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Keywords:

  • Antarctica;
  • endolithic lichens;
  • granitic rock;
  • ITS rDNA;
  • Lecidea;
  • ultrastructure

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  • • 
    Through the combined use of molecular and microscopy techniques, the endolithic lichens Lecidea cancriformis and Lecidea sp. were identified, even in the absence of fruiting bodies, and positioned under epilithic lichens. Cells of both algal and fungal symbionts were observed in fissures and cracks of the lithic substrate with no clear heteromerous structure. At the ultrastructural level, the two lichens differed in terms of their algal–fungal relationships.
  • • 
    Only one genotype of Trebouxia ITS sequence was identified from specimens of Lecidea sp., Umbilicaria aprina and Buellia frigida from the same zone, which could be mainly determined by low availability of alga in these extreme environments.
  • • 
    These lichens showed features typical of both chasmoendolithic and euendolithic microorganisms. Signs of biogeophysical and biogeochemical action on the substrate were detected close to fungal cells. This action seemed to be mainly conditioned by the local physico-chemical features of the substrate. Evidence for the biomobilization of elements by these endolithic lichens was found. L. cancriformis was observed to accumulate substantial amounts of calcium-rich biominerals.
  • • 
    The combined approach proposed is useful for mapping the distribution of endolithic lichens and analysing the processes that occur in their microscopic environment.

Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

The lithic substrate is being increasingly viewed as an important and largely unexplored part of the biosphere. Life underneath the rock surface is particularly common in the most extreme terrestrial habitats on Earth (Friedmann, 1980; Gross et al., 1998; Kidron, 2000). However, apart from its ubiquitous presence in this environment and the fact that its colonizers set off the soil stabilization process (Wynn-Williams, 1993) and participate in the elemental cycles of diverse ecosystems (Ferris & Lowson, 1997; Blum et al., 2002; Hoffland et al., 2002), our knowledge of the endolithic habitat is still at a rudimentary stage.

Fungi and algae living in symbiotic association as lichens are common internal colonizers of the lithic substrate (Friedmann, 1982; Friedmann et al., 1988; Ascaso et al., 1995, 1998; De los Ríos et al., 2002). The endolithic habitat shelters the lichen cells from both excessive drought and temperature extremes (Fry, 1922; Pinna et al., 1998; Kidron, 2000). Organisms that live within hard rocky substrates acquire specialized adaptive features, which in turn determine different endolithic ecological niches. These organisms are able to colonize existing cracks and fissures (chasmoendolithic), internal pores (cryptoendolithic) or penetrate actively into the rock (euendolithic) (Golubic et al., 1981). Water, light and nutrients can be rather sparse in the endolithic habitat, but in extreme environments its microclimate can be less severe than at the exposed rock surfaces where harsher environmental conditions exist. At this rock–atmosphere interface, lichenized alga and fungi, along with free-living microorganisms, are able to form different ‘biofilms’ (Warscheid & Braams, 2000; de los Ríos et al., 2002; Ascaso & Wierzchos, 2003). The biofilm organization provide protection to the resident microorganisms against the environmental conditions, since they can maintain conditions inside that are radically different to those of the external environment (Little et al., 1997; De los Ríos et al., 2003). In continental Antarctica, lithobiontic communities are the predominant forms of terrestrial life (Friedmann, 1982; Ascaso & Wierzchos, 2003; De la Torre et al., 2003; De los Ríos et al., 2003) and are thus an essential target for any evaluation of Antarctica's biodiversity. However, the study of lithobiontic microorganisms is met with numerous hurdles since these microorganisms are embedded in a hard substrate and are extremely difficult to culture. Some endolithic lichens can be observed with the naked eye when their mature fruiting organs appear at the rock surface, but in their absence endolithic lichens are not so easily determined (Fry, 1922). Besides, the factors determining colonization and succession in the lithic substrate are not completely known and it is difficult to foresee the presence of particular endolithic forms when there are no associated epilithic forms. The use of molecular methods to identify Antarctic endolithic microorganisms has recently been reported after their laboratory culture (Smith et al., 2000; Hughes & Lawley, 2003) or without the need for previous culture (De la Torre et al., 2003). These molecular studies are starting to provide insight into the existing biodiversity, yet the lack of microscopy studies carried out in parallel to these approaches means there is little information on the specific microsites inhabited by the microorganisms identified, or their interactions or organization. The aim of the present study was to apply both microscopy and molecular techniques in an effort to simultaneously identify and characterize endolithic lichen symbionts in rock samples from Granite Harbour (Antarctica).

Materials and Methods

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Materials

Pieces of granite rock sparsely colonized by lichens and mosses were collected from the Ross Sea coast, Granite Harbour (77°00′-S, 162°34′-E) across a range of altitudes from the coast to the summit of Discovery Bluff (5–500 m a.s.l). In this zone, epilithic forms of life were only present in places regularly exposed to melt water (Seppelt et al., 1995). Samples were collected under natural conditions and stored at −20°C until processing for microscopy, molecular, or microanalytical procedures.

Microscopy studies

The rock samples were prepared according to a procedure developed for observing the rock–microorganism interface by scanning electron microscopy with backscattered electron imaging (SEM–BSE, Wierzchos & Ascaso, 1994). In brief, the pieces of rock were fixed in glutaraldehyde and then in osmium tetroxide, dehydrated in a series of ethanol solutions, and embedded in LR-White resin. Blocks of resin-embedded rock samples were finely polished, carbon coated and observed using DMS 960 and DMS 940 A Zeiss SEM microscopes. Microprobe analyses were performed using an energy dispersive X-ray spectroscopy (EDS) instrument fitted with a Link ISIS microanalytical system during SEM observation. The microscopy and/or microanalytical operating conditions were as follows: 0° tilt angle, 35° take-off angle, 15 kV acceleration potential, 6 or 25 mm working distance and 1–5 nA specimen current. The EDS method allows qualitative and quantitative microanalysis by providing element spatial distribution maps. These maps indicate the relative concentrations of elements according to a colour scale: dark blue represents a concentration of absolute zero and white denotes the maximum absolute concentration of the corresponding pure-component spectrum.

In addition, endolithic lichen masses were removed from the same rock under the stereomicroscope using a sterile needle and blocked in 2% (w/v) agar. Small pieces of agar containing the microbial cells were fixed in glutaraldehyde and osmium tetroxide solutions, dehydrated in a graded ethanol series, and embedded in Spurr's resin following the protocol described by De los Ríos and Ascaso (2001). Several ultrathin sections (15–20) from the different samples were post-stained with lead citrate (Reynolds, 1963) and observed using a Zeiss EM910 transmission electron microscope.

Molecular studies

Total DNA was extracted from the epilithic lichens, fruiting bodies of endolithic lichens and microorganisms colonizing internal fissures of the rock, by the method of Cubero et al. (1999). In this last case, small fragments of the endolithic biofilm were selected under the stereomicroscope after fracturing the rock along the lines of fissure. Two methods were used to amplify endolithic fungal or algal rDNA. In the first method, small fragments of lithic substrate containing endolithic fungal masses were placed in a 0.5 ml-Eppendorf tube for direct PCR-amplification of the ITS regions of fungal rDNA, modified after Wolinski et al. (1999). In the second method, PCR was performed on total DNA isolated from identified apothecia and unidentified endolithic fungal masses. In total, 12 endolithic fungal ITS sequences corresponding to three different altitudes were determined. The primers used for amplification of the DNA from the fungal partner were ITS1F and ITS4 (White et al., 1990) and from the algal partner were ITS1T and ITS4T (Kroken & Taylor, 2000). In each case, the 50 µl PCR volume (10 mm Tris pH 8.3/50 mm KCl/1.5 mm MgCl2/50 µg gelatine) contained 1.25 units of Dynazyme Taq polymerase (Finnzymes, Espoo, Finland), 0.2 mm of each of the four dNTPs, 0.5 µm of each primer and c. 10–50 ng genomic DNA. Products were cleaned using the QiaQuick Spin kit (Qiagen, Vienna, Austria). Both complementary strands were sequenced using the BigDye Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems, Vienna, Austria) according to the manufacturer's instructions. Sequences were run on an ABI310 capillary sequencer (Applied Biosystems). The sequences are submitted to GenBank (nos AY667580, AY667581, AY667582 and AY667583).

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Epilithic thalli were scarce in the study area. Thalli of Buellia frigida were mainly observed in zones of lower altitude. A few thalli of Umbilicaria aprina and Caloplaca sp. were also growing on rocks. Epilithic black apothecia corresponding to endolithic lichens could be detected in samples collected across the entire range of altitudes. Detailed light microscopy analysis of these apothecia showed that these can be assigned to the species Lecidea cancriformis C.W. Dodge & G.E. Baker and Lecidea sp. (Øvstedal & Lewis Smith, 2001). While L. cancriformis showed a more or less brown hypothecium and small and thin ascospores (7–9 × 2–3 µm), the hypothecium was colourless and ascospores were broadly elliptic (14–17 × 6–9 µm) in Lecidea sp. The second species cannot be assigned to any of the species cited for Antarctica. Neither are the broadly elliptic ascospores of this Lecidea consistent with Lecidea sp. A (Øvstedal & Lewis Smith, 2001), which is described to have subglobose ascospores. No fruiting bodies were observed in further samples but endolithic growths visible to the naked eye as white masses were seen when the rocks were fractured along their fissures. The ITS regions of genomic DNA extracted from the two apothecia described above were amplified to yield two PCR products of approximate size 900 bp and 650 bp (lines 3 and 6 in Fig. 1). Products of similar size were also obtained by PCR amplification of single ITS regions of genomic DNA from different endolithic masses (with and without fruiting bodies) found in rock fissures across the altitude range (lines 2 and 5 in Fig. 1). We also tried direct PCR on very small rock fragments colonized by endolithic forms with comparable results to those obtained after DNA isolation (line 4 in Fig. 1). The analysis of the obtained sequences detected only two distinct fungal sequences corresponding to the two different product sizes. These sequences of similar length were almost identical, but differed from those obtained for the epilithic lichens found in the zone. Nevertheless, they were identical to those obtained through amplification of genomic DNA isolated from the two types of apothecium, allowing the identification of endolithic growths. Lecidea sp. was detected only in rock samples collected on the coast and in close proximity to the corresponding apothecia. In contrast, endolithic L. cancriformis were identified by molecular methods at coastal sites and in zones at 100 and 500 m a.s.l.

image

Figure 1. PCR products using fungal-specific primers of ITS ribosomal DNA regions of DNA isolated from Antarctic endolithic fungal forms. Lines 1 and 7: size marks; line 2: products of DNA isolated from fungal masses within a fissure in rock from a coastal region; line 3: products of DNA isolated from a Lecidea sp. apothecium; line 4: direct PCR products obtained from endolithic fungal cells. Line 5: products of DNA isolated from endolithic forms collected 500 m a.s.l.; Line 6: products of DNA isolated from a Lecidea cancriformis apothecium.

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Algal rDNA ITS regions from specimens of Lecidea sp., and of U. aprina and B. frigida from the same zone were also amplified and sequenced. However, we were only able to identify one Trebouxia sequence among the three species. The sequence coincides (98% similarity) with the denominated by Romeike et al. (2002) as Trebouxia sp. D11. The phylogenetic relationships of this ITS-variant, which has so far been found only in Continental Antarctica, are not still clarified. We did not manage to amplify algal rDNA from L. cancriformis, perhaps because of the high amounts of calcium oxalate in some of these samples.

We then applied the SEM–BSE method to examine how these endolithic forms were distributed in the rock substrate. Epilithic apothecia showed continuity within the lithic substrate. The Lecidea sp. apothecium is connected via a cord of densely packed hyphae to the endolithic part of the lichen (Fig. 2a). Figure 2b is an enlargement of this apothecium showing subhymenium and hymenium layers made up of paraphyses and asci containing ascospores at different developmental stages. Apothecia were also detected inside fissures (marked by arrowheads in Fig. 2c). Despite the fact that the apothecia observed inside fissures were not as well structured as the external ones, it is possible to distinguish their cup shape (outlined by a dotted line in Fig. 2c). Algal and fungal cells were observed in fissures close to the apothecium (Fig. 2d). Fragmentation of the mineral substrate was clearly visible in zones occupied by this lichen (Fig. 2a). The symbiont cells appeared to be closely associated with mineral fragments (Fig. 2e). In zones occupied by Lecidea sp., algal cells could be observed by SEM–BSE to be penetrated by fungal cells (arrows in Fig. 2f). On TEM examination, the interface between the algal and fungal cells of this endolithic lichen could be seen in more detail. Fungal haustoria penetrated the algal cell wall and plasma membrane (Fig. 3a), pushing on the cytoplasm. These intracellular haustoria were thin at their extreme ends (arrowhead in Fig. 3a,b) and appeared to be surrounded by a sheath (arrows in Fig. 3a). Most of the algal cells were penetrated and lipid globules were observed at the sites of penetration (Fig. 3b). Bacterial cells were detected close to these penetrated cells (black arrows in Fig. 3b).

image

Figure 2. (a–d) Scanning electron microscopy with backscattered electron imaging (SEM–BSE) images of Lecidea sp. (a) Lichen apothecium and a nearby network of fissures filled with fungal hyphae. (b) Detail of the apothecium shown in (a) in which the subhymenium (sh) and hymenium (h) layer can be distinguished. P (paraphyses); A (asci). (c) Apothecium within a fissure indicated by arrowheads and a dotted line. (d) Enlargement of the zone indicated by the black arrow in (a) showing algal and fungal symbiont cells. (e) TEM image showing endolithic Lecidea sp. fungal cells in close contact with mineral fragments (black arrows). (f) SEM–BSE image of Lecidea sp. showing photobiont cells penetrated by fungal haustoria (black arrows).

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image

Figure 3. (a–b) TEM images showing the fungal–algal interface of Lecidea sp. symbionts. Arrowheads point to the thinnest part of the haustoria. (a) Zone of algal cell penetrated by fungal haustoria. Black arrows indicate the haustoria sheath. (b) Trebouxia cell with numerous lipid bodies (L) in the zone of haustoria penetration. Black arrow indicates bacteria cells. (c–f) Scanning electron microscopy with backscattered electron imaging (SEM–BSE) images of Lecidea cancriformis. (c) General view of a network of fissures colonized by these species. (d) Detailed image of photobiont and mycobiont cells. (e) Fissure occupied by algal and fungal cells in which calcium oxalate deposits can be observed (white arrow). (f) Fissure colonized by mycobiont cells showing large deposits of presumptive biominerals in the form of calcium oxalate deposits.

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In rock samples from areas of higher altitude, L. cancriformis was frequently identified by ITS rDNA sequencing, even when epilithic apothecia were lacking. Our SEM–BSE study of these samples revealed internal fissures of the rock fully occupied by fungal and algal cells (Fig. 3c). This time, algal cells were not penetrated by the fungal partner (Fig. 3d). Several mineral deposits were observed closely associated with the cells in the fissures occupied by this lichen (arrow Fig. 3e). In some zones, these deposits covered extensive areas of the fissure (arrows in Fig. 3f), coinciding with zones lacking algal cells. EDS analysis revealed their basic composition was calcium, carbon and oxygen (black arrows in Fig. 4a) permitting their identification as calcium oxalate deposits. A further mineral alteration was observed close to these calcium oxalate deposits consisting of numerous holes (white arrows in Fig. 4a). This altered zone was shown by EDS to have a lower amount of calcium than nearby areas of unaltered substrate (white arrows in Fig. 4a), indicating calcium depletion. Potassium depletion in biotites was also observed in the proximity of this endolithic lichen, as shown in the EDS element distribution map in Fig. 4b. Chemical alterations were also noted in feldspar mineral grains found inside fissures (Fig. 4c). The EDS scan-line shown in this figure indicates a lower amount of calcium in peripheral than in the central areas of these grains.

image

Figure 4. (a) SEM–BSE and energy dispersive X-ray spectroscopy (EDS) elemental distribution maps of a zone colonized by Lecidea cancriformis. Black arrow indicates calcium oxalate deposits and white arrows point to an altered zone showing calcium depletion. (b) EDS potassium and iron distribution map showing biotites in areas of potassium depletion starting from the zone indicated by arrows. (c) Feldspar fragment found inside a granite fissure and EDS scan-line showing relative concentration changes in silicon (above) and calcium (below) along the line, indicating the depletion of calcium at the margins of the rock fragment. (d–f) SEM–BSE images of L. cancriformis detected under Umbilicaria aprina thalli in a coastal locality. (d) Zone close to the surface in which only fungal cells intermixed with several mineral fragments appeared. (e) Zone of the fissure showing the first algal cells. (f) Deeper zone showing numerous algal cells.

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Using molecular methods we were also able to identify endolithic lichens positioned under epilithic lichens. Fig. 4d–f show an endolithic form of L. cancriformis from the coast under an U. aprina thallus. The part of the fissure open to the surface was occupied by mineral fragments and fungal cells (Fig. 4d). Under this, a second zone could be identified in which the first algal cells appeared (Fig. 4e). Beneath this, the deeper cavities were seen to harbour many algal cells (Fig. 4f).

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

A link between epilithic apothecia and the endolithic growth form seems evident in many cases but is uncertain in others (Friedmann et al., 1988). This makes their identification using only morphological criteria almost impossible. However, the combined use of molecular and microscopy techniques enabled the identification of endolithic lichens (even in the absence of fruiting bodies), which can be precisely localized in the rock substrate and characterized at the ultrastructural level. High resolution microscopy procedures were needed to localize and characterize the lichen symbionts whereas molecular techniques served to identify them. Our approach also enabled us to simultaneously evaluate specific interrelations between the symbionts and the lithic substrate and deepen our understanding of the ecology of the biofilms formed at the interface of the substrate colonized by these lichens. In this way, the characterization of these endolithic biofilms can be complete and precise.

Lecidea cancriformis and Lecidea sp. were detected across an altitudinal range in Granite Harbour (Antarctica). The presence of L. cancriformis had been already reported in different Antarctic regions (Øvstedal & Lewis Smith, 2001). Several endolithic lichen thalli show a similar structure to ordinary epilithic forms when embedded in limestone (Fry, 1922; Pinna et al., 1998; Russell et al., 1998). However, in the endolithic lichens growing on granite examined here, it was not possible to discern all these layers. Indeed, the distribution of cells in the fissures and cracks of the granite substrate might hinder this kind of organization. However, while fungal cells occurred in all the fissures occupied by the lichen, algal cells only appeared in some zones. Different mycobiont–photobiont interfaces such as appressoria, and intraparietal and intracellular haustoria have been identified in the few endolithic lichen species studied so far (Plessl, 1963; Galun et al., 1971; Kushnir & Galun, 1977; Friedmann, 1982). These two endolithic Antarctic lichens differed in terms of their alga-fungus relationship. Frequent fungal penetration into the algal partner by means of intracellular haustoria was only observed in Lecidea sp. For endolithic growths, this ultrastructural trait was consistent with molecular identification.

It is not completely clear how these lichens colonize their endolithic habitat. Diversity and distribution patterns of lichen communities in Antarctica could be a combination of relict flora, long-distance dispersal and recent colonization (as suggested by Vincent, 2000; Romeike et al., 2002). The internal distribution of lichens in the lithic substrate can be the end result of several factors, differing on a small spatial scale. Local aspects such as rock composition, the presence and configuration of cracks, fissures or rock discolorations can alter the amount of light penetrating deeply, and among other factors, condition the presence of particular microorganisms in the substrate (Matthes et al., 2001). In addition, the endolithic development of certain organisms is also influenced by the presence of epilithic lichens or other endolithic microorganisms and their actions on the substrate, inducing the formation of microhabitats and chemical environments in rock substrates (De los Ríos et al., 2002). The observation of epilithic lichen thalli upon substrates colonized by endolithic Lecidea with the same photobiont species suggests that endolithic lichens may provide algal species that facilitate colonization by other lichens.

Pinna et al. (1998) attributed typical features of euendolithic organisms to several endolithic calcicolous lichens. The endolithic lichens of our study shared chasmoendolithic and euendolithic features. These lichens colonize fissures and cracks in the rock, but the alterations observed in the proximity of the fungal partner also indicates a perforating capacity. Adopting one particular endolithic ecological niche over another could depend of the features of the biofilm and the environmental conditions. The long-term survival of Antarctic endolithic communities is closely linked to weathering (Johnston & Vestal, 1993). The development of apothecia at the surface and penetration of fungal cells in fissures observed in the material examined can explain the biogeophysical action on the substrate related to both lichen species. Physical weathering by epilithic lichens is frequently associated with a biogeochemical action (Bjelland & Thorseth, 2002; De los Ríos et al., 2002). Several signs of biogeochemical substrate alterations were also detected in the vicinity of these endolithic lichens. Evidence of calcium and potassium biomobilization processes, as well as calcium oxalate accumulation, was observed around the mycobiont cells. Biomobilization has been frequently associated with the biogeochemical action of different lithobionts (Wierzchos & Ascaso, 1996, 1998; de los Ríos et al., 2003). Extensive calcium oxalate deposits were detected close to L. cancriformis thalli. However, it was not possible to attribute the presence of calcium oxalate to this species, since we found no such deposits in the vicinity of L. cancriformis detected on the coast. The presence of calcium oxalate deposits seemed here to be mainly related to the chemical composition of the substrate. Fungi colonizing granite with high amounts of feldspar plagioclase containing calcium were found to accumulate this biomineral. Bjelland et al. (2002) have also recently associated the presence of calcium oxalate with specific local geochemical features of the lichen-colonized substrate. The functions of calcium oxalate in lichens remain unclear. Calcium oxalate synthesis was interpreted by Wadsten & Moberg (1985) as a calcium detoxification mechanism, but this seems unlikely in these granite colonizers, since the amount of calcium in the substrate is low. Cells of endolithic biofilms densely accumulate in fissures and cavities where light is scarce. The presence of calcium oxalate in these biofilms could facilitate endolithic life if they act as radiation reflectors (Modenesi et al., 2000), especially in the presence of extracellular polymeric substances (EPS). It has been recently reported that biofilm organization and, more specifically, the ‘biofilm gel effect’ of EPS lead to a more efficient acquisition of solar energy (Decho et al., 2003).

Romeike et al. (2002) established ITS sequences in four Antarctic Umbilicaria collected from different sites. Their ITS rDNA sequence for Trebouxia, consistent with the sequence obtained in our study, corresponds to the only Trebouxia sequence that to our knowledge has been obtained from lichens collected around the Granite Harbour area. These results could indicate a low incidence of Trebouxia species in these harsh conditions and/or poor specificity of the mycobiont. Mycobionts less specific in their choice of photobiont are able to survive in conditions in which only some photobiont species exist. A low degree of selectivity towards the photobiont partner has been attributed to some Antarctic lichens for different reasons. High genetic diversity of Trebouxia was found in Umbilicaria species along a transect of the Antarctic peninsula (Romeike et al., 2002), while low diversity of Nostoc was reported in different cyanolichen species from maritime Antarctica (Wirtz et al., 2003). The authors related the low photobiont selectivity found in these mycobionts to the extreme environmental conditions. However, if the occurrence of algal species depends on environmental conditions, the availability of algae in the severe environment of this continental Antarctic region would be a more limiting factor for the mycobiont than its preferences for certain photobionts. It is likely that only certain resistant strains of Trebouxia are able to live in these peculiar environmental conditions. Only one dominant green alga rDNA sequence was also found by De la Torre et al. (2003) in cryptoendolithic communities of Beacon sandstone from the McMurdo Dry Valleys (Antarctica). Fully analysing these microbial communities and approaching the complex processes that take place within them involves identifying their members and understanding their functional relationships. We describe the use of molecular and microscopy tools to map the distribution of endolithic lichens and evaluate different aspects of their ecology. Across the range of altitudes examined, we identified two Lecidea lichens showing a different distribution pattern and established the existence of a close relationship with their immediate environment.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

The authors would like to thank Fernando Pinto and Sara Lapole for technical assistance, Dr M. Castello (Trieste, Italy) for helping with taxonomic identification and Ana Burton for reviewing the English. We are especially grateful to Prof. A.T.G. Green and Antarctica New Zealand for their logistic support and excellent field facilities. This study was funded by grants REN2003-07366-C02-02, BOS2003-02418 of the Plan Nacional I+D and Acciones Integradas funds (ÖAD/MCyT). MG acknowledges support by FWF P11998.

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  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
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