al., 2000) Poikilohydric Poikilohydric Homoiohydric Homoiohydric (except filmy ferns and other ‘resurrection ferns’) (Kessler and Siorak, 2007) Homoiohydric (except ‘resurrection plants’) Desiccation tolerance (Farrant et al., 2017) Rare, reported for Zygnema and Klebsormidium spp. (Herburger and Holzinger, 2015) Frequent, especially in mosses and leafy liverworts Uncommon (except Selaginella spp.) More frequent than in angiosperms Uncommon (Gaff and Oliver, 2013) Water-conducting cells (Ligrone et al., 2012) Absent Absent, perforate water conducting cells (hydroids). Protoplast lost at maturity (Kenrick and Crane, 1997). Surrounding stands of hydroids are leptoids, analogous to phoem Xylem (tracheids) present (Friedman and Cook, 2000) Xylem (mostly tracheids) present, highly reduced in filmy ferns Xylem (vessels) present (Carlquist and Schneider, 1997a, b) Open in new tab Table 2. Divergences in the main structural, chemical, anatomical, and physiological traits of vascular plants, bryophytes, and charophytes Traits . Charophytes . Bryophytes (liverworts, mosses, and hornworts) . Lycophytes . Ferns . Angiosperms . Thickness of cell walls 1 µm Zygnema sp., (Herburger and Holzinger, 2015) 0.67–3.4 µm (Carriquí et al., 2019) 0.2–0.5 µm (Veromann-Jürgenson et al., 2017; Carriquí et al., 2019) 0.17–0.81 µm (Tosens et al., 2015) 0.11–0.54 µm (Onoda et al. 2017) Composition of cell walls (Matsunaga et al., 2004; Sørensen et al., 2011; Roberts et al., 2012) Lack of lignins. Lignin–like polymers Lack of lignins. Only lignans and other phenolic compounds (Ligrone et al., 2008; Popper, 2008; Popper et al., 2011; Ligrone et al., 2012), recently linked to mechanical reinforcement (Brodribb et al., 2020) Lignins present in some groups (e.g. Selaginellales; Weng et al., 2010) Lignins abundant in the secondary cell wall Lignins abundant in the secondary cell wall High amount of mannans and uronic acids (Popper, 2003; Sarkar et al., 2009; Popper et al., 2011) Higher proportion of mannose in the pectin than in ferns and equal percentage of uronic acids (Silva et al., 2011) Mannans in lower proportion in the primary cell wall. Smaller amount of uronic acids. Arabinose-rich polymers in ‘resurrection plants’ (Moore et al., 2013) Poor in cellulose, with callose instead, which explains its outstanding flexibility upon desiccation (Holzinger et al., 2011; Karsten and Holzinger, 2012) Callose present in sieve elements (Burr and Evert, 1973; Warmbrodt and Evert, 1974) Callose generally sparsely produced (0.3–5% of the total cell wall content; Falter et al., 2015), mainly in plasmodesmata formation (Cui and Lee, 2016) In gametophyte generation, small amounts of RG-II-like polysaccharide Same amount of RG-II as in ferns and some angiosperms High amount of RG-II-like polysaccharides Cuticle (Bargel et al., 2004) Lack of cuticle Cuticle as a thin extracellular membrane Thin cuticle (Hübers et al., 2011) Cuticle present. Absent in filmy ferns Cuticle very variable (40–10 000 nm) (Jeffree, 2008) Stomata (Duckett and Pressel, 2017) Lack of stomata Lack of stomata in the gametophyte Stomata present. Passive control Stomata present. Passive control. Stomata absent in filmy ferns (Bravo et al., 2016) Stomata present. Controlled by abscisic acid Leaf structure (Duckett and Pressel, 2017) Unistratose, high transmittance Photosynthetic tissue one/two cell-layered (high transmittance) or with photosynthetic filaments (e.g. Polytrichaceae and Lunulariaceae). Hornworts and thalloid liverworts have multi-layered liquid/mucilage-filled intercellular spaces Pluristratose mesophyll, low transmittance Pluristratose mesophyll, low transmittance (except filmy ferns, which are unistratose with high transmittance; Ebihara et al., 2006) Pluristratose mesophyll, low transmittance Water control (Oliver et al., 2000) Poikilohydric Poikilohydric Homoiohydric Homoiohydric (except filmy ferns and other ‘resurrection ferns’) (Kessler and Siorak, 2007) Homoiohydric (except ‘resurrection plants’) Desiccation tolerance (Farrant et al., 2017) Rare, reported for Zygnema and Klebsormidium spp. (Herburger and Holzinger, 2015) Frequent, especially in mosses and leafy liverworts Uncommon (except Selaginella spp.) More frequent than in angiosperms Uncommon (Gaff and Oliver, 2013) Water-conducting cells (Ligrone et al., 2012) Absent Absent, perforate water conducting cells (hydroids). Protoplast lost at maturity (Kenrick and Crane, 1997). Surrounding stands of hydroids are leptoids, analogous to phoem Xylem (tracheids) present (Friedman and Cook, 2000) Xylem (mostly tracheids) present, highly reduced in filmy ferns Xylem (vessels) present (Carlquist and Schneider, 1997a, b) Traits . Charophytes . Bryophytes (liverworts, mosses, and hornworts) . Lycophytes . Ferns . Angiosperms . Thickness of cell walls 1 µm Zygnema sp., (Herburger and Holzinger, 2015) 0.67–3.4 µm (Carriquí et al., 2019) 0.2–0.5 µm (Veromann-Jürgenson et al., 2017; Carriquí et al., 2019) 0.17–0.81 µm (Tosens et al., 2015) 0.11–0.54 µm (Onoda et al. 2017) Composition of cell walls (Matsunaga et al., 2004; Sørensen et al., 2011; Roberts et al., 2012) Lack of lignins. Lignin–like polymers Lack of lignins. Only lignans and other phenolic compounds (Ligrone et al., 2008; Popper, 2008; Popper et al., 2011; Ligrone et al., 2012), recently linked to mechanical reinforcement (Brodribb et al., 2020) Lignins present in some groups (e.g. Selaginellales; Weng et al., 2010) Lignins abundant in the secondary cell wall Lignins abundant in the secondary cell wall High amount of mannans and uronic acids (Popper, 2003; Sarkar et al., 2009; Popper et al., 2011) Higher proportion of mannose in the pectin than in ferns and equal percentage of uronic acids (Silva et al., 2011) Mannans in lower proportion in the primary cell wall. Smaller amount of uronic acids. Arabinose-rich polymers in ‘resurrection plants’ (Moore et al., 2013) Poor in cellulose, with callose instead, which explains its outstanding flexibility upon desiccation (Holzinger et al., 2011; Karsten and Holzinger, 2012) Callose present in sieve elements (Burr and Evert, 1973; Warmbrodt and Evert, 1974) Callose generally sparsely produced (0.3–5% of the total cell wall content; Falter et al., 2015), mainly in plasmodesmata formation (Cui and Lee, 2016) In gametophyte generation, small amounts of RG-II-like polysaccharide Same amount of RG-II as in ferns and some angiosperms High amount of RG-II-like polysaccharides Cuticle (Bargel et al., 2004) Lack of cuticle Cuticle as a thin extracellular membrane Thin cuticle (Hübers et al., 2011) Cuticle present. Absent in filmy ferns Cuticle very variable (40–10 000 nm) (Jeffree, 2008) Stomata (Duckett and Pressel, 2017) Lack of stomata Lack of stomata in the gametophyte Stomata present. Passive control Stomata present. Passive control. Stomata absent in filmy ferns (Bravo et al., 2016) Stomata present. Controlled by abscisic acid Leaf structure (Duckett and Pressel, 2017) Unistratose, high transmittance Photosynthetic tissue one/two cell-layered (high transmittance) or with photosynthetic filaments (e.g. Polytrichaceae and Lunulariaceae). Hornworts and thalloid liverworts have multi-layered liquid/mucilage-filled intercellular spaces Pluristratose mesophyll, low transmittance Pluristratose mesophyll, low transmittance (except filmy ferns, which are unistratose with high transmittance; Ebihara et al., 2006) Pluristratose mesophyll, low transmittance Water control (Oliver et al., 2000) Poikilohydric Poikilohydric Homoiohydric Homoiohydric (except filmy ferns and other ‘resurrection ferns’) (Kessler and Siorak, 2007) Homoiohydric (except ‘resurrection plants’) Desiccation tolerance (Farrant et al., 2017) Rare, reported for Zygnema and Klebsormidium spp. (Herburger and Holzinger, 2015) Frequent, especially in mosses and leafy liverworts Uncommon (except Selaginella spp.) More frequent than in angiosperms Uncommon (Gaff and Oliver, 2013) Water-conducting cells (Ligrone et al., 2012) Absent Absent, perforate water conducting cells (hydroids). Protoplast lost at maturity (Kenrick and Crane, 1997). Surrounding stands of hydroids are leptoids, analogous to phoem Xylem (tracheids) present (Friedman and Cook, 2000) Xylem (mostly tracheids) present, highly reduced in filmy ferns Xylem (vessels) present (Carlquist and Schneider, 1997a, b) Open in new tab Hypothesis for bryophytes, charophytes and filmy ferns: hydric strategy Different chemical composition (summarized in Table 2) and simple photosynthetic structures, phyllids/thalli, have been reported for bryophytes, charophytes, and filmy ferns, all of them in some cases being composed of single-cell layers (Hennequin, 2003; Waite and Sack, 2010; Holzinger and Pichrtová, 2016). The higher amount of callose reported for bryophytes (Popper and Fry, 2003; Popper, 2008; Popper et al., 2011) has been suggested to play a role in increasing the elasticity of tissues in algae (Herburger and Holzinger, 2015). In addition, high leaf mass per area and leaf density have been correlated with ε across angiosperms (Niinemets, 2001; Sack et al., 2003), suggesting that high ε could at least partially arise from bulk tissue accumulation and increased robustness of the leaves. To illustrate the differences in the structures of photosynthetic organs, photomicrographs of a transverse section of the photosynthetic structures of representative species of each group are shown at the same scale in Fig. 2. It is apparent from these images that the robustness of leaves in, for example, Plagiomnium must be much greater than in, for example, Olea, despite Plagiomnium having much thicker cell walls, which strongly limit its photosynthesis (Carriquí et al., 2019). Fig. 2. Open in new tabDownload slide Macroscopic and microscopic images of photosynthetic structures (leaves, phyllids, and thalli) of an angiosperm (Olea europaea), a non-filmy fern (Nephrolepis exaltata), a spikemoss (Selaginella denticulata), a liverwort (Lunularia cruciata), and a moss (Plagiomnium undulatum). Microscopic images consist of a transverse semi-thin section of the photosynthetic structures stained with toluidine blue. e, epidermis; f, photosynthetic filaments; np, non-photosynthetic parenchyma; p, palisade parenchyma; s, spongy parenchyma; v, vascular bundle. Asterisks indicate intercellular spaces, which are liquid/mucilage-filled in L. cruciata. (This figure is available in colour at JXB online.) Fig. 2. Open in new tabDownload slide Macroscopic and microscopic images of photosynthetic structures (leaves, phyllids, and thalli) of an angiosperm (Olea europaea), a non-filmy fern (Nephrolepis exaltata), a spikemoss (Selaginella denticulata), a liverwort (Lunularia cruciata), and a moss (Plagiomnium undulatum). Microscopic images consist of a transverse semi-thin section of the photosynthetic structures stained with toluidine blue. e, epidermis; f, photosynthetic filaments; np, non-photosynthetic parenchyma; p, palisade parenchyma; s, spongy parenchyma; v, vascular bundle. Asterisks indicate intercellular spaces, which are liquid/mucilage-filled in L. cruciata. (This figure is available in colour at JXB online.) Bryophytes, Chara sp., and filmy ferns share a common characteristic related to their water control: they are poikilohydric plants, meaning that they exert no control over their water content (Proctor and Tuba, 2002; Holzinger and Pichrtová, 2016). Owing to the lack of cuticle and stomata (Paton and Pearce, 1957; Brodribb and McAdam, 2017), non-controlled gas exchange (and therefore water loss) occurs across the phyllid/thallus surface in poikilohydric species. In contrast, vascular plants can regulate water loss and carbon gain by modifying stomatal conductance (gs), creating a large resistance to the hydraulic system that tightly couples gs with leaf hydraulic conductance (Kleaf; Sack and Holbrook 2006; Brodribb et al., 2007, Scoffoni et al., 2016) and, therefore, with An (Flexas et al., 2013, 2018; Xiong et al., 2017; Lu et al., 2019). Interestingly, despite the lack of stomata, the relationship between An and Kleaf in bryophytes appears to be identical to that of vascular plants (including lycophytes) (Brodribb et al., 2007). Water-conducting cells analogous to xylem, termed hydroids, have been described in some gametophytes of bryophytes (Kenrick and Crane, 1997; Ligrone et al., 2012) and have been recently described as functionally equivalent to vascular tissues of tracheophytes, at least in some species (Brodribb et al., 2020), as they are able to transport water under the same range of tensions as those used by vascular plants to extract water from the soil without the collapse of transport cells and with a resistance to cavitation within the vascular plant spectrum. Filmy ferns also lack stomata but retain a simple vascular tissue (Russow, 1872; Härtel, 1940; Bravo et al., 2016). Recently, Xiong and Nadal (2020) reviewed the relationships between Kleaf, ɛ, and leaf water-storage capacitance (i.e. Cleaf) in vascular plants, showing that elastic leaves have high water transport efficiency and high Cleaf. As an explanation, it was proposed that a dynamic storage compartment of water can buffer the fluctuations of water potential in the leaf, which would be more frequent in plants with high evaporative demand (Sack and Tyree 2005; Tuzet and Perrier, 2008; Xiong and Nadal, 2020). This evolutionary constraint between hydraulic conductance and water storage could explain the trade-off shown by Nadal et al. (2018). However, bryophytes, charophytes, and filmy ferns show elevated values of Cleaf that are uncoupled from An (Fig. 3A). Furthermore, ɛ of bryophytes was invariable as Cleaf increased, in contrast to angiosperms and non-filmy ferns, in which more rigid leaves had higher Cleaf (Fig. 3B). In liverworts, species with a multi-cell-layer thallus, intercellular spaces are filled with liquid or mucilage (Duckett and Pressel, 2017), but most of the water storage is observed in non-photosynthetic tissues (see e.g. Lunularia cruciata in Fig. 2). Thus, in mosses the storage of water does not seem to carry any diffusional/hydraulic advantage, and Kleaf (and therefore An) could be directly limited by the passive gas exchange through the cell wall and mesophyll, explaining the disconnection between ɛ and An in poikilohydric groups. Fig. 3. Open in new tabDownload slide Relationship between leaf water-storage capacitance (Cleaf) and (A) net CO2 assimilation (An) and (B) bulk modulus of elasticity (ε) across the studied species. For angiosperms, non-filmy ferns, and Isoetes velata (indicated by arrows) only, logarithmic correlations were significant between Cleaf and An (y=6.78ln(x)+14.68, R2=0.461) or ε (y= –3.75ln(x)+12.51, R2=0.250). Fig. 3. Open in new tabDownload slide Relationship between leaf water-storage capacitance (Cleaf) and (A) net CO2 assimilation (An) and (B) bulk modulus of elasticity (ε) across the studied species. For angiosperms, non-filmy ferns, and Isoetes velata (indicated by arrows) only, logarithmic correlations were significant between Cleaf and An (y=6.78ln(x)+14.68, R2=0.461) or ε (y= –3.75ln(x)+12.51, R2=0.250). Hypothesis for lycophytes: uncoupled stomatal control and water supply The three studied species of lycophytes, H. squarrosa, S. denticulata, and I. velata, have a cuticle, vascular system, and passive stomatal control as in non-filmy ferns (Table 2). Therefore, despite the fact that non-isoetales lycophytes (H. squarrosa and S. denticulata) on the one hand, and I. velata on the other, lie in different corners of the An–ε relationship, no relevant structural differences in pluricellular photosynthetic tissues were observed. Furthermore, the cell wall composition of lycophytes has been reported to be similar to that of ferns, with the same proportion of mannose (Silva et al., 2011) and other glycosyl residues (Matsunaga et al., 2004). Interestingly, elastic tissues of H. squarrosa and S. denticulata also showed high Cleaf (2.9±0.3 and 4.8±2.8 mol m–2 MPa–1, respectively), lying close to the fitted model for the relationship between ε and Cleaf of angiosperms, non-filmy ferns, and I. velata (Fig. 3B), but uncoupled from An as in bryophytes (Fig. 3A). In the cladistic study of Kenrick and Crane (1997), several morphological differences in the vascular system of the two groups of lycophytes were summarized, such as the xylem strand shape (a unique anchor shape in Isoetes) and secondary growth (present only in Isoetes and the extinct genus Paralycopodites). However, the most interesting discrepancy between Isoetes and other lycophytes lies in its rootlet anatomy, which is characterized by a vascular bundle attached to one side of a large central cavity that is surrounded by cortex, instead of being simple and non-rhizomorphic as in Huperzia and Selaginella (Kenrick and Crane, 1997). Whether these differences could be relevant in explaining the disconnection between An and ε in some lycophytes remains unknown. Since the speed of gas exchange depends partially on the speed of water supply (Jones, 1978; Meinzer, 2002), we hypothesize that Kleaf is limited by simple roots (root water uptake) in Huperzia and Selaginella, instead of by cell wall and mesophyll as in bryophytes, or by stomata as in the rest of tracheophytes. According to this hypothesis, the high Cleaf and elasticity of Huperzia and Selaginella might not result from an evolutionary constraint to the evaporative demand of high photosynthetic tissues, as suggested for the rest of the tracheophytes (Xiong and Nadal, 2020), but instead result from the need to buffer an inefficient water supply combined with passive control of gs. The casual observation of apical growth in a Huperzia sp. individual despite having detached and rotten roots (Fig. 4) supports this hypothesis. This observation indicates that the high Cleaf of Huperzia tissues together with a hydraulically uncoupled gs allows some growth and gas exchange, with an unknown temporal viability, even when water uptake from the roots is interrupted. This hypothesis is not incompatible with the identical relationship between An and Kleaf in well-watered lycophytes, bryophytes, and angiosperms reported by Brodribb et al. (2007), as gas exchange depends mechanistically on gas conductance. However, lycophytes are then expected to be outliers in the Kleaf–Cleaf relationship reviewed by Xiong and Nadal (2020), as high Cleaf and elastic tissues are not supposed to sustain a high evaporative demand, but an inefficient water uptake. Testing this hypothesis would require accurate measurements of the water relations of lycophytes, including xylem (Kx) and outside xylem (Kox) conductance, foliar water uptake, and minimum conductance. Kx is then hypothesized to be more related to Kleaf than Kox in non-isoetales lycophytes, contrary to what has been reported for angiosperms (Xiong and Nadal, 2020). Fig. 4. Open in new tabDownload slide Photograph of Huperzia sp. growing in a wet shadowed greenhouse. Although not apparent in this image, the base of the stem was rotten and had been detached from the roots for at least 4 months, but green and hydrated leaves were sustained in the stem apex. (This figure is available in colour at JXB online.) Fig. 4. Open in new tabDownload slide Photograph of Huperzia sp. growing in a wet shadowed greenhouse. Although not apparent in this image, the base of the stem was rotten and had been detached from the roots for at least 4 months, but green and hydrated leaves were sustained in the stem apex. (This figure is available in colour at JXB online.) ε relates to the cessation of net photosynthesis during drought in all studied phyla Interestingly, although differences in their hydric strategy and hydraulic system make bryophytes, charophytes, filmy ferns, and some lycophytes behave as clear outliers in the An–ε relationship under optimum conditions, when considering dehydration dynamics, a common functionality of ε does emerge across all land plants: general regressions between the RWC at which An decreases to 0 (RWC0) and ε (Fig. 5A), and between RWC0 and the RWC at which cell turgor is lost (RWCTLP) (Fig. 5B), were found across all studied species when a progressive desiccation assay was applied to detached leaves/shoots/thalli. All species except the ferns Nephrolepis exaltata and Phlebodium aureum had significantly lower RWC0 than RWCTLP (Fig. 5B), indicating that photosynthesis generally ceases fully after passing the turgor loss point, or at least simultaneously in some ferns. In vascular plants, the RWC0–RWCTLP relationship could be explained by differential stomatal control. Stomata are often reported to be the main drivers of photosynthesis under decreasing RWC in vascular plants: abscisic acid synthesis is triggered upon turgor loss in angiosperms (McAdam and Brodribb, 2016), whereas turgor loss per se leads to stomatal closure in ferns and lycophytes (Brodribb and McAdam, 2011). Fig. 5. Open in new tabDownload slide Relationships along phylogeny between (A) bulk modulus of elasticity (ε) and relative water content at which An=0 (RWC0) (linear model: y=2.86x+26.74; r=0.739, P<0.0001 across seven bryophytes, three lycophytes, seven ferns, and 11 angiosperms); (B) relative water content at the turgor loss point (RWCTLP) and RWC0 (non-linear model: y=9.13×10–4x3–0.139x2+6.81x–79.25, r=0.901, P<0.0001); and (C) RWCTLP and ε (non-linear model: y=0.025ln(x)+4.078, r=0.791, P<0.0001). Data are means ±SE; n=4–5 for RWC0, n=6 for pressure–volume-derived parameters. Symbols are as in Figs 1 and 3. Fig. 5. Open in new tabDownload slide Relationships along phylogeny between (A) bulk modulus of elasticity (ε) and relative water content at which An=0 (RWC0) (linear model: y=2.86x+26.74; r=0.739, P<0.0001 across seven bryophytes, three lycophytes, seven ferns, and 11 angiosperms); (B) relative water content at the turgor loss point (RWCTLP) and RWC0 (non-linear model: y=9.13×10–4x3–0.139x2+6.81x–79.25, r=0.901, P<0.0001); and (C) RWCTLP and ε (non-linear model: y=0.025ln(x)+4.078, r=0.791, P<0.0001). Data are means ±SE; n=4–5 for RWC0, n=6 for pressure–volume-derived parameters. Symbols are as in Figs 1 and 3. RWCTLP was correlated with ε (Fig. 5C; r=0.791, P<0.000 when all phyla were pooled, but the relationship was not significant if only angiosperms were included). In vascular plants, high ε (i.e. sclerophylly) has been related to the maintenance of symplastic volume by the preservation of RWCTLP (Bartlett et al., 2012), at least above a threshold of rigidity. Below this threshold, low ε is possibly related to the ability to sustain moderate water loss, as is related to both high leaf capacitance and maintenance of rehydration capacity at lower RWC (John et al., 2018; Nadal et al., 2018). Likewise, RWC0 strongly correlated with ε across species (Fig. 5A; r=0.715, P<0.000). Highly elastic tissues would be able to sustain positive assimilation over a wider range of RWC, whereas rigid leaves will promptly cease assimilation at small changes in RWC. At one extreme, RWC0 in bryophytes and Hymenophyllum spp. is very low (from 18.5±8.1% in the liverwort Pellia endiviifolia to 38±3.9% in the moss T. tamariscinum), indicating non-hydraulic-dependent assimilation, where cessation of photosynthesis is indeed near the RWC point of loss of functionality of chlorophyll fluorescence reported in vascular plants (Trueba et al., 2019). This new RWC0–ε relationship adds to the possible roles of ε in leaf functionality. Here, elasticity is proposed to play a key role in the photosynthetic—and, possibly, hydraulic—response to water stress, determining the overall leaf strategy: low ε for an anisohydry-related response (and poikilohydry in the extreme case of bryophytes and filmy ferns) and high ε for an isohydric behavior. Thus, ε may provide a mechanistic basis for the continuum of leaf responses to water stress described by Klein (2014) and challenges our current paradigm on how water stress affects photosynthesis, that is, initial down-regulation associated with stomatal closure, followed by complete stomatal closure (and thus cessation of An) when turgor loss collapses the guard cells, with a secondary and later role for mesophyll conductance and biochemical limitations (Flexas et al., 2004, 2012; Nadal and Flexas, 2019). The fact that species lacking stomata fell within the same RWC0–ε relationship suggests instead a major role of tissue elasticity, at least in the severe water stress phase leading to full cessation of photosynthesis, which is fully independent not just of stomatal closure but even of the presence of stomata. Supplementary data The following supplementary data are available at JXB online. Fig. S1. Custom-made moss cuvette and ΔCO2 leaks for the measuring head 3010-S. Fig. S2. Variation of An with RWC or time during gas exchange measurement in Hymenophyllum dicranotrichum. Fig. S3. Correlation between TW measured by manually removing interstitial water and TW calculated from pressure–volume data. Fig. S4. Relationship between An of excised leaves or detached shoots/thalli and RWC. Acknowledgements AVPC was awarded a predoctoral fellowship (FPU-02054) supported by the Ministerio de Educación, Cultura y Deporte (MECD), Spain. MN was supported by a predoctoral fellowship (BES-2015-072578) from the Ministerio de Economía y Competitividad (MINECO), Spain, co-financed by the European Social Fund (ESF). The research of AVPC, MN, and JF is supported by project PGC2018-093824-B-C41 from the Ministerio de Ciencia, Innovación y Universidades (MICIU) and the European Regional Development Fund (ERDF/FEDER). We thank Dr Néstor Fernández-Del-Saz and Dr Rafael Coopman for their support in species collection. We are grateful to Dr Ana Losada and Dr Javier Martíınez-Abaigar for identifying most bryophytes. Author contributions AVPC, MN, and JF designed the study; AVPC and MN conducted the experiments; AVPC and MN performed the analyses; AVPC wrote the first draft of the manuscript; and all authors contributed to the final version of the manuscript. Data availability The data supporting the findings of this study are available from the corresponding author, Alicia V. Perera-Castro, upon request. 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TI - What drives photosynthesis during desiccation? Mosses and other outliers from the photosynthesis–elasticity trade-off
JF - Journal of Experimental Botany
DO - 10.1093/jxb/eraa328
DA - 2020-10-22
UR - https://www.deepdyve.com/lp/oxford-university-press/what-drives-photosynthesis-during-desiccation-mosses-and-other-2Zd5ylo5OB
SP - 6460
EP - 6470
VL - 71
IS - 20
DP - DeepDyve
ER -