Water isotopes in desiccating lichens
The stable isotopic composition of water is routinely used as a tracer to study water exchange processes in vascular plants and ecosystems. To date, no study has focussed on isotope processes in non-vascular, poikilohydric organisms such as lichens and bryophytes. To understand basic isotope exchange processes of non-vascular plants, thallus water isotopic composition was studied in various green-algal lichens exposed to desiccation. The study indicates that lichens equilibrate with the isotopic composition of surrounding water vapour. A model was developed as a proof of concept that accounts for the specific water relations of these poikilohydric organisms. The approach incorporates first their variable thallus water potential and second a compartmentation of the thallus water into two isotopically distinct but connected water pools. Moreover, the results represent first steps towards the development of poikilohydric organisms as a recorder of ambient vapour isotopic composition.
KeywordsCryptogams δ18O discrimination Isotope equilibration Modelling Vapour Water exchange
Crassulacean acid metabolism
Relative water content
List of symbols
Total conductance for water vapour of stomata and boundary layer
Air relative humidity corrected to lichen temperature
Hypothetical exchange coefficient
Isotopic composition of atmospheric water vapour
Isotopic composition of evaporated water
Isotopic equilibrium of water with water vapour
Isotopic composition of leaf water
Isotopic composition of maximum enrichment
Weight of the dry lichen
Leaf/Lichen water volume
Weight of the lichen at maximum water content
Atmospheric water mole fraction
Maximum water content
Saturation water mole fraction
Equilibrium fractionation factor
Kinetic fractionation factor
The isotopic composition of water is a natural tracer that is widely used in hydrological research in order to provide new insights into some of the underlying processes. Prominent examples are recharge rates of groundwater (Gibson et al. 2002) and river catchment hydrology (cf. Kendall and McDonnell 1998). Within hydrological research, isotopes give a new perspective on the water cycle (Gat 2000) and, for that matter, they also link the hydrological cycle with plant ecology. As plants transpire about 50% of the water on land, they are one of the major drivers of the water cycle (Dirmeyer et al. 2006). Because different pools of water in an ecosystem can easily be distinguished by water isotopes, this approach also helps to clarify the different strategies of plants to acquire water (e.g. Dawson 1998). Stable isotopes of water are routinely used, for example, to partition ecosystem water fluxes into evaporation and transpiration fluxes (Yakir and Wang 1996) and to determine water recycling rates on continental scales (Gat and Matsui 1991).
All applications rely on accurate assessment of the isotopic composition of the respective water pools. Techniques to sample and analyse liquid water are well established; however, it is much more difficult to adequately collect atmospheric water vapour. Commonly, air moisture is cryogenically trapped and later analysed in the laboratory (Helliker et al. 2002). Recent technological advances such as tuneable diode lasers or cavity ring-down spectroscopy now allow instantaneous measurements of water isotopes in the field (Lee et al. 2006). However, all of these methods involve laborious and costly equipment and require availability of liquid nitrogen, dry ice and electric power.
Thus, the possibility to use a natural tracer that reflects the vapour isotopic signal of its surroundings would highly facilitate the monitoring of ambient moisture isotopic compositions. Lai et al. (2008) proposed using the isotopic composition of leaf water just before sunrise as a measure of vapour isotopic composition. This is because steady-state leaf water enrichment comes to isotopic equilibrium with water vapour at very high relative humidity (Craig and Gordon 1965), which often occurs at night. The small numbers of simultaneous observations of leaf water enrichment and water vapour isotopes at night seem to support this view (Langendörfer et al. 2002; Lai et al. 2006). However, the degree of equilibration depends on the turnover rate of leaf water by water vapour, and hence on stomatal conductance at night and leaf water volume (cf. Cuntz et al. 2007). Leaf water enrichment is therefore no reliable measure of water vapour isotopic composition. It also only records the isotope value at the end of the night and not its diurnal variations, which may be several per mill depending for example on canopy structure and leaf area index (Lai et al. 2006). Recently, Helliker and Griffiths (2007) pointed out that any plant which only operates at high relative humidity could be used as a proxy for ambient vapour isotopic composition. They showed that the leaf water of the epiphytic crassulacean acid metabolism (CAM) plant Tillandsia usneoides resembles the isotopic ratio of liquid water in equilibrium with atmospheric vapour in a predictive manner since it is independent of soil water supply and opens its stomata only at night. Yet, T. usneoides took about 3–4 days to reach full isotopic equilibrium with ambient vapour. In addition, this approach can only assess the isotopic composition at night and/or in ecosystems that first reach high relative humidity and second do not show too large variations in vapour isotopic composition during consecutive nights. In this respect, lichens may provide a way to overcome the limitations of higher plants because of their specific inherent water relations (cf. Lakatos et al. 2007; Hartard et al. 2008). In contrast to homoiohydric vascular plants, non-vascular cryptogams such as cyanobacteria, algae, lichens and bryophytes follow an alternative strategy of hydration: (1) they have reasonably small absolute water contents and (2) their water status varies with surrounding conditions (e.g. Blum 1973). These poikilohydric organisms are desiccation-tolerant, surviving drought in an anabiotic state until water becomes available. This strategy has proven to be successful, particularly in ecosystems where vascular plants reach their ecophysiological limits due to adverse climatic conditions (Kappen 1988; Lange et al. 1992; Belnap et al. 2001). Lichens in particular present a worldwide distribution, even in the driest habitats. Their symbiotic relationship consists of a fungal partner, the ‘mycobiont’, and its photosynthetic associate, the ‘photobiont’. Approximately 45% of the lichen-forming fungi develop stratified thalli (heteromerous lichens) which are structured in various layers (Honegger 1993; also see Table 1a). Despite this internal stratification, lichens furthermore exhibit different growth forms, such as crustose (crust-forming), foliose (leaf-like) and fruticose (shrub-like) types (reviewed by Honegger 2006).
Lichens’ poikilohydric nature is based on a lack of vascular tissue and concurrent missing continuous water supply as well as a lack of specific features, e.g. stomata, to control water deficit. Their water status varies passively with surrounding conditions and, consequently, they are subject to frequent desiccation. During desiccation events, lichen relative water content (RWC) may drop down to 10–15% of its maximum value at water saturation (e.g. Blum 1973; Rundel 1988). Correspondingly, thallus’ water potential (Ψ) decreases and approaches that of the environment (e.g. Jonsson et al. 2008). In contrast to vascular plants, they are specialised to utilise a variety of different water sources such as soil water, precipitation, dew and fog. Particularly green algal lichens, which constitute about 85% of all lichens (Honegger 1997), and also some mosses are even able to reactivate photosynthesis by rehydration with water vapour only (Lange 1969; Lange et al. 2001).
The α’s (>1) are kinetic (αk) and equilibrium (α+) fractionation factors (Majoube 1971; Cappa et al. 2003) and hs is air relative humidity corrected to leaf temperature: hs = wa/wsat(TL) (wa: atmospheric water mole fraction, wsat(TL) saturation water mole fraction at temperature TL). RM tends towards Req, i. e. isotopic equilibrium with water vapour when relative humidity tends towards unity. (The expressions ‘isotopic equilibrium with water vapour’ and ‘is equilibrated isotopically with water vapour’ are used in the following synonymously with Req.) This maximum enrichment RM can also be observed in monocot leaves where the leaf tips tend towards RM (Farquhar and Gan 2003) or in so-called desert rivers, water streams that have no effluent (Fontes and Gonfiantini 1967). The time (constant) to reach equilibrium is the turnover time, τ, i.e. leaf water volume VL in relation to the exchange flux with the atmosphere. In vascular plants τ ≈ VL/gtwsat(TL), i.e. the leaf water volume VL is turned over by the one-way water flux from the leaf to the atmosphere, gtwsat(TL) [and not simply transpiration E (cf. Dongmann and Nürnberg 1974; Farquhar and Cernusak 2005)], with gt being the total conductance for water vapour of stomata and boundary layer. In contrast, in non-vascular plants this turnover is given as τ ≈ VL/gtwa. This means that the leaf water volume VL in non-vascular plants is turned over by the one-way water flux from the atmosphere into the leaf, exactly opposite to vascular plants (cf. Appendix 1). The Helliker and Griffiths (2007) single source model was developed for CAM epiphytes that lack constant water supply. So the model should consequently also apply to lichens though lichens should reach steady state much faster because of their lower water content.
Within the present study, (1) the mechanisms driving oxygen isotope variability in the thallus water of lichens are addressed by examining its δ18O performance in different green-algal lichens experiencing dehydration, and (2) the applicability of the single-source model (Helliker and Griffiths 2007) to describe water exchange processes of lichens is tested and evaluated.
Additionally, the 18O isotopic signal of CO2 that is released by the lichens was assessed. δ18O of water is passed onto CO2 via hydration of dissolved CO2. In the presence of carbonic anhydrase (CA)—an enzyme that catalyses the interconversion of CO2 and bicarbonate—this process is much accelerated (Silverman 1982). Intracellular as well as extracellular CA also frequently occurs in lichens (Palmqvist and Badger 1996). Previous studies on vascular plants with high CA activity showed that, while leaf water δ18O may be markedly heterogeneous (Yakir et al. 1989), any released CO2 is in isotopic equilibrium with water at the evaporative site of the leaf (Gillon and Yakir 2000; Cousins et al. 2006). Transferring this to lichens suggests that the isotopic signal of CO2 released from the thallus reflects the water located in surface structures, whereas directly measured thallus water δ18O may represent the more heterogeneous bulk thallus water (Lakatos et al. 2007; Hartard et al. 2008).
Materials and methods
Lichen material, collection, storage and preconditioning
Four distinct lichen species have been selected according to their differences in morphology and ecological preferences of micro-habitat. All species were collected in a Mediterranean sand dune ecosystem in Osso de Baleia, western Portugal (40°0′N, 08°54′W). For a more detailed description of the collection site see Hartard et al. (2008). Whereas Usnea filipendula Stirton s.str. and Parmelia caperata (L.) Ach. are corticolous (on bark growing) species exposed to rather xeric conditions, Cladonia convoluta (Lam.) Anders and Cladina arbuscula (Wallr.) Hale and W.Culb. are typical terricolous (ground growing) lichens influenced by soil moisture. This particularly applies to the latter C. arbuscula that—in its natural habitat—conglomerates in large cushion-like associations (Table 1b).
After collection, samples were air-dried and kept in the refrigerator at 4°C for up to 4 weeks before conducting laboratory experiments.
Assessment of the lichen morphology
Several studies have shown that the progress of water exchange in lichens strongly depends on morphological properties, in particular on the existence and size of different thallus structures such as dense layers with thick-walled fungal hyphae versus areas with more loosely associated cells and hydrophobic cell walls (cf. Blum 1973; Bewley 1979; Rundel 1988). Depending on the investigated species, these layers can be found either on the thallus surface (e.g. outer cortices and algal layer) or inside the thallus (e.g. medulla and central strands) (Table 1a). Consequently, water located in these distinct regions is more or less exposed to the surrounding atmosphere and to associated isotopic exchange processes. To investigate the effect of such morphologically distinct layers and their relative exposition to ambient air on the oxygen stable isotope composition of the thallus water, each species’ specific internal stratification was assessed. The thickness of each observed layer was measured and the relative volume of each layer was subsequently calculated in order to obtain an approximation of the size of the water pools present in each of the distinct thallus layers.
For this, cross sections of each species were made using a freezing microtome (Reichert-Jung, Heidelberg, Germany). The cross sections were investigated microscopically and thickness of the thallus as well as of each structurally distinct thallus layer was measured quantitatively using an ocular micrometer. Five thalli of each species were randomly selected and ten cross sections of 40 μm thickness were prepared from each thallus. Each cross section represents the mean of five pseudo-replications (measured at five different positions) and the mean of ten cross sections was used to represent one replicate, i.e. thallus.
To identify the mechanisms driving 18O variability in lichens, a laboratory experiment was conducted and the oxygen isotope processes of the thallus water of the studied lichen species during dehydration were monitored. For this, the acclimated lichen samples were submersed in tap water for 30 min prior to the experiment to ensure initial water-saturation. Subsequently, the samples were slightly blotted to remove any excess water, and fresh weight at maximum water content (WCmax) was determined using an analytical balance (MC1 Analytic, Sartorius, Epsom, UK; accuracy: 0.1 mg). Except for three samples, which served as control, all other samples were placed into a darkened Plexiglas container and closed with a perforated lid, to allow homogeneous dehydration similar to natural conditions. Due to the highly fragile nature of lichens and to create a natural boundary layer, the air inside the container was not stirred by a vent. The control samples were immediately transferred into small, tight plastic containers to avoid any further dehydration until the start of the isotopic analysis. To ensure constant environmental conditions, the dehydration experiment was conducted within a climate chamber which supplied constant temperature of an average 25°C and 60% relative humidity (RH). Humidity conditions inside the Plexiglas container were measured in pre-experiments using the same experimental set-up and processing and measuring relative humidity with a small microclimatic data logger (HoboPro, Onset, MA, USA) programmed to record RH data once every minute throughout the whole pre-experiment. The measurements showed that immediately after placing the water-saturated samples inside the container, RH increased up to approximately 85%. This high RH remained approximately constant for 2.5–3 h and subsequently steadily decreased, approaching humidity conditions of the surrounding air of the climate chamber (60%). However, throughout the experiment, short periods down to 60% RH also occurred after each sampling collection due to the opening of the lid. During the dehydration experiment, which lasted for 7 h, every hour a set of at least three samples of each species was removed from the container. Each sample was placed into an individual pre-weighed exetainer (Labco Limited, Buckinghamshire, UK) and sealed with a gastight Teflon septum cap. Exetainers were wrapped in aluminium foil to ensure dark respiration and flushed with CO2-free air. This was done by penetrating the exetainers’ Teflon septum with two needles and flushing them with atmospheric air directed through a soda lime column. The outgoing air was directed into an absolute CO2 infrared gas analyser (Binos-100, Leybold-Heraeus GmbH, Hanau, Germany). Once the air coming out proved to be CO2-free, the needles were removed and the lichens were left to respire for at least 10 min. A pre-experiment was conducted to investigate the lichens’ respiration rates. Since they were found to be highly species specific and mainly dependent on the prevailing water status, specific respiration times were adjusted according to the expected respiration rates of the individual samples. In this way, the amount of respired CO2 inside the exetainers was similar for all samples (359 ± 196 ppm) at the time of isotopic analysis. The effect of the flushing with CO2-free air on the isotopic composition of the thallus water was also tested. Prior to the experiment, individual samples were repeatedly flushed and each time subsequent isotopic analysis of the lichen’s respired CO2 were performed. These experiments did not show significant effects of the flushing on the CO2 isotopic composition of the samples (data not shown).
Isotopic analysis of respired CO2
For isotopic analysis of the respired CO2, sample exetainers were placed in a multiflow autosampler (GV, Manchester, UK) coupled to a stable isotope ratio mass spectrometer (IRMS, ISOPRIME, GV). A gas sample of the respired CO2 of each exetainer was entrained in the carrier gas He. CO2 was isolated via gas chromatography separation and, subsequently, carried to the mass spectrometer. At least every 10 samples, external standard gas samples of laboratory standard gas of 300 ppm CO2 (Messer, Griesheim, Germany) were measured to reconstruct any drift occurring over the measurement period. Using a linear regression of the standard gas values, the sample data was then re-calculated whenever the drift exceeded 0.05‰. Following the analysis, all exetainers were stored in a freezer until thallus water extractions took place.
Thallus water extractions
After assessing the lichens’ respired CO2 isotopic composition, the same samples were processed to obtain the thallus water 18O signal. The thallus water of each sample was extracted using cryogenic vacuum distillation. Each sample exetainer was opened and instantly placed into the extraction vessel of a vacuum line. The vessel was then immersed into liquid nitrogen to keep the sample frozen, attached to the vacuum line and evacuated. Subsequently, the vessel was heated up to 100°C and the vaporised water was collected into a cold finger. Once all moisture had been extracted from the sample, the cold finger was sealed under vacuum using a torch. The WC of each sample was determined by weighing the exetainers before and after water extraction. The RWC was then calculated as difference between the lichen’s actual weight VL and the weight of the dry lichen Vdry divided by the difference between the wet weight Vwet at maximum water content WCmax (Beckett 1995). Since distillation yields the absolute dry weight, there might be a slight difference of about 10–20% compared to conventionally observed dry weights achieved by drying at 60°C or with silica gel.
Isotopic analysis of extracted thallus water
Oxygen isotope ratios of the extracted water samples were determined by equilibration of water with CO2 (Epstein and Mayeda 1953). 200 μl of the water sample was placed into a gastight vial. An aliquot of pure CO2 was added and was allowed to equilibrate for 16 h at a constant temperature of 17°C in a water bath. The equilibrated CO2 was then analysed for its oxygen isotopic composition using a multiflow autosampler coupled with an IRMS (ISOPRIME, GV).
All δ18O ratios are reported relative to VSMOW (Vienna Standard Mean Ocean Water). The measurements were taken against calibrated reference gas (CO2 5.3, Linde AG, Pullach, Germany) cross-calibrated to ISO-TOP reference gas (CO2, Messer), which in turn was cross-calibrated to IAEA-CH-4 and IAEA-CH-6 standards (International Atomic Energy Agency, Vienna, Austria).
a Morphological scheme of fruticose (left) and foliose (right) lichens. b Illustrations and morphological characteristics of the four studied green-algal lichen species
To receive a rough estimate of the proportional amount of lichen matrix directly exposed to air, the ratio of the outermost layers (cortex and algal layer) versus the internal thallus structures (medullary layer and/or central strand or cylinder, respectively) was calculated from the relative volume of the various thallus structures. Regarding this, about 51 and 29% of the matrix of the fruticose species C. arbuscula and U. filipendula, respectively, is located inside the thallus, and 49% (C. arbuscula) and 71% (U. filipendula) of the structures are directly exposed to the surrounding atmosphere. In contrast, the internal thallus structures of the foliose species P. caperata and C. convoluta amount to approximately 79 and 70% of total thallus thickness, respectively, whereas their exposed upper cortex and algal layer at the thallus surface comprises only 21% (P. caperata) and 30% (C. convoluta).
Water relations of lichen species during the experiment
After full rehydration P. caperata and C. arbuscula showed higher WCmax (Vwet) of approximately 2.25 and 2.60 g(H2O) g(DW)−1, respectively, as compared to C. convoluta and U. filipendula which only achieved about 1.45 and 1.20 g(H2O) g(DW)−1, respectively (Table 1). During exposure to the experimental conditions for 7 h, the lichens dehydrated at varying rates down to 0.09 ± 0.083 g(H2O) g(DW)−1 on average (Fig. 1a). Species-specific absolute evaporation rates (Fig. 1b) were calculated using numerical derivation of the WC data. In order to illustrate species-specific differences in dehydration extent and velocity, Fig. 1c presents the decline in RWC (% water content relative to WCmax).
C. arbuscula showed the highest initial evaporation rate, loosing 95% of its total evaporated water within the first 3.5 h of the experiment. U. filipendula and P. caperata took approximately 5.5 and 6 h to evaporate 95% of their total water. In contrast, C. convoluta showed rather moderate but more continuous water loss throughout the course of the experiment and took 6.5 h to loose 95% of its total water. However, the similar WC (and RWC) values of all species at the end of the experiment confirm the lichens physical equilibration with the environment. This indicates that, in terms of expected isotopic processes, the observed initial differences in evaporation rates may become irrelevant once the lichens equilibrate with the environmental conditions.
δ18O of thallus water during dehydration
Correlating RWC and thallus water δ18O confirmed a general progressive enrichment with decreasing RWC in all studied lichens (Fig. 2b). Furthermore, the overall degree of δ18O shift also depended upon the lichen species: at the start of the experiment, in totally water-saturated samples (90–100% RWC), δ18O of thallus water was similar between the respective species starting about 1.5‰ more enriched compared to δ18O of the water source (−8 ± 0.5‰). Cernusak et al. (2003) observed that leaves separated from the petiole lose about 8.5% of their water content within the first 2–3 min after cutting and concluded that this leads to a bias in the leaf water isotopic analyses due to evaporative enrichment. For the present experiment, rehydrated lichen samples removed from the water were blotted and weighed before being placed into a sealed exetainer which required approx. 3–5 min. It is therefore inferred that the isotopic deviation of the water-saturated lichens’ thallus water δ18O from that of the source water derives from strong initial evaporation during sample treatment.
At the end of the desiccation, at RWC below 30%, the isotopic composition of the thallus water showed significant differences between the corticolous, more xeric species (U. filipendula and P. caperata) and the terricolous, more mesic species (C. arbuscula and C. convoluta;t test: t = 4.7; FG = 26: P < 0.0001). U. filipendula and P. caperata showed a stronger δ18O enrichment of 8.4 ± 1.1‰ (corresponding to a final absolute δ18O of +1.9 ± 1.1‰), whereas C. convoluta and C. arbuscula only achieved changes of 6.8 ± 0.4‰ (corresponding to a final δ18O of +0.3 ± 0.4‰) during dehydration.
Comparing δ18O of thallus water and respired CO2
The single-source model
The Ψ model
The modelled values of this so-called ‘Ψ model’ (dashed lines, Fig. 4) correlate closer to the measurements but are still significantly different, especially during the second half of the experiment. This may have different reasons which will be discussed in the following. One possibility may simply be the adoption of erroneous PV-curves. If, with decreasing RWC, thallus Ψ descends faster than suggested by our exponential fit, relative humidity hL would have already approached unity which would result in equilibration between lichen thallus water and atmospheric vapour.
Since the box was not ventilated due to the fragile nature of lichens, a third possibility may be the existence of a large gradient in box relative humidity. Relative humidity was measured at the top of the box. Thus a gradient from high relative humidity near the evaporating lichens to low relative humidity near the box top may result in discrepancies between actual and measured relative humidity conditions. However, this difference should decrease with time because of very low evaporation rates at the end of the experiment and probably does not explain the high end values.
The Ψ2 model
In summary it can be said that, considering our modelling results the study shows first appropriate steps towards a better understanding of oxygen isotope processes of lichens during water exchange. Nevertheless, single aspects remain open that have to be affirmed conclusively. The results of the dehydration experiment turned out to be somewhat ambiguous. The lichens’ fragile nature created rather tricky experimental conditions. For example, the air inside the chamber could not be circulated effectively, which probably caused an inhomogeneous distribution of air moisture and water isotopes within the chamber. In addition, relevant parameters such as air moisture isotope composition and thallus water potential could not be measured. As a result, some of the input parameters of the model had to be determined indirectly. At that, the experimental approach pointed out that the aforementioned, imperative parameters need to be assessed in future studies. Continuous recording of these essential factors will then enable a precise survey and a more accurate assessment of the identified crucial mechanisms. Ultimately, detailed understanding will allow the transfer of these results to similar systems. Our results suggest that the proposed modelling approach may be applied to other non-vascular organisms such as bryophytes, terrestrial algae and cyanobacteria. Moreover, the rapid equilibration process indicates the potential value of lichens to be used as natural tracer for ambient vapour isotopic composition. Particularly their universal distribution, not only on a global scale but also within single ecosystems, points out the practicability of this approach.
A novel isotope enrichment model was demonstrated that can be used to explain oxygen isotope processes taking place in green-algal lichens subject to progressive dehydration. This model takes into account the specific poikilohydric nature of these cryptogams. As such, the water status as well as the thallus water potential of these organisms constantly varies along with surrounding conditions. Accordingly, it could be shown that their remaining thallus water rapidly equilibrates isotopically with surrounding atmospheric vapour. This is in contrast to vascular, homoiohydric plants, where leaf water is generally isotopically enriched compared to that in equilibrium with ambient water vapour.
Whereas previous studies have shown the suitability of stable carbon isotopes of poikilohydric cryptogams to serve as, e.g. long-term tracers for carbon acquisition, environmental change of CO2 sources and global change (Máguas and Brugnoli 1996; Lakatos et al. 2007), or as paleo-CO2 proxies (Fletcher et al. 2005), to date no work has focussed on the ability of oxygen stable isotope ratios of poikilohydric organisms to function as long-term indicator of water sources. Considering that the 18O signal of plant organic material of T. usneoides may be used to reconstruct the isotope ratio of atmospheric water for large spatial and temporal scales (Helliker and Griffiths 2007), and pre-eminently in view of the here presented results, poikilohydric epiphytes may serve as prospective long-term proxies for different environmental water sources.
We are particularly thankful to the stable isotope laboratory of Christiane Werner (University Bielefeld) which provided for parts of the isotopic analysis. We thank Howard Griffiths (University Cambridge) for scientific support and discussions. In addition, several people helped to conduct the laboratory studies. Special thanks to Rodrigo Maia (University Lisbon), Barbara Teichner (University Bielefeld), Beatrix Weber and Silvia Kühner (University Kaiserslautern), for their assistance. M.L. was granted in the frame of the EC-program NETCARB (HPRN-CT-1999-59) and of the ESF-program SIBAE (62561 EXGC EX03). B.H., C.M. and M.L. were supported by the PPP project of DAAD/GRICES (D/04/42019) and the Portuguese Science Foundation, FCT (POCTI/BIA-BDE/60140/2004).
- Belnap J, Büdel B, Lange OL (2001) Biological soil crusts: characteristics and distribution. In: Belnap J, Lange OL (eds) Biological soil crusts: structure, function and management (ecological studies 150). Springer, Berlin, pp 3–30Google Scholar
- Blum OB (1973) Water relations. In: Ahmadjian V, Hale ME (eds) The lichens. Academic Press, New York, pp 381–400Google Scholar
- Büdel B, Scheidegger C (1996) Thallus morphology and anatomy. In: Nash TH III (ed) Lichen biology. Cambridge University Press, Cambridge, pp 37–64Google Scholar
- Cernusak LA, Wong SC, Farquhar GD (2003) Oxygen isotope composition of phloem sap in relation to leaf water in Ricinus communis. Funct Plant Biol 30:1059–1070Google Scholar
- Craig H, Gordon LI (1965) Deuterium and oxygen 18 variations in the ocean and the marine atmosphere. In: Tongiorgi E (ed) Stable isotopes in oceanographic studies and paleotemperatures. Laboratorio di Geologia Nucleare, Pisa, pp 9–130Google Scholar
- Fletcher BJ, Beerling DJ, Brentnall SJ, Royer DL (2005) Fossil bryophytes as recorders of ancient CO2 levels: experimental evidence and a Cretaceous case study. Glob Biogeochem Cycles 19:GB3012Google Scholar
- Gibson JJ, Aggarwal P, Hogan J, Kendall C, Martinelli LA, Stichler W, Rank D, Goni I, Choudhry M, Gat J, Bhattacharya S, Sugimoto A, Fekete B, Pietroniro A, Maurer T, Panarello H, Stone D, Seyler P, Maurice-Bourgoin L, Herczeg A (2002) Isotope studies in large river basins: a new global research focus. Eos Trans AGU 83(52):613–617CrossRefGoogle Scholar
- Goebel K (1926) Ein Beitrag zur Biologie der Flechten. Ann Jard Bot Buitenz 36:1–83Google Scholar
- Helliker BR, Griffiths H (2007) Towards a plant-based proxy for the isotope ratio of atmospheric water vapor. Glob Change Biol 13:723–733Google Scholar
- Honegger R (1996) Morphogenesis In: Nash TH III (ed) Lichen biology. Cambridge University Press, Cambridge, pp 65–87Google Scholar
- Honegger R (1997) Metabolic interaction at the mycobiont-photobiont interface in lichens. In: Carroll GC, Tudzynski P (eds) Plant relationships. The Mycota V part A. Springer, Berlin, pp 209–221Google Scholar
- Honegger R (2006) Water relations in lichens. In: Gadd GM, Watkinson SC, Dyer P (eds) Fungi and the environment Lichen Biology. Cambridge University Press, Cambridge, pp 185–200Google Scholar
- Jahns HM (1984) Morphology, reproduction and water relations—a system of morphogenetic interactions in Paremlia saxatilis. In: Hertel H, Oberwinkler F (eds) Beiträge zur Lichenologie. Festschrift J. Poelt. Beiheft zur Nova Hedwigia 79. J. Cramer, Vaduz, pp 715–737Google Scholar
- Kappen L (1988) Ecophysiological relationships in different climatic regions. In: Galun M (ed) Handbook of lichenology, vol 2. CRC Press, Boca Raton, pp 37–100Google Scholar
- Kendall C, McDonnell JJ (eds) (1998) Isotope tracers in catchment hydrology. Elsevier, AmsterdamGoogle Scholar
- Lai C-T, Riley W, Owensby C, Ham J, Schauer A, Ehleringer J (2006) Seasonal and interannual variations of carbon and oxygen isotopes of respired CO2 in a tallgrass prairie: measurements and modeling results from three years with contrasting water availability. J Geophys Res 111:D08S06Google Scholar
- Lakatos M, Hartard B, Máguas C (2007) The stable isotopes δ13C and δ18O of lichens can be used as tracers of microenvironmental carbon and water sources. In: Dawson TE, Siegwolf RTW (eds) Stable isotopes as indicators of ecological change. Terrestrial Ecology Series, Elsevier, Oxford, pp 73–88Google Scholar
- Langendörfer U, Cuntz M, Ciais P, Peylin P, Bariac T, Milyukova I, Kolle O, Naegler T, Levin I (2002) Modelling of biospheric CO2 gross fluxes via oxygen isotopes in a spruce forest canopy: a 222Rn calibrated box model approach. Tellus 54B:476–496Google Scholar
- Lee X, Smith R, Williams J (2006) Water vapour 18O/16O isotope ratio in surface air in New England USA. Tellus 58B:293–304Google Scholar
- Majoube M (1971) Fractionnement en oxygène 18 et en deutérium entre l’eau et sa vapeur. J Chim Phys 68:1423–1436Google Scholar
- Nash TH III (1996) Photosynthesis, respiration, productivity and growth. In: Nash TH III (ed) Lichen biology. Cambridge University Press, Cambridge, pp 88–120Google Scholar
- Rawlins SL (1972) Theory of thermocouple psychrometers for measuring plant and soilwater potential. In: Brown RW, van Haveren BP (eds) Psychrometry in water relations research. University Logan Utah Agricultural Experiment Station, Utah State, USA, pp 43–50Google Scholar
- Rundel PW (1988) Water relations. In: Galun M (ed) Handbook of lichenology, vol 2. CRC Press, Boca Raton, pp 17–36Google Scholar
- Tyree MT, Jarvis PG (1981) Water in tissue and cells. In: Lange OL, Nobel PS, Osmond CB, Ziegler H (eds) Encyclopedia of plantphysiology. New series. Physiological plant ecology, vol 12B. Springer, Berlin, pp 35–77Google Scholar