The Biology of Arid Soils 9783110419047, 9783110419986

Citation preview

Blaire Steven (Ed.) The Biology of Arid Soils Life in Extreme Environments

Life in Extreme Environments

| Edited by Dirk Wagner

Volume 4

The Biology of Arid Soils |

Editor Blaire Steven Department of Environmental Sciences Connecticut Agricultural Experiment Station 123 Huntington Street New Haven, CT 06511, USA [email protected]

ISBN 978-3-11-041998-6 e-ISBN (PDF) 978-3-11-041904-7 e-ISBN (EPUB) 978-3-11-041914-6 ISSN 2197-9227 Library of Congress Cataloging-in-Publication Data A CIP catalog record for this book has been applied for at the Library of Congress. Bibliographic information published by the Deutsche Nationalbibliothek The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliografie; detailed bibliographic data are available on the Internet at http://dnb.dnb.de. © 2017 Walter de Gruyter GmbH, Berlin/Boston Cover image: Medioimages/Photodisc/thinkstock Typesetting: le-tex publishing services GmbH, Leipzig Printing and binding: CPI books GmbH, Leck ♾ Printed on acid-free paper Printed in Germany www.degruyter.com

Preface When Dr. Dirk Wagner asked me to edit an edition in the series “Life in Extreme Environments” on the topic of arid soils, I was a little surprised. Other books in the series discussed life in the deep ocean, caves, and Earth’s thermal vents. Studies where scientists require large field campaigns, submersible vehicles, and potential personal risk to collect samples. In contrast, many people could collect a sample of arid soil in a brisk walk from wherever they may be reading this. In this regard, arid soils did not seem to be such an “extreme” of an environment. Yet, arid soils are united by a common characteristic, namely water scarcity, which limits the diversity and productivity of these systems. Furthermore, arid ecosystems also occur in both the hottest and coldest regions of the planet and therefore may experience a multitude of other severe environmental conditions. So, in many respects arid soils may be as harsh of an environment as more treacherous locals. Soil has been described as one of nature’s most complex ecosystems. Thus, any scientist that takes on the study of soil biology faces a daunting task. By the virtue of arid soil organisms existing at the low water availability to support life, these communities tend to be simplified compared to more temperate soils. The collection of papers in this volume highlight the work of researchers that are employing arid soils to understand the limits of life under low water availability, the functioning of soil ecosystems, and predicting how these systems will respond to an altered climate. In putting together this volume I called in favors from collaborators, met new colleagues, and learned more about arid soils than I knew before. I was also able to include photographs taken by my father on his various travels (see Figure 1.1). He has always been a hobbyist, but can know say he is a published photographer. Congratulations dad. The list of contributing authors to this volume highlights the international scope of arid land research and the broad disciplines involved. Like any good work of science I hope this work raises as may questions for future research as it answers for those with the curiosity to read it. Blaire Steven

Volumes published in the series Volume 1 Jens Kallmeyer, Dirk Wagner (Eds.) Microbial Life of the Deep Biosphere ISBN 978-3-11-030009-3

Volume 2 Corien Bakermans (Ed.) Microbial Evolution under Extreme Conditions ISBN 978-3-11-033506-4

Volume 3 Annette Summers Engel (Ed.) Microbial Life of Cave Systems ISBN 978-3-11-033499-9

Contents Preface | V Contributing authors | XI Blaire Steven 1 An Introduction to Arid Soils and Their Biology | 1 1.1 The Definition and Extent of Arid Ecosystems | 1 1.2 Characteristics of Arid Soils | 2 1.3 Soil Habitats in Arid Regions | 2 1.3.1 Refugia Sites Associated with Rocks | 3 1.3.2 Shrubs as Islands of Fertility | 3 1.3.3 Biological Soil Crusts | 5 1.4 The Pulse Reserve Paradigm of Arid Ecosystems | 6 1.5 Response of Arid Ecosystems to Disturbance | 7 1.6 Arid Ecosystems as a Model for Soil Biology | 7 1.7 Summary | 7 Carlos Garcia, J.L.Moreno, T. Hernandez, and F. Bastida 2 Soils in Arid and Semiarid Environments: the Importance of Organic Carbon and Microbial Populations. Facing the Future | 15 2.1 Introduction | 15 2.2 Climate Regulation and Soil Organic Carbon in Arid-Semiarid Zones | 16 2.3 Land Use and Soil Organic Carbon in Arid-Semiarid Zones | 17 2.4 Soil Restoration in Arid-Semiarid Zones: Amendments Based on Exogenous Organic Matter | 18 2.5 Microbial Biomass and Enzyme Activity in Arid-Semiarid Zones | 19 2.6 Organic Carbon, Macro and Microaggregates, and C Sequestration in Arid-Semiarid Zones | 22 2.7 Conclusion | 23 Gary M. King 3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange in Arid and Semiarid Ecosystems | 31 3.1 Introduction | 31 3.2 Water Potential and Water Potential Assays | 32 3.3 Limits of Growth and Metabolic Activity | 35 3.4 Water Potential and Trace Gas Exchanges | 37 3.5 Conclusions | 41

VIII | Contents

Thulani P. Makhalanyane, Storme Z. de Scally, and Don A. Cowan 4 Microbiology of Antarctic Edaphic and Lithic Habitats | 47 4.1 Introduction | 47 4.2 Classification of Antarctic soils | 48 4.2.1 McMurdo Dry Valley Soils | 49 4.2.2 Antarctic Peninsula Soils | 50 4.3 Bacterial Diversity of Soils in the MDVs and Antarctic Peninsula | 51 4.4 Cryptic Niches in Antarctic Environments | 54 4.4.1 Hypoliths | 55 4.4.2 Epiliths | 56 4.4.3 Endoliths | 57 4.5 Biogeochemical Cycling in Antarctic Environments | 59 4.6 Viruses in Antarctic Edaphic Ecosystems | 59 4.7 Conclusions and Perspectives | 60 Matthew A. Bowker, Burkhard Büdel, Fernando T. Maestre, Anita J. Antoninka, and David J. Eldridge 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences | 73 5.1 Overview | 73 5.1.1 Moss, Liverwort, and Lichen Biology | 73 5.2 Global Diversity and Characteristic Taxa | 74 5.2.1 Global Species Pool | 74 5.2.2 Global Characteristic Taxa and β Diversity | 75 5.3 Determinants of Moss, Liverwort, and Lichen Diversity on Arid Soils | 78 5.3.1 Geographic Isolation and Biogeography | 78 5.3.2 Climatic Gradients and Climate Change | 79 5.3.3 Calcicole–Calcifuge Dichotomy and Soil pH Gradients | 80 5.3.4 The Special Case of Gypsiferous Soils | 81 5.4 Consequences of Moss, Liverwort, and Lichen Diversity on Arid Soils | 82 5.4.1 Contribution of Biocrust Lichens and Bryophytes to Arid Ecosystem Function | 82 5.4.2 Biodiversity–Ecosystem Functioning Relationship | 83 5.4.3 Effects of Species Richness, Turnover, and Evenness on Ecosystem Functions | 84 5.4.4 Multifunctionality | 87 5.4.5 Functional Redundancy or Singularity? | 88 5.5 Summary and Conclusions | 89

Contents | IX

Andrea Porras-Alfaro, Cedric Ndinga Muniania, Paris S. Hamm, Terry J. Torres-Cruz, and Cheryl R. Kuske 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils | 97 6.1 Spatial Heterogeneity of Fungal Communities in Arid Lands | 97 6.1.1 Biocrusts | 100 6.1.2 Plant Associated Fungi in Deserts | 103 6.2 Roles in Nutrient Cycling and Effects of Climate Change on Fungal Communities | 107 6.3 Extremophiles in Deserts | 108 6.3.1 Thermophilic and Thermotolerant Fungi | 109 6.3.2 Rock Varnish and Microcolonial Fungi in Deserts | 109 6.4 Human Pathogenic Fungi in Desert Ecosystems | 111 6.4.1 Coccidioides immitis and C. posadasii | 112 6.4.2 Dematiaceous and Keratinolytic Fungi in Deserts | 112 6.4.3 Eumycetoma | 113 6.4.4 Mycotoxins | 114 6.5 Importance of Fungal Biodiversity in Arid Lands | 115 T.G. Allan Green 7 Limits of Photosynthesis in Arid Environments | 123 7.1 Introduction | 123 7.2 Photosynthetic Responses to Environmental Factors, a Background | 124 7.2.1 Rates, Chlorophyll and Mass | 124 7.2.2 Response of Net Photosynthesis (NP) to Light (PPFD, μmol m−2 s−1 ) | 126 7.2.3 Response of Net Photosynthesis to Temperature | 127 7.2.4 Response of Net Photosynthesis to Thallus Water Content (WC) | 127 7.2.5 Response of Net Photosynthesis to CO2 Concentration | 129 7.3 Optimal Versus Real Photosynthetic Rates | 129 7.4 Limits to Photosynthesis in Arid Areas | 131 7.4.1 Length of Active Time | 131 7.4.2 Limits When Active – External Limitation Through Light and Temperature | 132 7.4.3 Limits When Active – Internal Limitation Through Thallus Hydration | 132 7.4.4 Catastrophes | 133 7.5 Flexibility – an Often Overlooked Factor | 134 7.6 Summary | 134

X | Contents

Blaire Steven, Theresa A. McHugh, and Sasha Reed 8 The Response of Arid Soil Communities to Climate Change | 139 8.1 Overview | 139 8.2 Biological Responses to Elevated Atmospheric CO2 | 140 8.3 Biological Responses to Increased Temperature | 142 8.4 Biological Responses to Changes in Precipitation | 143 8.4.1 Natural Precipitation Gradients | 145 8.4.2 Precipitation Manipulation Studies | 147 8.5 Interactions Between Temperature and Soil Moisture | 149 8.6 Conclusion | 150 Doreen Babin, Michael Hemkemeyer, Geertje J. Pronk, Ingrid Kögel-Knabner, Christoph C. Tebbe, and Kornelia Smalla 9 Artificial Soils as Tools for Microbial Ecology | 159 9.1 Introduction | 159 9.2 Soil Definition | 160 9.3 History of Artificial Soil Experiments | 162 9.4 Methods in Soil Microbial Ecology and Soil Science | 164 9.5 Insights into Microbial Communities from Artificial Soil Studies | 166 9.5.1 Establishment and Structuring of Soil Microbial Communities | 166 9.5.2 Functioning of Soil Microbial Communities | 169 9.6 Artificial Soils for Arid Soil Research | 174 9.7 Concluding Remarks | 175 Index | 181

Contributing authors Anita J. Antoninka School of Forestry Northern Arizona University Flagstaff, Arizona, 86011, USA e-mail: [email protected]

Doreen Babin Julius Kühn-Institut – Federal Research Centre for Cultivated Plants (JKI) Institute for Epidemiology and Pathogen Diagnostics Braunschweig, Germany e-mail: [email protected]

Felipe Bastida Department of Soil and Water Conservation, CEBAS-CSIC Campus Universitario de Espinardo Murcia, Spain. e-mail: [email protected]

Matthew A. Bowker School of Forestry Northern Arizona University Flagstaff, Arizona, 86011, USA e-mail: [email protected]

Burkhard Büdel Plant Ecology & Systematics Faculty of Biology University of Kaiserslautern Kaiserslautern, Germany e-mail: [email protected]

Don A. Cowan Centre for Microbial Ecology and Genomics Department of Genetics, Natural Sciences 2 University of Pretoria Hatfield, Pretoria, USA e-mail: [email protected]

Storme Z. de Scally Centre for Microbial Ecology and Genomics Department of Genetics, Natural Sciences 2 University of Pretoria Hatfield, Pretoria, 0028 e-mail: [email protected] David J. Eldridge Centre for Ecosystem Studies School of Biological, Earth and Environmental Sciences University of New South Wales Sydney, Australia e-mail: [email protected] Carlos García Department of Soil and Water Conservation CEBAS-CSIC, Campus Universitario de Espinardo Murcia, Spain e-mail: [email protected] T. G. Allan Green Departamento de Vegetal II, Farmacia Facultad Universidad Complutense 28040, Madrid, Spain e-mail: [email protected] Paris S. Hamm Department of Biological Sciences Western Illinois University Macomb, Illinois, USA e-mail: [email protected] Michael Hemkemeyer Thünen Institute of Biodiversity Federal Research Institute for Rural Areas, Forestry and Fisheries Braunschweig, Germany Present address: Faculty of Life Sciences Rhine-Waal University of Applied Sciences Kleve, Germany e-mail: [email protected]

XII | Contributing authors Teresa Hernández Department of Soil and Water Conservation CEBAS-CSIC, Campus Universitario de Espinardo Murcia, Spain e-mail: [email protected] Gary M. King Department of Biological Sciences Louisiana State University Baton Rouge, Louisiana 70803, USA e-mail: [email protected] Ingrid Kögel-Knabner Lehrstuhl für Bodenkunde, Technische Universität München Freising-Weihenstephan, Germany Institute for Advanced Study, Technische Universität München Garching, Germany e-mail: [email protected] Cheryl R. Kuske Bioscience Division Los Alamos National Laboratory Los Alamos, New Mexico, USA e-mail: [email protected] Fernando T. Maestre Departamento de Biología y Geología, Física y Química Inorgánica Escuela Superior de Ciencias Experimentales y Tecnología Universidad Rey Juan Carlos Móstoles, Spain e-mail: [email protected] Thulani P. Makhalanyane Centre for Microbial Ecology and Genomics Department of Genetics, Natural Sciences 2 University of Pretoria Hatfield, Pretoria, USA e-mail: [email protected] Theresa A. Mchugh Southwest Biological Science Center U.S. Geological Survey Moab, Utah, USA e-mail: [email protected]

José Luis Moreno Department of Soil and Water Conservation CEBAS-CSIC, Campus Universitario de Espinardo Murcia, Spain e-mail: [email protected]

Cedric Ndinga Muniania Department of Biological Sciences Western Illinois University Macomb, Illinois, USA e-mail: [email protected]

Andrea Porras-Alfaro Department of Biological Sciences Western Illinois University Macomb, Illinois, USA e-mail: [email protected]

Geertje J. Pronk Lehrstuhl für Bodenkunde, Technische Universität München Freising-Weihenstephan, Germany Institute for Advanced Study, Technische Universität München Garching, Germany Present address: Ecohydrology Research Group, University of Waterloo Waterloo, Ontario, Canada e-mail: [email protected]

Sasha Reed Southwest Biological Science Center U.S. Geological Survey Moab, Utah, USA e-mail: [email protected]

Kornelia Smalla Julius Kühn-Institut – Federal Research Centre for Cultivated Plants (JKI) Institute for Epidemiology and Pathogen Diagnostics Braunschweig, Germany e-mail: [email protected]

Contributing authors

Blaire Steven Department of Environmental Sciences Connecticut Agricultural Experiment Station New Haven, CT, USA e-mail: [email protected] Christoph C. Tebbe Thünen Institute of Biodiversity Federal Research Institute for Rural Areas, Forestry and Fisheries Braunschweig, Germany e-mail: [email protected]

Terry J. Torres-Cruz Department of Biological Sciences Western Illinois University Macomb, Illinois, USA e-mail: [email protected]

|

XIII

Blaire Steven

1 An Introduction to Arid Soils and Their Biology 1.1 The Definition and Extent of Arid Ecosystems When one invokes the terms arid ecosystem or dryland it is often assumed that the term refers to a desert. However, there are regional differences in the concept of a “desert” as well as differences in terms for describing and classifying arid lands. The one characteristic that unites all arid lands is a lack of water availability, generally due to low precipitation. Yet, lack of precipitation is not the only factor that limits water availability. Water can be lost from the landscape through evaporation and transpiration, and the evaporative loss of water from plants. Together these processes are referred to as evapotranspiration [1]. Thus, the “dryness” of a region can be determined by calculating the net difference between precipitation and water losses through evapotranspiration, also referred to as the Aridity Index [2–4]. These metrics have been a useful tool to generate a standardized method to categorize and define drylands. The aridity index, as well as other metrics such as the dominant vegetation and climate, have been used to classify arid lands into three main categories (󳶳 Fig. 1.1): Hyperarid zone (arid index 0.03 or below): Dryland areas of scant or no vegetation. Annual rainfall is low, rarely exceeding 100 mm. Precipitation events are infrequent and irregular, with dry periods lasting up to several years. Hyperarid regions cover ∼ 8% of the Earth’s surface [5]. Examples: Atacama Desert, South America; Namib Desert and Sahara Desert, Africa; and Lut Desert, Iran. Arid zone (arid index 0.03–0.20): Vegetation consists of sparsely distributed patches of annual or perennial grasses, patchily distributed shrubs, cacti, or small trees. Maximum precipitation varies from 100–300 mm per year. Arid zones cover ∼ 16% of the planet’s land surface. Examples: Chihuahuan Desert, U.S.A. and Simpson Desert, Australia.

(a)

(b)

(c)

Fig. 1.1: Examples of different arid zone landscapes. (a) Hyperarid zone. Namib Desert, South Africa. Photo courtesy Don Cowan. (b) Arid zone. Saguaro National Park, Arizona, U.S.A, (c) Semiarid zone. Witfontein Nature Reserve grassland, South Africa. Photos b and c courtesy Douglas Steven. DOI 10.1515/9783110419047-001

2 | 1 An Introduction to Arid Soils and Their Biology

Semiarid zone (arid index 0.20–0.50): Vegetation is more diverse and may cover the surface. For instance, semiarid grasslands or steppes are common. Annual precipitation can reach 800 mm per year and may occur in distinct dry and wet seasons. Semiarid zones cover ∼ 18% of the Earth. Examples: Great Plains U.S.A, Kenyan Savanah, and Mongolian Steppes. It is important to note that not all arid soils occur in regions classified as drylands. Isolated patches of arid soils can occur in otherwise temperate regions, for example, alpine tundra or volcanic cinders [6, 7].

1.2 Characteristics of Arid Soils Arid soils possess unique characteristics that distinguish them from soils from more humid regions. Arid systems are generally limited in biological activity and thus contain low levels of organic carbon. This lack of organic carbon is a large driver in the structuring and function of arid soils and is the focus of Chapter 2. Extended periods of water deficiencies also slow the elimination or leaching of soluble salts, which are further accumulated due to high rates of evaporation [8]. Thus, arid soils tend to accumulate calcium carbonate, gypsum, or silica [9]. Despite similarities in soil genesis, the different climates, geology, and vegetation of arid lands create unique soil characteristics, so that the morphology and soil characteristics vary between different drylands [10]. The water holding capacity of a soil depends on its physical characteristics, including texture, structure, and soil depth [11]. This leads to large differences in the available water for biology between different soils. The critical importance in water potential is discussed in Chapter 3. So soil characteristics play an integral role in determining the composition and function of arid soil biological communities. In fact, soil parent material and chemistry have been found to play a large role in shaping arid soil biology [12, 13]. In this respect, local edaphic factors need to be included in any study of arid soil biology.

1.3 Soil Habitats in Arid Regions A characteristic of arid regions is reduced biological diversity. This has been well documented for vegetation (e.g., [13–16]) and other macro fauna [18]. Similar patterns have emerged for soil bacterial and fungal communities [19, 20]. In fact, a global survey of drylands worldwide found that the diversity of soil bacteria and fungi was linearly correlated to the aridity of the ecosystem [21]. In this regard, aridity is a large predictor of the diversity of soil communities. However, drylands are not homogenous regions experiencing low precipitation. Arid regions are patchy at a variety of scales. The vegetation is sparse, soil edaphic factors vary, the terrain is uneven, and precipitation and temperature vary erratically [22–25]. In this respect, not every patch of arid soil

1.3 Soil Habitats in Arid Regions

| 3

is created equally. Certain niches in drylands differ in their ability to support biological communities. For example, aspects of the landscape such as slope or shading that may alter water retention of the soil have the potential to alter the abundance and diversity of the communities the soil can support [26]. This results in distinct ecological niches, some of which are discussed below.

1.3.1 Refugia Sites Associated with Rocks In hyperarid deserts, the shelter provided within the shade of a rock can be the difference between life and death. These lithic associated communities often inhabit regions so devoid of moisture that a significant portion of their water requirements is met by fog rather than precipitation [27, 28]. Rocks in deserts can support a number of different communities. These include: hypolithic communities inhabiting the basal surface of rocks [29, 30], endolithic communities that live inside rocks or pores between mineral grains [31–34], and chasmolithic communities under rock flakes produced by weathering [35, 36]. Rocks provide the soil microbiota physical stability, increased water retention by shading, protection from ultraviolet radiation, and micronutrients from the mineral components of the rock material [37]. Translucent rocks allow for light transmission to a depth sufficient to support phototrophs, such as mosses or cyanobacteria. A common cyanobacteria occurring in hypolithic niches is Chroococcidiopsis sp. [38], which has been detected in deserts worldwide [39]. These phototrophic populations fix carbon, which can then feed heterotrophic populations, resulting in relatively complex ecosystems [35, 40]. Thus, these communities act as a source of organic carbon, which is a valuable commodity in otherwise nearly barren soils [41]. Additionally, the presence of active biology can accelerate the weathering of the rocks. This can occur either by metabolic activity of the communities, scavenging nitrogen or phosphorous from the rock material, which has been shown to increase the weathering rate of rock by up to three orders of magnitude, or by physical infiltration into rock crevices and the mechanical disruption of porous stones [42–44]. These communities can also increase weathering by encouraging grazing and the associated scraping of rock surfaces by predatory invertebrates [45]. So beyond fixing organic carbon, rock associated communities can also release limiting nutrients supporting the growth of multiple trophic levels. In this respect, even the interspersed rocks in the desert can act as abiotic oases for soil biology.

1.3.2 Shrubs as Islands of Fertility In arid ecosystems where plants are sparse, a shrub is often a conspicuous aspect of the ecosystem. As wind moves across the landscape the canopy of the shrub can dis-

4 | 1 An Introduction to Arid Soils and Their Biology

rupt currents, collecting dust [46]. Later, precipitation moving through the canopy of the shrub can pick up this deposited dust and other plant litter, transporting this material to the under canopy soils [47]. Analyses of fall water have shown that it contains up to ten times more nutrients than bulk precipitation occurring outside of the shrub canopy [48]. Thus this material can act to fertilize soils in the canopy zone of the shrub. Additionally, shrubs supply nest sites, shade, and food resources for animal populations, which can enrich the local soils through feces, discarded carcasses, and nest materials [49]. Shrubs are also important in the interception, infiltration, and storage of water, thereby increasing soil moisture [50]. Finally, the shrub itself contributes to the enrichment of soil nutrients. In addition, litter production, root exudates, and deadfall all contribute to enriching the soils in the vicinity of the shrub [51]. Thus shrubs in drylands are potent collectors of resources and [52, 53] are often referred to as “islands of fertility” [54]. Shrubs also act as a cradle for biological diversity, protecting the communities from ultraviolet radiation and decreasing evaporation through shading [55]. Nutrients in the shrub root zone are vertically distributed with the majority of nutrients being a few millimeters under the surface [53, 56]. This suggests a low mixing of the soils and implicates litter production as a large source of the resource accumulation [57]. Shrub canopy zone soils support increased microbial activity, as soil respiration rates are generally higher in shrub root zone soils than in interspace soils (e.g., [57–59]). This effect seems to be specific to shrubs as similar increases are not apparent in the vicinity of annual grasses [59]. Despite consistent findings of increased metabolic activity in under shrub soils, the characteristics of the biologic communities in shrub zones versus interspace soils are not as uniform. Shrub zone soils tend to support a higher abundance of macroinvertebrates and nematodes [61–63], although shrub zone soils may harbor similar or even decreased levels of insect diversity [64]. For soil bacteria and fungi, studies have found an increased [65–67] or no effect [68] on their abundance, although the composition of the communities between the two habitat types generally differs [69]. More recently, studies employing replicated sequencing datasets have shown that the differences between the shrub associated communities and interspaces were primarily due to a difference in the abundance of the species rather than the membership of the communities (󳶳 Fig. 1.2 [68, 70]). In other words, shrub canopy soils harbor roughly the same bacteria and fungi as interspace soils, but the structure of the community differs. This has two important implications. First, it suggests that the bacteria and fungi that are well adapted to inhabiting arid soils may be ubiquitous across the landscape, even in habitat patches that show different characteristics. Secondly, there may be a relatively small number of bacterial and fungal species that need to be accounted for to understand biogeochemical cycles and functioning of arid soils.

1.3 Soil Habitats in Arid Regions

0

5

Shared Biocrusts

Root zones

Roots Biocrusts

Roots Biocrusts

5

B. Fungal OTUs % of sequence reads 30 20 10 0 10 20 30 40 50 60

10

Shared

A. Bacterial OTUs % of sequence reads 25 20 15 10 5

|

Biocrusts

Root zones

Fig. 1.2: Similarity in membership of bacteria and fungi between dryland habitats. Each panel denotes the relative abundance of either bacterial of fungal operational taxonomic units (OTUs) in biocrusts or the root zones of creosote bushes. The OTUs are split into three categories, OTUs shared between the habitat patches, those unique to biocrusts, and those unique to the root zones. For both the bacteria and fungi the most abundant OTUs were shared between the habitats, suggesting a similar membership for the communities in both habitats, although the abundance of those same OTUs varied widely between the two habitats. Thus the membership of the communities is similar, although the structure may vary. Figure adapted from [68].

1.3.3 Biological Soil Crusts The surface soils between rocks and plants of arid regions are not devoid of life. In fact, some of the most diverse arid soil communities occur in plant interspaces of arid and semiarid lands as communities colonizing surface soils. These communities form a surface crust that has been variously referred to as cryptogamic, microbiotic, cryptobiotic, or microphytic [71]. More inclusively, the term biological soil crusts (shortened to biocrusts for this chapter) has been used to refer to the biological crusts that inhabit a multitude of arid lands [72, 73]. In some arid lands, biocrusts cover up to 60–70% of the surface soils [74]. Biocrusts have been identified on every continent on Earth and are a conspicuous feature of drylands worldwide [75]. The keystone species of most biocrusts are cyanobacteria [76–78]. Filamentous species of cyanobacteria, predominantly in the order Oscillatoriales, such as Microcoleus vaginatus form the structural component of the biocrusts [79]. These organisms bind soil particles together and produce fixed carbon for other community members [80]. Some of this carbon is in the form of extracellular polymeric substances that act as the glue to bind the soil together and the matrix to create the surface crust biofilm [81]. Other cyanobacteria in the biocrusts fix atmospheric nitrogen or produce pigments, such as scytonemin, that protect the crust organisms from ultraviolet radiation [82–84]. Beyond cyanobacteria, biocrusts harbor mosses, lichens, fungi, algae, a variety of heterotrophic bacteria, and archaea [85–89]. This also leads to an enrichment of other soil fauna, as nematode populations are more abundant and diverse in mature biocrusts [88]. Because the dominant species of biocrusts are phototrophic,

6 | 1 An Introduction to Arid Soils and Their Biology

the biomass of the crusts is concentrated in the upper few millimeters of soil, but leaching of these nutrients can enrich surrounding and underlying soils [56]. In this regard, biocrusts are a complex and diverse ecosystem that support multiple trophic levels and enrich the surrounding soils. Biocrusts perform a multitude of ecological services. The pinnacled and roughened surface of biocrusts trap dust, collecting nutrients and aiding in water retention [90, 91]. The physical binding of soil particles increases aeration and reduces soil erosion by wind and water [92–95]. Biocrusts are a significant source of fixed carbon and nitrogen in a landscape where plants are sparse [96]. The presence of well developed biocrusts can elevate the amount of organic carbon by 3000% compared to surrounding bare soils [75]. Similarly, biocrusted soils have been found to enrich nitrogen by a factor of 200%, the majority of which is rapidly leached into surrounding soils [97–99]. This nutrient trapping and leaching may also assist in the establishment and development of desert plants [100–102]. Some evidence even suggests that there may be fungal nutrient bridges that allow for the passage of nutrients between biocrusts and plants [103, 104]. In this respect, biocrusts are not isolated soil patches of increased soil fertility but are an integral component to dryland ecosystem function.

1.4 The Pulse Reserve Paradigm of Arid Ecosystems Dryland ecosystems are not just defined by a lack of water; precipitation occurs as episodic events. Therefore, an essential resource (water) is only available in pulses, with large intervening periods of limitation. In this respect, it is not enough to consider the amount of available water only but also the size, duration, and periodicity of precipitation events. In 1973, Noy-Meir [105] proposed the “pulse reserve” model of production in arid systems. Conceptually the model proposes that a pulse of water, provided through a precipitation event, stimulates the initiation of biological activity (generally photosynthesis). After a period of activity the organism builds reserves of energy to sustain it through the following dry period and to the next pulse. This model was developed for dryland plants but it has also been shown to be applicable to mosses [106] and cyanobacteria [107]. A central aspect of this model is that precipitation events need to be “biologically meaningful,” in that the water needs to of sufficient amount and duration to stimulate biological activity [108]. This sets up a hierarchical response to precipitation events. Small precipitation events will stimulate soil cyanobacteria or algae but are inadequate to initiate plant activity [109]. For example, it has been estimated that ∼ 2 mm precipitation events are generally adequate to activate soil cyanobacteria within a few minutes, whereas plants may require in the range of 3–5 mm of precipitation with soil moisture lasting for at least an hour [11]. In this respect, understanding dryland ecosystems extends beyond just considering the limitation of water and must consider the magnitude, duration, and timing of precipi-

1.7 Summary

|

7

tation events. The factors in drylands that act to limit photosynthesis, thus constraining the buildup of reserves, are discussed in Chapter 7.

1.5 Response of Arid Ecosystems to Disturbance Arid lands are under threat from a variety of sources. Human impact due to agriculture, recreation, and mineral extraction all dramatically affect arid lands worldwide [110, 111]. Changes in climate are warming drylands and changing precipitation patterns [112]. Because arid soil communities survive at the lower thresholds of water availability to support life, even small disturbances have the potential to alter the composition and function of arid soil communities dramatically. As a consequence of the low biodiversity of arid soils there are generally lower levels of functional redundancy in the community [113]. Thus the loss of a community member may result in a tipping point at which the community may not easily recover. Experimental manipulations testing the effects of chronic physical disturbance and climate change perturbations have been conducted in drylands and show that the structure and functioning of arid soil communities can be severely altered by even relatively small perturbations [106, 107]. Chapter 8 investigates how dryland communities respond to perturbations, particularly those associated with climate change.

1.6 Arid Ecosystems as a Model for Soil Biology As mentioned previously, arid soils generally harbor less diverse soil communities than other soils. Further, arid soils also often show a characteristic of trophic simplicity, the communities of arid soils are generally composed of only a limited number of trophic levels, and these levels generally become more simple as the environment becomes more extreme [35]. This relatively low biodiversity and complexity allows researchers to disentangle the biologic, climatic, and environmental factors that drive the composition and functioning of ecosystems more easily. Thus, arid soil systems have been proposed as a system to understand biodiversity ecosystem function relationships better [114]. In Chapter 9 artificial soil microcosms and their contribution to understanding soil biological processes are discussed.

1.7 Summary The Earth’s drylands are a diverse patchwork of systems united by a common feature of limited water availability. While the differences between drylands are numerous, certain aspects of limited moisture lead to predictable patterns in the diversity, energetics, and composition of soil communities. The purpose of this book is to document

8 | 1 An Introduction to Arid Soils and Their Biology

what is known about these patterns and to try to disentangle the biotic and abiotic factors that shape the distinct, unique, and often overlooked soil communities of arid lands.

References [1] [2] [3] [4] [5] [6] [7]

[8] [9] [10] [11] [12]

[13]

[14] [15] [16] [17]

[18] [19]

Sellers WD. Potential Evapotranspiration in Arid Regions. J Appl Meteorol 1964, 3:98–104. Girvetz EH, Zganjar C. Dissecting indices of aridity for assessing the impacts of global climate change. Clim Change 2014, 126:469–83. Tsakiris G, Vangelis H. Establishing a drought index incorporating evapotranspiration. Eur Water 2005, 9:3–11. Levin NE, Cerling TE, Passey BH, Harris JM, Ehleringer JR. A stable isotope aridity index for terrestrial environments. Proc Natl Acad Sci 2006, 103:11201–5. Tucker CJ, Newcomb WW, Dregne HE. AVHRR data sets for determination of desert spatial extent. Int J Remote Sens 1994, 15:3547–65. Taylor RV, Seastedt TR. Short- and long-term patterns of soil moisture in alpine tundra. Arct Alp Res 1994, 26:14. Weber CF, King GM. Distribution and diversity of carbon monoxide-oxidizing bacteria and bulk bacterial communities across a succession gradient on a Hawaiian volcanic deposit, CO oxidizer diversity across a succession gradient. Environ Microbiol 2010, 12:1855–67. Ewing SA, Sutter B, Owen J, et al. A threshold in soil formation at Earth’s arid–hyperarid transition. Geochim Cosmochim Acta 2006, 70:5293–322. Skujins J. Genesis and Classification of Arid Region Soils. In: Semiarid Lands and Deserts, Soil Resource and Reclamation. CRC Press, 1991, 33. Bronick CJ, Lal R. Soil structure and management: a review. Geoderma 2005, 124:3–22. Austin AT, Yahdjian L, Stark JM, et al. Water pulses and biogeochemical cycles in arid and semiarid ecosystems. Oecologia 2004, 141:221–35. Steven B, Gallegos-Graves LV, Belnap J, Kuske CR. Dryland soil microbial communities display spatial biogeographic patterns associated with soil depth and soil parent material. FEMS Microbiol Ecol 2013, 86:101–13. Deng H, Yu Y-J, Sun J-E, et al. Parent materials have stronger effects than land use types on microbial biomass, activity and diversity in red soil in subtropical China. Pedobiologia 2015, 58:73–9. Qian H, Ricklefs RE. A latitudinal gradient in large-scale beta diversity for vascular plants in North America. Ecol Lett 2007, 10:737–44. von Hardenberg J, Meron E, Shachak M, Zarmi Y. Diversity of vegetation patterns and desertification. Phys Rev Lett 2001, 87:198101. Kreft H, Jetz W. Global patterns and determinants of vascular plant diversity. Proc Natl Acad Sci 2007, 104:5925–30. Davenport ML, Nicholson SE. On the relation between rainfall and the Normalized Difference Vegetation Index for diverse vegetation types in East Africa. Int J Remote Sens 1993, 14:2369– 89. Abramsky Z, Rosenzweig ML. Tilman’s predicted productivity–diversity relationship shown by desert rodents. Nature 1984, 309:150–1. Dunbar J, Takala S, Barns SM, Davis JA, Kuske CR. Levels of bacterial community diversity in four arid soils compared by cultivation and 16S rRNA gene cloning. Appl Environ Microbiol 1999, 65:1662–9.

References | 9

[20] [21] [22] [23] [24] [25] [26] [27]

[28]

[29] [30] [31] [32] [33] [34] [35] [36] [37]

[38] [39]

[40]

[41]

Whitford WG. The importance of the biodiversity of soil biota in arid ecosystems. Biodivers Conserv 1996, 5:185–95. Maestre FT, Delgado-Baquerizo M, Jeffries TC, et al. Increasing aridity reduces soil microbial diversity and abundance in global drylands. Proc Natl Acad Sci 2015, 112:15684–89. Huenneke LF, Clason D, Muldavin E. Spatial heterogeneity in Chihuahuan Desert vegetation, implications for sampling methods in semi-arid ecosystems. J Arid Environ 2001, 47:257–70. Aguiar MR, Sala OE. Patch structure, dynamics and implications for the functioning of arid ecosystems. Trends Ecol Evol 1999, 14:273–7. Kéfi S, Rietkerk M, Alados CL, et al. Spatial vegetation patterns and imminent desertification in Mediterranean arid ecosystems. Nature 2007, 449:213–7. Maestre FT, Cortina J. Spatial patterns of surface soil properties and vegetation in a Mediterranean semi-arid steppe. Plant Soil 2002, 241:279–91. Burke A. Properties of soil pockets on arid Nama Karoo inselbergs–the effect of geology and derived landforms. J Arid Environ 2002, 50:219–34. Warren-Rhodes KA, McKay CP, Boyle LN, et al. Physical ecology of hypolithic communities in the central Namib Desert, The role of fog, rain, rock habitat, and light. J Geophys Res Biogeosciences 2013, 118:1451–60. Cáceres L, Gómez-Silva B, Garró X, Rodríguez V, Monardes V, McKay CP. Relative humidity patterns and fog water precipitation in the Atacama Desert and biological implications. J Geophys Res 2007, 112(G4). Chan Y, Lacap DC, Lau MCY, et al. Hypolithic microbial communities, between a rock and a hard place, Hypolithic microbial communities. Environ Microbiol 2012, 14:2272–82. Cowan DA, Khan N, Pointing SB, Cary SC. Diverse hypolithic refuge communities in the McMurdo Dry Valleys. Antarct Sci 2010, 22:714–20. Friedmann EI. Endolithic Microorganisms in the Antarctic Cold Desert. Science 1982, 215:1045–53. Friedmann EI. Endolithic Microbial Life in Hot and Cold Deserts. In: Ponnamperuma C, Margulis L (eds). Limits of Life. Dordrecht, Springer Netherlands, 1980, 33–45. Omelon CR. Endolithic microbial communities in polar desert habitats. Geomicrobiol J 2008, 25:404–14. Wierzchos J, Ascaso C, McKay CP. Endolithic cyanobacteria in halite rocks from the hyperarid core of the Atacama Desert. Astrobiology 2006, 6:415–22. Cary SC, McDonald IR, Barrett JE, Cowan DA. On the rocks, the microbiology of Antarctic Dry Valley soils. Nat Rev Microbiol 2010, 8:129–38. Cowan DA, Tow LA. Endangered Antarctic Environments. Annu Rev Microbiol 2004, 58:649– 90. Cowan DA, Pointing SB, Stevens MI, Craig Cary S, Stomeo F, Tuffin IM. Distribution and abiotic influences on hypolithic microbial communities in an Antarctic Dry Valley. Polar Biol 2011, 34:307–11. Grilli Caiola M, Ocampo-Friedmann R, Friedmann EI. Cytology of long-term desiccation in the desert cyanobacterium Chroococcidiopsis (Chroococcales). Phycologia 1993, 32:315–22. Pointing SB, Warren-Rhodes KA, Lacap DC, Rhodes KL, McKay CP. Hypolithic community shifts occur as a result of liquid water availability along environmental gradients in China’s hot and cold hyperarid deserts. Environ Microbiol 2007, 9:414–24. Lacap DC, Warren-Rhodes KA, McKay CP, Pointing SB. Cyanobacteria and chloroflexi-dominated hypolithic colonization of quartz at the hyper-arid core of the Atacama Desert, Chile. Extremophiles 2011, 15:31–8. Cowan DA, Sohm JA, Makhalanyane TP, et al. Hypolithic communities, important nitrogen sources in Antarctic desert soils. Environ Microbiol Rep 2011, 3:581–6.

10 | 1 An Introduction to Arid Soils and Their Biology

[42]

[43] [44] [45] [46] [47] [48] [49] [50] [51] [52] [53] [54] [55] [56]

[57] [58] [59] [60]

[61]

[62]

Banfield JF, Barker WW, Welch SA, Taunton A. Biological impact on mineral dissolution, application of the lichen model to understanding mineral weathering in the rhizosphere. Proc Natl Acad Sci 1999, 96:3404–11. Viles H. Ecological perspectives on rock surface weathering, Towards a conceptual model. Geomorphology 1995, 13:21–35. Bennett PC, Rogers JR, Silicates WJ, Silicate weathering, and microbial ecology. Geomicrobiol J 2001, 18:3–19. Danin A, Garty J. Distribution of cyanobacteria and lichens on hillsides of the Negev Highlands and their impact on biogenic weathering. Flora Israel 1983, 27:423–44. Coppinger KD, Reiners WA, Burke IC, Olson RK. Net erosion on a sagebrush steppe landscape as determined by cesium-137 distribution. Soil Sci Soc Am J 1991, 55:254. Martinez-Meza E, Whitford WG. Stemflow, throughfall and channelization of stemflow by roots in three Chihuahuan desert shrubs. J Arid Environ 1996, 32:271–87. Whitford WG, Anderson J, Rice PM. Stemflow contribution to the “fertile island” effect in creosotebush, Larrea tridentata. J Arid Environ 1997, 35:451–7. Dean WRJ, Milton SJ, Jeltsch F. Large trees, fertile islands, and birds in arid savanna. J Arid Environ 1999, 41:61–78. Nulsen RA, Bligh KJ, Baxter IN, Solin EJ, Imrie DH. The fate of rainfall in a mallee and heath vegetated catchment in southern Western Australia. Aust J Ecol 1986, 11:361–71. Butterfield BJ, Briggs JM. Patch dynamics of soil biotic feedbacks in the Sonoran Desert. J Arid Environ 2009, 73:96–102. Garcia-Moya E, McKell CM. Contribution of shrubs to the nitrogen economy of a desert-wash plant community. Ecology 1970, 51:81. Charley JL, West NE. Plant-induced soil chemical patterns in some shrub-dominated semidesert ecosystems of Utah. J Ecol 1975, 63:945. Schlesinger WH, Reynolds JF, Cunningham GL, et al. Biological feedbacks in global desertification. Science 1990, 247:1043–8. Berg N, Steinberger Y. Role of perennial plants in determining the activity of the microbial community in the Negev Desert ecosystem. Soil Biol Biochem 2008, 40:2686–95. Garcia-Pichel F, Johnson SL, Youngkin D, Belnap J. Small-scale vertical distribution of bacterial biomass and diversity in biological soil crusts from arid lands in the Colorado Plateau. Microb Ecol 2003, 46:312–21. Zaady E, Groffman PM, Shachak M. Litter as a regulator of N and C dynamics in macrophytic patches in Negev desert soils. Soil Biol Biochem 1996, 28:39–46. Conant RT, Klopatek JM, Malin RC, Klopatek CC. Carbon pools and fluxes along an environmental gradient in northern Arizona. Biogeochemistry 1998, 43:43–61. Su Y, Zhao H, Li Y, Cui J. Carbon mineralization potential in soils of different habitats in the semiarid Horqin Sandy Land, a laboratory experiment. Arid Land Res Manag 2004, 18:39–50. Dossa EL, Khouma M, Diedhiou I, et al. Carbon, nitrogen and phosphorus mineralization potential of semiarid Sahelian soils amended with native shrub residues. Geoderma 2009, 148:251–60. Liu R, Zhao H, Zhao X, Drake S. Facilitative effects of shrubs in shifting sand on soil macrofaunal community in Horqin Sand Land of Inner Mongolia, Northern China. Eur J Soil Biol 2011, 47:316–21. Doblas-Miranda E, Sánchez-Piñero F, González-Megías A. Different microhabitats affect soil macroinvertebrate assemblages in a Mediterranean arid ecosystem. Appl Soil Ecol 2009, 41:329–35.

References |

[63]

[64] [65] [66] [67]

[68]

[69]

[70]

[71] [72] [73] [74]

[75] [76]

[77]

[78]

[79]

[80]

[81]

11

Yong-zhong S, Xue-fen W, Rong Y, Xiao Y, Wen-jie L. Soil fertility, salinity and nematode diversity influenced by Tamarix ramosissima in different habitats in an arid desert oasis. Environ Manage 2012, 50:226–36. Yeates GW, Schipper LA, Smale MC. Site condition, fertility gradients and soil biological activity in a New Zealand frost-flat heathland. Pedobiologia 2004, 48:129–37. Bachar A, Soares MIM, Gillor O. The Effect of resource islands on abundance and diversity of bacteria in arid Soils. Microb Ecol 2012, 63:694–700. Housman DC, Yeager CM, Darby BJ, et al. Heterogeneity of soil nutrients and subsurface biota in a dryland ecosystem. Soil Biol Biochem 2007, 39:2138–49. Ewing SA, Southard RJ, Macalady JL, Hartshorn AS, Johnson MJ. Soil microbial fingerprints, carbon, and nitrogen in a Mojave Desert creosote-bush ecosystem. Soil Sci Soc Am J 2007, 71:469. Steven B, Gallegos-Graves LV, Yeager CM, Belnap J, Kuske CR. Common and distinguishing features of the bacterial and fungal communities in biological soil crusts and shrub root zone soils. Soil Biol Biochem 2014, 69:302–12. Kuske CR, Ticknor LO, Miller ME, et al. Comparison of soil bacterial communities in rhizospheres of three plant species and the interspaces in an arid grassland. Appl Environ Microbiol 2002, 68:1854–63. Steven B, Gallegos-Graves LV, Starkenburg SR, Chain PS, Kuske CR. Targeted and shotgun metagenomic approaches provide different descriptions of dryland soil microbial communities in a manipulated field study. Environ Microbiol Rep 2012, 4:248–56. Belnap J. The world at your feet, desert biological soil crusts. Front Ecol Environ 2003, 1:181–9. Belnap J, Büdel B, Lange OL. Biological soil crusts, characteristics and distribution. Springer, 2003. Steven B, Lionard M, Kuske CR, Vincent WF. High bacterial diversity of biological soil crusts in water tracks over permafrost in the high Arctic Polar Desert. PLoS ONE 2013, 8:e71489. Ustin SL, Valko PG, Kefauver SC, Santos MJ, Zimpfer JF, Smith SD. Remote sensing of biological soil crust under simulated climate change manipulations in the Mojave Desert. Remote Sens Environ 2009, 113:317–28. Pointing SB, Belnap J. Microbial colonization and controls in dryland systems. Nat Rev Microbiol 2012, 10:551–62. Garcia-Pichel F, López-Cortés A, Nübel U. Phylogenetic and morphological diversity of Cyanobacteria in soil desert crusts from the Colorado Plateau. Appl Environ Microbiol 2001, 67:1902–10. Steven B, Gallegos-Graves LV, Yeager CM, Belnap J, Evans RD, Kuske CR. Dryland biological soil crust cyanobacteria show unexpected decreases in abundance under long-term elevated CO2 . Environ Microbiol 2012, 14:3247–58. Belnap J, Phillips SL, Witwicki DL, Miller ME. Visually assessing the level of development and soil surface stability of cyanobacterially dominated biological soil crusts. J Arid Environ 2008, 72:1257–64. Langhans TM, Storm C, Schwabe A. Community assembly of biological soil crusts of different successional stages in a temperate sand ecosystem, as assessed by direct determination and enrichment techniques. Microb Ecol 2009, 58:394–407. Billings S, Schaeffer S, Evans R. Nitrogen fixation by biological soil crusts and heterotrophic bacteria in an intact Mojave Desert ecosystem with elevated CO2 and added soil carbon. Soil Biol Biochem 2003, 35:643–9. Mazor G, Kidron GJ, Vonshak A, Abeliovich A. The role of cyanobacterial exopolysaccharides in structuring desert microbial crusts. FEMS Microbiol Ecol 1996, 21:121–30.

12 | 1 An Introduction to Arid Soils and Their Biology

[82]

Bowker MA, Reed SC, Belnap J, Phillips SL. Temporal variation in community composition, pigmentation, and Fv/Fm of desert cyanobacterial soil crusts. Microb Ecol 2002, 43:13–25. [83] Yeager CM, Kornosky JL, Morgan RE, et al. Three distinct clades of cultured heterocystous cyanobacteria constitute the dominant N2 -fixing members of biological soil crusts of the Colorado Plateau, USA. FEMS Microbiol Ecol 2007, 60:85–97. [84] Gao Q, Garcia-Pichel F. Microbial ultraviolet sunscreens. Nat Rev Microbiol 2011, 9:791–802. [85] Nagy ML, Pérez A, Garcia-Pichel F. The prokaryotic diversity of biological soil crusts in the Sonoran Desert (Organ Pipe Cactus National Monument, AZ). FEMS Microbiol Ecol 2005, 54:233–45. [86] Gundlapally SR, Garcia-Pichel F. The community and phylogenetic diversity of biological soil crusts in the Colorado Plateau studied by molecular fingerprinting and intensive cultivation. Microb Ecol 2006, 52:345–57. [87] Martínez I, Escudero A, Maestre FT, de la Cruz A, Guerrero C, Rubio A. Small-scale patterns of abundance of mosses and lichens forming biological soil crusts in two semi-arid gypsum environments. Aust J Bot 2006, 54:339. [88] Darby BJ, Neher DA, Belnap J. Soil nematode communities are ecologically more mature beneath late- than early-successional stage biological soil crusts. Appl Soil Ecol 2007, 35:203–12. [89] Bates ST, Garcia-Pichel F. A culture-independent study of free-living fungi in biological soil crusts of the Colorado Plateau, their diversity and relative contribution to microbial biomass. Environ Microbiol 2009, 11:56–67. [90] Eldridge D, Zaady E, Shachak M. Infiltration through three contrasting biological soil crusts in patterned landscapes in the Negev, Israel. Catena 2000, 40:323–6. [91] Bowker MA, Belnap J, Davidson DW, Phillips SL. Evidence for micronutrient limitation of biological soil crusts, importance to arid-lands restoration. Ecol Appl 2005, 15:1941–51. [92] Belnap J, Gillette DA. Vulnerability of desert biological soil crusts to wind erosion, the influences of crust development, soil texture, and disturbance. J Arid Environ 1998, 39:133–42. [93] Belnap J, Gillette DA. Disturbance of biological soil crusts, impacts on potential wind erodibility of sandy desert soils in southeastern Utah. Land Degrad Dev 1997, 8:355–62. [94] Eldridge DJ, Leys JF. Exploring some relationships between biological soil crusts, soil aggregation and wind erosion. J Arid Environ 2003, 53:457–66. [95] Bowker MA, Belnap J, Bala Chaudhary V, Johnson NC. Revisiting classic water erosion models in drylands, the strong impact of biological soil crusts. Soil Biol Biochem 2008, 40:2309–16. [96] Yeager CM, Kornosky JL, Housman DC, Grote EE, Belnap J, Kuske CR. Diazotrophic community structure and function in two successional stages of biological soil crusts from the Colorado Plateau and Chihuahuan Desert. Appl Environ Microbiol 2004, 70:973–83. [97] Johnson SL, Neuer S, Garcia-Pichel F. Export of nitrogenous compounds due to incomplete cycling within biological soil crusts of arid lands. Environ Microbiol 2007, 9:680–9. [98] Evans RD, Ehleringer JR. A break in the nitrogen cycle in aridlands? Evidence from δ 15 N of soils. Oecologia 1993, 94:314–7. [99] Johnson SL, Budinoff CR, Belnap J, Garcia-Pichel F. Relevance of ammonium oxidation within biological soil crust communities. Environ Microbiol 2005, 7:1–12. [100] Harper KT, Belnap J. The influence of biological soil crusts on mineral uptake by associated vascular plants. J Arid Environ 2001, 47:347–57. [101] Su Y-G, Li X-R, Cheng Y-W, Tan H-J, Jia R-L. Effects of biological soil crusts on emergence of desert vascular plants in North China. Plant Ecol 2007, 191:11–9. [102] Langhans TM, Storm C, Schwabe A. Biological soil crusts and their microenvironment, Impact on emergence, survival and establishment of seedlings. Flora Morphol Distrib Funct Ecol Plants 2009, 204:157–68.

References | 13

[103] Green LE, Porras-Alfaro A, Sinsabaugh RL. Translocation of nitrogen and carbon integrates biotic crust and grass production in desert grassland, translocation between crust and grass. J Ecol 2008, 96:1076–85. [104] Porras-Alfaro A, Herrera J, Natvig DO, Lipinski K, Sinsabaugh RL. Diversity and distribution of soil fungal communities in a semiarid grassland. Mycologia 2011, 103:10–21. [105] Noy-Meir I. Desert ecosystems, environment and producers. Annu Rev Ecol Syst 1973, 4:25– 51. [106] Reed SC, Coe KK, Sparks JP, Housman DC, Zelikova TJ, Belnap J. Changes to dryland rainfall result in rapid moss mortality and altered soil fertility. Nat Clim Change 2012, 2:752–5. [107] Steven B, Kuske CR, Gallegos-Graves LV, Reed SC, Belnap J. Climate change and physical disturbance manipulations result in distinct biological soil crust communities. Appl Environ Microbiol 2015, 81:7448–59. [108] Ogle K, Reynolds JF. Plant responses to precipitation in desert ecosystems, integrating functional types, pulses, thresholds, and delays. Oecologia 2004, 141:282–94. [109] Schwinning S, Sala OE. Hierarchy of responses to resource pulses in arid and semi-arid ecosystems. Oecologia 2004, 141:211–20. [110] Pointing SB, Belnap J. Disturbance to desert soil ecosystems contributes to dust-mediated impacts at regional scales. Biodivers Conserv 2014, 23:1659–67. [111] Evans J, Geerken R. Discrimination between climate and human-induced dryland degradation. J Arid Environ 2004, 57:535–54. [112] Dore MHI. Climate change and changes in global precipitation patterns, what do we know? Environ Int 2005, 31:1167–81. [113] Wall DH, Virginia RA. Controls on soil biodiversity, insights from extreme environments. Appl Soil Ecol 1999, 13:137–50. [114] Bowker MA, Maestre FT, Escolar C. Biological crusts as a model system for examining thebiodiversity–ecosystem function relationship in soils. Soil Biol Biochem 2010, 42:405–17.

Carlos Garcia, J.L.Moreno, T. Hernandez, and F. Bastida

2 Soils in Arid and Semiarid Environments: the Importance of Organic Carbon and Microbial Populations. Facing the Future Abstract: Drylands occupy 47% of the Earth’s land area and accumulate 35–42 t carbon (C) ha−1 . In comparison to other biomes, the natural depletion of C content in arid and semiarid lands harbors a high potential for carbon sequestration. We provide a comprehensive review of carbon biogeochemistry, the associated microbial communities and strategies for soil restoration in drylands under the scope of global change. In these areas, the biogeochemistry of organic carbon is governed by climate conditions. Photodegradation, water availability and temperature, overcontrol microbial activity and hence carbon cycling. Under limited water availability, microbial activity is diminished and hence the organic matter accumulation in soil increases, but the development of a sustainable plant cover is not promoted. Soil degradation as a consequence of low carbon content can be avoided by organic amendments consisting of biosolids (composts, sludges, etc.). Organic amendments promote an increase of soil organic matter and microbial activity, which are linked to a rise in soil fertility. Appropriate management practices in cropland and shrub lands, which have deep soil profiles with low organic carbon saturation, seem to be a win–win option for sequestering carbon and improving soil productivity. This fundamental research is needed to balance soil fertility and carbon sequestration, particularly under the global change scenario.

2.1 Introduction Drylands occupy 6.31 × 109 ha or 47% of the Earth’s land area (UNEP 1992) and are distributed among four climate zones: hyperarid (1.0 × 109 ha), arid (1.62 × 109 ha), semiarid (2.37 × 109 ha), and dry subhumid (1.32 × 109 ha). Arid and semiarid or subhumid zones are characterized by low and erratic rainfall, periodic droughts, and different associations of vegetative cover and soils. The annual rainfall varies from up to 350 mm in arid zones to 700 mm in semiarid areas. Desertification is the main problem that arid and semiarid lands face. Within the context of Agenda 21, desertification is defined as “land degradation in arid, semi-arid and dry subhumid areas resulting from climatic variations and human activities” [1]. Either due to human induced actions or natural conditions, the loss of soil organic matter (SOM) is strongly linked to soil degradation and desertification in arid and semiarid areas, and causes a decline in agronomical productivity and failure of soil ecosystem services. Although arid and semiarid ecosystems have less vegetation and, DOI 10.1515/9783110419047-002

16 | 2 Soils in Arid and Semiarid Environments

hence, lower carbon accumulation than boreal or tropical areas, they are estimated to contain 20% of the global soil C pool (organic plus inorganic) in continental areas [2]. Lal et al. (2004) [3] concluded that the predicted amounts of carbon in drylands are 159–191 billion tons, with a density of 35–42 (t C ha−1 ). If we compare the latter value with the values estimated for boreal (247–344 t C ha−1 ), tropical (121–113 t C ha−1 ), and tundra (121–127 t C ha−1 ) ecosystems, it is clear that soils under this climate are depleted in carbon, both for “natural” or “anthropogenic” reasons. The hypothesis is that these soils still have capacity for carbon sequestration, which would increase soil quality, ensure food security, and mitigate global change [3]. The organic matter content of soils is subjected to strong and complex physical, chemical, biochemical, and biological controls that are ultimately responsible for carbon stabilization and its mineralization [4, 5]. An alteration of such equilibriums due to land use (i.e., tillage) [6, 7] and climate pressures may alter the C stocks in soils and potentially cause soil degradation, hence affecting the sustainability of the planet. The degradation of soils due to carbon losses in many arid and semiarid areas of the planet cannot be afforded in the future for two reasons: 1. Many of these areas are located in extensive agricultural zones (i.e., California, Israel, southeastern Spain, southern Italy, Greece, etc.) and must provide enough food for a growing population. 2. The need for global change mitigation by C sequestration, where these soils can play a key role. Considering that, ultimately, the dynamics of organic carbon are governed by biochemical and microbiological processes, we aim to present the main findings and trends concerning the biogeochemistry of organic carbon and the intrinsic dynamics of microbial communities in soils developed under arid and semiarid conditions. The role of organic matter, the significance of the microbial biomass, and the structure of microbial communities will be highlighted, with special emphasis on soil restoration strategies and the application of methods that provide novel knowledge. Finally, we reflect on the main gaps in our knowledge that should be addressed in order to increase the ecological value of soils located in arid and semiarid areas in the future.

2.2 Climate Regulation and Soil Organic Carbon in Arid-Semiarid Zones Climate change is a special concern regarding the control of SOM. Variations in temperature and precipitation may alter both biotic and abiotic factors that control carbon immobilization in semiarid areas. The positive microbial community feedback in response to elevated CO2 concentration and warming can accelerate the microbial decomposition of SOM and potentially lead to soil C losses [8]. However, at the global

2.3 Land Use and Soil Organic Carbon in Arid-Semiarid Zones |

17

level, the effects of temperature on the decomposition of SOM are less clear [9]. Some studies have indicated that global emissions of CO2 as a consequence of SOM decomposition would increase as a response to rising temperatures [10]. In contrast, it has been suggested that dryland soils would most likely sequester C with a future increase in precipitation but release C with a decrease in precipitation [11]. Episodic water availability clearly affects element cycling in arid and semiarid ecosystems [12]. High temperatures and erratic moisture inputs impose a pulsed pattern on biological activities [13], which, in turn, will determine the C and N turnover; so, organic matter tends to accumulate during dry periods when plant and microbial growth are restricted [14]. Moreover, drought affects the quality and composition of humic acids, which – biologically and chemically – are the most active fraction of SOM [15]. Thus, losses of aliphatic and polysaccharide-like structures, secondary amides, polycondensed aromatic systems of large molecular size, and other unsaturated bond systems such as carbonyl and carboxyl groups were observed in semiarid soil humic acids after a long drought [14]. Soil processes in arid lands are controlled principally by water availability but the photodegradation of above ground litter and the overriding importance of spatial heterogeneity are modulators of the biotic responses to water availability [16]. Microbiological soil properties are negatively affected by drought since soil moisture plays a key role in the survival and activity of soil microorganisms [14]. Mechanisms such as the retarded diffusion of soluble substrates and/or reduced microbial mobility (and consequent access to substrates) could explain the low microbial biomass found in soils with low water content [17]. Liu et al. (2009) [18] suggested that soil water availability was more important than temperature in regulating the soil microbial respiration and microbial biomass in a semiarid temperate steppe. Accordingly, some authors have found that organic matter stocks are progressively preserved with the increasing duration and intensity of droughts [19]. Conversely, an experimental field study about the impact of climate change on desertification along a Mediterranean arid transect demonstrated that the SOM content decreased with aridity [20].

2.3 Land Use and Soil Organic Carbon in Arid-Semiarid Zones Adequate land use management helps to control the global stocks of organic carbon in drylands and fight against soil desertification [11, 21]. Despite the extensive number of studies aiming to evaluate the effects of land use on organic C stocks, there are still some discrepancies. For instance, the conversion of ecosystems from natural conditions to agricultural use generally results in decreased carbon stocks in arid and semiarid climates [22, 23]. Disturbance by shrub removal and/or livestock grazing significantly reduced the amount of organic matter in an Australian semiarid woodland [24]. However, other studies did not find any significant effect of land management on soil organic carbon (SOC) [22, 25]. As stated by Booker et al. (2013) [26], car-

18 | 2 Soils in Arid and Semiarid Environments

bon uptake in arid and semiarid areas is most often controlled by abiotic factors that are not easily changed by management or vegetation. In this sense, photodegradation, which is highly intense in arid ecosystems, exerts a dominant control on above ground litter decomposition [27]. Losses through photochemical reactions may represent a short circuit in the carbon cycle, with a substantial fraction of the carbon fixed in plant biomass being lost directly to the atmosphere without cycling through soil organic matter pools [27]. More studies based on the prevention of photodegradation should be carried out to promote carbon sequestration in soil and climate change mitigation. For instance, the placement of a wide vegetation cover may reduce the effects of photodegradation and enhance soil moisture. Reforestation may influence carbon balances, increase soil carbon stocks and serves for fighting against desertification in many arid and semiarid regions [28, 29]. In general, soils in arid and semiarid conditions depict a positive relationship between the organic carbon content and plant cover [30, 31]. Nevertheless, the spatial heterogeneity of plant cover in semiarid shrublands is the principal cause of the spatial heterogeneity of the SOC content, which is associated with the development of islands of fertility under shrubs [32].

2.4 Soil Restoration in Arid-Semiarid Zones: Amendments Based on Exogenous Organic Matter The scant vegetation of the soils in arid and semiarid zones, which is mainly a result of low productivity and subsequent abandonment, causes the inputs of organic matter into the soil to be low. Hence, together with the usual soil erosive processes and high photodegradation rates, many soils have a low organic matter level, which compromises their functionality and the provision of ecosystem services and can even end in intense degradation phenomena. Since the Kyoto Protocol of 1992, which identified soils as a possible sink for carbon, there has been much progress. A report on organic matter and biodiversity within the European Thematic Strategy [33] mentions that exogenous organic matter, that is, organic materials added to a degraded soil in order to improve harvests or restore it for subsequent use, constitutes an invaluable source of organic matter and contributes to the fixation of C in the soil, thus partially diminishing the greenhouse effect derived from the release of CO2 to the atmosphere. The application of organic materials enhances the nutrient status of soil by serving as a source of macro and micronutrients, and improves its physical properties by increasing soil porosity and water retention because of the presence of humic-like substances, known as a polycondensed macromolecular structure. In addition, one of the beneficial effects of humic substances is that soil enzymes bound to humic fractions remain protected in the long term against denaturalization by proteolysis attacks

2.5 Microbial Biomass and Enzyme Activity in Arid-Semiarid Zones

|

19

in soil. The use of organic amendments to improve soil quality and restore degraded lands has been widespread [34–36]. Application of organic amendments usually improves soil aggregation [37] and, hence, the physical structure of the soil [38, 39]. Furthermore, organic amendment generates a better nutritional scenario for progressive plant growth [40, 41]. Plant inputs to soil promote the development of the microbial biomass and its activity, which raises soil fertility in the long term [36, 42, 43]. Different types of organic amendments have been applied in arid and semiarid environments: crop residues, pig slurry, farmyard manure, municipal solid waste, olive mill waste, sewage sludge, etc. However, the addition of organic amendments to soil has to be carried out carefully since it does not always lead to an increase in soil quality. For instance, Tejada et al. (2007) [44] reported that the application of fresh beet vinasse worsened the physical and biological properties due to its content of sodium ions. In addition to the carbon inputs arising from the above ground development after amendment, the organic amendments themselves provide exogenous carbon that may persist in the soil. The stability and nature of the amendment can determine the residence time of the added organic carbon [45, 46]. In dryland ecosystems, due to the high potential for carbon sequestration, the stabilization of SOM is believed to be controlled more by the quantity of the inputs and its interaction with the soil matrix (i.e., texture) than by the quality of the organic amendment [47, 48]. It is thought that fine soil particles have a critical role in C fixation. Some authors observed an increase in the carbon fixation into fine particles (clay or silt) after organic amendment [48, 49], while others did not find any variation in the organic carbon content of the fine fractions in the long term [22]. Recent studies based on carbon stable isotope probing have also suggested a protective role of clays [50, 51], even concluding that there is major fixation of carbon in clay soils despite the highly labile nature of added carbon (i.e., 13 C-glucose) [50]. Regardless of the fact that part of the added carbon probably persists in soil physically linked to soil particles, a clear benefit of organic amendment derives from the improvement in the nutritional conditions of the soil – which enhances subsequent plant growth (󳶳 Fig. 2.1). Plant development provides organic matter to the soil, benefits its structure, and avoids soil erosion, a very important issue in sloping areas [36].

2.5 Microbial Biomass and Enzyme Activity in Arid-Semiarid Zones As stated above, the microbial biomass is largely responsible for soil carbon cycling. The microbial biomass of semiarid soils is usually constrained by the low amounts of plant inputs and water availability. The evaluation of microbial biomass by phospholipid fatty acids (PLFAs) analysis revealed that the total PLFAs ranged between 2.2 and 100 nmol fatty acids g−1 soil in arid and semiarid areas [41, 52–55]. Nevertheless, the interpretation of PLFA patterns in extremely arid ecosystems must be done carefully [52]. Water activity below a certain threshold may protect cellular remains from

20 | 2 Soils in Arid and Semiarid Environments

18 months after organic amendment restoration

Fig. 2.1: Field experiment in Spain: soil restoration.

degradation [56]. Hence, the results obtained following treatment might be biased by the previous viable microbial community. Generally, the level of biomass correlates well with the amount of organic carbon and is closely related to the moisture content of dryland soils. For instance, various authors have observed changes in the microbial biomass linked to the organic carbon after a change in land use [57, 58]. Similarly, the restoration of soil quality by addition of organic waste byproducts increases the microbial biomass 1.6–3 times [41]. The microbial biomass also responds to plant growth and the parallel increase in SOM [52, 55]. In detail, Ben-David et al. (2011) [52] found that the fatty acid 16:1w7, indicative of cyanobacteria [59], increased in intershrub soils of the Negev Desert (Israel); this suggests an increase in the relative abundance of cyanobacteria, which are known to be the primary colonizers of biological crusts in drylands [60]. Dry periods may have a deleterious effect on bacterial communities through starvation, induced osmotic stress, and resource competition, which affects the structure and functioning of soil bacterial communities and leads to a slowing down of N and C mineralization [14, 61]. For soils that have not received recent organic matter additions, wet–dry cycles initially stimulate C and net N mineralization and diminish the microbial biomass during drying but stimulate microbial growth after wetting, and the wet–dry cycle itself results in higher net N and C mineralization when compared to continuously moist soils [62, 63]. Accumulation of inorganic N usually occurs during dry periods because diffusion of ions is severely restricted in the thin water films of dry soil and because sinks of inorganic N are limited by reduced microbial growth and limited plant uptake [14, 64]. A portion of the microbial biomass is killed under dry conditions [65]; this is readily decomposed by surviving organisms when the soil

2.5 Microbial Biomass and Enzyme Activity in Arid-Semiarid Zones

|

21

is rewetted. This dead microbial biomass, with its low C:N ratio, becomes available for microbial activity and leads to high N mineralization, large pulses of CO2 and gaseous fluxes of N, and a pulse of increased C and N availability. In principle, as stated by Entry et al. (2004) [57], Gram positive biomarkers would be expected to increase in desiccated or degraded soils due to their sporulation capacity under harsh conditions. However, this trend is usually not found [14, 41, 54, 57]. Perhaps, the relatively fast response of soils to nutrient or water pulses might be taken into consideration, and the measurement of PLFAs at a particular time has to be discussed carefully. Moreover, only a fraction of the microbial biomass survives both the dry season in arid environments and the osmotic shock associated with the rapid increase in moisture after the first rainfall [66]. The microbial biomass is responsible for the production of enzymes that are excreted into the extracellular microenvironment, where they can be protected by immobilization in humic and clay colloids [67, 68]. The basic importance of enzyme activity in soil lies in the fact that ecosystem functioning cannot be totally understood without the participation of enzymatic processes and their catalytic reactions related to nutrient cycling [69]. Extracellular enzymes are closely related to organic matter decomposition, and key enzymatic reactions include those involved in the degradation of cellulose and lignin, those that hydrolyze reservoirs of organic N such as proteins, chitin, and peptidoglycan, and those that mineralize P from nucleic acids, phospholipids, and other ester phosphates [70]. Extracellular enzyme activity (EEA) mediates microbial nutrient acquisition from organic matter, and these activities are commonly interpreted as indicators of microbial nutrient demand and soil quality [69, 71]. In general, enzymes are associated with viable, proliferating cells, but they can be excreted from a living cell or released into the soil solution from dead cells. Once enzymes have left the shelter of the cell, they are exposed to an inhospitable environment in which nonbiological denaturalization, adsorption, inactivation, and degradation by proteolytic microorganisms all conspire to harm the enzymes, unless they survive due to the new protection afforded by the mineral and/or humic association, which is more resistant to proteolysis than the free enzymes. In arid and semiarid environments, the soil EEA has been used to examine the functional responses of the soil microbial biomass to factors such as increased nutrient deposition [72], heavy metal contamination [73], organic amendment [36, 41, 74], soil management [75–77], plant diversity [78], type of agroecosystem [79], and climate change [80]. More than any other factor, OM dynamics are closely related to the regulation of enzyme activity. In arid and semiarid areas, the potential activities of enzymes that decompose proteins (e.g., aminopeptidase) and recalcitrant C compounds such as lignin and humic substances (e.g., phenol oxidases) exceed those of mesic soils by more than an order of magnitude in both absolute terms and in relation to the activities of enzymes that break down cellulose, which generally dominate the EEA of mesic soils [81]. The pH is a strong regulator of EEA, with important consequences for

22 | 2 Soils in Arid and Semiarid Environments

SOM dynamics. Because of carbonate accumulation, the pH of arid soils can reach 8 or above, which is optimal for phenol oxidase enzymes [82]. In contrast, the pH optima of glycosidases (e.g., cellulase, chitinase) generally range from 4 to 6. Soil texture and moisture also determine the enzyme activity, by influencing the microbial biomass and by controlling the substrate availability. When the soil moisture is low, the EEA is also low. Prolonged droughts are likely to decrease enzyme production, resulting in lower measured activities when moisture returns [83]. Because rewetting sometime results in a pulse of microbial biomass turnover [84, 85], many intracellular enzymes may be released into the soil, creating a temporary increase in EEA. Prolonged precipitation can result in increased EEA in arid or semiarid soils [80], although this may be at least partially due to enhanced plant growth and rhizodeposition [86].

2.6 Organic Carbon, Macro and Microaggregates, and C Sequestration in Arid-Semiarid Zones Converting forest to cultivated areas reduces soil organic carbon mainly through the reduction of biomass inputs into the soil and the stimulation of soil organic matter mineralization, thus increasing soil erosion rates [87]. There is evidence that the magnitude of this loss of soil organic carbon through cultivation could be greater in semiarid areas than in more humid areas [88]; this impact decreases with depth. The analysis of environmental control factors suggests a negative effect on soil organic carbon in a climatic change scenario with increased temperature and a decrease in rainfall, as is expected in semiarid areas. Some data indicate that this negative impact on soil organic carbon would be greater in soil surface than in the soil subsurface. For this reason, a strategy for C sequestration should be focused on subsoil sequestration. Appropriate management practice in cropland and shrubland, which have deep soil profiles with low organic carbon saturation, seems to be a win–win option for sequestering atmospheric organic carbon and improving soil productivity. Some studies confirm that the potential sequestration of C in semiarid reforested areas depends largely on the techniques used for reforestation. The C stocks in reforested ecosystems are directly proportional to the amount of biomass produced, which, in turn, is determined by the productivity of the soil. For this reason, methods that improve the productivity of the soil must be used. The addition of organic amendments to the soil, prior to planting, could be very effective in terms of C sequestration [87, 89]. In semiarid areas, studies on degraded soil rehabilitation have proved that the addition of organic amendments to these soils increases the percentage of both soil macroaggregates and microaggregates within macroaggregates, as well as the concentration of organic C in these soil fractions [90]. This is of great interest since microaggregation formation is crucial for the storage and stabilization of soil C in the long

2.7 Conclusion |

23

term [91, 92]. Other authors have reported an increase of C concentration in fine soli particles (silt and clay) with the addition of organic amendment to semiarid degraded soils [49, 93]. In semiarid and arid soils, the chemical stabilization of organic carbon, through the formation of complexes with silt and clay particles and their physical protection in microaggregates formed within macroaggregates, could be the main mechanism of C sequestration in these soils, in both agricultural and forest areas. The physical protection of soil organic carbon could be promoted by the changes, both qualitative and quantitative, in plant contributions to soil. In both forested and agricultural areas in semiarid climates and where a green cover has been incorporated, an increase in the labile pool of soil organic carbon occur [94]. Fresh plants induce the formation of macroaggregates both directly, by acting as a binding agent between soil particles, and indirectly, by activating the production of microbially derived binding agents. The establishment of these new macroaggregates can increase the formation of microaggregates that occlude organic matter inside and make it inaccessible to the microorganisms [90, 95]. In the agricultural soils in semiarid and arid areas, minimum tillage seems necessary, since it promotes the incorporation of plant material into deeper layers, promoting the formation of aggregates and, therefore, organic carbon occlusion within them [94]. A strong positive correlation between basal soil respiration and the percentage of microaggregates within macroaggregates has been found in reforested soils, while this correlation was negative in degraded shrubland [96]. This suggests that the formation of microaggregates, which are rich in organic carbon, could be a self defense mechanism of the soil to protect organic carbon from increased microbiological activity [96]; for these reasons, these correlations could serve as indicators of processes of improvement (positive correlations) or degradation (negative correlation) of the soil.

2.7 Conclusion Soil degradation due to aggressive human action or passive climate pressure must be avoided in order to conserve soils that have a high ecological value for the future. The fragility of these soils contrasts with their intense response to soil restoration programs, which include the addition of organic matter and their potential capacity for carbon sequestration. Organic amendments help to preserve and improve the quality and fertility of the soils in these areas, which could be particularly important under a global change scenario. The biogeochemical and microbiological information on arid and semiarid soils is abundant but perhaps more limited than that for other climates. Nevertheless, such studies are widespread across the planet and numerous research groups are focused on the topic. This fact will increase our knowledge of the biogeochemistry of carbon,

24 | 2 Soils in Arid and Semiarid Environments

as well as our capacity for managing the cycling of elements and the sustainability of arid and semiarid soils in the future. However, if we aim to increase such an “ecological capital,” soil science must necessarily move on and search for answers to new, more focused questions: 1. Which biochemical processes are responsible for carbon fixation and humus formation? 2. Are we able to “control” the microbial populations and carbon related biochemical reactions of these soils? The mutual benefits of microbial activity, carbon sequestration, and plant growth are clear in terms of sustainability. To enhance the physicochemical protection of soil organic carbon the stability of microaggregates should be maximized, while ensuring a suitable rate of macroaggregate turnover that will allow the fixation of new organic carbon. This could be promoted by minimum tillage, an increase of plant inputs, particularly root inputs (by modifying residue amount and quality, altering mycorrhizal associations and vegetal species), etc. It can promote the formation of new macroaggregates that can increase the formation of microaggregates that occlude organic matter inside and make them inaccessible to the microorganisms. However, fundamental research is needed to balance soil fertility and carbon sequestration with economic or environmental needs. Managing soil conditions or designing “à la carte” organic amendments, which promote a punctual rise in fertility when needed (i.e., an increase in agricultural productivity) or foster carbon sequestration for environmental purposes in abandoned lands at a particular moment, would definitively increase the ecological value of arid and semiarid soils in the coming era. Acknowledgment: F. Bastida thanks the Spanish Government for his “Ramón y Cajal” contract (RYC-2012-10666) and FEDER founding. The authors are grateful to the Fundación Séneca of Murcia Region (19896/GERM/15). The authors thank the Spanish Ministry for the CICYT projects AGL2014-55269-R and AGL2014-54636.

References [1]

[2] [3] [4]

UNCED. Managing fragile ecosystems: Combating desertification and drought (Rio de Janeiro, 3–14 June 1992), Report of the United Nations Conference on Environment and Development, General A/CONF.151/26 (Vol. II), Chapter 12, (http://www.unccd.ch/). Rasmussen C, Southard RJ, Howarth WR. Mineral control of organic carbon mineralization in a range of temperate conifer forest soils. Global Change Biol 2006, 12:834–47. Lal R. Soil carbon sequestration impacts on global climate change and food security. Science 2004, 304:1623–26. Six J, Conant RT, Paul EA, Paustian K. Stabilization mechanisms of soil organic matter: Implications for C-saturation of soils. Plant Soil 2002, 241:155–76.

References |

[5]

[6]

[7]

[8] [9] [10]

[11]

[12]

[13]

[14]

[15]

[16] [17] [18]

[19] [20] [21]

[22] [23]

25

von Lutzow M, Koegel-Knabner I, Ekschmitt K, Matzner E, Guggenberger G, Marschner B, Flessa H. Stabilization of organic matter in temperate soils: mechanisms and their relevance under different soil conditions – a review. Eur J Soil Sci 2006, 57:426–45. Kandeler E, Stemmer M, Klimanek EM. Response of soil microbial biomass, urease and xylanase within particle size fractions to long-term soil management. Soil Biol Biochem 1999, 31:261–73. Conant RT, Six J, Paustian K. Land use effects on soil carbon fractions in the southeastern United States. II. Changes in soil carbon fractions along a forest to pasture chronosequence. Biol Fertil Soils 2004, 40:194–200. Nie M, Pendall E, Bell C, Gasch CK, Raut S, Tamang S, Wallenstein MD. Positive climate feedbacks of soil microbial communities in a semi-arid grassland. Ecol Lett 2013, 16:234–41. Giardina CP, Ryan MG. Evidence that decomposition rates of organic carbon in mineral soil do not vary with temperature. Nature 2000, 404:858–61. Jones C, McConnell C, Coleman K, Cox P, Fallon P, Jenkinson D, Powlson. Global climate change and soil carbon stocks; predictions from two contrasting models for the turnover of organic carbon in soil. Global Change Biol 2005, 11:154–66. Albaladejo J, Ortiz R, García-Franco N, Ruiz-Navarro A, Almagro M, García-Pintado J, MartínezMena M. Land use and climate change impacts on soil organic carbon stocks in semi-arid Spain. J Soil Sediment 2013, 13:265–77. Austin AT, Yahdjian L, Stark JM, Belnap J, Porporato A, Norton U, Ravetta DA, Schaeffer SM. Water pulses and biogeochemical cycles in arid and semiarid ecosystems. Oecologia 2004, 141:221–35. Collins SL, Sinsabaugh RL, Crenshaw C, Green L, Porras-Alfaro A, Sutrsova M, Zegkin LH. Pulse dynamics and microbial processes in aridland ecosystems. Journal of Ecology 2008, 96:413– 20. Hueso S, García C, Hernández T. Severe drought conditions modify the microbial community structure, size and activity in amended and unamended soils. Soil Biol Biochem 2012, 50:167– 73. Buurman P, Nierop KGJ, Kaal J, Senesi N. Analytical pyrolysis and thermally assisted hydrolysis and methylation of EUROSOIL humic acid samples – A key to their source. Geoderma 2009, 150:10–22. Austin AT. Has water limited our imagination for aridland biogeochemistry? Trends Ecol Evol 2011, 26:229–35. van Meeteren MJM, Tietema A, van Loon EE, Verstraten JM. Microbial dynamics and litter decomposition under a changed climate in a Dutch heathland. Appl Soil Ecol 2008, 38:119–27. Liu W, Zhang Z, Wan S. Predominant role of water in regulating soil and microbial respiration and their responses to climate change in a semiarid grassland. Global Change Biol 2009, 15:184–95. Borken W, Matzner E. Reappraisal of drying and wetting effects on C and N mineralization and fluxes in soils. Global Change Biol 2009, 15:808–24. Lavee H, Imeson AC, Sarah P. The impact of climate change on geomorphology and desertification along a Mediterranean-arid transect. Land Degrad Dev 1998, 9:407–22. de Baets S, Meersmans J, Vanacker V, Quine TA, van Oost K. Spatial variability and change in soil organic carbon stocks in response to recovery following land abandonment and erosion in mountainous drylands. Soil Use Manage 2012, 29:65–76. Steffens M, Kölbl A, Totsche KU, Kögel-Knabner I. Grazing effects on soil chemical and physical properties in a semiarid steppe of Inner Mongolia (P.R. China). Geoderma 2008, 143:63–72. Pérez-Quezada JF, Delpiano CA, Snyder KA, Johnson DA, Franck N. Carbon pools in an arid shrubland in Chile under natural and afforested conditions. J Arid Environ 2011, 75:29–37.

26 | 2 Soils in Arid and Semiarid Environments

[24] Daryanto S, Eldridge DJ, Throop HL. Managing semi-arid woodlands for cabon storage: Grazing and shrub effects on above- and belowground carbon. Agr Ecosyst Environ 2013, 169:1–11. [25] Seddaiu G, Porcu G, Ledda L, Roggero PP, Agnelli A, Corti G. Soil organic matter content and composition as influenced by soil management in a semi-arid Mediterranean agro-silvopastoral system. Agr Ecosyst Environ 2013, 167:1–11. [26] Booker K, Huntsinger L, Bartolome JW, Sayre NF, Stewart W. What can ecological science tell us about opportunities for carbon sequestration on arid rangelands in the United States? Glob Environ Change 2013, 23:240–51. [27] Austin AT, Vivanco. Plant litter decomposition in a semi-arid ecosystem controlled by photodegradation. Nature 2006, 442:555–58. [28] Harper RJ, Okom AEA, Stilwell AT et al. Reforesting degraded agricultural landscapes with Eucalypts: Effects on carbon storage and soil fertility after 26 years. Agr Ecosyst Environ 2010, 163:3–13. [29] Hu YL, Zeng DH, Chang SX, Mao R. Dynamics of soil and root C stocks following afforestation of croplands with poplars in a semi-arid region in northeast China. Plant Soil 2013, 368:619–27. [30] García C, Hernández T, Roldán A, Martín A. Effect of plant cover decline on chemical microbiological parameters under Mediterranean climate. Soil Biol Biochem 2002, 34:635–42. [31] García C, Roldán A, Hernández T. Ability of different plant species to promote microbiological processes in semiarid soil. Geoderma 2005, 124:193–202. [32] Schlesinger WH, Raikks JA, Hartley AE, Cross AF. On the spatial pattern of soil nutrients in desert ecosystems. Ecology 1996, 77:364–74. [33] van Camp L, Bujarrabal B, Gentile AR et al. Reports of the Technical Working Groups Established under the Thematic Strategy for Soil Protection. EUR 21319 EN/3. Luxembourg, Office for Official Publications of the European Communities, 2004, 1–872. [34] García C, Hernández T, Costa F. Variation in some chemical parameters and organic matter in soils regenerated by the addition of municipal solid-waste. Environ Manage 1992, 16:763–68. [35] Tejada M, Hernández MT, García C. Application of two organic amendments on soil restoration: Effects on the soil biological properties. J Environ Qual 2006, 35:1010–17. [36] Bastida F, Moreno JL, Garcia C, Hernandez T. Addition of urban waste to semiarid degraded soil: Long-term effect. Pedosphere 2007, 17:557–67. [37] Albiach R, Canet R, Pomares F, Ingelmo F. Organic matter components and aggregate stability after the application of different amendments to a horticultural soil. Bioresour Technol 2001, 76:125–29. [38] Albaladejo J, Castillo V, Díaz E. Soil loss and runoff on semiarid land as amended with urban solid refuse. Land Degr Develop 2000, 16:551–59. [39] Caravaca F, Masciandaro G, Ceccanti B. Land use in relation to soil chemical and biochemical properties in a semiarid Mediterranean environment. Soil Tillage Res 2002, 68:23–30. [40] García C, Hernández T, Albaladejo J, Castillo V, Roldán A. Revegetation in semiarid zones: influence of terracing and organic refuse on microbial activity. Soil Sci Soc Am J 1998, 62:670–76. [41] Bastida F, Kandeler E, Moreno JL, Ros M, Garcia C, Hernandez T. Application of fresh and composted organic wastes modifies structure, size and activity of soil microbial community under semiarid climate. Appl Soil Ecol 2008, 40:318–29. [42] Ros M, Hernández MT, García C. Soil microbial activity after restoration of a semiarid soil by organic amendments. Soil Biol Biochem 2003, 35:463–69. [43] Bastida F, Hernández T, Albaladejo J, García C. Phylogenetic and functional changes in the microbial community of long-term restored soils under semiarid climate. Soil Biol Biochem 2013, 65:12–21.

References |

27

[44] Tejada M, Moreno JL, Hernández MT, García C. Application of two beet vinasse forms in soil restoration: Effects on soil properties in an and environment in southern Spain. Agr Ecosyst Environ 2007, 119:289–98. [45] Kiem R, Kögel-Knabner I. Contribution of lignin and polysaccharides to the refractory carbon pool in C-depleted arable soils. Soil Biol Biochem 2003, 35:101–18. [46] Abiven S, Menasseri S, Chenu C. The effects of organic inputs over time on soil aggregate stability – A literature analysis. Soil Biol Biochem 2009, 41:1–12. [47] Gentile R, Vanlauwe B, Six J. Litter quality impacts short- but not long-term soil carbon dynamics in soil aggregate fractions. Ecol Appl 2011, 21:695–703. [48] Nicolás C, Hernández T, García C. Organic amendments as strategy to increase organic matter in particle-size fractions of a semi-arid soil. Appl Soil Ecol 2012, 57:50–58. [49] García E, García C, Hernández T. Evaluation of the suitability of using large amounts of urban wastes for degraded arid soil restoration and C fixation. Eur J Soil Sci 2012, 63:650–58. [50] Bastida F, Torres IF, Hernández T, Bombach P, Richnow HH, García C. Can the labile carbon contribute to carbon immobilization in semiarid soils? Priming effects and microbial community dynamics. Soil Biol Biochem 2013, 57:892–902. [51] Helgason BL, Gregorich EG, Janzen HH, Ellert BH, Lorenz N, Dick RP. Long-term microbial retention of residue C is site-specific and depends on residue placement. Soil Biol Biochem 2014, 68:231–40. [52] Ben-David EA, Zaady E, Sher Y, Nejidat A. Assessment of the spatial distribution of soil microbial communities in patchy arid and semi-arid landscapes of the Negev Desert using combined PLFA and DGGE analyses. FEMS Microbiol Ecol 2011, 76:492–503. [53] Cotton J, Acosta-Martínez V, Moore-Kucera J, Burow G. Early changes due to sorghum biofuel cropping systems in soil microbial communities and metabolic functioning. Biol Fertil Soils 2012, 49:403–13. [54] Drenovsky RE, Steenwerth KL, Jackson LE, Scow KM. Land use and climatic factors structure regional patterns in soil microbial communities. Glob Ecol Biogeogr 2010, 19:27–39. [55] Hortal S, Bastida F, Armas C, Lozano YM, Moreno JL, García C, Pugnaire FI. Soil microbial community under a nurse-plant species changes in composition, biomass and activity as the nurse grows. Soil Biol Biochem 2013, 64:139–46. [56] Lester ED, Satomi M, Ponce A. Microflora of extreme arid Atacama Desert soils. Soil Biol Biochem 2007, 39:704–08. [57] Entry JA, Fuhrmann JJ, Sojka RE, Shewmaker GE. Influence of irrigated agriculture on soil carbon and microbial community structure. Environ Manage 2004, 33:363–73. [58] Jia GM, Zhang PD, Wang G, Cao J, Han JC, Huang YP. Relationship between microbial community and soil properties during natural succession of abandoned agricultural land. Pedosphere 2010, 20:352–60. [59] Potts M, Olie JJ, Nickels JS, Parsons J, White DC. Variation in Phospholipid Ester-Linked Fatty Acids and Carotenoids of Desiccated Nostoc commune (Cyanobacteria) from Different Geographic Locations. Appl Environ Microbi 1987, 53:4–9. [60] Belnap J, Lange OL. Biological Soil Crust: Structure, Function, and Management. Berlin, Springer-Verlag 2001, 5–12. [61] Griffiths RI, Whiteley AS, O’Donnell AG, Bailey MJ. Physiological and community responses of established grassland bacterial populations to water stress. Appl Environ Microb 2003, 69:6961–68. [62] Fierer N, Schimel JP. Effects of drying-rewetting frequency on soil carbon and nitrogen transformations. Soil Biology and Biochemistry 2002, 34:777–787. [63] Huxman TE, Snyder KA, Tissue D et al. Precipitation pulses and carbon fluxes in semiarid and arid ecosystems. Oecologia 2004, 141:254–68.

28 | 2 Soils in Arid and Semiarid Environments

[64] Stark JM, Firestone MK. Mechanisms for soil moisture effects on activity of nitrifying bacteria. Appl Environ Microb 1995, 61:218–21. [65] Bottner P. Response of microbial biomass to alternate moist and dry conditions in a soil incubated with 14 C- and 15 N-labelled plant material. Soil Biol Biochem 1985, 17:329–37. [66] Kieft TL, Soroker E, Firestone MK. Microbial biomass response to a rapid increase in water potential when dry soil is wetted. Soil Biol Biochem 1987, 19:119–26. [67] Ceccanti B, Nannipieri P, Cerveli S, Sequi P. Fractionation of humus-urease complexes. Soil Biol Biochem 1978, 10:39–45. [68] Bastida F, Jindo K, Moreno JL, Hernández T, García C. Effects of organic amendments on soil carbon fractions, enzyme activity and humus-enzyme complexes under semi-arid conditions. Eur J Soil Biol 2012, 53:94–102. [69] Nannipieri P, Grego S, Ceccanti B. Ecological significance of the biological activity in soils. In: Bollag JM, ed. Stotzky G 2nd edn. New York, Marcel Dekker, 1990, 293–355. [70] Sinsabaugh RL, Lauber CL, Weintraub MN et al. Stoichiometry of soil enzyme activity at global scale. Ecol Lett 2008, 11:1252–64. [71] Bastida F, Moreno JL, Hernández T, García C. Microbiological degradation index of soils in a semiarid climate. Soil Biol Biochem 2006, 38:3463–73. [72] Sinsabaugh RL, Gallo ME, Lauber CL, Waldrop M, Zak DR. Extracellular enzyme activities and soil carbon dynamics for northern hardwood forests receiving simulated nitrogen deposition. Biogeochemistry 2005, 75:201–15. [73] Moreno JL, Hernández T, García C. Effects of a cadmium-contaminated sewage sludge compost on dynamics of organic matter and microbial activity in an arid soil. Biol Fertil Soils 1999, 28:230–37. [74] Pascual JA, García C, Hernández T, Ayuso M. Changes in the microbial activity of an arid soil amended with urban organic wastes. Biol Fertil Soils 1997, 24:429–34. [75] Madejon E, Moreno F, Murillo JM, Pelegrin F. Soil biochemical response to long-term conservation tillage under semi-arid Mediterranean conditions. Soil Till Res 2007, 94:346–52. [76] Moreno B, García-Rodríguez S, Cañizares R, Castro J, Benítez E. Rainfed olive farming in southeastern Spain: Long-term effect of soil management on biological indicators of soil quality. Agr Ecosyst Environ 2009, 131:333–39. [77] Melero S, Lopez-Bellido RJ, Lopez-Bellido L et al. Stratification ratios in a rainfed Mediterranean Vertisol in wheat under different tillage, rotation and N fertilisation rates. Soil Till Res 2012, 119:7–12. [78] González-Polo M, Austin AT. Spatial heterogeneity provides organic matter refuges for soil microbial activity in the Patagonian steppe, Argentina. Soil Biol Biochem 2009, 41:1348–51. [79] Acosta-Martinez V, Acosta-Mercado D, Sotomayor-Ramirez D, Cruz-Rodriguez L. Microbial communities and enzymatic activities under different management in semiarid soils. Appl Soil Ecol 2008, 38:249–60. [80] Henry HAL. Soil extracellular enzyme dynamics in a changing climate. Soil Biol Biochem 2012, 47:53–59. [81] Stursova M, Sinsabaugh RL. Stabilization of oxidative enzymes in desert soil may limit organic matter accumulation. Soil Biol Biochem 2008, 40:550–53. [82] Sinsabaugh RL, Carreiro MM, Repert DA. Allocation of extracellular enzymatic activity in relation to litter composition, N deposition, and mass loss. Biogeochemistry 2002, 60:1–24. [83] Burns RG, DeForest JL, Marxsen J et al. Soil enzymes in a changing environment: Current knowledge and future directions. Soil Biol Biochem 2013, 58:216–34. [84] Fierer N, Schimel JP. A proposed mechanism for the pulse in carbon dioxide production commonly observed following the rapid rewetting of a dry soil. Soil Sci Soc Am J 2003, 67:798– 805.

References | 29

[85] Schimel J, Balser TC, Wallenstein M. Microbial stress-response physiology and its implications for ecosystem function. Ecology 2007, 88:1386–94. [86] Bell TH, Henry HAL. Fine scale variability in soil extracellular enzyme activity is insensitive to rain events and temperature in a mesic system. Pedobiologia 2011, 54:141–46. [87] Albaladejo J, Ortiz R, Garcia-Franco N, Ruiz-Navarro A. Almagto M, Garcia-Pintado J, MartinezMena M. Land use and climate change impacts on soil organic carbon stock in semiarid spain. J Soil Sediments, 2012, 13:265–277. [88] Martinez-Mena M, Lopez J, Almagro M Boix-Fayos C Albaladejo J. Effect of water erosion and cultivation on the soil carbon stock in a semiarid area of South-East Spain. Soil till Res 2008, 99:119–129. [89] Maestre FT, Cortina J. Are Pinus halepensis plantations useful as a restoration tool in semiarid Mediterranean areas? Forest Ecol Manag 2004, 198:303–317. [90] Nicolás C, Kennedy JN, Hernández T, García C, Six J. Soil aggregation in a semiarid soil amended with composted and non-composted sewage sludge- A field experiment. Geoderma 2014, 219–220:24–31. [91] Six J, Elliot ET, Paustian K, Doran JW. Aggregation and soil organic matter accumulation in cultivated and native grassland soils. Soil Sci Soc Am J 1998, 62:1367–1377. [92] Gale WJ, Cambardella CA, Bailey TB. Root-derived carbon and the formation and stabilization of aggregates. Soil Sci Soc Am J 2000, 64:201–207. [93] Caravaca F, Lax A, Albaladejo J. Soil aggregate stability and organic matter in clay and fine silt fractions in urban refuse-amended semiarid soils. Soil Sci Soc Am J 2001, 65:1235–1238. [94] Lopez-Garrido R, Madejon E, Leon-Camacho M, Giron I, Moreno F, Murillo JM. Reduced tillage as an alternative to no tillage under Mediterranean conditions: a case study. Soil Till Res 2014, 140:40–47. [95] Six J, Bossuyt H, Degryze S, Denef K. A history of research on the link between (micro) aggregates, soil biota and soil organic matter dynamics. Soil Till Res 2004, 79:7–31. [96] Garcia-Franco N. Carbon sequestration mechanisms in semiarid soils according to lnad use and management practices. Doctoral Thesis, Murcia University (Spain) 2014, 186 pp.

Gary M. King

3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange in Arid and Semiarid Ecosystems Abstract: Soil water status strongly affects qualitative and quantitative aspects of soil– atmosphere trace gas exchange. Soil water status is most often expressed in terms of gravimetric water contents, which can be particularly useful when translated to gas filled pore space. Gas filled pore space has predictive value for both gas transport rates and the types of processes involved in gas production and consumption. However, water potential offers deeper insights that reflect the physiological responses of cells, while also providing a basis for comparing activities among different soil types and across wetting and drying events. Nonetheless, relatively few studies have incorporated water potential measurements with analyses of trace gas fluxes. Results for atmospheric methane uptake suggest similar sensitivities to water potential for arid soils and forest soils, with strong inhibition below −0.5 MPa. Atmospheric CO uptake in forest soils shows sensitivities similar to those of methane uptake, but recent evidence suggests that CO oxidizers in arid and saline soils might maintain activity at remarkably low potentials. Advances in sensor design should facilitate much more extensive analyses of water potential, more mechanistic models of trace gas exchange, and a better understanding of the controls trace gas dynamics.

3.1 Introduction Water plays a profoundly important role in soil–atmosphere gas exchange [1–6]. Water shapes plant communities; litter development; the presence and characteristics of soil horizons; soil organic matter content; microbial community composition, structure and activity; soil texture, porosity and gas transport [7]. All of these variables interact with water regimes to determine rates of gas emission to, or uptake from, the atmosphere. This is no truer for tropical rainforests than it is for arid ecosystems, the characteristics of which often reflect long term climate change, and not just contemporary hydrologic regimes. For example, the playa soils of the northwestern United States are mostly remnants of extensive Pleistocene lakes that disappeared as a consequence of global climate change (e.g., Lake Bonneville), leaving behind fine grained sediment beds that progressively evolved in response to sparse plant colonization and strongly seasonal patterns of temperature and precipitation [8]. Although water limitations often lead to relatively low rates of gas exchange per m2 , soils in arid and semiarid ecosystems can still play significant roles in some global DOI 10.1515/9783110419047-003

32 | 3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange

trace gas budgets; this is because they account for roughly one third of the total terrestrial surface area [9]. For example, the global soil methane sink is substantially less than it would be if uptake rates in arid systems were equivalent to those in grasslands and forests. Likewise, global uptake of atmospheric carbon monoxide is reduced by the combination of low uptake rates in some arid soils, and emissions from others [10, 11]. Gas exchange in arid and semiarid ecosystems is sensitive to natural and anthropogenic disturbances, many of which affect water regimes and related variables [12– 17]. Climate change, for instance, may result in increased thermal stress and prolonged periods of drought punctuated by extreme precipitation. Irrigation for agriculture has resulted in soil salinization, in some cases rendering soils unsuitable for crop production, and changing local biogeochemical dynamics [18]. While many variables obviously contribute to rates and patterns of gas exchange in arid systems, soil water potential is arguably the most important. Water potential, which is a measure of water availability, affects gas production and consumption at the level of cells, and elicits immediate responses as it changes through its impact on cell physiology [19]. However, in spite of its importance, relationships between trace gas dynamics and water potential have not been characterized extensively. An overview of these relationships and recent observations are summarized here.

3.2 Water Potential and Water Potential Assays Although several weight or volume based indices provide convenient measures of soil water content (e.g., [20]), and are useful in the context of variables such as gas diffusion and advection (e.g., [21, 22]), they provide little insight about the physiological responses of microbes to soil water status, and often cannot be directly compared among systems [23]. In contrast, soil water status can be more completely specified using physical chemical terms (e.g., [19, 24, 25]). The rationale for using a physical chemical description of water as an alternative to volumetric measures is simple. The direction of water movement across cell membranes cannot be predicted on the basis of weight or volumetric measures of water content, but can be predicted using measures of the energy status of water, and water potential in particular. Water potential calculations begin with the mole fraction of water in a solution: Nw = nw /(nw + n i ) , with nw representing number of moles of water kg−1 of solvent (= molality, about 55.51 mol kg−1 or 55.51 m) and n i representing the moles of solute kg−1 of solvent. Since solutions are often not ideal in a thermodynamic context, an activity coefficient, γ, specific for a given solute, is applied, yielding a definition for water activity: aw = γNw .

3.2 Water Potential and Water Potential Assays

|

33

Water activity is often used as a temperature independent measure of water availability, and water activity values will be presented below when relevant for specific discussions. Where appropriate, a water potential equivalent will be presented for a temperature of 25°C. Though there are some advantages to a temperature independent measure of water status, water activity itself does not necessarily predict directions of water flow, and it is inadequate for complex, multiphase systems such as soil. Water potential provides a more complete measure of water availability. Water potential is defined in energetic terms as the partial molal free energy of a solution of water under specified conditions of solute composition, temperature, pressure and gravitational potential: μw = (∂G/∂nw )n i ,T,P,h , where G represents Gibbs free energy, n i is solute concentration, P is pressure, and h is height (ignored in most biogeochemical contexts [23]). This yields a working expression for the chemical potential of water: μw = μ 0w + RT ln aw + Vw P , where μ 0w represents the chemical potential of water in a standard reference state; R, T (in Kelvin) and P represent the gas law constant, temperature, and pressure, respectively, and Vw is the partial molal volume of water (about 1.8×10−5 m3 mol−1 at 25°C). Rearranging yields: (μ w − μ 0w )/Vw = RT ln aw /Vw + P , where the left hand expression is a chemical potential difference per molal volume and is designated water potential, ψ: ψ = RT ln aw /Vw + P . This expression indicates that water potential in a solution can be subdivided into a pressure term (taken as a departure from 1 atm) and a solute dependent term. As applied to soils, the total water potential, Ψ, is typically distributed among three terms: Ψ = ψs + ψp + ψm , where ψs , ψp , ψm are the potentials due to solutes, pressure, and the soil matrix, respectively. The total water potential for any solution is < 0 and is expressed in units of bars or pascals (N m−2 ). Unlike water activity or other measures of water status, Ψ provides a complete description that can be compared among systems and used to predict the direction of water flows, for example into or out of cells. The matric potential term, ψm , is especially relevant in soils. This potential arises as a result of the interaction of water at surfaces in a porous matrix, and has been described by analogy to the behavior of water inside a capillary tube immersed in pure water. The force associated with the rise of water a distance h in a capillary is related

34 | 3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange to the matric potential within the capillary (= hρg, where ρ is water density [kg m−3 ] and g is the gravitational constant [m sec−2 ]); the height of capillary rise is inversely proportional to the capillary radius, r. Soil is essentially a porous matrix in which the matric potential is related to pore size (i.e., pore radius) and the distribution of water among pores (a function of water content). When all pores are filled (water saturation), the matric potential is zero. The matric potential decreases with desaturation due to the loss of water from larger pores and retention in smaller pores. Progressive loss leaves the remaining water in smaller pores at progressively lower potentials. The relationship between water potential and soil pore size distribution has a number of important consequences, especially for gas exchange. With decreasing water content and matric potential, gas transport increases [22, 26, 27], which can accelerate some gas transformations as well as exchanges with the atmosphere. However, water potentials lower than about −0.5 MPa typically inhibit many bacterial activities due to physiological stresses, physical constraints on substrate transport, cell movement, and the thickness of films available for bacterial immersion. This limitation is especially relevant for arid soils, which often experience water potentials much less than −0.5 MPa. Soil water content can be measured readily using relatively simple gravimetric methods [28]. Modifications of these methods yield additional indices of soil pore space, which can aid analyses of soil–atmosphere gas exchange. Several methods and associated instrumentation are also available for analyses of the water potential. However, the choice of method depends greatly on the application. Methods suitable for use in a laboratory context often are unsuited for field use and vice versa. It is also important to understand whether solute potentials, matric potentials, or both need to be measured, since this influences method selection. Finally, the range of expected water potentials must be considered. For arid soils, the range can potentially exceed limits for any one analytical system, since values can approximate zero during wet seasons or immediately after precipitation events, but fall below −100 MPa with drying. For laboratory measurements and water potentials from about −2 kPa to −500 kPa, a pressure plate apparatus can be used (e.g., [29]). Pressure plates essentially apply pressure to a soil sample and drive excess water out through a porous ceramic plate. At equilibrium, the water potential is assumed to equal the applied pressure. The water content of the soil sample is then measured. A set of water content determinations at different pressures is then used to construct a moisture release curve that, in turn, is used to estimate sample potentials at their initial water contents. Other than its simplicity, this approach has little to recommend it, since other methods offer greater accuracy, broader ranges, and more convenience. Tensiometers, which make direct contact with the soil liquid phase, find use in both laboratory and field contexts [30]. These instruments use a porous ceramic reservoir containing pure water (∼ 0 MPa) in contact with a headspace and a pressure transducer or vacuum gauge. When placed in soil with water at lower potential, water flowing from the reservoir results in a reduced headspace pressure equivalent to the

3.3 Limits of Growth and Metabolic Activity

| 35

soil water potential. Since flows are reversible, tensiometers can function as piezometers in some configurations. Though inexpensive and typically rugged, their dynamic range (> −1 kPa to about −100 kPa) substantially limits applications in arid systems. However, a new microtensiometer might greatly extend these limits [31]. An alternative approach that is well suited for laboratory applications measures the energy status of water in a vapor phase equilibrated with a soil sample. Dew point hygrometry has found a wide range of applications, since it is suitable for samples with water potentials from about −0.1 MPa to < −100 MPa [32, 33]. As implemented by Decagon Instruments (Pullman, WA) WP4-T, dew point hygrometry covers water potential values common in arid soils, and does so with good accuracy. However, the approach and the WP4-T have found limited use in the field due to constraints on temperature control. In addition to the WP4-T, Decagon Instruments also offers sensors suitable for field deployment in arid soils [34]. These sensors, e.g., MPS-6, are based on a ceramic substrate with a known moisture release curve. The sensors can be buried in soil, where they record both temperature and water potential changes as the water content of the ceramic substrate varies. The stated measurement range is from −0.01 MPa to −100 MPa. MPS-6 sensors measure the matric potential and thus are not suitable for saline soils or other systems with significant solute potentials. In addition, their utility has not been established for surface soils (e.g., 0–5 cm) that vary substantially over a diurnal cycle.

3.3 Limits of Growth and Metabolic Activity The effects of water availability (most often expressed as aw ) on microbial growth have been given considerable attention in the context of food preservation [35]. Numerous studies have led to general estimates of lower growth limits for a variety of bacteria and fungi that commonly occur in processed foods or that contribute to spoilage. In general, Gram negative bacteria (e.g., Proteobacteria and Bacteroidetes) do not grow at aw < about 0.95 (−7.06 MPa), while Gram positive bacteria (e.g., Actinobacteria and Firmicutes) do not grow with aw < about 0.90 (−14.49 MPa) [19]. There are exceptions, of course. Pontibacillus sp. AS2 and Salinicola sp. LC26 (Firmicutes and Proteobacteria, respectively) grow at aw = 0.775 (−35.06 MPa), and the actinobacterium Mycobacterium parascrofulaceum LAIST_NPS017 grows at aw = 0.800 (−31.93 MPa at 37 °C) (36). Members of the euryarchaeal Halobacteriaceae, typically grow at aw = 0.755 (−40.60 MPa at 40°C), but limits as low as 0.611 (−67.76 MPa) have been extrapolated from growth data [36]. Many fungi grow at aw = 0.700−0.900 (−49.06 MPa to −14.49 MPa), but lower limits of 0.611 have also been extrapolated for a few exceptional strains [36]. Though studies on water activity collectively represent a reasonably broad survey of some economically important taxa, they have nonetheless explored relatively few

36 | 3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange

species from relatively few phyla (mostly Actinobacteria, Euryarchaea, Firmicutes, and Proteobacteria), and have been limited by the need to use cultivable isolates. Thus, water activity limits are essentially unknown for a large percentage of Bacteria, Archaea, and Eucarya, and for members of soil microbial communities in particular. Perhaps more importantly, growth limitation by water availability is largely understood in the context of solute potentials (ψs ), yet matric potentials (ψm ) often determine water availability in soils. While one might propose that the effects of low water potential on macromolecules, especially DNA, would be the same regardless of the mechanism by which water potential is lowered, the ability of cells to respond physiologically to water stress may depend greatly on the relative contribution of solutes versus pore based capillarity (e.g., [37]). Where solutes dominate total water potential, Ψ, intracellular water potentials can be adjusted to osmoconformers via solute transport. When matric potentials dominate Ψ, the ability of cells to adjust may be constrained by solute availability and by the energy required to synthesize intracellular compatible solutes. This has not been explored systematically, but studies with isolates have shown differential responses to ψs versus ψm (e.g., [38, 39]). Nonetheless, relatively little is known about the growth or activity responses of specific isolates to matric potential. Addressing this knowledge gap should be a research priority, particularly since changing precipitation regimes in the future will be accompanied by changing soil water potential regimes. Work by Schnell and King [40] with methanotrophs provides an example of the potential significance of solute versus matric potentials. They used NaCl as a readily transported solute, and sucrose as an impermeable solute to adjust Ψ in growth media. While not directly equivalent to a matric potential, a solute potential arising from an impermeable solute can mimic the effect of matric potentials on cells. Schnell and King [40] observed that both growth and methane uptake rates were inhibited with decreasing water potential to a greater degree with sucrose than with NaCl. This suggests that water potential limits for growth might be lower when solutes dominate Ψ. This is especially relevant for semiarid and arid soils that experience matric potential extremes well below growth limits due to solute potentials. How do the members of soil microbial communities cope with such extremes? While growth certainly provides an exquisitely sensitive index of the ability of microbes to tolerate extreme conditions, metabolic activity can continue beyond the limits for growth. Analyses of metabolic activity as a function of temperature have indicated that maintenance and survival metabolism occur at subzero temperatures well below those at which growth ceases [41]. These results are relevant for understanding relationships between water availability and metabolism, since bacterial activity in ice occurs within solutions that have low ψs . However, lower limits for activity have not been explored systematically as a function of ψs or Ψ for either isolates or mixed populations in natural systems. This is yet another knowledge gap that should be addressed. Price and Sowers [41] have suggested that there is no evidence for a minimum temperature for metabolism, but this might not hold true for water potential.

3.4 Water Potential and Trace Gas Exchanges |

37

3.4 Water Potential and Trace Gas Exchanges Methane. Water content has a profound and well documented impact on soil–atmosphere methane exchanges. At saturation, anoxic conditions can develop, which promote methanogenesis and methane emission. Numerous variables affect the extent to which methanogenic activity occurs, including soil organic matter content and electron acceptor availability. While water potential has not been specifically addressed as a variable for soil methanogenesis, it is clear that some methylotrophic methanogens tolerate solute potentials as low as −40 MPa, since they can produce methane in salt saturated sediments or solutions [42]. Nonetheless, in most cases where methanogens are active, water potentials are high due to low solute concentrations and the absence of matric potentials. Furthermore, there are relatively few arid or semiarid soils for which methanogenesis would have any relevance, since these soils are unsaturated and methanogenesis is inhibited by molecular oxygen, regardless of water potential regimes. Atmospheric methane consumption by methanotrophic bacteria obviously occurs far more commonly in arid and semiarid soils than does methanogenesis. Due to the significance of soil methanotrophs for the atmospheric methane budget (e.g., [43]), numerous studies have addressed the role of variables such as water content, pH, temperature, soil texture, nitrogen content, and land use [6, 44–49]. The effects of water content have largely been understood in the context of gas transport, with high water contents inhibiting uptake from the atmosphere due low diffusion fluxes, and low water contents inhibiting activity presumably due to undefined water stresses. Water potential effects per se have been addressed to only a limited extent. Schnell and King [40] showed that atmospheric methane uptake was very sensitive to water potential in a forest soil. Extreme potentials (e.g., to −10 MPa) in the “O” and “A” horizons that developed during summer appeared to strongly inhibit uptake and constrain activity to lower depths, the effect of which was to reduce area based rates year round. Combined analyses of water content and water potential also showed that interactions between soil gas exchange, methane concentration, and water stress determined uptake rates and responses to water potential. In particular, decreasing water content at high water potentials (> −0.2 MPa) increased gas transport and methane uptake, even though methanotrophs experienced water stress. However, continued decreases in water content led to increased stress and decreased methane uptake (󳶳 Fig. 3.1). Addition of exogenous methane to a concentration of 200 ppm minimized gas transport limitation and revealed that water stress inhibition developed at Ψ ≥ −0.2 MPa (󳶳 Fig. 3.1). Isolates were similarly sensitive to water stress, whether it was imposed as a solute stress or through a mimic of the matric potential. The patterns observed in Maine forest soils (USA) were confirmed by Bradford et al. [47] for UK temperate forests, and by Gulledge and Schimel [46] for boreal soils. Water stress sensitivity observed for surface soils in these studies likely occurs in surface soils of arid and semiarid systems, which might explain the subsurface localization of

(a)

3.0

0.055

2.5

0.050

2.0

0.045

1.5

0.040

1.0

0.035

0.5

0.030 –1.0

0.00

–0.20 Water potential (MPa)

0.060

Methane uptake rate (nmol gdw–1 h–1 )

Methane uptake rate constant (h–1 gdw–1)

38 | 3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange

–0.40

–0.60

–0.80

0.0 –0.80

–0.60

–0.40

–0.20

Water potential (MPa)

–1.00

0.0

15

(b)

20

25

30

35

40

Water content (%)

Fig. 3.1: (a) Methane uptake rate constants with atmospheric methane and methane uptake rates at 200 ppm methane versus soil water potential for Maine forest soils. From Schnell and King (40). (b) Water potential versus water content for the same soils.

a process that depends on an atmospheric substrate (e.g., [44]). If surface soils were not inhibitory in some manner, they would be the locus of greatest uptake activity, since the supply of methane is greatest there. However, the lack of parallel, time varying depth specific water potential and methane uptake data limit extrapolations. Even so, it is clear that extreme water potentials develop in the surface soils of arid systems, and that soils most conducive to active methanotrophy occur primarily in deeper horizons (e.g., > 10 cm). Seasonal studies have also shown that the highest methane uptake rates in arid soils are associated with precipitation events, albeit with a lag, which indicates that water stress tolerant methanotrophs likely do not occur at substantial levels. Though models of climate change impacts on soil methane fluxes include relationships between water potential and inhibition of methane uptake (e.g., [50– 52]), one such relationship predicts significant uptake at water potential values ≪ −10 MPa [50], an outcome that has not been verified empirically for soils in general, let alone for arid and semiarid soils. Given the lack of spatial coverage by direct studies of atmospheric methane uptake, simulation models offer a potentially valuable tool for developing estimates of global uptake rates. However, to be fully useful, the water potential uptake rate relationship should be established empirically for multiple soil types and systems and for wetting and drying cycles to evaluate hysteresis effects. Carbon monoxide. By regulating hydroxyl radical concentrations to a great degree, CO plays a critical role in tropospheric chemistry [53]. Hydroxyl radical is the primary oxidant in the troposphere, and as such is responsible for chemical oxidation of atmospheric methane and other organic gases. Since it contributes significantly to atmospheric CO dynamics, uptake by soils has been the focus of multiple studies, which have addressed rates, controls, and some aspects of CO microbiology [54, 55]. Although CO transformations in soil have been explored much less than methane

3.4 Water Potential and Trace Gas Exchanges |

39

transformations, several studies have established dependencies on soil water content [56]. Patterns somewhat analogous to those for methane oxidation have emerged, with lower rates of CO uptake at high water contents, and increasing uptake rates as gas transport increases with lower water contents; at relatively low water contents, uptake ceases due to water stress, and net CO emission can sometimes be observed. Relationships between water potential and atmospheric CO uptake have received little attention. Weber and King [57] examined controls of CO uptake by unvegetated and vegetated volcanic cinders on Hawai’i Island (USA). Though not in an arid or semiarid climate, water availability oscillated dramatically on a diurnal basis (between 0 and −60 MPa) for unvegetated cinders due to their very limited water retention capacity, which resulted from low organic contents. In contrast, water potential for nearby cinders at a vegetated site with high organic concentrations varied very little (0 to −0.1 MPa). During a moderate drying event (from 0 to −1.7 MPa), atmospheric CO consumption by intact cores from the unvegetated site decreased 2.7-fold, indicating a strong dependence on water potential. In laboratory assays, maximum potential CO oxidation rates decreased by 40 and 60%, respectively, when water potentials were lowered from 0 to −1.5 MPa, confirming sensitivity observed in the field, but also indicating that CO oxidizing communities at the two sites were not differentially adapted to water stress. Additional analyses revealed that even after desiccation to −150 MPa for 63 days, CO oxidation by unvegetated cinders resumed within a few hours of rehydration, which indicated that CO oxidizers were able to survive extended water stress. Samples from both sites that were exposed to multiple wetting–drying cycles (from 0 to −80 MPa) lost significant activity after the first cycle, but uptake quickly stabilized and was similar after repeated cycles [57]. This suggested that CO oxidizers at both sites were relatively resistant and resilient to water stress. CO oxidizers in arid and semiarid soils must be similarly resistant and resilient to water stress, however empirical studies that establish this point are lacking. Nonetheless, pilot studies of atmospheric CO uptake by playa soils from the Alvord Basin (Oregon, USA) during July 2014 and 2015 (GM King, unpublished) revealed activity at water potentials between approximately −30 MPa to −50 MPa for sites that had experienced water potentials between −200 MPa and −300 MPa (consistent with ambient relative humidity). This clearly documents a substantial capacity for tolerance of extreme water stress. The possibility that atmospheric CO can be consumed at water potentials as low as −50 MPa also distinguishes the capabilities of playa soil CO oxidizers from those of forest soils and cinders, and suggests that arid and semiarid soils might play a greater role in the global soil methane sink than some have previously assumed [58]. There are, of course, numerous unanswered questions about CO oxidation at such low water potentials: What organisms are involved? What mechanisms promote their activity? How do they respond to diurnal and seasonal variations in water availability? How does activity in arid and semiarid soils vary among systems and soil types? Recent results from saline soils near the Bonneville Salt Flats (Utah, USA) have provided some insights for a few of these questions. King [59] observed atmospheric

40 | 3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange 300

Core headspace CO (ppb)

250

200

150

100

50 Thershold, 60.6 parts per billion 0 0

5

15

10

20

25

Time (h) Fig. 3.2: Atmospheric CO uptake by triplicate intact cores from saline soils adjacent to the Bonneville Salt Flats; water potentials were approximately −41 MPa. Data are the means of triplicate assays with 1 standard error indicated. The dashed line indicates the uptake threshold concentration. From King [59].

CO uptake by intact cores of saline soils with surface water potentials of about −40 MPa (󳶳 Fig. 3.2). Depth profiles of CO uptake potential and water potential revealed an inverse relationship, with the highest uptake potential at the lowest water potential. This suggested that a CO oxidizing community was adapted to water stress regimes dominated by the presence of salts. Additional analyses revealed CO oxidizing extreme halophiles (Euryarchaeota) that could consume atmospheric CO while growing in halite saturated brines [59, 60]. These results further established the potential for CO uptake under conditions of low water potential and extended activity to saline soils. They also indicated that novel euryarchaeotes might be the active agents when potentials are poised by solutes versus matric stresses. Obviously, a great deal remains to be learned. Other gases. Soils are globally important sources and/or sinks for many other trace gases, few of which have been evaluated in the context of water potential or water stress [61, 62]. Disregarding CO2 , a trace gas that should be treated separately (e.g., [5, 48, 63–65]), perhaps the most thoroughly studied gases other than methane include nitrous oxide and NO. Both play roles in radiative forcing. Nitrous oxide is well known for its contribution to stratospheric ozone depletion and for its greenhouse properties [62]. NO is well known as an important reactant in tropospheric chemistry, and it contributes to formation of tropospheric ozone, which is a potent greenhouse

3.5 Conclusions | 41

gas that also causes substantial losses of plant production in agriculture and damage to human health [62]. Nitrous oxide and NO dynamics depend substantially on soil water regimes. High water contents and low water potentials favor nitrous oxide production from denitrification, since it is oxygen sensitive. However, denitrification is often nitrate limited and dependent on nitrification, an aerobic process [66]. Nitrification is favored at lower water contents, but it is also very sensitive to water potentials of less than about −0.1 MPa [67, 68]. In addition, nitrification (ammonia oxidation in particular) can form both NO and nitrous oxide. The outcome of these relationships is that nitrous oxide and NO emissions tend to be maximized at intermediate water contents, and presumably intermediate water potentials, though the latter have seldom been measured during flux studies [69–71]. In arid and semiarid soils, nitrogen gas fluxes often depend on water pulses in the form of episodic precipitation, which can drastically and rapidly alter microbial community activity, resulting in short term bursts of metabolism that include nitrification and denitrification, and elevated, but time varying nitrous oxide and NO emissions (e.g., [1, 4. 17, 72,73,74]). Though water contents have been routinely measured in precipitation or wetting studies, water potential has not. Given the possibility of hysteresis effects in water potential–water content relationships, and different relationships for different soil types [75], water potential analyses could promote a greater understanding of the mechanisms and variables that control nitrogen gas transformations, while also facilitating comparisons among systems. Water content and water potential also play important roles in the dynamics of nitrogen oxide emission from biological soil crusts (BSC), which can represent significant NOx sources during wetting events (e.g., [70, 76, 77]). Although BSC behavior is certainly very sensitive to water potential [78], water content has been most commonly measured in studies of BSC photosynthesis or other activities (e.g., [2]). Nonetheless, Potts and Friedman [38] showed that matric and solute stresses elicit different responses from cyanobacteria, and that responses to a given stress differ among cyanobacteria. These findings suggest that responses to water stress by BSC may vary across space or time as community composition varies. Given the global extent and significance of BSC, and their sensitivity to climate change, a greater emphasis on water potential and not just water content is essential for an improved mechanistic understanding and for model projections of responses to change.

3.5 Conclusions Soil water potential is a master variable that to a large degree determines the patterns and rates of trace gas exchanges between soils and the atmosphere. Soil water potential varies with volumetric water content, but the relationship is nonlinear and varies among soil types. In addition, water potential, but not water content, offers a mech-

42 | 3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange

anistic understanding of trace gas production and consumption at a cellular level. For example, decreasing water contents can enhance the physical process of gas exchange, but the accompanying decreases in water potential typically inhibit trace gas production and consumption physiologically. Improved designs for small, relatively inexpensive systems that can measure in situ water potentials at < −10 MPa, and even < −100 MPa, offer new possibilities for more extensive water potential monitoring in semiarid and arid soil systems. More routine application of these technologies will greatly improve predictive models for trace gas dynamics, especially in the context of changing climate regimes and increased frequencies of extreme events.

References [1] [2]

[3]

[4] [5]

[6]

[7] [8] [9] [10] [11] [12] [13]

[14] [15]

McLain JET, Martens DA. Moisture controls on trace gas fluxes in semiarid riparian soils. Soil Sci Soc Am J 2006, 70:367. Grote EE, Belnap J, Housman DC, Sparks JP. Carbon exchange in biological soil crust communities under differential temperatures and soil water contents: implications for global change. Global Change Biol 2010, 16:2763–74. Wu X, Yao Z, Brüggemann N, Shen ZY, Wolf B, Dannenmann M, et al. Effects of soil moisture and temperature on CO2 and CH soil–atmosphere exchange of various land use/cover types in a semi-arid grassland in Inner Mongolia, China. Soil Biol Biochem 2010, 42:773–87. Harms TK, Grimm NB. Responses of trace gases to hydrologic pulses in desert floodplains. Journal of Geophysical Research: Biogeosci 2012, 117:doi:10.1029/2011JG001775. Moyano FE, Vasilyeva N, Bouckaert L, Cook F, Craine J, Curiel Yuste J, et al. The moisture response of soil heterotrophic respiration: interaction with soil properties. Biogeosci 2012, 9:1173–82. Luo GJ, Kiese R, Wolf B, Butterbach-Bahl K. Effects of soil temperature and moisture on methane uptake and nitrous oxide emissions across three different ecosystem types. Biogeosci 2013, 10:3205–19. Porporato A, Daly E, Rodriguez-Iturbe I. Soil water balance and ecosystem response to climate change. Am Nat 2004, 164:625–632. Oviatt CG. Lake Bonneville fluctuations and global climate change. Geol 1997, 25:155–158. Galbally IE, Kirstine WV, Meyer CP, Wang YP. Soil–atmosphere trace gas exchange in semiarid and arid zones. J Environ Qual 2008, 37:599. Conrad R, Seiler W. Arid soils as a source of atmospheric carbon monoxide. Geophys Res Lett 1982, 9:1353–56. Conrad R, Seiler W. Influence of temperature, moisture, and organic carbon on the flux of H2 and CO between soil and atmosphere: field studies in subtropical regions. 1985, 90:5699–709. Billings SA, Schaeffer SM, Evans RD. Trace N gas losses and N mineralization in Mojave desert soils exposed to elevated CO2 . Soil Biol Biochem 2002, 34:1777–84. Pérez MVA, Castañeda JG, Frías-Hernández JT, Franco-Hernández O, Van Cleemput O, Dendooven L, et al. Trace gas emissions from soil of the central highlands of Mexico as affected by natural vegetation: a laboratory study. Biol Fertil Soils 2004, 40:252–9. McLain JET, Martens DA, McClaran MP. Soil cycling of trace gases in response to mesquite management in a semiarid grassland. J Arid Environ 2008, 72:1654–65. Dijkstra FA, Morgan JA, LeCain DR, Follett RF. Microbially mediated CH4 consumption and N2 O emission is affected by elevated CO2 , soil water content, and composition of semi-arid grassland species. Plant Soil 2009, 329:269–81.

References | 43

[16] Singh JS. Anticipated effects of climate change on methanotrophic methane oxidation. Climate Change Environ Sustain 2013, 1:20. [17] Homyak PM, Sickman JO. Influence of soil moisture on the seasonality of nitric oxide emissions from chaparral soils, Sierra Nevada, California, USA. J Arid Environ 2014, 103:46–52. [18] Ladeiro B. Saline agriculture in the 21st century: using salt contaminated resources to cope with food requirements. J Bot 2012, doi:10.1155/2012/310705. [19] Brown AD. Microbial water stress physiology: principles and perspectives. 1990, Wiley & Sons, NY. [20] Tate RL III. Soil microbiology, 2nd edn. 2000, Wiley & Sons, NY. [21] Castro MS, Steudler PA, Bowden RD. Factors controlling atmospheric methane consumption by temperate forest soils. Glob Biogeochem Cyc 1995, 9:1–10. [22] Moldrup P, et al. Predicting the gas diffusion coefficient in undisturbed soil from soil water characteristics. Soil Sci Soc Am J 2000, 64:94–100. [23] Fenchel T, King GM, Blackburn TH. Bacterial biogeochemistry: the ecophysiology of mineral cycling. 2012,Academic Press, New York. [24] Griffin DM. Water and microbial stress. Adv Microb Ecol 1981, 5:91–136. [25] Nobel PS. Physiochemical and environmental plant physiology, 2nd edition. 1999, Academic Press, New York. 489 p. [26] Skopp J. Oxygen uptake and transport in soils: analysis of the air-water interfacial area. Soil Sci Soc Am J 1985, 49:1327–31. [27] Skopp J, Jawson MD, Doran JW. Steady-state aerobic microbial activity as a function of soil water content. Soil Sci Soc Am J 1990, 54:1619–25. [28] Jarrell WM, Armstrong DE, Grigal DF, Kelly EF, Monger HC, Wedin DA. Soil water and temperature status. In: Robertson GP, Coleman DC, Bledsoe CS, Sollins P (eds). Standard soil methods for long-term ecological research. Oxford Univ. Press, Oxford, 1999, 55–73. [29] Bittelli M, Flury M. Errors in water retention curves determined with pressure plates. Soil Sci Soc Am J 2009, 73:1453–60. [30] Whalley WR, Ober ES, Jenkins M. Measurement of the matric potential of soil water in the rhizosphere. J Exp Bot 2013, 64:doi:10.1093/jxb/ert044. [31] Pagay V, Santiago M, Sessoms DA, Huber EJ, Vincent O, Pharkya A, Corso TN, Lakso AN, Stroock AD. A microtensiometer capable of measuring water potentials below −10 MPa. Lab Chip 2014, 14:142806–17. [32] Fonteyn PJ, Schlesinger WH, Marion GM. Accuracy of soil thermocouple hygrometer measurements in desert ecosystems. Ecol 1987, 68:1121–24. [33] Mantri S, Bulut R. Evaluating performance of a chilled mirror device for soil total suction measurements. Geotechnical Special Publication 2014, doi:10.1061/9780784478509.008. [34] Nolz R, Kammerer G, Cepuder P. Calibrating water potential sensors integrated into a wireless network. Ag Wat Manage 2013, 116:12–20. [35] Jay JM. Modern food microbiology, 5th edn. 2012 Springer Science & Business Media. [36] Stevenson A, Burkhardt J, Cockell CS, Cray JA, Dijksterhuis J, Fox-Powell M, et al. Multiplication of microbes below 0.690 water activity: implications for terrestrial and extraterrestrial life. Environ Microbiol 2015, 17:257–77. [37] Cytryn EJ, Sangurdekar DP, Streeter JG, Franck WL, Chang WS, Stacey G, et al. Transcriptional and physiological responses of Bradyrhizobium japonicum to desiccation-induced stress. J Bacteriol 2007, 189:6751–62. [38] Potts M, Imre-Friedman E. Effects of water stress on cryptoendolithic cyanobacteria from hot desert rocks. Arch Microbiol 1981, 130:267–71.

44 | 3 Water Potential as a Master Variable for Atmosphere–Soil Trace Gas Exchange

[39] Johnson DR, Coronado E, Moreno-Forero SK, Heipieper HJ, van der Meer JR. Transcriptome and membrane fatty acid analyses reveal different strategies for responding to permeating and non-permeating solutes in the bacterium Sphingomonas wittichii. BMC Microbiol 2011, 11:250. [40] Schnell S, King GM. Responses of methanotrophic activity in soils and cultures to water stress. Appl Environ Microbiol 1996, 62:3203–09. [41] Price PB, Sowers T. Temperature dependence of metabolic rates for microbial growth, maintenance, and survival. Proc Natl Acad Sci USA 2004, 101:4631–6. [42] Giani D, Jannsen D, Schostak V, Krumbein W. Methanogenesis in a saltern in the Bretagne (France). FEMS Microbiol Ecol 1989, 62:143–50. [43] King GM. Ecological aspects of methane oxidation, a key determinant of global methane dynamics. Adv Microbial Ecol 1992, 12:431–468. [44] Striegl RG, McConnaughey TA, Thorstenson DC, Weeks EP, Woodward JC. Consumption of atmospheric methane by desert soils. Nature 1992, 357:145–7. [45] Ball BC, Smith KA, Klemedtsson L, Brumme R, Sitaula BK, Hansen S, et al. The influence of soil gas transport properties on methane oxidation in a selection of northern European soils. J Geophys Res 1997, 102:23309. [46] Gulledge J, Schimel JP. Moisture control over atmospheric CH4 consumption and CO2 production in diverse Alaskan soils. Soil Biol Biochem 1998, 30:1127–32. [47] Bradford MA, Wookey PA, Ineson P, Lappin-Scott HM. Controlling factors and effects of chronic nitrogen and sulphur deposition on methane oxidation in a temperate forest soil. Soil Biol Biochem 2001, 33:93–102. [48] Davidson EA, Ishida FY, Nepstad DC. Effects of an experimental drought on soil emissions of carbon dioxide, methane, nitrous oxide, and nitric oxide in a moist tropical forest. Glob Change Biol 2004, 10:718–30. [49] Norton U, Mosier AR, Morgan JA, Derner JD, Ingram LJ, Stahl PD. Moisture pulses, trace gas emissions and soil C and N in cheatgrass and native grass-dominated sagebrush-steppe in Wyoming, USA. Soil Biol Biochem 2008, 40:1421–31. [50] Curry CL. Modeling the soil consumption of atmospheric methane at the global scale. Global Biogeochem Cyc 2007, 21:4. [51] Curry CL. The consumption of atmospheric methane by soil in a simulated future climate. Biogeosci 2009, 6:2355–67. [52] Nazaries L, Murrell JC, Millard P, Baggs L, Singh BK. Methane, microbes and models: fundamental understanding of the soil methane cycle for future predictions. Environ Microbiol 2013, 15:2395–417. [53] Crutzen PJ, Gidel LT. A two-dimensional photochemical model of the atmosphere. 2: The tropospheric budgets of the anthropogenic chlorocarbons, CO, CH4 , CH3Cl and the effect of various NOx sources on tropospheric ozone. J Geophys Res 1983, 88:6641–61. [54] Conrad R. Soil microorganisms as controlers of atmospheric trace gases (H2 , CO2 , CH4 , OCS, N2O , NO). Microbiol Rev 1996, 60:609–640. [55] King GM. Characteristics and significance of atmospheric carbon monoxide consumption by soils. Chemosphere: Global Change Sci 1999, 1:53–63. [56] King GM. Attributes of atmospheric carbon monoxide oxidation in Maine forest soils. Appl Environ Microbiol 1999, 65:5257–64. [57] Weber CF, King GM. Water stress impacts on bacterial carbon monoxide oxidation on recent volcanic deposits. ISME J 2009, 3:1325–34. [58] Potter CS, Davidson EA, Verchet LV. Estimation of global biogeochemical controls and seasonality in soil methane consumption. Chemosphere 1996, 32:2219–46. [59] King GM. Carbon monoxide as a metabolic energy source for extremely halophilic microbes: Implications for microbial activity in Mars regolith. Proc Natl Acad Sci USA 2015, 112:4465–70.

References |

45

[60] McDuff S, King GM, Neupane S, Myers M. Isolation and characterization of extremely halophilic CO-oxidizing Euryarchaeota from hypersaline cinders, sediments and soils, and description of a novel CO oxidizer, Haloferax namakaokahaiae Mke2.3T , sp. nov. FEMS Microbiol Ecol 2016, 92:doi:10.1093/femsec/fiw028. [61] Mooney HA, Vitousek PM, Matson PA. Exchange of materials between terrestrial ecosystems and the atmosphere. Science 1987, 238:926–32. [62] Monson RK, Holland EA. Biospheric trace gas fluxes and their control over tropospheric chemistry. Annu Rev Ecol Syst 2001, 32:547–76. [63] Davidson EA, Verchot LV, Cattanio JH, Ackerman IL, Carvalho JEM. Effects of soil water content on soil respiration in forests and cattle pastures of eastern Amazonia. Biogeochem 2000, 48:53–69. [64] Fierer N, Schimel JP. A proposed mechanism for the pulse in carbon dioxide production commonly observed following the rapid rewetting of a dry soil. Soil Sci Soc Am J 2003, 67:798– 805. [65] Jassal RS, Black TA, Novak MD, Gaumont-Guay D, Nesic Z. Effect of soil water stress on soil respiration and its temperature sensitivity in an 18-year-old temperate Douglas-fir stand. Global Change Biol 2008, 14:1305–18. [66] Bateman EJ, Baggs EM. Contributions of nitrification and denitrification to N2 O emissions from soils at different water-filled pore space. Biol Fertil Soils 2005, 41:379–88. [67] Stark JM, Firestone MK. Mechanisms for soil moisture effects on activity of nitrifying bacteria. Appl Environ Microbiol 1995, 61:218–21. [68] Gleeson DB, Herrmann AM, Livesley SJ, Murphy DV. Influence of water potential on nitrification and structure of nitrifying bacterial communities in semiarid soils. Appl Soil Ecol 2008, 40:189–94. [69] Bargsten A, Falge E, Pritsch K, Huwe B, Meixner FX. Laboratory measurements of nitric oxide release from forest soil with a thick organic layer under different understory types. Biogeosci 2010, 7:1425–41. [70] Weber B, Wu D, Tamm A, Ruckteschler N, Rodriguez-Caballero E, Steinkamp J, et al. Biological soil crusts accelerate the nitrogen cycle through large NO and HONO emissions in drylands. Proc Natl Acad Sci USA 2015, 112:15384–9. [71] Vourlitis GL, DeFotis C, Kristan W. Effects of soil water content, temperature and experimental nitrogen deposition on nitric oxide (NO) efflux from semiarid shrubland soil. J Arid Environ 2015, 117:67–74. [72] Fierer N, Schimel JP, Holden PA. Influence of drying-rewetting frequency on soil bacterial community structure. Microb Ecol 2003, 45:63–71. [73] Austin AT, Yahdjian L, Stark JM, Belnap J, Porporato A, Norton U, et al. Water pulses and biogeochemical cycles in arid and semiarid ecosystems. Oecol 2004, 141:221–35. [74] Steenwerth K, Jackson L, Calderon F, Scow K, Rolston D. Response of microbial community composition and activity in agricultural and grassland soils after a simulated rainfall. Soil Biol Biochem 2005, 37:2249–62. [75] Royer JM, Vachaud G. Field determination of hysteresis in soil-water characteristics. Soil Sci Soc Am J 1975, 39:221–223. [76] Barger NN, Belnap J, Ojima DS, Mosier A. NO Gas loss from biologically crusted soils in Canyonlands National Park, Utah. Biogeochem 2005, 75:373–91. [77] Abed RM, Lam P, de Beer D, Stief P. High rates of denitrification and nitrous oxide emission in arid biological soil crusts from the Sultanate of Oman. ISME J 2013, 7:1862–75. [78] Rajeev L, da Rocha UN, Klitgord N, Luning EG, Fortney J, Axen SD, et al. Dynamic cyanobacterial response to hydration and dehydration in a desert biological soil crust. ISME J 2013, 7:2178–91.

Thulani P. Makhalanyane, Storme Z. de Scally, and Don A. Cowan

4 Microbiology of Antarctic Edaphic and Lithic Habitats 4.1 Introduction The Antarctic atmosphere has recently exceeded the nominal barrier of 400 ppm CO2 [1]. Climate models designed to predict future temperature regimes over the Antarctic continent are complicated by the interactions between the atmosphere, ocean, and ice in lower latitude regions [2]. Nevertheless, these models consistently predict a long term increase in average surface temperatures [3], where southern polar regions may experience average temperature increases of between 0.3–4.8°C by the end of the twenty first century [4]. The projected upper range temperature increases are likely to substantially influence biological community composition and functional processes in a range of nonmarine Antarctic ecosystems, including lakes and ponds [5, 6], permafrost [7, 8], ice shelves [9, 10], glaciers and meltwater streams [11–13], and soils (and their associated cryptic and refuge niches) [14–16]. However, feedback of soil ecosystems to climate change remain unclear, despite the fact that more carbon is stored in these systems than in plant and atmospheric pools [17, 18]. For instance, carbon stored in Arctic and Antarctic permafrost alone may significantly intensify climate change through carbon–climate feedback [19]. We therefore argue, as have others [20–22], that a comprehensive understanding of the terrestrial microbiota of the Antarctic continent is essential in order to appreciate the impacts of projected future climate changes. The majority of the Antarctic continent is covered by an extensive ice sheet, with less than 3% of the total land surface comprised of ice free regions [23, 24]. These regions include mountain ranges, nunataks and coastal arid soils, but are mostly restricted to coastal areas. Ice free soils may only represent a very small fraction of the total land area of the continent, but they harbor considerable numbers and diversity of microbial taxa that survive in these extremely challenging environmental conditions [25]. The development of modern metagenomic methods has, as elsewhere, helped to reveal the true extent of microbial diversity in a diverse range of Antarctic habitats, including oligotrophic, copiotrophic, psychrophilic, and thermophilic soils. In this chapter, we review the status of current microbiology research on Antarctic soil communities and the associated cryptic niche habitats (hypoliths, endoliths and epiliths). We have not focused extensively on permafrost and biological soil crust habitats, both of which have been the subjects of recent reviews [16, 26].

DOI 10.1515/9783110419047-004

48 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

4.2 Classification of Antarctic soils Studies on Antarctic soils began in the early 1900s and were based on genetic (pedogenic processes) and taxonomic (soil properties) classification schemes [27]. Jensen (1916) was the first to propose that Antarctic soils cannot be classified as “typical” due to the lack of the organic layer typically associated with soils in other environments (󳶳 Fig. 4.1a). Loosely arranged unconsolidated Antarctic terrestrial sediments, most of which lack higher life forms (e.g., plants), also failed to adhere to accepted soil taxonomy classification guidelines (󳶳 Fig. 4.1b) [27]. However, studies during the 1960s led to the recognition of a range of soil forming or pedogenic processes within the ice free regions of the Antarctic continent [28–31], and the recognition that Antarctic soil development is influenced by a number of common pedogenic factors including time, climate and the parent material. The accepted conclusion is that the unconsolidated gray materials were valid soils [27]. The initial Antarctic soil classification scheme, introduced in 1966, led to the categorization of six groups [32]. These included the ahumic soils (low organic matter content), evaporate soils (containing substances left after the evaporation of a body of water), regosols (weakly developed, loose mineral soils), lithosols (soil containing mostly weathered rock fragments), protoranker soils (colonized by moss and lichens) and ornithogenic soils (influenced by birds) [27]. Further soil classifications were introduced by Campbell and Claridge (1977), with the subdivision of the six groups into zonal, intrazonal and azonal categories. Ahumic soils are considered zonal as they are strongly influenced by climate and are, therefore, further subdivided on the basis of moisture availability, soil development and parent material composition [33]. Regosols are considered azonal, whereas evaporate, protoranker and ornithogenic soils are intrazonal [33].

(a)

(b)

Fig. 4.1: (a) Antarctic Dry Valley soils showing the typical pavement structure where mineral soils are overlain by stones (typically quartz), with the typical organic layer absent. (b) An ice free Antarctic Dry Valley region showing terrestrial soils that are loosely arranged and lack higher terrestrial life forms.

4.2 Classification of Antarctic soils

|

49

Early investigations revealed that chemical weathering and ionic migration also occurred within Antarctic soils, shaping their formation and characteristics [34, 35]. The determination of soil properties, as well as the introduction of the soil classification schemes, led to an alternative definition of soil, which was recognized and approved (Soil Survey Staff, 1999). The new definition described soil as “a natural body comprised of solids, liquids and gases organized into horizons readily distinguishable from the initial starting material as a result of addition, losses, transfers and transformation of energy and matter” [36]. Based on this new definition, Antarctic soils could be classified according to pedogenic processes affected by factors such as time and climate, as well as soil properties. Climatic conditions and physiochemical properties differ markedly across the ice free regions of the Antarctic continent, such as the McMurdo Dry Valleys (MDVs) and the Antarctic Peninsula, resulting in unique soil biotopes in each region [27].

4.2.1 McMurdo Dry Valley Soils The MDVs, occurring within the South Victoria Land zone (roughly from 77° S to 78° S), represent the largest ice free region of Antarctica [37]. The MDVs are characterized as cold hyperarid desert regions [38] and are subject to extreme climatic conditions including very low temperatures [39, 40], low atmospheric moisture levels and water availability [41], high levels of UV radiation [37] and strong katabatic winds [42]. The MDVs have a mean precipitation rate of less than 10 cm yr−1 [43], mostly in the form of snow that sublimes rather than melts, allowing very little moisture to reach the soil subsurface [37, 38]. Average annual air temperatures range from −15°C to −30°C [44], although surface soil temperatures can reach a maximum of around 15°C for short periods in the summer months [44, 45]. Frequent freeze–thaw cycles occur in MDV soils, where fluctuations of −15°C to > +20°C have been observed within a single day [39, 40]. The Dry Valleys contain both ephemerally wetted soils from glacial melt exposure and depauperate mineral soils [46, 47]. The mineral soils within the MDVs are mostly alkaline, with pH values ranging from 7 to almost 10 in some valley regions [48–51]. MDV soils are often saline and may contain high concentrations of soluble salts such as calcium, magnesium, sodium, chloride, nitrate and sulfate [37, 41, 50]. Soluble nitrogen and phosphorus concentrations vary widely, with ranges of 0–1250 µg g−1 and 0.01–120 µg g−1 , respectively [48]. Organic matter content is typically very low, with a mean percentage carbon level of less than 0.1% in many soils [52]. The percentage of sand is markedly higher than the percentage of clay and silt (usually less than 15% combined) within MDV soils [27]. MDV soils are influenced by both chemical and physical parameters, perhaps more so than other soils [27]. The predominant pedogenic processes in this region include salinization and desert pavement formation [53]. These mineral soils contain

50 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

a layer of cemented permafrost, although the depth of this layer may vary [8]. The taxonomic classification of MDV soils into two suborders of the order Gelisols, namely Turbels and Orthels, is based on the characteristics and proximity of permafrost to the mineral soil surface [27]. Turbels contain ice cemented permafrost within 70 cm of the soil surface and are generally cryoturbated, indicating that materials from different soil horizons were mixed due to freeze–thaw cycles [27]. Orthels, in contrast, contain dry permafrost and little cryoturbation [27]. Based on these classifications, the dominant soil types within the MDVs are Typic Haploturbels, Typic Anhyturbels and Typic Anhyorthels, where haplo refers to simple and anhy refers to low levels of moisture or precipitation [54]. The depth of the permafrost layer and the degree of permafrost melting may be important factors in water availability to surface and shallow subsurface microbial communities.

4.2.2 Antarctic Peninsula Soils The Antarctic Peninsula, in contrast to the MDVs, experiences less severe environmental conditions. Nutrient and moisture availability is generally much greater, with many soils within this region being copiotrophic [24, 55]. The more temperate conditions of the Peninsula support the development of higher life forms, such as plants, which then sustain other animals, such as birds [56]. The nutrient inputs from these organisms alter the physiochemical characteristics of the soil, thereby leading to the alternative, well developed soil biotopes present on the Antarctic Peninsula and surrounding islands [57]. The greater soil taxonomic diversity within the peninsula is due to the diverse soil characteristics as well as the number of soil forming processes in this region [58, 59]. The main pedogenic processes occurring within the maritime Antarctic include rubification, carbonation, humification, podsolization, phosphatization and cryoturbation [53]. The common soil orders within the Antarctic Peninsula, as classified by soil taxonomy, include the entisols (soils that are extremely underdeveloped), inceptisols (soils that are weakly developed) and histosols (soils that contain organic matter) [54]. Within these, the two suborders, Typic Gelorthents and Typic Gelaquents, are the most common, although Turbic Dystrogelepts, Turbic Humigelepts and Saprists also occur within the peninsula [60]. Ornithogenic soils, which are common on the Antarctic Peninsula, are characterized as continuous or historical nutrient inputs from birds, particularly guano (bird excrement) [27]. As a consequence, ornithogenic soils are highly enriched in nutrients such as phosphorus, inorganic nitrogen and organic carbon [61]. This external nutrient input also results in high ammonium levels (up to 5% of the dry weight of soil) due to the conversion of uric acid to ammonia [62]. Ornithogenic soils are typically acidic (pHs ranging from 3.9 to 5.1) due to the high concentrations of organic acids and ammonia [61]. Nitrate concentrations are much lower, with ranges of 0–130 µg g−1 previously reported on Marion Island [63]. Ornithogenic soils also harbor high moisture

4.3 Bacterial Diversity of Soils in the MDVs and Antarctic Peninsula |

51

content, with up to 30% water saturation by weight [62]. Despite the high nutrient and moisture status of these soils, the high percentage of soluble salts limits the growth of plants, lichens and mosses [62]. Fellfield soils occur mainly within more temperate Antarctic regions, such as the peninsula and surrounding subantarctic islands, for example Signy and Marion Islands. Fellfield soils are placed in two categories: (i) moist and nutrient rich, with a high silt content [64], (ii) dry and nutrient poor, containing high sand content [65]. The first class of fellfield soils contrasts substantially to the desiccated, mostly sandy soils of the MDVs [66]. For example, fellfield soils on Signy Island may contain as much as 20% (wt) of soil water content [66]. Maritime Antarctic fellfield soils are prone to leaching and, therefore, are much less saline than MDV mineral soils [64]. Cryptogams, which include mosses and lichens, provide a common but discontinuous vegetative distribution within fellfield soils [64]. However, cryptograms are not well anchored to the underlying soils and are, therefore, highly unstable habitats. Nevertheless, the presence of cryptogams in fellfield soils increases the abundance of key nutrients [24]. For example, within coastal Antarctic fellfield soils the soluble phosphorus, nitrate and ammonium concentrations range from 4–45 µg g−1 , 1–20 µg g−1 and 15–20 µg g−1 , respectively [34]. Fellfield soils therefore contain substantially higher nutrient and organic matter levels than the depauperate MDV mineral soils [34]. The Antarctic continent harbors a wide array of soil biotopes due to its nonhomogeneous structure and characteristics, as well as the presence of higher life forms such as plants and birds in some continental regions. Although the different Antarctic soil biotopes reflect the diverse nature of the continent, its diversity is also impacted by the presence of specialized cryptic or refuge niches [67–69].

4.3 Bacterial Diversity of Soils in the MDVs and Antarctic Peninsula Studies surveying microbial diversity within Antarctica were originally based on the determination of bacterial cell densities through ATP, lipid or DNA quantification [70], the culturing of active microorganisms [71] and microscopic analysis [72]. Microbial biomass detected within the nutrient rich ornithogenic and fellfield soils of the Peninsula are in the range of 107 –1010 prokaryotic cells g−1 [73, 74]. Surprisingly, microbial biomass counts within the MDVs are only slightly lower, with a range of 106 – 108 prokaryotic cells g−1 detected [70]. Microbial cell densities within Antarctic soils were positively correlated with soil water content and negatively correlated with salinity [75]. Culture dependent studies on Antarctic soils identified the presence of mostly aerobic heterotrophic microorganisms with limited anaerobic bacteria. The bacterial

52 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

phylotypic diversity was rather limited, consisting mainly of Actinobacteria and Firmicutes [76–81]. Culture independent phylogenetic and metagenomic techniques, which are based on the analysis of total community DNA extracted directly from environmental samples, avoid any bias induced by the requirement for microbial growth and, therefore, may provide truer estimates of microbial diversity [81–83]. Phylogenetic fingerprinting methods, such as terminal restriction fragment length polymorphism (TRFLP), autosomal ribosomal intergenic spacer analysis (ARISA) and denaturing gradient gel electrophoresis (DGGE) have provided estimates of the dominant members of microbial community structures within these regions [81, 84]. However, metagenomic sequencing, using either large insert libraries, shotgun or amplicon sequencing, identifies the “entire” microbial community composition within a specific sample [82, 83]. Taken together, these techniques have resulted in the detection of a much greater microbial diversity within Antarctic niches than originally predicted. However, it should be noted that even with the use of modern phylogenetic marker sequencing technologies, microbial taxa are typically only identified down to the genus level (in most cases) and that the true microbial diversity at species and strain levels within Antarctic niches is, therefore, still largely unclassified [85]. Interestingly, the large number of uncultured microbial representatives commonly detected in surveys of microbial diversity within Antarctica may also include novel species (particularly members of the family Actinobacteria) that may have important applications in biotechnology [24]. Overall, studies have shown that bacterial diversity in Antarctic terrestrial environments is highly heterogeneous, but with some phyla consistently maintained across many Antarctic soil environments [86–88]. Smith et al. (2006) used DGGE to analyze the microbial diversity of mineral soils from three different MDV sites. The samples were dominated by Actinobacteria, Acidobacteria, Cyanobacteria and Bacteroidetes, and included Verrucomicrobia, Chloroflexi, Alphaproteobacteria and Betaproteobacteria at lower abundances. Actinobacteria occurred ubiquitously in all samples, possibly due to the dispersal capabilities and high abundance of this phylum within soils (󳶳 Tab. 4.1) [79, 89–100]. A similar study on soils within the more northern (and drier) McKelvey Valley identified additional taxa such as Gemmatimonadetes and the desiccation tolerant Deinococcus–Thermus and Rubrobacter [87]. In contrast, the more nutrient rich soils of the Peninsula (including both vegetated and fellfield soils) are dominated by Proteobacteria (including representatives of the Alpha, Beta, Gamma and Delta Proteobacteria), with lower abundances of Actinobacteria and Bacteroidetes [39, 76, 88]. Other studies focused on the bacterial diversity of Antarctic soil biotopes have investigated the factors responsible for driving differences in community structure [50, 76, 101]. Lee et al. (2012) used a combination of pyrosequencing and DGGE to determine microbial community structure within soils from four geographically isolated MDVs [50]. Only a limited number of phylotypes were identified at each of the four sites (typically members of the Actinobacteria and Bacteroidetes), with much of the bacte-

4.3 Bacterial Diversity of Soils in the MDVs and Antarctic Peninsula | 53

rial diversity identified being specific to one or more sites. Regional differences were also observed from other MDV sites: for example, the usually dominant Acidobacteria were found to occur at very low abundances within the Miers Valley and at Battleship Promontory. These differences were found to be significantly driven by altitude (specifically, altitude related temperature) and by soil salt content. Studies on soil biotopes within the Antarctic Peninsula have shown similar community patterns [88, 101]. Yergeau et al. (2006) assessed the microbial diversity of soils along an environmental gradient within the Antarctic Peninsula, Falkland Island and Signy Island using DGGE [101]. This study showed that microbial abundance was significantly and positively influenced by vegetation related factors such as nitrogen and carbon, and soil water content. Microbial community structure was also significantly correlated with location and latitude, including specific factors such as mean temperature, nitrate and pH. These communities were influenced by the complex relationship between vegetation and latitude, where latitude had less of an effect in the presence of vegetation. Similarly, it has been shown using 16S rRNA gene amplicon sequencing that bacterial diversity declines with increasing latitude for fellfield but not vegetated soils within the Antarctic Peninsula [88]. Mineral soil bacterial community structure has also been shown to be markedly different from ornithogenic soils [58, 76]. Aislabie et al. (2008) used RFLP methods to analyze microbial diversity in four different mineral soils and one ornithogenic soil [76]. The mineral soils were found to contain similar bacterial phyla, dominated by Acidobacteria, Actinobacteria, Firmicutes, Cyanobacteria, Proteobacteria, Bacteroidetes and Deinococcus–Thermus. No difference in microbial diversity was found between soil taxonomic classifications of the mineral soils but was rather found according to physiochemical parameters, such as pH. The ornithogenic soils were found to contain an abundance of endospore formers such as Oceanobacillus, Clostridium and Sporosarcina, probably reflecting to the high number of Firmicutes found in the gut and fecal deposits of Antarctic penguins [58]. The microbial diversity within rhizosphere soils of two native vascular plants from the Antarctic Peninsula was recently assessed [58]. Surprisingly, in contrast to other peninsula soils [88, 101], the dominant bacterial phylotypes identified were the Firmicutes, Actinobacteria and Proteobacteria, with Acidobacteria, occurring rarely and at a low abundance. Firmicutes were also identified as the dominant phylum, while Proteobacterial diversity was comparatively low, in contrast to other vegetated and fellfield peninsula soils [88, 101]. The high abundance of anaerobic spore formers (such as the Firmicutes) may be due to the higher levels of moisture within the rhizosphere, or the adaptation of these communities to nutrient (e.g., carbon) limiting conditions during the winter [58]. This study highlights the importance of local environmental and physiochemical properties on bacterial community structure within Antarctic soil biotopes.

54 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

4.4 Cryptic Niches in Antarctic Environments The ice free regions of the Antarctic continent provide extensive expanses of exposed rocky substrate. The microbial colonization of rock substrates is a particular feature of these regions. Lithic associated microhabitats are referred to as lithobiontic niches, with their communities termed lithobionts [102]. Previous studies have shown that lithobionts [also referred to as soil rock surface communities (SRSCs)] are ubiquitously distributed in both hot and cold deserts [103–105]. In the most hyperarid regions, lithobionts are often the only visible forms of life (󳶳 Fig. 4.2a–d) and are thought to contribute significantly to the ecology of these regions [51, 68, 105]. The three major lithobiontic niches, which are based largely on the mode of colonization of the mineral substrate, are all prevalent in Antarctic ice free regions. Hypoliths (microbial assemblages found on the ventral surfaces of translucent rocks, mostly marble and quartz stones) are probably the most studied of the three niches. Epiliths (organisms populating the surface of stable rock substrata; the subcategory of chasmoliths inhabits cracks in rocks) occur on various igneous rock surfaces, while endoliths (communities colonizing the interior of rocks) are usually restricted to porous sandstones and weathered granitic rocks [67, 68]. In all three niches, micro-

(a)

(b)

(c)

(d)

Fig. 4.2: Examples of four lithobiont communities/cryptic soil niches, dominated by Cyanobacteria. (a) A hypolithon with the green biofilm layer, which is distinctive of Cyanobacteria dominated hypoliths. (b) An endolithon, which has been exposed, showing microbial colonization within the green under layer. (c) A cryptoendolith occurring along the crack within the rock, showing visible Cyanobacteria colonization (thin green line along the crack). (d) Endolithic colonization by Cyanobacteria.

4.4 Cryptic Niches in Antarctic Environments

|

55

bial colonization is limited by the availability of photosynthetically active radiation (PAR), which tends to favor the development of photoautotrophs [24, 69].

4.4.1 Hypoliths Hypolithic microbial communities (hypolithons) have been studied within several of the MDVs and are present wherever suitable mineral substrates (such as quartz pebbles) are available [87, 92, 97, 106]. While these communities are present at most altitudes, colonization of such substrates does not occur at high altitudes (such as University Valley; DA Cowan, personal observation) where little or no seasonal permafrost melt occurs. Hypolith communities may be highly similar to, or distinct from, the surrounding soil communities, depending on whether they occur in low or high altitude regions, respectively [87, 92]. Microclimate conditions occurring at different altitudes, such as variations in temperature and moisture availability, which decrease at higher altitudes, may account for these differences [106]. Where both open soil and hypolithic communities are found to be similar in composition, it has been suggested that hypoliths recruit microbial communities directly from the surrounding soil [107]. Interestingly, hypolithic communities show some variation in gross morphotypic structure: while most are physically (and visually) dominated by Cyanobacterial biofilms, a small proportion of quartz hypoliths support moss (Hennendiella spp.) dominated communities [106]. Hypoliths are thought to be the dominant autotrophic communities in some Antarctic terrestrial soil environments (i.e., those where suitable translucent mineral substrates are present in the desert pavement). They are probably the key primary producers in those Antarctic Dry Valleys that lack high productivity lake systems [97]. A number of recent studies have provided substantial insights into the compositions and functional diversity of hypolithic microbial communities [108–111]. A combination of microscopy and culture independent studies showed that Cyanobacteria, dominated by filamentous Oscillatorian morphotypes, were prevalent in MDV hypoliths [38, 112]. Microcoleus, Phormidium and Oscillatoria phylotypes were also recently identified in MDV hypoliths [111] using 16S rRNA gene pyrosequencing. In the Vestfold Hills, Oscillatorian Cyanobacterial morphologies were dominant, typically associated with Lyngbya/Phormidium/Plectonema groups, together with coccoid cells similar to Chroococcidiopsis [112]. Other dominant bacterial phyla identified in hypolithic communities include Actinobacteria, α and β Proteobacteria, Planctomycetes, Firmicutes, Acidobacteria and Verrumicrobia [87, 110, 111, 113]. The diversity of fungal phylotypes in Antarctic (particularly Dry Valley) soils is typically much lower than that of bacteria [114–116], and is dominated by Ascomycetes lineages [108, 109]. Members of the genera Acremonium, Stromatonectria and Verrucaria were most commonly identified [108]. Ascomycetes were initially reported as the

56 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

only fungal taxa present in hypolithic communities [97]. However, a recent study reported the presence of Basidiomycetes in hypoliths and soils [117] although they occur at low abundance. The low moisture availability in desert soils may explain the low fungal diversity [118]. Other lower eukaryotes, particularly protists, have been identified in Antarctic Miers Valley hypolithic communities [117]. The relative abundances of Amoebozoa and Cercozoa phylotypic signals were linked to the sample type (i.e., hypolith type) [106]. Interestingly, the presence of these protists appeared to be unique to the hypolithic environment, and these organisms have not been identified in nearby open soils. Clearly, their presence in this habitat has implications for the structure and functioning of food webs in Antarctic soils and requires further examination.

4.4.2 Epiliths In Antarctic regions, epilithic colonization is probably the least extensive of all rock associated habitats. However, studies of the microbial communities present on mineral surfaces from other (non-Antarctic) environments [119], particularly rock varnishes [120], suggest that Antarctic epilithic microbial communities may be more widespread and complex than previously considered. A possible role for shallow subsurface endolithic microbial populations in the genesis of Antarctic rock varnish layers has been proposed [121]. In Antarctic regions, surface rock communities are limited by the combination of extremely low temperatures, freeze–thaw cycles, katabatic wind episodes and high ultraviolet radiation levels [122]. However, in general very little is known regarding the microbiology of epiliths, in comparison to other lithobionts (endoliths and hypoliths) [67]. Early studies suggested that epilithic colonization is primarily associated with moss and lichen communities [123]. Both lichens and mosses synthesize a wide range of secondary metabolites, which may act as protectants against some environmental stressors (such as desiccation and UV damage), explaining their dominance in these niches [124, 125]. Moreover, epiliths are typically found where the rock substrata have access to moisture [103, 126]. As such, epilithic lichens are widespread across the coastal regions of Antarctica but decrease toward the interior [126, 127]. Recent studies indicate widespread prevalence of black meristematic fungi in the coastal northern and southern Victoria Land regions of Antarctica [128]. Black fungi may be crucial in the hydration or protection of photobionts by dissipating excess sunlight [129]. In contrast, epiliths from the Princess Elizabeth Land and Mawson Rock regions are dominated by Chroococcidiopsis spp. [130, 131]. Chroococcidiopsis are dominant in both hypolithic and endolithic niches and may support the epilithic “genesis” theory [121]. A comprehensive analysis assessing the dominance of other bacterial phyla in epiliths may validate this proposal.

4.4 Cryptic Niches in Antarctic Environments

| 57

4.4.3 Endoliths Endolithic microbial communities are defined as those existing inside lithic strata, but are classified into various subniches [102, 132–134]. Chasmoendoliths (also known as chasmoliths) are found in interstitial cracks and fissures, while cryptoendoliths are found in the pores between mineral grains [102, 113, 135, 136]. Like all lithobionts, endoliths are dominated by Cyanobacteria [67, 68, 87, 136–138]. Early microscopic analyses of endoliths suggested that the Cyanobacteria co-existed with lichens [91] (mostly Gloeocapsa, Hormathonema–Gloeocapsa and Chroococcidiopsis communities). More recent molecular analyses have largely concurred with these studies [126, 139]. Endolithic habitats may impart a degree of species selection; for example, a highly novel cyanobacterium, a Chloroglea sp., was detected in endoliths from Alexander Island [133], although a range of different Cyanobacterial phylotypes have been identified in various studies on endolithic microbial communities. Plectonema species have been identified in 16S rRNA gene clone libraries generated from Dry Valley cryptoendolithic samples [89]. Studies within the Taylor Valley have identified Nostoc, Cyanothece, and Chroococcidiopsis species in endoliths [140–142]. Endoliths in McKelvey Valley have been shown to be dominated by Nostocales and Chroococcidiopsis-like phylotypes [87]. The drivers for selection of the different cyanobacterial phylotypes in different endolithic habits are not understood, although community structures have been shown to vary along a lateral transect within the Miers Valley, which is probably a result of the different microclimatic conditions of north facing (warmer and wetter) and south facing (colder and drier) slopes [143]. Although all samples were dominated by Leptolyngbya, the north facing slopes contained the highest microbial diversity with a relatively high abundance of Synechococcus-like phylotypes while, in contrast, the south facing slopes contained Chroococcidiopsis-like phylotypes [143]. It is tempting to speculate that resistance to extremes, particularly extremes of desiccation, is a factor in the selection of the dominant photoautotroph. Cyanobacteria in endoliths form consortia with heterotrophic phyla, which vary in taxonomic composition depending on their location [72]. MDV cryptoendolithic communities, analyzed by microscopy, consisted of heterotrophic assemblages consisting primarily of Alphaproteobacteria (some members of which are potentially capable of photosynthesis) and Deinococcus–Thermus phylotypes, a group of organisms with known resistance to desiccation stress. Unlike open soil populations, Actinobacteria occur at a comparatively low abundance [89]. In contrast, Acidobacteria and Actinobacteria were the dominant endolithic heterotrophs in samples from the north facing slopes of the Miers Valley, whereas Deinococcus–Thermus dominated the colder south facing slopes [143]. Chasmoliths and endoliths from the McKelvey Valley contained high abundances of Bacteroidetes, Actinobacteria, and Gammaproteobacteria, with Acidobacteria, Deinococcus–Thermus and Alphaproteobacteria at lower abundances [87].

58 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats Hypolith

(a) Cyanobacteria Bacteriodetes Actinobacteria

Endolith

(b) Acidobacteria Proteobacteria Verrucomicrobia

Cyanobacteria Bacteriodetes Actinobacteria

Open soil

(c) Acidobacteria Proteobacteria DeinococcusThermus

Cyanobacteria Bacteriodetes Actinobacteria Acidobacteria Proteobacteria

DeinococcusThermus Chloroflexi Gemmatimonadetes Verrucomicrobia

Fig. 4.3: (a) Phylum level classification of bacterial diversity from Antarctic hypolithic communities. Data is based on the percentage of 16S rRNA gene sequences and tRFLP signatures identified for each phylum [87, 97], where data was obtained from Pointing et al. (2009) and Khan et al. (2011). (b) Phylum level classification of bacterial diversity from Antarctic endolithic communities. Data is based on the percentage of phylum abundances identified from tRFLP fingerprints [87] and was obtained from Pointing et al. (2009). (c) Phylum level classification of bacterial diversity from Antarctic MDV mineral soils. Data is based on the number of 16S rRNA gene sequences present following analysis from MDV soil samples [38] as determined by Cary et al. (2010).

In comparison to hypoliths and open soils, endoliths appear to harbor higher bacterial diversity (󳶳 Fig. 4.3) [87]. In general, all lithobiont microbial communities are more similar to each other than to those of open soils [87, 113, 143], although significant differences in microbial community structures exists between endolithic and hypolithic communities [87, 142]. Lithobionts are Cyanobacteria dominated, whereas open soil microbial communities consist of a majority of heterotrophic bacterial phylotypes (󳶳 Fig. 4.3) [87, 143]. Differences between endoliths and hypoliths have been shown within the McKelvey Valley, where the dominant phylotypes were shown to be Chroococcidiopsis and Leptolyngbya, respectively [87]. Although both endoliths and hypoliths are dominated by cyanobacteria, endoliths contain a higher diversity of heterotrophic microorganisms relative to hypoliths [87]. Although multiple abiotic factors may drive the differences in bacterial community structure in different Antarctic soil biotopes [50, 58, 88], differences are also observed when comparing open soil and cryptic niches [87]. The differences seen between refuge niches such as hypoliths and endoliths and the open soil are partly due to the protection that refuge niches provide from environmental stressors [51], and the increased availability of moisture and nutrients within xeric, nutrient limiting habitats [87]. These factors, and the environmental conditions occurring at different altitudes and latitudes, have been shown to drive the differences in microbial community structures between cryptic niches and the open soil [87].

4.6 Viruses in Antarctic Edaphic Ecosystems |

59

4.5 Biogeochemical Cycling in Antarctic Environments Antarctic soils are generally oligotrophic and have generally low nutrient status in comparison to those from more temperate biomes [50]. Nonetheless, these soils demonstrate a high capacity for functional processes [108, 109, 144–146]. For example, soils in the Sør Rodane Mountains, located in the Dronning Maud Land (DML) region of Antarctica, harbored both autotrophic and phototrophic bacteria [146]. Soils in this region contained a high diversity of puf M genes (which encode a subunit of the type 2 photochemical reaction center found in anoxygenic phototrophic bacteria) and bchL/chlL sequences (genes implicated in bacterio-chlorophyll synthesis). The majority of puf M sequences were related to those previously found in Proteobacteria, while the origin of the bchL/chlL was linked to Cyanobacteria. Another study based on clone libraries of the large subunit of ribulose-1,5-biphosphate carboxylase/oxygenase (RuBisCO) genes (cbbL, cbbM) and dinitrogenase-reductase (nif H) genes also identified Cyanobacteria (mostly Nostocales lineages) as the primary photoautotrophs in DML soils [146]. Surprisingly, these soils lack signatures for alternate energy acquiring processes, such as rhodopsin genes, suggesting that Cyanobacteria in Antarctic regions may have evolved to efficiently cycle C and N. In contrast to soils in the DML region, biogeochemical cycling in MDV soils is apparently driven by microbial communities linked to cryptic niche habitats, as indicated by recent GeoChip based analyses [109, 111, 147]. These studies have indicated that while cryptic niches have higher biomass, with autotrophs being more diverse in these systems, open soil communities are more diverse in terms of diazotrophic guilds [147]. In addition, both soils and cryptic niches were highly abundant in functional genes linked to Archaea (mostly Halobacteria). Interestingly, most genes implicated in metabolic pathways linked to carbon transformations in soils were attributed to fungi [147].

4.6 Viruses in Antarctic Edaphic Ecosystems Recent metagenomic studies have demonstrated the presence of high levels of viral diversity in a range of environments [148–151]. In Antarctic environments, the majority of studies have focused on viruses found in freshwater ponds and lake ecosystems [152–156]. These studies have provided key insights into the influence of environmental extremes on viral diversity, and the role of viruses in biogeochemical cycles. For instance, a study by Yau and colleagues (2010) highlighted virophages as crucial regulators of host–virus interactions, a finding that has consequences for carbon flux dynamics in lake ecosystems [154]. Surprisingly, comparatively little is known of the role of viruses in Antarctic soil ecosystems. Given the high amount of carbon stored in these soils, the interactions between viruses and bacteria may be crucial feedback

60 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

mechanisms on carbon cycling. The diversity and ecology of viruses in Antarctic soils have been reviewed recently [157]. Isolation methods, and analyses using electron microscopy, have shown that Antarctic soils are dominated by tailed viruses (mostly belonging to the family Myoviridae) and spherical viruses (mostly of the family Levividae) [158]. Direct counts using epifluorescence of extractable and extracellular virus particles suggests that Antarctic soils may have the highest recorded virus-to-bacteria ratios [159]. A study by Williamson and colleagues showed that the abundance of viruses increased relative to bacteria as water and organic content decreased [159]. While the impacts of climate change and the melting of previously buried ice has not been assessed for viral communities, this finding does suggests enhanced roles for viral communities as a consequence of these perturbations.

4.7 Conclusions and Perspectives In Antarctic microbiology, two of the revelations of the past two decades are that bacterial diversity of Antarctic edaphic niches is much greater than previously thought, and that specialized cryptic niche communities in cold desert soils may play an important role in ecosystem processes [24] (󳶳 Tab. 4.1). The presence of substantial populations of Cyanobacteria, Chloroflexi and Proteobacteria suggests that these organisms contribute to primary productivity in depauperate Antarctica desert soils [87, 106], and that the presence of diverse heterotrophic organisms (including both bacteria and fungi) along with viruses [160], macroinvertebrate grazers [161] and predators [162] suggests the presence of a fully functional trophic hierarchy [24]. However, the global microbial community is familiar with the concept that predicting organismal or community functions from taxonomic identity is extremely weak, providing, at best, limited but testable information on functional processes [163]. An assessment of the diversity (and frequency) of key functional genes within a sample, and relating such data to taxonomic identity, is a step closer to understanding community function [109], but ultimately should be verified through the determination of real process rates. Despite the recent surge of research activity and publications on the structure, and to some extent, function of Antarctic edaphic microbial communities, we lack a comprehensive understanding of the finer details: the nature of community interactions in food web structures, the interactive roles of hosts and predators, and the balance between abiotic and biotic factors in controlling community function. Such understanding is important for many reasons, not least understanding how changing climate conditions may impact microbial communities in Antarctic terrestrial environments. It is well known that cyanobacteria are essential mediators of biogeochemical processes in many habitats, and it is argued that their role in Antarctic soils may be even

4.7 Conclusions and Perspectives |

61

Table 4.1: Microbial diversity from various Antarctic niches. Domain Archaea

Bacteria

Identity

Niche Soil Epilith

Endolith

Archaea Crenoarcheota Euryarchaeota

Hypolith *

Acidobacteria Actinobacteria Arthrobacter Brevibacterium Demetria Gordonia Janibacter Kocuria Lapillicoccus Leifsonia Marisediminicola Micromonospora Mycobacterium Nocardiodetes spp. Patulibacter Rhodococcus Unclass. Intrasporangiaceae Unclass. Microbacteria Uncultured Pseudonocardia

* * * * * * * * *

*

* * *

*

* * *

*

Aquificae Bacteroidetes Unclass. Flexibacteraceae Unclass. Saprospiraceae Unclass. Sphingobacteriales Cyanobacteria Acaryochloris spp. Anabaena spp. Chroococcidiopsis spp. Cylindrospermum spp. Gloeocapsa spp. Hormathonema spp. Leptolyngbya spp. Lyngbya spp. Microcoleus spp. Nostoc spp. Oscillatoria spp. Phormidium spp. Plectonema spp. Synechococcus spp.

* * *

* * *

* * * * * * *

* * *

* *

* * *

*

* * * *

* *

62 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

Table 4.1 (cont.): Microbial diversity from various Antarctic niches. Domain

Identity

Niche Soil Epilith

Endolith

Hypolith

Chloroflexi

*

*

*

Deinococcus/Thermus Deinococcus

*

Firmicutes Unclass. Bacillaceae Unclass. Clostridiales Staphylococcus Sporosarcina Trichoccus Erysipelothrix Atopostipes

* * * * * * *

Plactomycetes

Fungi

Proteobacteria Alkanindiges Dokdonella Lysobacter Psychrobacter Rhodanobacter Lysobacter Unclass. Xanthamonadeaceae Unclass. Pseudomonadaceae Unclass. Rhizobiales

* * * * * * * * *

Verrumicrobia

*

Ascomycota Alternaria Antarctomyces Cadophora spp. Candida spp. Cladosporium Debaryomyces Geomyces spp. Leuconeurospora Nadsonia Nectriaceae Onygenales Penicillium Phaeosphaeria Phoma Pseudeurotium Thelebolus Thielavia Theobolaceae

* * * * * * * * * * * * * * * * * * *

*

*

*

4.7 Conclusions and Perspectives |

63

Table 4.1 (cont.): Microbial diversity from various Antarctic niches. Domain

Identity

Niche Soil Epilith

Basidiomycota Bensingtonia Bulleromyces Cryptococcus spp. Leucosporidiella Rhodotorula

* * * * *

Zygomycota Mortierellaceae Mortierella

* *

Endolith

Hypolith

Data was compiled from several resources [38, 48, 69, 76, 86, 87, 89, 90, 92–100].

more critical in the absence of higher eukaryotic phototrophs. Modern metagenomics provides a set of tools that, at least, give ready access to information of an organism’s potential capacity to respond to change. For instance, a cyanobacterial genome sequence provides some insight into the organism’s stress response capacity, which can be verified using ex situ culture dependent stress experiments. However, the technical challenges associated with the isolation of slow growing cold active cyanobacterial cultures have posed a considerable challenge [164, 165]. A novel approach to (partially) overcoming this challenge may be to sequence “mixed” cyanobacterial cultures and implement genome binning approaches, which are increasingly used in the field of environmental metagenomics [166–168]. Metagenomic binning approaches have yielded insights on the ecology of other extreme habitats [169] and have the capacity to contribute a greater understanding of community interactions in Antarctic soils. A note of caution, relating specifically to issues of “legacy DNA”, must be added. Conditions in the driest and coldest soils of the Antarctic continent, particularly the McMurdo Dry Valleys, are not inconsistent with those used routinely by microbiologists for the preservation of biological material: i.e., freeze drying [170]. It is, therefore, instructive to contemplate the impacts on metagenomic DNA dependent phylotypic surveys of these extreme habitats due to the presence of a legacy of dead cells and even residual genomic DNA [171]. A recent study by Fierer’s group [172] suggests that legacy (relic) DNA forms a significant proportion of metagenomic DNA extracted from temperate soils, suggesting that at least some of the published surveys of Antarctic soil microbial diversity might reflect both historical and extant community compositions. It is well accepted by the microbial ecology community that RNA-based phylogenetic surveys, which assess the “functioning” fraction of the microbial community, are more reliable and informative. However, the extreme technical difficulties of extracting usable quantities of RNA from low biomass, low activity environments such as the cold desert soils of Antarctica makes this an objective rather than a current reality.

64 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

Acknowledgment: The authors wish to thank the University of Pretoria, Antarctica New Zealand, and the South African National Research Foundation (SANAP program) for supporting field and laboratory research programs.

References [1] [2]

[3] [4]

[5] [6] [7]

[8]

[9] [10] [11]

[12] [13]

[14]

[15]

[16]

Glikson A. Cenozoic mean greenhouse gases and temperature changes with reference to the Anthropocene. Glob Chang Biol 2016, 22:3843–3858. Flato G, Marotzke J, Abiodun B, et al. Evaluation of Climate Models. In: Stocker TF, Qin D, Plattner GK, et al. eds. Climate Change 2013: The physical science basis. Contribution of Working Group I to the Fifth Assessment Report of the Intergovernmental Panel on Climate Change. Cambridge, Cambridge University Press, 2013, 741–866. Vaughan DG, Marshall GJ, Connolley WM, et al. Recent rapid regional climate warming on the Antarctic Peninsula. Clim Change 2003, 60:243–74. Christensen JH, Kanikicharla KK, Marshall G, Turner J. Climate phenomena and their relevance for future regional climate change. In: Pauline M, ed. Climate Change 2013: The physical science basis. Contribution of Working Group I to the fifth Assessment of the Intergovernmental Panel on Climate Change. Cambridge, Cambridge University Press, 2013, 1217–1308. Spaulding SA. Antarctic Lakes. Arct, Antarc, and Alp Res 2015, 47:401–2. Cavicchioli R. Microbial ecology of Antarctic aquatic systems. Nature Rev Microbiol 2015, 13:691–706. Gooseff MN, McKnight DM, Welch KA, Lyons WB. Stream biogeochemical and suspended sediment responses to permafrost degradation in stream banks in Taylor Valley, Antarctica. Biogeosciences 2016, 13:1723. Stomeo F, Makhalanyane TP, Valverde A, et al. Abiotic factors influence microbial diversity in permanently cold soil horizons of a maritime-associated Antarctic Dry Valley. FEMS Microbiol Ecol 2012, 82:326–40. Christner BC, Priscu JC, Achberger AM, et al. A microbial ecosystem beneath the West Antarctic ice sheet. Nature 2014, 512:310–3. Boetius A, Anesio AM, Deming JW, Mikucki JA, Rapp JZ. Microbial ecology of the cryosphere: sea ice and glacial habitats. Nature Rev Microbiol 2015, 13:677–90. Kohler TJ, Van Horn DJ, Darling JP, Takacs-Vesbach CD, McKnight DM. Nutrient treatments alter microbial mat colonization in two glacial meltwater streams from the McMurdo Dry Valleys, Antarctica. FEMS Microbiol Ecol 2016, 92:fiw049. Stanish LF, O’Neill SP, Gonzalez A, et al. Bacteria and diatom co-occurrence patterns in microbial mats from polar desert streams. Environ Microbiol 2013, 15:1115–31. Archer SD, McDonald IR, Herbold CW, Cary SC. Characterisation of bacterioplankton communities in the meltwater ponds of Bratina Island, Victoria Land, Antarctica. FEMS Microbiol Ecol 2014, 89:451–64. Colesie C, Allan Green TG, Haferkamp I, Budel B. Habitat stress initiates changes in composition, CO2 gas exchange and C-allocation as life traits in biological soil crusts. ISME J 2014, 8:2104–15. Caruso T, Chan Y, Lacap DC, Lau MC, McKay CP, Pointing SB. Stochastic and deterministic processes interact in the assembly of desert microbial communities on a global scale. ISME J 2011, 5:1406–13. Makhalanyane TP, Van Goethem MW, Cowan DA. Microbial diversity and functional capacity in polar soils. Curr Opin Biotechnol 2016, 38:159–66.

References | 65

[17] [18] [19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29] [30] [31] [32] [33] [34] [35] [36] [37] [38] [39] [40]

Zhang X, Johnston ER, Li L, Konstantinidis KT, Han X. Experimental warming reveals positive feedbacks to climate change in the Eurasian Steppe. ISME J 2017, 11:885–895. Scharlemann JP, Tanner EV, Hiederer R, Kapos V. Global soil carbon: understanding and managing the largest terrestrial carbon pool. Carbon Manag 2014, 5:81–91. Schuur EA, Bockheim J, Canadell JG, et al. Vulnerability of permafrost carbon to climate change: Implications for the global carbon cycle. BioScience 2008, 58:701–14. Walther G-R, Post E, Convey P, et al. Ecological responses to recent climate change. Nature 2002, 416:389–95. Arneth A, Harrison SP, Zaehle S, et al. Terrestrial biogeochemical feedbacks in the climate system. Nat Geosci 2010, 3:525–32. Convey P, Bindschadler R, Di Prisco G, et al. Antarctic climate change and the environment. Antarct Sci 2009, 21:541–63. Convey P, Chown SL, Clarke A, et al. The spatial structure of Antarctic biodiversity. Ecol Monogr 2014, 84:203–44. Cowan DA, Makhalanyane TP, Dennis PG, Hopkins DW. Microbial ecology and biogeochemistry of continental Antarctic soils. Front Microbiol 2014, 5:154. Cowan DA. Antarctic Terrestrial Microbiology: Physical and Biological Properties of Antarctic Soils. Heidelberg, Berlin, Springer-Verlag, 2014. Jansson JK, Taş N. The microbial ecology of permafrost. Nature Rev Microbiol 2014, 12:414–25. Ugolini FC, Bockheim JG. Antarctic soils and soil formation in a changing environment: a review. Geoderma 2008, 144:1–8. Ugolini F. Soil investigations in Lower Wright Valley, Antarctica. Proceedings of an International Conference on Permafrost 1963; 1966, 55–61. Ugolini F. A study of pedogenic processes in Antarctica: Final report to the National Science Foundation. New Brunswick, NJ, Rutgers University, 1964. Ugolini FC, Bull C. Soil development and glacial events in Antarctica, Ohio State University, Institute of Polar Studies, 1965. Ugolini F, Starkey R. Soils and micro-organisms from Mount Erebus, Antarctica. Nature 1966, 211:440–441. Tedrow J, Ugolini F. Antarctic soils. In: Tedrow JC, ed. Antarctic soils and soil forming processes. Washington DC, American Geophysical Union, 1966, 161–77. Campbell I, Claridge G. A classification of frigic soils-the zonal soils of the Antarctic continent. Soil Sci 1969, 107:75–85. Ugolini FC, Anderson DM. Ionic migration and weathering in frozen Antarctic soils. Soil Sci 1973, 115:461–70. Jackson M, Lee S, Ugolini F, Helmke P. Age and uranium content of soil micas from Antarctica by the fission particle track replica method. Soil Sci 1977, 123:241–8. Bockheim J. Properties of a chronosequence of ultraxerous soils in the Trans-Antarctic Mountains. Geoderma 1982, 28:239–55. Horowitz N, Cameron RE, Hubbard JS. Microbiology of the dry valleys of Antarctica. Science 1972, 176:242–5. Cary SC, McDonald IR, Barrett JE, Cowan DA. On the rocks: the microbiology of Antarctic Dry Valley soils. Nat Rev Micro 2010, 8:129–38. Aislabie JM, Chhour K-L, Saul DJ, et al. Dominant bacteria in soils of Marble Point and Wright Valley, Victoria Land, Antarctica. Soil Biol and Biochem 2006, 38:3041–56. Barrett JE, Virginia RA, Wall DH, Adams BJ. Decline in a dominant invertebrate species contributes to altered carbon cycling in a low-diversity soil ecosystem. Glob Chang Biol 2008, 14:1734–44.

66 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

[41]

[42] [43] [44] [45] [46]

[47]

[48] [49]

[50] [51]

[52]

[53] [54] [55] [56]

[57]

[58] [59] [60]

Witherow RA, Lyons WB, Bertler NA, et al. The aeolian flux of calcium, chloride and nitrate to the McMurdo Dry Valleys landscape: evidence from snow pit analysis. Antarct Sci 2006, 18:497–505. Nylen TH, Fountain AG, Doran PT. Climatology of katabatic winds in the McMurdo Dry Valleys, Southern Victoria Land, Antarctica. J Geophys Res Atmos 2004, 109:D03114. Doran PT, McKay CP, Fountain AG, et al. Hydrologic response to extreme warm and cold summers in the McMurdo Dry Valleys, East Antarctica. Antarct Sci 2008, 20:499–509. Doran PT, Priscu JC, Lyons WB, et al. Antarctic climate cooling and terrestrial ecosystem response. Nature 2002, 415:517–20. Barrett J, Virginia R, Wall D, et al. Persistent effects of a discrete warming event on a polar desert ecosystem. Glob Chang Biol 2008, 14:2249–61. Niederberger TD, Sohm JA, Tirindelli J, et al. Diverse and highly active diazotrophic assemblages inhabit ephemerally wetted soils of the Antarctic Dry Valleys. FEMS Microbiol Ecol 2012, 82:376–90. Simmons B, Wall D, Adams B, Ayres E, Barrett J, Virginia R. Long-term experimental warming reduces soil nematode populations in the McMurdo Dry Valleys, Antarctica. Soil Biol and Biochem 2009, 41:2052–60. Cowan DA, Ah Tow L. Endangered antarctic environments. Annu Rev Microbiol 2004, 58:649–90. Toner JD, Sletten RS, Prentice ML. Soluble salt accumulations in Taylor Valley, Antarctica: Implications for paleolakes and Ross Sea Ice Sheet dynamics. J Geophys Res: Earth Surf 2013, 118:198–215. Lee CK, Barbier BA, Bottos EM, McDonald IR, Cary SC. The inter-valley soil comparative survey: the ecology of Dry Valley edaphic microbial communities. ISME J 2012, 6:1046–57. Makhalanyane TP, Valverde A, Velázquez D, et al. Ecology and biogeochemistry of cyanobacteria in soils, permafrost, aquatic and cryptic polar habitats. Biodivers Conserv 2015, 24:1–22. Matsumoto G, Chikazawa K, Murayama H, Torii T, Fukushima H, Hanya T. Distribution and correlation of total organic carbon and mercury in Antarctic dry valley soils, sediments and organisms. Geochem J 1983, 17:241–6. Bockheim JG, Ugolini FC. A review of pedogenic zonation in well-drained soils of the southern circumpolar region. Quat Res 1990, 34:47–66. Bockheim J, McLeod M. Soil distribution in the McMurdo Dry Valleys, Antarctica. Geoderma 2008, 144:43–9. Hopkins D, Sparrow A, Elberling B, et al. Carbon, nitrogen and temperature controls on microbial activity in soils from an Antarctic dry valley. Soil Biol and Biochem 2006, 38:3130–40. Otero X, Fernández S, de Pablo Hernandez M, Nizoli E, Quesada A. Plant communities as a key factor in biogeochemical processes involving micronutrients (Fe, Mn, Co, and Cu) in Antarctic soils (Byers Peninsula, maritime Antarctica). Geoderma 2013, 195:145–54. Bokhorst S, Huiskes A, Convey P, Van Bodegom P, Aerts R. Climate change effects on soil arthropod communities from the Falkland Islands and the Maritime Antarctic. Soil Biol and Biochem 2008, 40:1547–56. Teixeira LC, Peixoto RS, Cury JC, et al. Bacterial diversity in rhizosphere soil from Antarctic vascular plants of Admiralty Bay, maritime Antarctica. ISME J 2010, 4:989–1001. Niederberger TD, McDonald IR, Hacker AL, et al. Microbial community composition in soils of Northern Victoria Land, Antarctica. Environ Microbiol 2008, 10:1713–24. Blume H, Bölter M. Soils and soil scapes. In: Beyer L, Bölter M (eds). Geoecology of Antarctic Ice-Free Coastal Landscapes. Heidelberg, Berlin, Springer-Verlag, 2002, 91–113.

References | 67

[61] [62] [63]

[64] [65] [66]

[67]

[68] [69] [70] [71] [72] [73] [74] [75] [76] [77]

[78] [79] [80]

[81]

Schaefer CEGR, Pereira C, Torres T, et al. Soils and landforms at Hope Bay, Antarctic Peninsula: formation, classification, distribution, and relationships. Soil Sci Soc Am J 2015, 79:175–84. Speir T, Cowling J. Ornithogenic soils of the Cape Bird adelie penguin rookeries, Antarctica. Polar Biol 1984, 2:199–205. Sanyika TW, Stafford W, Cowan DA. The soil and plant determinants of community structures of the dominant actinobacteria in Marion Island terrestrial habitats, Sub-Antarctica. Polar Biol 2012, 35:1129–41. Wynn-Williams DD. Ecological aspects of Antarctic microbiology. In: Marshall KC, ed. Advances in microbial ecology. NY, Springer US, 1990, 71–146. Block W, Lewis Smith R, Kennedy A. Strategies of survival and resource exploitation in the Antarctic fellfield ecosystem. Biol Rev 2009, 84:449–84. Yergeau E. Fell-Field Soil Microbiology. In: Cowan D, ed. Antarctic Terrestrial Microbiology: Physical and Biological Properties of Antarctic Soils. Heidelberg, Berlin, Springer-Verlag, 2014, 115–29. Makhalanyane TP, Pointing SB, Cowan DA. Lithobionts: Cryptic and Refuge Niches. In: Cowan D, ed. Antarctic Terrestrial Microbiology: Physical and Biological Properties of Antarctic Soils. Heidelberg, Berlin, Springer-Verlag, 2014, 163–79. Pointing SB. Hypolithic Communities. In: Weber B, Büdel B, Belnap J (eds). Biological Soil Crusts: An Organizing Principle in Drylands. Springer International Publishing 2016, 199–213. Chan Y, Lacap DC, Lau MC, et al. Hypolithic microbial communities: between a rock and a hard place. Environm Microbiol 2012, 14:2272–82. Cowan D, Russell N, Mamais A, Sheppard D. Antarctic Dry Valley mineral soils contain unexpectedly high levels of microbial biomass. Extremophiles 2002, 6:431–6. Vishniac H. The microbiology of Antarctic soils. In: Friedmann EL, ed. Antarctic microbiology. NY, Wiley-Liss, 1993, 297–341. de los Ríos A, Wierzchos J, Sancho LG, Ascaso C. Exploring the physiological state of continental Antarctic endolithic microorganisms by microscopy. FEMS Microbiol Ecol 2004, 50:143–52. Ramsay AJ, Stannard RE. Numbers and viability of bacteria in ornithogenic soils of Antarctica. Polar Biol 1986, 5:195–8. French D, Smith V. Bacterial populations in soils of a subantarctic island. Polar Biol 1986, 6:75–82. Cameron RE, King J, David CN. Soil microbial and ecological studies in Southern Victoria Land. Antarct J US 1968, 3:121–3. Aislabie JM, Jordan S, Barker GM. Relation between soil classification and bacterial diversity in soils of the Ross Sea region, Antarctica. Geoderma 2008, 144:9–20. Giudice AL, Brilli M, Bruni V, De Domenico M, Fani R, Michaud L. Bacterium–bacterium inhibitory interactions among psychrotrophic bacteria isolated from Antarctic seawater (Terra Nova Bay, Ross Sea). FEMS Microbiol Ecol 2007, 60:383–96. Nicolaus B, Marsiglia F, Esposito E, et al. Isolation of five strains of thermophilic eubacteria in Antarctica. Polar Biol 1991, 11:425–9. Babalola OO, Kirby BM, Le Roes-Hill M, et al. Phylogenetic analysis of Actinobacterial populations associated with Antarctic Dry Valley mineral soils. Environ Microbiol 2009, 11:566–76. Bottos EM, Scarrow JW, Archer SD, McDonald IR, Cary SC. Bacterial community structures of Antarctic soils. In: Cowan D, ed. Antarctic Terrestrial Microbiology: Physical and Biological Properties of Antarctic Soils. Heidelberg, Berlin, Springer-Verlag, 2014, 9–33. Kirk JL, Beaudette LA, Hart M, et al. Methods of studying soil microbial diversity. J Microbiol Methods 2004, 58:169–88.

68 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

[82] Zhou J, He Z, Yang Y, Deng Y, Tringe SG, Alvarez-Cohen L. High-throughput metagenomic technologies for complex microbial community analysis: open and closed formats. mBio 2015, 6:e02288–14. [83] Thomas T, Gilbert J, Meyer F. Metagenomics–a guide from sampling to data analysis. Microb Inform Exp 2012, 2:3. [84] Tytgat B, Verleyen E, Obbels D, et al. Bacterial diversity assessment in Antarctic terrestrial and aquatic microbial mats: a comparison between bidirectional pyrosequencing and cultivation. PloS One 2014, 9:e97564. [85] Pearce DA, Newsham KK, Thorne MA, et al. Metagenomic analysis of a southern maritime antarctic soil. Front Microbiol 2012, 3:403. [86] Smith JJ, Tow LA, Stafford W, Cary C, Cowan DA. Bacterial diversity in three different Antarctic cold desert mineral soils. Microb Ecol 2006, 51:413–21. [87] Pointing SB, Chan Y, Lacap DC, Lau MC, Jurgens JA, Farrell RL. Highly specialized microbial diversity in hyper-arid polar desert. Proc Natl Acad Sci USA 2009, 106:19964–9. [88] Yergeau E, Newsham KK, Pearce DA, Kowalchuk GA. Patterns of bacterial diversity across a range of Antarctic terrestrial habitats. Environ Microbiol 2007, 9:2670–82. [89] de le Torre J, Goebel BM, Friedmann EI, Pace NR. Microbial diversity of cryptoendolithic communities from the McMurdo Dry Valleys, Antarctica. Appl Environ Microbiol 2003, 69:3858–67. [90] de Scally S, Makhalanyane T, Frossard A, Hogg I, Cowan D. Antarctic microbial communities are functionally redundant, adapted and resistant to short term temperature perturbations. Soil Biol and Biochem 2016, 103:160–70. [91] Friedmann EI, Hua M, Ocampo-Friedmann R. Cryptoendolithic lichen and cyanobacterial communities of the Ross Desert, Antarctica. Polarforschung 1988, 58:251–9. [92] Wood SA, Rueckert A, Cowan DA, Cary SC. Sources of edaphic cyanobacterial diversity in the Dry Valleys of Eastern Antarctica. ISME J 2008, 2:308–20. [93] Wood SA, Mountfort D, Selwood AI, Holland PT, Puddick J, Cary SC. Widespread distribution and identification of eight novel microcystins in Antarctic cyanobacterial mats. Appl Environ Microbiol 2008, 74:7243–51. [94] Bahl J, Lau MCY, Smith GJD, et al. Ancient origins determine global biogeography of hot and cold desert cyanobacteria. Nature Commun 2011, 2:163. [95] Cowan DA, Sohm JA, Makhalanyane TP, et al. Hypolithic communities: important nitrogen sources in Antarctic desert soils. Environ Microbiol Rep 2011, 3:581–6. [96] Taton A, Grubisic S, Brambilla E, De Wit R, Wilmotte A. Cyanobacterial diversity in natural and artificial microbial mats of Lake Fryxell (McMurdo Dry Valleys, Antarctica): a morphological and molecular approach. Appl Environ Microbiol 2003, 69:5157–69. [97] Khan N, Tuffin M, Stafford W, et al. Hypolithic microbial communities of quartz rocks from Miers Valley, McMurdo Dry Valleys, Antarctica. Polar Biol 2011, 34:1657–68. [98] Wong FK, Lacap DC, Lau MC, Aitchison JC, Cowan DA, Pointing SB. Hypolithic microbial community of quartz pavement in the high-altitude tundra of central Tibet. Microb Ecol 2010, 60:730–9. [99] Jungblut AD, Hawes I, Mountfort D, et al. Diversity within cyanobacterial mat communities in variable salinity meltwater ponds of McMurdo ice shelf, Antarctica. Environ Microbiol 2005, 7:519–29. [100] Cowan DA, Pointing SB, Stevens MI, Cary SC, Stomeo F, Tuffin IM. Distribution and abiotic influences on hypolithic microbial communities in an Antarctic Dry Valley. Polar Biol 2011, 34:307–11.

References | 69

[101] Yergeau E, Bokhorst S, Huiskes AH, Boschker HT, Aerts R, Kowalchuk GA. Size and structure of bacterial, fungal and nematode communities along an Antarctic environmental gradient. FEMS Microbiol Ecol 2006, 59:436–51. [102] Golubic S, Friedmann I, Schneider J. The lithobiontic ecological niche, with special reference to microorganisms. J Sediment Res 1981, 51:475–8. [103] Pointing SB, Belnap J. Microbial colonization and controls in dryland systems. Nature Rev Microbiol 2012, 10:551–62. [104] Pointing SB, Belnap J. Disturbance to desert soil ecosystems contributes to dust-mediated impacts at regional scales. Biodivers Conserv 2014, 23:1659–67. [105] Makhalanyane TP, Valverde A, Gunnigle E, Frossard A, Ramond JB, Cowan DA. Microbial ecology of hot desert edaphic systems. FEMS Microbiol Rev 2015, 39:203–21. [106] Cowan DA, Khan N, Pointing SB, Cary SC. Diverse hypolithic refuge communities in the McMurdo Dry Valleys. Antarct Sci 2010, 22:714–20. [107] Makhalanyane TP, Valverde A, Birkeland N-K, Cary SC, Tuffin IM, Cowan DA. Evidence for successional development in Antarctic hypolithic bacterial communities. ISME J 2013, 7:2080– 90. [108] Le PT, Makhalanyane TP, Guerrero LD, Vikram S, Van de Peer Y, Cowan DA. Comparative metagenomic analysis reveals mechanisms for stress response in hypoliths from extreme hyperarid deserts. Genome Biol Evol 2016, 8:2737–47. [109] Chan Y, Van Nostrand JD, Zhou J, Pointing SB, Farrell RL. Functional ecology of an Antarctic dry valley. Proc Natl Acad Sci USA 2013, 110:8990–5. [110] Gunnigle E, Ramond JB, Guerrero LD, Makhalanyane TP, Cowan DA. Draft genomic DNA sequence of the multi-resistant Sphingomonas sp. strain AntH11 isolated from an Antarctic hypolith. FEMS Microbiol Lett 2015, 362:fnv037. [111] Wei STS, Lacap-Bugler DC, Lau MCY, et al. Taxonomic and functional diversity of soil and hypolithic microbial communities in Miers Valley, McMurdo Dry Valleys, Antarctica. Front Microbiol 2016, 7:1642. [112] Smith MC, Bowman JP, Scott FJ, Line MA. Sublithic bacteria associated with Antarctic quartz stones. Antarct Sci 2000, 12:177–84. [113] Van Goethem MW, Makhalanyane TP, Valverde A, Cary SC, Cowan DA. Characterization of bacterial communities in lithobionts and soil niches from Victoria Valley, Antarctica. FEMS Microbiol Ecol 2016, 92:fiw051. [114] Rao S, Chan Y, Lacap D, Hyde K, Pointing S, Farrell R. Low-diversity fungal assemblage in an Antarctic Dry Valleys soil. Polar Biol 2011, 35:567–74. [115] Arenz BE, Held BW, Jurgens JA, Farrell RL, Blanchette RA. Fungal diversity in soils and historic wood from the Ross Sea Region of Antarctica. Soil Biol and Biochem 2006, 38:3057–64. [116] Arenz B, Blanchette R. Distribution and abundance of soil fungi in Antarctica at sites on the Peninsula, Ross Sea Region and McMurdo Dry Valleys. Soil Biol and Biochem 2011, 43:308–15. [117] Gokul J, Valverde A, Tuffin M, Cary S, Cowan D. Micro-eukaryotic diversity in hypolithons from Miers Valley, Antarctica. Biology 2013, 2:331–40. [118] Dreesens LL, Lee CK, Cary SC. The distribution and identity of edaphic fungi in the McMurdo Dry Valleys. Biology 2014, 3:466–83. [119] Uroz S, Kelly LC, Turpault M-P, Lepleux C, Frey-Klett P. The mineralosphere concept: mineralogical control of the distribution and function of mineral-associated bacterial communities. Trends Microbiol 2015, 23:751–62. [120] Kuhlman K, Fusco W, La Duc M, et al. Diversity of microorganisms within rock varnish in the Whipple Mountains, California. Appl Environ Microbiol 2006, 72:1708–15.

70 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

[121] Mergelov N, Goryachkin S, Shorkunov I, Zazovskaya E, Cherkinsky A. Endolithic pedogenesis and rock varnish on massive crystalline rocks in East Antarctica. Eurasian Soil Sci 2012, 45:901–17. [122] Edwards HG, Newton EM, Wynn-Williams DD, Coombes SR. Molecular spectroscopic studies of lichen substances 1: parietin and emodin. J Mol Struct 2003, 648:49–59. [123] Howard-Williams C, Vincent WF. Microbial communities in southern Victoria Land streams (Antarctica) I. Photosynthesis. In: Vincent WF, Ellis-Evans JC (eds). High Latitude Limnology. Springer Netherlands, 1989, 27–38. [124] Grube M, Cernava T, Soh J, et al. Exploring functional contexts of symbiotic sustain within lichen-associated bacteria by comparative omics. ISME J 2015, 9:412–24. [125] Erxleben A, Gessler A, Vervliet-Scheebaum M, Reski R. Metabolite profiling of the moss Physcomitrella patens reveals evolutionary conservation of osmoprotective substances. Plant Cell Rep 2012, 31:427–36. [126] Zucconi L, Onofri S, Cecchini C, et al. Mapping the lithic colonization at the boundaries of life in Northern Victoria Land, Antarctica. Polar Biol 2016, 39:91–102. [127] Wynn-Williams D. Cyanobacteria in Deserts – Life at the Limit? In: Whitton BA, Potts M (eds). The Ecology of Cyanobacteria. Springer Netherlands, 2002, 341–66. [128] Selbmann L, Grube M, Onofri S, Isola D, Zucconi L. Antarctic epilithic lichens as niches for black meristematic fungi. Biology 2013, 2:784–97. [129] Selbmann L, De Hoog G, Mazzaglia A, Friedmann E, Onofri S. Fungi at the edge of life: cryptoendolithic black fungi from Antarctic desert. Stud Mycol 2005, 51:1–32. [130] Broady PA. The ecology of sublithic terrestrial algae at the Vestfold Hills, Antarctica. British Phycological Journal 1981, 16:231–40. [131] Broady PA. Ecological and taxonomic observations on subaerial epilithic algae from Princess Elizabeth Land and Mac. Robertson Land, Antarctica. Br Phycol J 1981, 16:257–66. [132] De Los Rios A, Wierzchos J, Sancho LG, Green TA, Ascaso C. Ecology of endolithic lichens colonizing granite in continental Antarctica. Lichenol 2005, 37:383–95. [133] Hughes KA, Lawley B. A novel Antarctic microbial endolithic community within gypsum crusts. Environ Microbiol 2003, 5:555–65. [134] Weber B, Büdel B. Endoliths. In: Reitner J, Thiel V (eds). Encyclopedia of Geobiology. Springer Netherlands, 2011, 348–55. [135] Nienow J, Friedmann E, Ocamp-Friedmann R. Endolithic microorganisms in arid regions. In: Encyclopedia of environmental microbiology. NY, John Wiley & Sons Inc, 2003, 2:1100–12. [136] De Los Ríos A, Grube M, Sancho LG, Ascaso C. Ultrastructural and genetic characteristics of endolithic cyanobacterial biofilms colonizing Antarctic granite rocks. FEMS Microbiol Ecol 2007, 59:386–95. [137] Friedmann EI. Endolithic microbial life in hot and cold deserts. Orig Life 1980, 10:223–35. [138] Pointing SB, Warren-Rhodes KA, Lacap DC, Rhodes KL, McKay CP. Hypolithic community shifts occur as a result of liquid water availability along environmental gradients in China’s hot and cold hyperarid deserts. Environ Microbiol 2007, 9:414–24. [139] Archer SD, de los Ríos A, Lee KC, et al. Endolithic microbial diversity in sandstone and granite from the McMurdo Dry Valleys, Antarctica. Polar Biol 2016, doi:10.1007/s00300-016-2024-9. [140] Büdel B, Bendix J, Bicker FR, Allan Green T. Dewfall as a water source frequently activates the endolithic cyanobacterial communities in the granites of Taylor Valley, Antarctica. J Phycol 2008, 44:1415–24. [141] Büdel B, Schulz B, Reichenberger H, Bicker F, Green T. Cryptoendolithic cyanobacteria from calcite marble rock ridges, Taylor Valley, Antarctica. Algol Stud 2009, 129:61–9. [142] Jungblut AD, Neilan BA. NifH gene diversity and expression in a microbial mat community on the McMurdo Ice Shelf, Antarctica. Antarct Sci 2010, 22:117–22.

References | 71

[143] Yung CC, Chan Y, Lacap DC, et al. Characterization of chasmoendolithic community in Miers Valley, McMurdo Dry Valleys, Antarctica. Microb Ecol 2014, 68:351–9. [144] Choi A, Cho H, Kim S-H, Thamdrup B, Lee S, Hyun J-H. Rates of N2 production and diversity and abundance of functional genes associated with denitrification and anaerobic ammonium oxidation in the sediment of the Amundsen Sea Polynya, Antarctica. Deep Sea Res Part II Top Stud Oceanogr 2016, 123:113–25. [145] Goordial J, Davila A, Greer C, et al. Comparative activity and functional ecology of permafrost soils and lithic niches in a hyper-arid polar desert. Environ Microbiol 2016, 19:443–58. [146] Tahon G, Tytgat B, Stragier P, Willems A. Analysis of cbbL, nif H, and puf LM in soils from the Sør Rondane Mountains, Antarctica, reveals a large diversity of autotrophic and phototrophic bacteria. Microb Ecol 2016, 71:131–49. [147] Wei ST, Fernandez-Martinez M-A, Chan Y, et al. Diverse metabolic and stress-tolerance pathways in chasmoendolithic and soil communities of Miers Valley, McMurdo Dry Valleys, Antarctica. Polar Biol 2015, 38:433–43. [148] Edwards RA, Rohwer F. Viral metagenomics. Nature Rev Microbiol 2005, 3:504–10. [149] Dinsdale EA, Edwards RA, Hall D, et al. Functional metagenomic profiling of nine biomes. Nature 2008, 452:629–32. [150] Schoenfeld T, Liles M, Wommack KE, Polson SW, Godiska R, Mead D. Functional viral metagenomics and the next generation of molecular tools. Trends Microbiol 2010, 18:20–9. [151] Fancello L, Trape S, Robert C, et al. Viruses in the desert: a metagenomic survey of viral communities in four perennial ponds of the Mauritanian Sahara. ISME J 2013, 7:359–69. [152] Wilson WH, Lane D, Pearce DA, Ellis-Evans JC. Transmission electron microscope analysis of virus-like particles in the freshwater lakes of Signy Island, Antarctica. Polar Biol 2000, 23:657–60. [153] Zawar-Reza P, Argüello-Astorga GR, Kraberger S, et al. Diverse small circular single-stranded DNA viruses identified in a freshwater pond on the McMurdo Ice Shelf (Antarctica). Infect, Genet and Evol 2014, 26:132–8. [154] Yau S, Lauro FM, DeMaere MZ, et al. Virophage control of antarctic algal host–virus dynamics. Proc Natl Acad Sci USA 2011, 108:6163–8. [155] Laybourn-Parry J, Anesio AM, Madan N, Säwström C. Virus dynamics in a large epishelf lake (Beaver Lake, Antarctica). Freshwater Biol 2013, 58:1484–93. [156] Le Romancer M, Gaillard M, Geslin C, Prieur D. Viruses in extreme environments. Rev Environ Sci Bio 2007, 6:17–31. [157] Zablocki O, Adriaenssens EM, Cowan D. Diversity and ecology of viruses in hyperarid desert soils. Appl Environ Microbiol 2016, 82:770–7. [158] Hopkins D, Swanson M, Taliansky M. What do we know about viruses in terrestrial Antarctica? In: Cowan D, ed. Antarctic Terrestrial Microbiology: Physical and Biological Properties of Antarctic Soils. Heidelberg, Berlin, Springer-Verlag, 2014, 79–90. [159] Williamson KE, Radosevich M, Smith DW, Wommack KE. Incidence of lysogeny within temperate and extreme soil environments. Environ Microbiol 2007, 9:2563–74. [160] Zablocki O, van Zyl L, Adriaenssens EM, et al. High diversity of tailed phages, eukaryotic viruses and virophage-like elements in the metaviromes of Antarctic soils. Appl Environ Microbiol 2014, 80:6888–97. [161] Hogg ID, Stevens MI, Wall DH. Invertebrates. In: Cowan D, ed. Antarctic Terrestrial Microbiology: Physical and Biological Properties of Antarctic Soils. Heidelberg, Berlin, Springer-Verlag, 2014, 55–78. [162] Boveng PL, Hiruki LM, Schwartz MK, Bengtson JL. Population growth of Antarctic fur seals: limitation by a top predator, the leopard seal? Ecology 1998, 79:2863–77.

72 | 4 Microbiology of Antarctic Edaphic and Lithic Habitats

[163] Xu Z, Malmer D, Langille MG, Way SF, Knight R. Which is more important for classifying microbial communities: who’s there or what they can do. ISME J 2014, 8:2357–9. [164] Rampelotto PH. Extremophiles and extreme environments. Life 2013, 3:482–5. [165] Olsson-Francis K, de la Torre R, Cockell CS. Isolation of novel extreme-tolerant cyanobacteria from a rock-dwelling microbial community by using exposure to low Earth orbit. Appl Environ Microbiol 2010, 76:2115–21. [166] Sharon I, Banfield JF. Genomes from metagenomics. Science 2013, 342:1057–8. [167] Albertsen M, Hugenholtz P, Skarshewski A, Nielsen KL, Tyson GW, Nielsen PH. Genome sequences of rare, uncultured bacteria obtained by differential coverage binning of multiple metagenomes. Nat Biotechnol 2013, 31:533–8. [168] Chatterji S, Yamazaki I, Bai Z, Eisen JA. CompostBin: A DNA composition-based algorithm for binning environmental shotgun reads. In: Vingron M, Wong L (eds). Annual International Conference on Research in Computational Molecular Biology. Heidelberg, Berlin, SpringerVerlag, 2008, 17–28. [169] Lewin A, Wentzel A, Valla S. Metagenomics of microbial life in extreme temperature environments. Curr Opin Biotechnol 2013, 24:516–25. [170] Cowan DA, Chown SL, Convey P, et al. Non-indigenous microorganisms in the Antarctic: assessing the risks. Trends in Microbiol 2011, 19:540–8. [171] Nielsen KM, Johnsen PJ, Bensasson D, Daffonchio D. Release and persistence of extracellular DNA in the environment. Environ Biosafety Res 2007, 6:37–53. [172] Carini P, Marsden PJ, Leff JW, Morgan EE, Strickland MS, Fierer N. Relic DNA is abundant in soil and obscures estimates of soil microbial diversity. Nature Microbiol 2016, 2:16242.

Matthew A. Bowker, Burkhard Büdel, Fernando T. Maestre, Anita J. Antoninka, and David J. Eldridge

5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences 5.1 Overview Arid regions are distinct from most other biomes in that vascular plant cover is discontinuous, allowing light to reach the soil surface. Thus, a niche exists for the photosynthetic organisms that together comprise biological soil crusts (biocrusts). Biocrusts are a feature of arid regions worldwide, in both hot and cold climates, where they are a permanent component of successionally mature ecosystems [1]. Biocrusts are a continuous soil aggregate of the uppermost millimeters of the soil, distinguishable from other types of soil crust in that they are engineered by biota [2]. They harbor a wide variety of organisms (archaea, fungi, and bacteria – notably cyanobacteria [3–5]), in addition to mosses, liverworts and lichens, the subject of this chapter.

5.1.1 Moss, Liverwort, and Lichen Biology Mosses and liverworts are often grouped as “bryophytes”, although current understanding regards these as a polyphyletic group [6]. We will use the term bryophyte here for convenience to collectively refer to both mosses and liverworts. Both are true plants, of the kingdom Plantae, which lack the lignified vascular tissue characteristic of tracheophytes [7]. Without these tissues, their size is constrained, confining them to the soil surface, often beneath and in between vascular plants. Bryophytes are older than vascular plants, and are first encountered on land in the middle Ordovician period (∼ 470 mya), prior to the formation and breakup of the supercontinent Pangea [8]. Perhaps not surprisingly, they are found on all continents. Both mosses and liverworts may have impressive desiccation tolerance strategies to cope with low water availability, and are commonly found on arid soils as well [9]. Bryophytes do not reproduce by seed, but instead produce spores as a result of sex, dispersed by the sporophyte. Although spores can be dispersed long distances, including from continent to continent [10], many dominant bryophytes of arid regions produce no or few sporophytes [11, 12], constraining their dispersal, and possibly generating local adaptation. Bryophytes are generally capable of vegetative reproduction from any type of tissue [13], and may or may not also have specialized asexual propagules [14]. Lichens are a symbiosis of at least two primary bionts, a fungal partner (mycobiont; generally an ascomycete), and a photosynthetic partner (photobiont; a green DOI 10.1515/9783110419047-005

74 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

alga or cyanobacterium). Though they are often grouped together with bryophytes as nonvascular “plants,” they do not belong to the kingdom Plantae; rather they are classified as fungi and named based upon the mycobiont [15]. Despite lacking taxonomic relatedness, lichens do share some characteristics with bryophytes, including reproduction by spores and the lack of specialized water conductance mechanisms, which is related to small size and desiccation tolerance. Lichens are apparently younger than bryophytes, dating to ∼ 415 mya (the Devonian period) [16], but have controversially been proposed to date over 100 mya earlier [17]. Lichens are found on all continents, are small in stature and confined near to surfaces such as soils. Spores are the product of sex in the fungal biont and can be a long-distance dispersal agent [18], but to form a lichen must encounter a compatible photobiont upon germination [19]. Many lichens also reproduce vegetatively from propagules that contain both mycobiont fungal cells and photobiont cells, including specialized propagules such as soredia, isidia, or unspecialized thallus fragments [20]. Bryophytes and lichens are found throughout the world, from arctic tundra, to temperate tree trunks, to rock outcrops, to arid zone biocrusts. In drylands, at local scales, they may comprise a substantial amount of the eukaryotic diversity present [21, 22]. The purpose of this chapter is to summarize the dimensions of their biodiversity on arid soils, outline some of the major determinants of their biodiversity, and summarize the effects of bryophyte and lichen biodiversity on arid soil function.

5.2 Global Diversity and Characteristic Taxa 5.2.1 Global Species Pool The diversity distribution of biocrust organisms around the world is incompletely known. As a first approach to quantify this, we defined seven geographical regions spanning arid and semiarid areas, as well as polar deserts and initial soils of the temperate, boreal, and arctic climatic zones, which are characterized by a very sparse cover of vascular plants (Asia, Africa, North America, including Central America and Greenland, South America, Antarctica, Europe, and the Pacific region, i.e., Australia and New Zealand). In total 323 bryophyte (68 liverworts, 255 mosses) and 553 lichen species (88 cyanolichens, 465 chlorolichens) have been identified explicitly as biocrust components all globally presently being unevenly distributed amongst the different geographical regions (continents and subcontinents) partly due to differing research effort in different parts of the world [5, 23–35] (󳶳 Fig. 5.1). Among all geographical regions differentiated here, South America is the least known in terms of biocrust presence and their diversity and taxonomic composition. Only recently have research activities emerged, investigating biocrusts of several regions of this understudied continent [36–38].

5.2 Global Diversity and Characteristic Taxa |

Cyanolichens

Chlorolichens

Liverworts

75

Mosses

50

7,960,000 km2 10,180,000 km2

14,000,000 km2

100

17,840,000 km2

150

24,709,000 km2

200

30,521,532 km2

Species number

250

33,579,000 km2

300

c Pa ci fi

Af ric a N Am orth er ic a So u Am t er h An ica ta rc tic a Eu ro pe

As ia

0

Geographical region, decreasing size Fig. 5.1: Species numbers per geographical region (N-America includes Central America and Greenland; Pacific includes Australia and New Zealand); regions are arranged according to size.

Biocrust lichens are well known for all regions except South America, while biocrust bryophytes are well known only for Europe, North America, and the Pacific region (󳶳 Fig. 5.1). The highest species numbers found so far have been in Europe, followed by North America and Asia. In Europe and North America there are many scientists working on this topic, while in Asia this is true for Russia and China only.

5.2.2 Global Characteristic Taxa and β Diversity No bryophyte or lichen species occurs in biocrusts in all of the seven geographical regions defined here. However, 20 species (17 lichens, 3 mosses) occurred in at least four out of the seven geographical regions (󳶳 Tab. 5.1). These can be thought of as the more cosmopolitan, characteristic taxa. Two lichens, but no bryophytes, are documented in biocrusts of all regions except Antarctica. While it is notable that a few species are so widely distributed, the wider pattern suggests that most species are confined to only one or a few regions. With 287 bryophyte (60 liverworts, 227 mosses) and 411 lichen species (64 cyanolichens, 347 chlorolichens), the bulk of species from biocrusts is restricted to only one of the seven geographical regions (󳶳 Fig. 5.2). In two of the seven regions we found 26 bryophytes and 95 lichens, whereas in three of seven regions the number declined to 7 bryophytes and 30 lichens. For further details, see 󳶳 Fig. 5.2 and 󳶳 Tab. 5.1. While it is true that a

76 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

× × × × × ×

× × × × ×

× × ×

× ×

× × × × ×

× × × × × × × ×

× ×

× × × × × × ×

×

× ×

×

×

× × × × × ×

×

× × ×

× ×

× × × × ×

×

× × × ×

× × × × × × × × × × × ×

×

×

Pacific2

× × ×

Antarctica

× × × × × × × × × × × × × ×

Europe

× × × × ×

S-America

× × × × × × × ×

N-America1

Lichens Heppia despreauxii (Mont.) Tuck. Placidium squamulosum (Ach.) Breuss Collema tenax (Sw.) Ach. Diploschistes diacapsis (Ach.) Lumbsch Diploschistes muscorum (Scop.) R. Sant. Endocarpon pusillum Hedw. Peltula patellata (Bagl.) Swinsc. & Krog Placidium lacinulatum (Ach.) Breuss Placidium pilosellum (Breuss) Breuss Psora decipiens (Hedw.) Hoffm. Toninia sedifolia (Scop.) Timdal Cladonia fimbriata (L.) Fr. Cladonia furcata (Huds.) Schrad. Collema coccophorum Tuck. Fulgensia fulgens (Sw.) Elenkin Heppia adglutinata (Kremp.) A. Massal. Heppia lutosa (Ach.) Nyl. Acarospora nodulosa (Dufour) Hue Buellia epigaea (Hoffm.) Tuck. Buellia punctata (Hoffm.) A. Massal. Candelariella vitellina (Hoffm.) Müll. Arg. Cetraria islandica (L.) Ach. Cladonia cervicornis (Ach.) Flot. Cladonia foliacea (Huds.) Willd. (including C. convoluta) Cladonia pocillum (Ach.) O. J. Rich. Cladonia pyxidata (L.) Hoffm. Cladonia verticillata (Hoffm.) Schaer. Collema crispum var. crispum (Huds.) Weber ex F. H. Wigg. Fulgensia bracteata ssp. bracteata (Hoffm.) Räsänen Fulgensia desertorum f. macrospora Llimona Fulgensia subbracteata (Nyl.) Poelt Gypsoplaca macrophylla (Zahlbr.) Timdal Heppia solorinoides (Nyl.) Nyl. Peccania fontqueriana P. P. Moreno & Egea Peltula obscurans (Nyl.) Gyelnik Peltula radicata Nyl. Phaeorrhiza nimbosa (Fr.) H. Mayrhofer & Poelt Placynthium nigrum (Huds.) Grey Psora crenata (Taylor) Reinke

Africa

Species

Asia

Table 5.1: List of the 56 lichen and bryophyte species recorded from at least three out of the seven geographical regions defined here [23–35]. Species are arranged first according to their frequency and second alphabetically.

× ×

× × × × × × × × × × × × × ×

×

× × × × × × × × × × × × × ×

×

× × × ×

× ×

5.2 Global Diversity and Characteristic Taxa |

77

× × × × × × ×

Psora lurida Ach. Rinodina terrestris Tomin Squamarina cartilaginea (With.) P. James Squamarina lentigera (Weber) Poelt Toninia aromatica (Turner) A.Massal Toninia lutosa (Ach.) Timdal Toninia ruginosa (Tuck.) Herre Bryophytes Bryum argenteum Hedw. Bryum caespiticium Hedw. Ceratodon purpureus (Hedw.) Brid. Weissia controversa Hedw. Crossidium crassinerve (De Not.) Jur. Didymodon cf. rigidulus Hedw. Riccia lamellosa Raddi Riccia sorocarpa Bisch. Syntrichia ruralis (Hedw.) F.Weber & D.Mohr Trichostomum brachydontium Bruch ex F. Muell. 2

×

×

×

×

×

Species number

400

Cyanolichens Chlorolichens

300 200 100

250

× × × × × × ×

×

× ×

×

× × ×

× × × ×

× × ×

Liverworts Mosses

150 100 50 0

1g e 2 ogr ge . r eg o 3 gr. ion ge re gi o 4 gr. ons ge re gi o 5 gr. ons ge re gi o 6 gr. ons ge re og gio r. n re s g Al ion lr eg s io ns

1g e 2 ogr ge . r eg o 3 gr. ion ge re gi o 4 gr. ons ge re gi o 5 gr. ons ge re gi o 6 gr. ons ge re og gio r. n re s gi Al o l r ns eg io ns (a)

(b)

Fig. 5.2: Frequency of lichen (a) and bryophyte (b) species across seven geographic regions.

× × × ×

×

200

0

Pacific2

× × × ×

× × × ×

Europe

Antarctica

S-America

× ×

including Central America and Greenland. Australia, New Zealand

Species number

1

× ×

×

×

N-America1

Species

Africa

Asia

Table 5.1 (cont.): List of the 56 lichen and bryophyte species recorded from at least three out of the seven geographical regions defined here [23–35]. Species are arranged first according to their frequency and second alphabetically.

× × ×

78 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

lack of detection does not mean that a taxon is truly absent from a region, these data suggest a considerable amount of species turnover from continent to continent. More sampling effort is necessary to fill in current distribution gaps.

5.3 Determinants of Moss, Liverwort, and Lichen Diversity on Arid Soils 5.3.1 Geographic Isolation and Biogeography At large scales, dispersal limitations likely shape the bryophyte and lichen β diversity of major landmasses, the genetic diversity and distinctiveness, and α diversity of arid soil bryophyte and lichen communities. Bryophytes and lichens can disperse spores over long distances, e.g., from continent to continent [10, 18]. However, many dryland species may rely more upon vegetative propagules, e.g., tissue fragments, which are much more dispersal limited due to their larger size, possibly allowing for geographic isolation. At the global scale, we might expect that the mode of reproduction dictates the distribution of species, and we can hypothesize that this mechanism arranges arid soil bryophytes and lichens into groups based on dispersal limitation. The less dispersal limited group, which might abundantly produce spores, and in the case of lichens, also associate with a widely distributed photobiont, would be expected to be widespread or possibly cosmopolitan. An exemplar might be the moss Ceratodon purpureus, which is a prolific sporophyte producer, present on all continents (though not always in arid soil biocrusts) [10]. For lichens, long distance dispersal of spores is not sufficient in and of itself, because the spores must encounter a compatible photobiont. The lichen Psora decipiens is a broadly distributed lichen, which may reduce this problem by associating with multiple photobionts [39]. There are limits to spore distribution, therefore, even among cosmopolitan species. Genetic distance and floristic dissimilarity among populations may increase as connectivity via wind or geographic proximity decreases [18]. Other species are dispersal limited, due to a lack of successful reproduction via spores, and may either be widespread (found on several continents) or restricted in range (found on one or a few continents). Widespread dispersal limited species may be hypothesized to be relatively old, predating the breakup of the supercontinents. Such species might exhibit a strong degree of interspecific variation, and local adaptation, for example chemical races of lichens (Culberson 1986). Widespread dispersal limited species could be either common or rare. Common ones might include species found in arid regions of multiple land masses, but only rarely reproduce sexually. The lichen Gypsoplaca macrophylla may be an example of a rare species that falls within this group. Currently it has a wide distribution on three continents, including arid gypsiferous soils of southwestern US [22], in addition to Greenland, the Alps, and a

5.3 Determinants of Moss, Liverwort, and Lichen Diversity on Arid Soils

|

79

few localities in Asia [40]. It is always a rare community member. Perhaps this strange distribution arose through extinction of a formerly widespread taxon. Geographically restricted and dispersal limited species might be found only within a single major land mass, or a portion of one. These endemic community components might be hypothesized to represent evolutionarily younger species that arose after the breakup of the continents and have remained isolated due to longdistance dispersal limitation. The lichen genus Xanthoparmelia originated after the breakup of the continents [41] and has multiple species that have adopted a reliance on dispersal of vagrant, unattached thalli as propagules [42]. This reliance on local dispersal may explain the large degree of local endemism in this genus [42].

5.3.2 Climatic Gradients and Climate Change Climate is a major global driver of biocrust α and β diversity and composition in drylands. Rainfall, potential evapotranspiration and temperature all combine to determine the type of biocrust communities that can be supported. These effects vary with spatial scale, from continental and landscape scales, down to the scale of meters or less. Simultaneously dry and very cold environments may be at the physiological limits for some species to survive. Water may be scarce due to rarity of precipitation, or infrequency of thawing temperatures. For example, there are no liverworts or cyanolichens known from Antarctica (󳶳 Fig. 5.1). We may hypothesize that chlorolichens and mosses are more able to survive given the rarity of liquid water or are able to activate photosynthesis with less water. Within less extreme climates in the temperate and tropical regions, biocrust lichen and moss richness is correlated with soil moisture across large precipitation gradients [43]. Cooler habitats appear to support a large diversity and biomass of lichen taxa [44], possibly because the balance of photosynthesis and respiration between the symbiotic partners maximizes the opportunity to form complex thallus structures. Similarly, higher rainfall has been correlated with increasing richness and changes in biocrust composition [45]. Rainfall seasonality can also have marked effects on biocrust composition [27, 46]. In Australia, for example, biocrust lichens are restricted to winter rainfall dominant areas, where they are able to avoid hydration of the thallus during extremely hot weather [47]. Despite the preference for winter rainfall, very cold temperatures are not necessarily preferred. Areas in the northwestern United States (a winter rainfall region) with warmer winter temperatures have been shown to be more conducive to crust development than areas with colder winters [48]. Biocrust species richness and composition are also known to vary with altitude, which is usually a surrogate for increasing precipitation and decreasing temperature [26]. Castillo-Monroy et al. [37] showed that biocrust species richness in an Ecuadorian dryland increased with increasing elevation, with clear differences in composition along the elevational

80 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

gradient. These altitudinal differences can be attributed to the redistribution of runoff and differences in soil texture, which largely drive soil moisture availability, and consequently, competition from vascular plants and available niches for biocrust taxa. Changes in soil moisture availability at more local scales can also alter biocrust cover and composition. For example, the two major patch types in drylands (resource shedding water runoff zones and resource accumulating water runon zones) that result from the redistribution of water, support different taxa at small scales. Lichens and cyanobacteria typically dominate resource shedding areas, whereas microsites where resources accumulate are often dominated by bryophytes [49, 50]. The mechanism behind this distribution may relate to the need for bryophytes to access free water to reproduce but is also related to competition with vascular plants (e.g., 51,52]. At the microsite scale, the distribution of biocrust taxa is strongly dependent on soil moisture [22, 53–55] and the availability of suitable niches for establishment. These microsites are often areas that receive slightly more moisture, are cooler and sheltered from temperature extremes [56, 57]. Biocrusts lichens and mosses have been predicted to mediate any substantial effects on ecosystem functioning due to climate change [58–60]. However, there are also likely to be substantial changes in biocrust composition and richness resulting from a changing climate. For example, Ferrenberg et al. [61] showed that an increase in small summer rainfall events changed biocrust composition from moss dominated (Syntrichia caninervis) to cyanobacteria dominated (Microcoleus vaginatus) communities [61], and Maestre et al. (2015) reported up to a 45% decline in lichen dominated biocrusts with warming after 4 years [62].

5.3.3 Calcicole–Calcifuge Dichotomy and Soil pH Gradients Biocrust β diversity, particularly that of lichens, is known to be strongly influenced by soil pH, which in turn is strongly influenced by the concentrations of calcium (Ca) carbonate and other carbonates in the soil [27, 28, 48, 63–65]. The relationship between lichen taxa and soil pH is so pronounced that lichens have been classified into two broad functional groups according to their response to soil pH. Calciphiles, which include the majority of soil lichens in drylands, are strongly associated with soils of high pH. Conversely, calcifuges have a low tolerance to high pH soils [66] and appear to be more common in mesic soils. This dichotomy recurs in many locations around the world, dictating both biocrust abundance and community composition. In drylands in the western USA and Ecuadorian dry mountain shrublands, biocrusts reach their greatest development on neutral to acidic soils [37, 48]. In other dryland areas of the USA, Spain, Australia, and Israel, biocrust lichens and bryophytes are more diverse and occupy a greater cover in areas of high pH (e.g., [17, 47, 63, 67, 68]). Lichens inhabiting Ca rich soils are thought to have greater concentrations of Ca oxalate on the outer surface of the thallus, reducing the concentration of Ca in the immediate area where

5.3 Determinants of Moss, Liverwort, and Lichen Diversity on Arid Soils

|

81

the lichen attaches [69]. Magnesium, manganese, and other nutrients have also been shown to be highly correlated with crust cover and composition [28, 43, 56, 56, 66, 70], but the exact mechanisms behind their effects on biocrust taxa are still not fully understood and may relate to pH or carbonate gradients.

5.3.4 The Special Case of Gypsiferous Soils Occasionally, dryland soils have high levels of Ca in the form of gypsum [71]. Gypsum content is one of the edaphic factors most influential on taxonomic richness and species turnover of soil mosses, liverworts, and lichens in a given region [72–74]. For example, on the Colorado Plateau (USA), out of eight different soil types, gypsiferous soils had the greatest species richness (∼ 21 species per site), supported the second greatest species evenness, and supported eight indicator species out of a total of 19 [22]. In this case study, the gypsiferous soils had a disproportionately large effect on diversity at both local scales and within the entire study area. Higher taxonomic and functional richness of both mosses and lichens is also reported in Europe and Australia on gypsum soils [28, 72, 73, 75]. Gypsiferous arid soils of the Northern hemisphere and Australia often appear to be dominated by well distributed gypsophile lichen taxa such as Diploschistes spp., Psora decipiens, Fulgensia spp., Acarospora nodulosa, and Squamarina lentigera, among others [22, 28, 72, 76–78]. Where gypsum soils are rare in the landscape, these species may be rare or narrowly distributed within a region, despite local abundance and wide distribution globally. Gypsiferous soils also appear to harbor a larger number of endemics compared to other soils, a phenomenon also observed in vascular plants [79]. Perhaps this is because the specific edaphic preferences of the lichens, coupled with dispersal limitations, lead to narrow distributions. One example is Lecanora gypsicola, described in 1998 and known only from sporadically occurring gypsiferous soils of the western United States [80]. Dominant mosses of gypsiferous arid soils appear to differ more than lichens from region to region and may be generalist species rather than gypsum specialists [22, 78]. Widespread, but usually subdominant gypsophile species include Aloina bifrons, and a few Crossidium spp. [22, 73]. There are clear gypsum endemic mosses, however, including the North American endemic Didymodon nevadensis, which was only discovered in the 1990s [81]. Guerra et al. [73] list seven rare gypsophile species, known only from the Iberian Peninsula, including a rare gypsum tolerating liverwort, Riccia crustata. Why are gypsum soils such a distinct habitat? Bogdanović et al. [82] showed that two moss species with no reported preference for gypsum were able to tolerate its presence. Thus, the ability to grow on gypsum might be widespread in mosses, and this might contribute to high α diversity, but would not explain high species turnover from gypsiferous habitats to nongypsiferous habitats nearby. Rather, true gypsophiles must

82 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

either derive a benefit from growing in the habitat type, or resist its specific stresses better than most species. Gypsum contains Ca and sulfur, both essential nutrients. The fact that some gypsophiles also are found on soil rich in Ca carbonate might suggest a high demand for, or tolerance of Ca. A recent study of vascular plant endemism detected accumulations of Ca oxalate in plant tissues of gypsophiles, and hypothesized that this is a mechanism for coping with excess Ca [83]. This may be an intriguing clue, since lichen pruina are composed of Ca oxalates and most lichens preferentially growing on gypsum abundantly produce pruina. Nonetheless, soils rich in Ca carbonate but not gypsum often have different floras [22, 84], suggesting that Ca alone is an unlikely explanation of unique lichen and bryophyte assemblages on gypsiferous soils.

5.4 Consequences of Moss, Liverwort, and Lichen Diversity on Arid Soils 5.4.1 Contribution of Biocrust Lichens and Bryophytes to Arid Ecosystem Function Biocrust mosses and lichens play major roles in nutrient cycling, and in building and maintaining soil fertility. Lichen and bryophyte dominated biocrusts are an important part of the global carbon (C) budget, taking up from 1 to 37 g C m−2 yr−1 in arid lands, depending on the species composition, amount of cover and water availability [85– 87]. This is a substantial contribution to productivity in arid lands, accounting for as much as 3.7–13.9% of net primary productivity [88]. Likewise, lichens and bryophytes play key roles in regulating terrestrial nitrogen (N) cycling. N is commonly the most limiting nutrient in terrestrial ecosystems [89]. Many lichens house N fixing cyanobacterial symbionts within their thallus, and, likewise, biocrust mosses, are known to host N fixing symbionts on their leaves [90, 91]. Enzyme activity is high in lichen and moss dominated biocrusts, and is dependent on species composition, which is important for N, C and phosphorous cycling [92]. Microbial N fixation and N transformation activity is known to be stimulated within biocrusts [93], and these combined activities can account for the majority of available N input to arid systems [88, 94]. They also capture dust, which helps to promote ecosystem productivity by addition of both soil and nutrients to the ecosystem [95]. Because mosses and lichens bind the soil together with filamentous structures such as hyphae, rhizines, and rhizoids, they aggregate soil, reducing soil loss due to wind and water erosion [96, 97]. This is true even during inactivity because lichens and bryophytes of biocrusts have remarkable desiccation tolerance [98, 99], and the physical structure of the biocrust persists. Due to the physical structure of the biocrusts, mosses and lichens have complex effects on soil hydrology, which are largely dependent on biocrust composition, rainfall intensity, ambient temperature, and soil texture [50, 100, 101]. Lichens can have mixed effects, either generating runoff or promoting infiltration, depending upon the

5.4 Consequences of Moss, Liverwort, and Lichen Diversity on Arid Soils

|

83

surface connectivity of the lichen thallus, whereas mosses have greater surface roughness and high water absorbing capacity, at 100−1000× their dry mass, enhancing infiltration [101, 102]. Sinuous microtopography of well developed lichen and moss biocrusts can slow down the movement of water, enhancing infiltration compared to smoother cyanobacterial biocrusts, but many lichen biocrusts can generate runoff at high rainfall events [97, 103, 104]. Well developed crusts also influence water retention by reducing evaporation [104, 105]. All of these factors influence water availability for vascular plants and the soil food web. Finally, biocrusts composed of bryophytes and lichens support a vibrant soil food web in the top millimeters of soil because they leak much of the C and N that they fix back into the soil [106]. Recent work has demonstrated that microbes specialize on specific biocrust excretions, allowing the C and N to be recycled and re-assimilated [107]. Lichens and bryophytes produce a number of secondary compounds that provide protection from harmful ultraviolet radiation [108–110]. Surface bryophyte and lichen community resilience is critical for protecting biocrust community members that lack UV protection (e.g., light cyanobacteria).

5.4.2 Biodiversity–Ecosystem Functioning Relationship Understanding the links between biodiversity and those processes that determine the functioning of ecosystems (biodiversity–ecosystem functioning relationship) has been a major research topic in community and ecosystem ecology over the last two decades [111–114]. During this period, several hundred biodiversity–ecosystem functioning relationship studies have been conducted with a wide variety of organisms, such as vascular plants, algae and soil, fauna, and ecosystem processes, including primary productivity, nutrient cycling, or water quality (see [112, 113] for reviews). Biocrusts have not been an exception to this, and multiple observational and experimental studies have explored how changes in the diversity of biocrust constituents, such as lichens and mosses, affect ecosystem functioning [115, 116, 118, 121, 126–128]. Indeed, some attributes of biocrusts, such as small size and the ease of transplant and/or culturing their constituents, make them particularly suitable for biodiversity and ecosystem functioning research, and their use by researchers on this topic is being encouraged [132]. Most studies on the biodiversity–ecosystem function relationship to date have focused on particular ecosystem processes, such as productivity, and on species richness as a focal aspect of biodiversity [111, 113]. These studies provide ample evidence of positive richness function relationships in nature. As an example, Cardinale et al. [113] found that the relationship between producer diversity and biomass was best described by some form of a positive but decelerating curve in 79% (of 272) studies, while linear relationships were found in only 13% of cases. Similar results were found when looking at functions such as nutrient uptake (89% positive but decelerating

84 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

curve, 9% linear relationship, 47 studies) or decomposition (61% positive but decelerating curve, 19% linear relationship, 36 studies; 113). Biocrusts have proven to be no exception to the positive relationship between biodiversity and ecosystem functioning reported with other organisms; however, they more commonly exhibit approximately linear relationships between the number of macroscopic species (bryophytes and lichens) and various indicators of nutrient cycling, hydrological, and soil development and retention functions. Positive richness function relationships are supported in multiple observational field studies conducted in drylands [115, 116], although sometimes negative effects or no effects are reported [117]. Moisture availability also plays a role in determining biodiversity–ecosystem functioning relationships. Mulder et al. (2001) experimentally tested the relationships between species diversity and productivity using mosses and liverworts [118]. They found that biomass increased with species richness, but only when communities were subject to experimental drought. Rixen and Mulder [119] exposed arctic tundra moss communities of varying richness to two drought and density levels, and found that productivity was increased in the species rich communities, particularly in the low density plots, but only when plots were watered regularly. They also found that moisture retention improved at high species richness levels, as a result of the positive effects that biomass had on moisture conditions. Other studies have explored how the diversity of microbes associated with biocrusts affect ecosystem functioning. For example, Hu et al. (2002) observed that artificial biocrusts composed of multiple cyanobacterial species aggregated soil more strongly than biocrusts formed by single species [120]. It would be reasonable to believe that some apparent effects of bryophyte and lichen diversity are actually mediated by community properties of associated bacteria and fungi. Nonetheless, CastilloMonroy et al. [121] found that lichen richness, rather than bacterial richness, was directly related to multiple ecosystem functions related to nutrient cycling. More studies on this topic will help partition the relative influence of bryophyte lichen and microbial diversity on ecosystem functions.

5.4.3 Effects of Species Richness, Turnover, and Evenness on Ecosystem Functions Despite biodiversity encompassing multiple components, most studies on the biodiversity–ecosystem functioning relationship conducted to date have targeted species richness, or α diversity, as the main biodiversity descriptor [113]. However, there is growing evidence suggesting that other components of biodiversity, such as species evenness, β diversity (species turnover), trait diversity, functional group diversity, phylogenetic diversity, and within species genetic diversity, have the potential to influence ecosystem processes [122–125]. Only some of these elements of biodiversity have been investigated using biocrusts. In 󳶳 Tab. 5.2, we compile results from the literature on the frequency of effects of biocrust lichen and bryophyte α diversity,

5.4 Consequences of Moss, Liverwort, and Lichen Diversity on Arid Soils

|

85

β diversity

% positive

evenness

% positive

Dataset

α diversity

Table 5.2: Percentage of cases in which α diversity, evenness, and β diversity of biocrust bryophytes and/or lichens have a detectable effect on an indicator of ecosystem function. In the case of α diversity and evenness, the proportion of these effects that are positive is also reported. We report main effects only; in some cases interactive effects are detected. White filled cells indicate no data. Black filled cells indicate that an effect on multifunctionality was reported. Mean reflects the average proportion of ecosystem function indicators affected per dataset. Frequency reflects the percentage of datasets in which there are > 0 effects on ecosystem function indicators detected.

Single site, Alicante, Spain [117]

0

Single site, Cuenca, Spain [117]

80

25

0

100

100

100

0

magnetic susceptibility

50

100

50

100

surface roughness, soil aggregate stability

Many sites, Utah, USA [115]

100

50

0

Single site, Communidad de Madrid, Spain [36, 115, 133]

33

100

0

Single site, Communidad de Madrid, Spain [50]

0

Many sites, Utah, USA [115] Many sites, Arizona, USA [115]

80

25

Function indicators bulk density, respiration, organic C, total N, soil aggregate stability bulk density, respiration, organic C, total N, soil aggregate stability

magnetic susceptibility, surface roughness 100

phosphatase, β-glucosidase, urease

100

Steady state infiltration

Many sites, Central & Southern Spain (gypsum soils) [116, 128]

83.3 100

16.7 100

66.7a “C cycling”, respiration, phosphatase, total N, urease, multifunctionality

Many sites, Central & Southern Spain (calcareous soils) [116, 128]

42.9

14.3 100

33.3a organic C, β-glucosidase, respiration, phosphatase, total N, urease, multifunctionality

Constructed biocrusts composition experiment (surface) [126, 134]

20

a

66.7

0

10

ammonium, nitrate, organic C, total N, β-glucosidase, phosphatase, urease, N-fixation, multifunctionality, microbial catabolic profile

Bowker et al. 2013 [116] did not address β diversity. Bowker et al. 2011 [128] analyzed β diversity effects on individual functions but not on multifunctionality.

86 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

Constructed biocrusts composition experiment (subsurface) [126]

80

80

Constructed biocrusts evenness experiment (surface) [126, 134]

10

100

Constructed biocrusts evenness experiment (subsurface) [126]

60

β diversity

% positive

evenness

% positive

Dataset

α diversity

Table 5.2 (cont.): Percentage of cases in which α diversity, evenness, and β diversity of biocrust bryophytes and/or lichens have a detectable effect on an indicator of ecosystem function. In the case of α diversity and evenness, the proportion of these effects that are positive is also reported. We report main effects only; in some cases interactive effects are detected. White filled cells indicate no data. Black filled cells indicate that an effect on multifunctionality was reported. Mean reflects the average proportion of ecosystem function indicators affected per dataset. Frequency reflects the percentage of datasets in which there are > 0 effects on ecosystem function indicators detected.

Function indicators

60

organic C, total N, β-glucosidase, phosphatase, multifunctionality

0

20

ammonium, nitrate, organic C, total N, β-glucosidase, phosphatase, urease, N-fixation, multifunctionality, microbial catabolic profile

0

40

organic C, total N, β-glucosidase, phosphatase, multifunctionality

Single site, Baja California, Mexico [129]

100

CO2 gas exchange

Single site, Communidad de Madrid, Spain [92]

100

organic C, hexoses, phenols, respiration, total N, microbial biomass N, amino acids, proteins, dissolved inorganic p, phosphatase

Mean Frequency

50.7 84.6

33.3

68.6 90.9

26.1 50.0

65.0 66.3 80.0 100.0

evenness, or β diversity on ecosystem functioning. Our rules for inclusion required an explicit manipulation or measurement of one of these elements of biodiversity, a focus on biocrusts of dryland soils, and a measurement of at least one indicator of ecosystem function. We excluded measurements of activity or physiology of isolated biocrust organisms, focusing instead on the functions of biocrust communities. Finally, in our consideration of β diversity, we included comparisons of biocrusts

5.4 Consequences of Moss, Liverwort, and Lichen Diversity on Arid Soils

|

87

dominated by a particular species, but excluded comparisons of biocrust types and effects of turnover among morphological groups, because species compositions were not explicitly measured. Overall, available evidence suggests that, as in several other communities, species richness commonly exerts positive effects on ecosystem functioning in biocrusts. In 85% of cases meeting our inclusion criteria, at least one α diversity relationship was detected with ecosystem function (󳶳 Tab. 5.2). On average, about half of the ecosystem function indicators were affected by α diversity, over two thirds of which were positive. The magnitude and sign of these effects depend on the characteristics of the biocrust community (abundance, spatial pattern), the ecosystem function considered, environmental conditions, and the interactions among these factors. Species richness has been found to be a better indicator of ecosystem functioning than the richness of a priori functional groups, perhaps because our limited knowledge of the functional traits of biocrust constituents does not properly group species according to their impacts on ecosystem functioning [51, 90]. Alternatively, it may mean that biocrust moss and lichen species tend to have unique suites of functional traits [84, 115], and perhaps a trait diversity index would prove to be even more informative than species richness. Biocrust evenness is less commonly related to ecosystem functioning; at least one evenness–function relationship occurs in about half of cases, and about a quarter of functional indicators were influenced by evenness (󳶳 Tab. 5.2). As with α diversity, most of these relationships were positive. Despite the lower frequency of main effects, evenness is sometimes influential in interaction with other biocrust properties (e.g., spatial patterning) [115, 126, 127]. Beta diversity was most the most consistent influence on ecosystem functioning. Relationships between β diversity and at least one ecosystem function were detected in all available studies meeting our criteria, and two thirds of ecosystem function indicators examined were influenced by β diversity (󳶳 Tab. 5.2) These effects extend to hydrology [50, 115], nutrient cycling [126, 128], and production [129]. While the number of studies conducted to date precludes us making strong inferences, the mounting available evidence suggests that species richness and β diversity are among the most influential biocrust attributes driving biodiversity–ecosystem functioning relationships. These biodiversity effects are as strong as or stronger than those of community attributes such as total cover or spatial patterning [117, 126].

5.4.4 Multifunctionality Increasingly, ecologists are moving beyond considering single ecosystem functions, such as productivity, to multifunctionality, defined as the simultaneous performance of multiple ecosystem functions [122]. Delgado-Baquerizo et al. [60] conducted a survey on three continents to assess how biocrust forming mosses affect multifunctionality, as measured with multiple soil variables related to carbon, nitrogen and phos-

88 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

phorus cycling and storage. Compared with soil surfaces lacking biocrusts, biocrust forming mosses enhanced multifunctionality in semiarid and arid environments, but not in humid and dry subhumid ones. They also found that the relatively positive effects of biocrust forming mosses on multifunctionality compared with bare soil increased with increasing aridity. Thus the presence of biocrusts does seem to enhance ecosystem multifunctionality. The next logical question is whether the diversity of biocrusts exerts an effect upon multifunctionality as it does for single ecosystem functions. Lefcheck et al. [114] conducted a meta-analysis of the effects of species richness on multifunctionality using a comprehensive database of 94 experiments, manipulating species richness across a wide variety of taxa, trophic levels, and habitat. Two key results from this study were: (i) multifunctionality was enhanced as species richness increased, and (ii) the overall effect of species richness on multifunctionality grew stronger as more functions were considered. To date, two studies have suggested that a greater number of biocrust species promotes greater multifunctionality and that a greater number of species is required to sustain multiple functions than a single function (󳶳 Tab. 5.2) [116, 126]. The few studies available indicate that diversity of biocrust mosses and lichens is highly important to maintain ecosystem multifunctionality in drylands, and that biocrusts follow the general trend exhibited by other communities.

5.4.5 Functional Redundancy or Singularity? Given that mosses, liverworts, and lichens are all poikilohydric, and desiccation and stress tolerant primary producers, it would be logical to suspect that they tend toward functional redundancy [130]. Redundant species are essentially interchangeable, and the loss of one such species would not be expected to reduce ecosystem function, although it has been suggested that redundancy may bolster an ecosystem’s ability to maintain function under differing conditions [131]. There are two reasons why we doubt that biocrust bryophytes and lichens are functionally redundant. First, if biocrust mosses, liverworts, and lichens were redundant, we would expect ecosystem function or multifunctionality to asymptote at relatively low levels of species richness; this is not so. Relationships between biocrust richness and their functionality are much closer to linear relationships than asymptotic ones, suggesting that at least across the range of observed values, an increase in richness leads to an increase in a given function or in multifunctionality [115, 132]. This observation might relate to variation in response to environment, for example different ideal combinations of water and light availability and temperature for maximal photosynthetic rate among species [129]. A multispecies community with different environmental optima would be more likely to maintain high productivity, regardless of the conditions at a given moment. The other reason to believe that individual species are fundamentally different is that individual species abundances can be tied to high values of particu-

5.5 Summary and Conclusions

| 89

lar functional indicators, suggesting distinct ecological roles [128, 133]. For example, biocrust communities rich in the lichen Squamarina lentigera exhibited higher phosphatase activity, when compared to communities dominated by Diploschistes diacapsis [128]. Likewise, mosses and lichens exhibit fundamentally different effects on hydrology, with mosses often acting as infiltration promoters, but lichens acting to generate runoff [50]. Different mosses and lichens are also known to have distinct functional traits. For example, only a subset of lichens is known to have the ability to fix nitrogen (e.g., Collema, Leptogium, Heppia, Peltula, Peltigera). Lichen and moss species also have a wide chemical diversity, and many of the chemicals likely affect other community members that may impact ecosystem processes [42, 92, 108]. We suggest that the perception of redundancy disappears when more than one function is considered. Functional profiles of 23 biocrust forming organisms in Spain were tabulated along with all of their documented effects on ecosystem functions [128]. Over half of them had a unique set of effects, even though many species exerted some of the same effects. When considering biodiversity loss, this suggests that at low levels of biodiversity, communities may have different functional attributes based on the particular species present. As more species are added, it becomes more likely that most functions are being conducted by at least one species, and, therefore, multifunctionality is more likely to be sustained at higher richness [116, 126].

5.5 Summary and Conclusions Biocrust lichens and bryophytes shape the landscape in all areas where vascular plant development is limited, including arid regions, occupying the soil surface and providing important ecosystem functions. Biocrust lichens and bryophytes are documented from all continents, and some species are widespread among land masses. The majority of species are restricted to one or a few geographic areas, a pattern that may partly be determined by dispersal limitations. Within major landmasses, α and β-diversity are largely determined by climatic gradients such as aridity, or edaphic factors such as pH or gypsum content of the soil. Depending on these factors, different community assemblages are formed, with resulting impacts on ecosystem function. In general, ecosystem function increases with higher biocrust species richness, for individual ecosystem functions as well as for ecosystem multifunctionality. Changes in community composition have also been linked to differences in ecosystem function or multifunctionality. Because of this, and evidence that some ecosystem functions are tied to particular species traits, it is important to consider individual biocrust moss and bryophyte species as singularly important, rather than functionally redundant. Climate change and land use practices are already impacting the function and diversity of biocrust communities. Management and conservation efforts should focus on maintaining viable biocrust habitat (especially that of endemics) aiding dispersal, and restoring biocrust communities in degraded habitat.

90 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

References [1]

Bowker MA. Biological soil crust rehabilitation in theory and practice: an underexploited opportunity. Restor Ecol 2007, 15:13–23. [2] Jones CG, Lawton JT, Shachack M. Organisms as ecosystem engineers. Oikos 1994, 69:373– 86. [3] Garcia-Pichel F, Loza V, Marusenko Y, Mateo P, Potrafka R. Temperature drives the continental scale distribution of key microbes in topsoil communities. Science 2013, 340:1574–7. [4] Steven B, Kuske CR, Reed SC, Belnap J. Climate change and physical disturbance manipulations result in distinct biological soil crust communities. Appl Env Microbiol 2015, 81:7448– 59. [5] Bowker MA, Belnap J, Büdel B, Sannier C, Pietrasiak N, Eldridge DJ, Rivera-Aguilar V. Controls on distribution patterns of biological soil crusts at micro- to global scales. In: Weber B, Büdel B, Belnap J (eds). Biological soil crusts: an organizing principle in drylands. Berlin, SpringerVerlag, 2016, 173–97. [6] Mishler BD, Lewis LA, Buchheim MA, Renzaglia KS, Garbary DJ, Delwiche CF, ZechmanFW, Kantz TS, Chapman RL. Phylogenetic relationships of the “green algae” and “bryophytes.” Ann Mo Bot Gard 1994, 81:451–83. [7] Graham LE, Cook ME, Busse JS. The origin of plants: body plan changes contributing to a major evolutionary radiation. Proc Nat Acad Sci USA 2000, 97:4535–40. [8] Rubinstein CV, Gerrienne P, de la Puente GS, Astini RA, Steemans P. Early middle Ordovician evidence for land plants in Argentina (eastern Gondwana). New Phytol 2010, 188:365–9. [9] Oliver MJ, Velten J, Mishler BD. Desiccation Tolerance in Bryophytes : A Reflection of the Primitive Strategy for Plant Survival in Dehydrating Habitats. Integr Comp Biol 2005, 45:789–99. [10] McDaniel SF, Shaw AJ. Selective sweeps and intercontinental migration in the cosmopolitan moss Ceratodon purpureus (Hedw.) Brid. Mol Ecol 2005, 14:1121–32. [11] Stark LR, Castetter RC. A gradient analysis of bryophyte populations in a desert mountain range. Memoirs of the New York Botanical Garden, 1987, 45:186–97. [12] Stark LR, Mishler BD, McLetchie DN. The cost of realized sexual reproduction : and sporophyte abortion in a desert moss. Am J Bot 2000, 87:1599–1608. [13] La Farge C, Williams KH, England JH (2013). Regeneration of Little Ice Age bryophytes emerging from a polar glacier with implications of totipotency in extreme environments. Proc Nat Acad Sci USA 2013, 110:9839–44. [14] Glime, Janice M. 2007 Bryophyte Ecology. Volume 1. Physiological Ecology. Houghton, Michigan, USA, Michigan Technological University and the International Association of Bryologists, 2007 (ebook accessed on 12 December 2015 at http://www.bryoecol.mtu.edu/). [15] Tehler A. Systematics, phylogeny and classification. In: Nash III, TH, ed. Lichen Biology. Cambridge, UK, Cambridge University Press, 1996, 217–39. [16] Honegger R, Edwards D, Axe L. The earliest records of internally stratified cyanobacterial and algal lichens from the lower Devonian of the Welsh borderland. New Phytol 2013, 197:264–75. [17] Retallack GJ. Ediacaran life on land. Nature 2013, 493:89–92. [18] Muñoz J, Felicísimo ÁM, Cabezas F, Burgaz AR, Martínez I. Wind as a Long-Distance dispersal vehicle in the southern hemisphere. Science 2004, 304:1144–7. [19] Seymour FA, Crittenden PD, Dyer PS. Sex in the extremes: lichen forming fungi. Mycologist 2005, 19:51–8. [20] Fahselt D. Individuals and populations of lichens. In: Nash TH, III, ed, Cambridge University Press, Cambridge, 2008, 252–73.

References | 91

[21]

[22] [23]

[24]

[25]

[26] [27] [28] [29] [30] [31]

[32]

[33] [34] [35]

[36]

[37] [38]

[39]

Rosentreter R. Compositional patterns within a rabbitbrush (Chrysothamnus) community of the Idaho Snake River Plain. In: McArthur D, Durant E, Welch BL (eds). Proceedings, Symposium on the biology of Artemisia and Chrysothamnus. Ogden, Utah, US Department of Agriculture, 1986, 273–7. Bowker MA, Belnap J. A simple classification of soil types as habitats of biological soil crusts on the Colorado Plateau, USA. J Veg Sci 2008, 19:831–40. Belnap J, Büdel B, Lange OL. Biological soil crusts: characteristics and distribution. In: Belnap J, Lange OL, ed. Biological soil crusts: structure, function, and management. Berlin, Springer, 2003, 3–30. Büdel B, Darienko T, Deutschewitz K, Dojani S, Friedl T, Mohr KI, Salisch M, Reisser W, Weber B. Southern African biological soil crusts are ubiquitous and highly diverse in drylands, being restricted by rainfall frequency. Microb Ecol 2009, 57:229–47. De los Rios A, Raggio J, Pérez-Ortega S, Vivas M, Pintado A, Green TGA, Ascaso C, Sancho LG. Anatomical, morphological and ecophysiological strategies in Placopsis pycnotheca (lichenized fungi, Ascomycota) allowing rapid colonization of recently deglaciated soils. Flora 2011, 206:857–64. Dettweiler-Robinson E, Bakker JD, Grace JB. Controls of biological soil crust cover and composition shift with succession in sagebrush shrub-steppe. J Arid Envir 2013, 94:96–104. Eldridge DJ. Distribution and floristics of terricolous lichens in soil crusts in arid and semi-arid New South Wales, Australia. Aust J Bot 1996, 44:581–599. Eldridge DJ, Tozer ME. Environmental factors relating to the distribution of terricolous bryophytes and lichens in semi-arid Eastern Australia. Bryologist 1997, 100:28–39. Eldridge DJ, Koen TB. Cover and floristics of microphytic soil crusts in relation to indices of landscape health. Plant Ecol 1998, 137:101–14. Frey W, Herrnstadt I, Kürschner H. Verbreitung und Soziologie terrestrischer Bryophytengesellschaften in der Jüdäischen Wüste. Phytocoenologia 1990, 19:233–65. Haarmeyer DH, Luther-Mosebach J, Dengler J, Schmiedel U, Finckh M et al. (2010) Biodiversity in southern Africa. Vol. 1: Patterns at local scale – the BIOTA observatories. Göttingen & Windhoek, Klaus Hess Publishers, 1–801. Hawkes CV, Flechtner VR Biological soil crusts in a xeric Florida shrubland: Composition, abundance, and spatial heterogeneity of crusts with different disturbance histories. Microb Ecol 2002, 43:1–12. Rogers RW. Soil surface lichens on a 1500 kilometre climatic gradient in subtropical eastern Australia. Lichenologist 2006, 38:565–75. McCune B, Rosentreter R. Biotic soil crust lichens of the Columbia Basin. Corvallis, Oregon, Northwest Lichenologists, 2007, 1–105. Williams W, Büdel B. Species diversity, biomass and long-term patterns of biological soil crusts with special focus on Cyanobacteria of the Acacia aneura Mulga Lands of Queensland, Australia. Algol Studies 2012, 140:23–50. Castillo-Monroy AP, Maestre FT. La costra biológica del suelo: Avances recientes en el conocimiento de su estructura y función ecológica. Revista Chilena de Historia Natural 2011, 84:1–21. Castillo-Monroy A, Benítez A, Reyes-Bueno F, Donoso D, Cueva A. Biocrust structure responds to soil variables along a tropical scrubland elevation gradient. J Arid Environ 2016, 124:31–38. Raggio J, Green TGA, Crittenden PD, Pintado A, Vivas M, Péres-Ortega S, De los Rios A, Sancho LG. Comparative ecophysiology of three Placopsis species, pioneer lichens in recently exposed Chilean glacial forelands. Symbiosis 2012, 56:55–66. Ruprecht U, Brunauer G, Türk R. High photobiont diversity in the common European soil crust lichen Psora decipiens. Biodivers Conserv 2014, 23:1771–85.

92 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

[40] Timdal E. Gypsoplacaceae and Gypsoplaca, a new family and genus of squamiform lichens. Bibl Lichenol 1990, 38:419–27. [41] Amo de Paz G, Cubas P, Divakar PK, Lumbsch HT, Crespo A. Origin and Diversification of Major Clades in Parmelioid Lichens (Parmeliaceae, Ascomycota) during the Paleogene Inferred by Bayesian Analysis. PLoS ONE 2011, 6:e28161. [42] Galloway DJ. Lichen biogeography. In: Nash III TH, ed. Lichen biology. Cambridge, UK, Cambridge University Press, 2008, 317–37. [43] Bowker MA, Belnap J, Davidson DW, Phillips SL. Evidence for micronutrient limitation of biological soil crusts: potential to impact aridlands restoration. Ecol Appl 2005, 15:1941–51. [44] Eversman S. Lichens of alpine meadows on the Beartooth Plateau, Montana and Wyoming, U.S.A. Arct Alp Res 1995, 27:400–6. [45] Concostrina-Zubiri L, Martínez I, Rabasa SG, Escudero A. The influence of environmental factors on biological soil crust: from a community perspective to a species level approach. J Veg Sci 2014, 25:503–13. [46] Zedda L, Grongroft A, Schultz M, Petersen A, Mills A, Rambold G. Distribution patterns of soil lichens across the principal biomes of southern Africa. J Arid Environ 2011, 75:215–20. [47] Rogers RW. Soil surface lichens in arid and subarid southeastern Australia. III. The relationship between distribution and environment. Aust J Bot 1972, 20:301–16. [48] Ponzetti J, McCune B. Biotic soil crusts of Oregon’s shrub steppe: community composition in relation to soil chemistry, climate, and livestock activity. Bryologist 2001, 104:212–25. [49] Maestre FT, Huesca MT, Zaady E, Bautista S, Cortina J. Infiltration, penetration resistance and microphytic crust composition in contrasted microsites within a Mediterranean semi-arid steppe. Soil Biol Biochem 2002, 34:895–898. [50] Eldridge DJ, Bowker MA, Maestre FT, Alonso P, Mau RL, Papadopoulos J, Escudero A. Interactive effects of three ecosystem engineers on infiltration in a semi-arid Mediterranean grassland. Ecosystems 2010, 13:499–510. [51] Eldridge DJ. Dynamics of moss- and lichen-dominated soil crusts in patterned Callitris glaucophylla woodlands in eastern Australia. Acta Oecol 1999, 20:159–70. [52] Eldridge DJ. Biological soil crusts of Australia. In: Belnap J, Lange OJ, Berlin, Springer-Verlag, 2003, 119–132. [53] George DB, Davidson DW, Schleip KC, Patrell-Kim LJ. Microtopography of microbiotic crusts on the Colorado Plateau, and the distribution of component organisms. Wes Nor Amer Nat 2000, 60:343–54. [54] Proctor M. The bryophyte paradox: tolerance of desiccation, evasion of drought. Plant Ecol2000, 151:41–9. [55] Raabe S, Müller J, Manthey M, Dürhammer O, Teuber U, Göttlein A, Förster B, et al. Drivers of bryophyte diversity allow implications for forest management with a focus on climate change. For Ecol Manage 2010, 260:1956–64. [56] Belnap J, Lange OL. Biological Soil Crusts: Structure, Function, and Management. SpringerVerlag, Berlin, 2003. [57] Maestre FT, Bowker MA, Canton Y, Castillo-Monroy AP, Cortina J, Escolar C, Escudero A, Lazaro R, Martinez I. Ecology and functional roles of biological soil crusts in semi-arid ecosystems of Spain. J Arid Environ 2011, 75:1282–91. [58] Reed SC, Coe KK, Sparks JP, Housman DC, Zelikova TJ, Belnap J. Changes to dryland rainfall result in rapid moss mortality and altered soil fertility. Nat. Clim. Change 2012, 2:752–55. [59] Maestre FT, Escolar C, de Guevara ML, Quero JL, Lazaro R, Delgado-Baquerizo M, Ochoa V, Berdugo M, Gozalo B, Gallardo A. Changes in biocrust cover drive carbon cycle responses to climate change in drylands. Global Change Biology 2013, 19:3835–47.

References |

93

[60] Delgado-Baquerizo M, Maestre FT, Eldridge DJ, Bowker MA, Ochoa V, Gozalo B, Berdugo M, Val J, Singh BK. Biocrust-forming mosses mitigate the negative impacts of increasing aridity on ecosystem multifunctionality in drylands. New Phytol 2016, doi:10.1111/nph.13688. [61] Ferrenberg S, Reed SC, Belap J. Climate change and physical disturbance cause similar community shifts in biological soil crusts. Proc Nat Acad of Sci USA 2015, 112:12116–21. [62] Maestre FT, Escolar C, Bardgett R, Dungait JAD, Gozalo B, Ochoa V. Warming reduces the cover and diversity of biocrust-forming mosses and lichens, and increases the physiological stress of soil microbial communities in a semi-arid Pinus halepensis plantation. Front Microbiol 2015, 6:865. [63] McCune B, Rosentreter R. Field key to soil lichens of central and eastern Oregon. Unpublished report. 1995, Oregon State University and USDI BLM. [64] Hauck M, Jürgens S-R, Willenbruch K, Huneck S, Leuschner C. Dissociation and metal-binding characteristics of yellow lichen substances suggest a relationship with site preferences of lichens. Ann Bot 2009, 103:13–22. [65] Rivera-Aguilar V, Godınez-Alvarez H, Moreno-Torres R, Rodrıguez-Zaragoza S. Soil physicochemical properties affecting the distribution of biological soil crusts along an environmental transect at Zapotitlan drylands, Mexico. J Arid Environ 2009, 73:1023–8. [66] Bowker MA, Belnap J, Davidson DW, Goldstein H. Correlates of biological soil crust abundance across a continuum of spatial scales: support for a hierarchical conceptual model. J Appl Ecol 2006, 43:152–63. [67] Ochoa-Hueso R, Hernandez RR, Pueyo JJ, Manrique E. Spatial distribution and physiology of biological soil crusts from semi-arid central Spain are related to soil chemistry and shrub cover. Soil Biol and Biochem 2011, 43:1894–1901. [68] Downing AJ, Selkirk PM. Bryophytes on the calcareous soils of Mungo National Park, and arid area of southern central Australia. Great Basin Naturalist 1993, 53:13–23. [69] Syers JK, Iskandar IK. The pedogenetic significance of lichens. In: Ahmadjian V, Hale ME (eds). The Lichens. Academic Press, New York, 1973, 225–48. [70] Thompson DB, Walker LR, Landau FH, Stark LR. The influence of elevation, shrub species, and biological soil crust on fertile islands in the Mojave Desert, USA J Arid Environ2005, 61:609– 29. [71] Ullmann I, Büdel B. Biological soil crusts on a landscape scale. In: Belnap J, Lange OJ. Biological soil crusts: structure, function, and management. Berlin, Springer-Verlag, 2003, 203–13. [72] Nimis PL, Poelt J, Tretiach M. Lichens from the gypsum Park of the northern Apennines (N Italy). Cryptogamie Bryol L1996, 17:23–38. [73] Guerra J, Ros R, Cano M, Casares M. Gypsiferous outcrops in SE Spain, refuges of rare, vulnerable and endangered bryophytes and lichens. Cryptogamie Bryol L 1995, 16:125–35. [74] Anderson DC, Rushforth SR. The cryptogam flora of desert soil crusts in southern Utah, USA. Nova Hedwig 1976, 28:691–729. [75] Casares-Porcel M, Gutiérrez-Carretero L. Síntesis de la vegetación liquénica gipsícola termo- y mesomediterránea de la Península Ibérica. Cryptogamie. Bryol L 1993, 14:361–88. [76] Jafari M, Tavili A, Zargham N, Heshmati GA, Zare Chahouki M, Shirzadian S, Sohrabi M. Comparing some properties of crusted and uncrusted soils in Alagol Region of Iran. Pakistan J Nut 2004, 3:273–7. [77] Lázaro R, Cantón Y, Solé-Benet A, Bevan J, Alexander R, Sancho LG, Puigdefábregas J. The influence of competition between lichen colonization and erosion on the evolution of soil surfaces in the Tabernas badlands (SE Spain) and its landscape effects. Geomorphology 2008, 102:252–66.

94 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

[78] Martínez I, Escudero A, Maestre F. Small-scale patterns of abundance of mosses and lichens forming biological soil crusts in two semi-arid gypsum environments. Aust J Bot 2006, 54:339–48. [79] Meyer SE. The ecology of gypsophile endemism in the Eastern Mojave Desert. Ecology 1986, 67:1303–13. [80] Rajvanshi F, St. Clair LL, Webb BL, Newberry CC. The terricolous lichen flora of the San Rafael Swell, Emery County, Utah, U.S.A. In: Glenn M, Cole M, Dirig R, Harris R (eds). Lichenographia Thomsoniana: North American lichenology in honor of John W. Thomson. Ithaca, New York, USA, Mycotaxon, LTD, 1998, 399–406. [81] Zander RH, Stark LR, Marrs-Smith G. Didymodon nevadensis, a new species for North America, with comments on phenology. Bryologist 1995, 98:590–5. [82] Bogdanović M, Sabovljević M, Sabovljević A, Grubišić D. The influence of gypsiferous substrata on bryophyte growth: are there obligatory gypsophilous bryophytes? Botan Serbica 2009, 33:75–82. [83] Palacio S, Aitkenhead M, Escudero A, Montserrat-Martí G, Maestro M, Robertson AHJ. Gypsophile chemistry unveiled: Fourier transform infrared (FTIR) spectroscopy provides new insight into plant adaptations to gypsum soils. PLoS ONE 2014, 9:e107285. [84] Concostrina-Zubiri L, Pescador DS, Martínez I, Escudero A. Climate and small scale factors determine functional diversity shifts of biological soil crusts in Iberian drylands. Biodivers Conserv 2014, 23:1757–70. [85] Belnap J, Welter W, Grimm NB, Barger NN, Ludwig JA. Linkages between microbial and hydrologic processes in arid and semiarid watersheds. Ecology 2005, 86:298–307. [86] Li XR, Zhang P, Su YG, Jia RL. Carbon fixation by biological soil crusts following revegetation of sand dunes in arid desert regions of China: a four-year field study. Catena 2012, 97:119–26. [87] Porada P, Weber B, Elbert W, Poscl U, Keidon A. Estimating impacts of lichens and bryophytes on global biogeochemical cycles. Global Biogeochem Cycles, 2013, 28:71–85. [88] Elbert W, Weber B, Burrows S, Steinkamp J, Budel B, Andreae M, Poschl U. Controbutions of cryptogamic covers to the global cycles of carbon and nitrogen. Nat Geosci 2012, 5:459–462. [89] Vitousek PM, Howart RW. Nitrogen limitation on land and in the sea: how can it occur? Biogeochemistry 1991, 13:87–115. [90] Bowker MA, Belnap J, Davidson DW. Microclimate and propagule availability are equally important for rehabilitation of dryland N-fixing lichens. Restor Ecol 2010, 18:30–33. [91] Rousk J, DeLuca TH, Rousk J. The cyanobacterial role in the resistance of feather mosses to decomposition – toward a new hypothesis. PLOS One, 2013, 4:e62058. [92] Delgado-Baquerizo M, Gallardo A, Covelo F, Prado-Comesaña A, Ochoa V, Maestre FT. Differences in thallus chemistry are related to species-specific effects of biocrust-forming lichens on soil nutrients and microbial communities. Func Ecol 2015, 29:1087–98. [93] Delgado-Baquerizo M, Morillas L, Maestre FT, Gallardo A. Biocrusts control the nitrogen dynamics and microbial functional diversity of semi-arid soils in response to nutrient additions. Plant Soil 2013, 372:643–54. [94] Evans RD, Erlinger JR. A break in the nitrogen cycle in Aridlands? Evidence from δ15N of Soils. Oecologia, 1993, 94:314–7. [95] Chaudhary VB, Bowker MA, O’Dell TE, Grace JB, Redman AE, Johnson NC, Rillig MC. Untangling the biological controls on soil stability in semi-arid shrublands. Ecol Appl 2008, 40:2309– 2316. [96] Eldridge DJ, Leys JF. Exploring some relationships between biological soil crusts, soil aggregation and wind erosion. J. Arid Environ 2003, 53:457–66.

References |

[97]

[98]

[99] [100] [101]

[102]

[103] [104] [105] [106] [107]

[108] [109] [110]

[111]

[112]

[113]

[114]

[115]

95

Rodríguez-Caballero E, Aguilar MA, Castilla YC, Chamizo S, Aguilar FJ. Swelling og biocrusts upon wetting induces changes in surface microtopography. Soil Biol Biochem 2015, 82:107–11. Stark LR, Brinda JC, McLetchie DN, Oliver MJ. Extended periods of hydration do not elicit dehardening to desiccation tolerance in regeneration trials of the moss Syntrichia caninervis. Int J Plant Sci 2012, 173:333–343. Kranner I, Beckett R, Hochman A, Nash TH. Desiccation tolerance in lichens: a review. Bryologist, 2008, 111:576–93. Tighe M, Harling RE, Flavel RJ, Young IM. Ecological succession, hydrology and carbon acquisition of biological soil crusts measured at the micro-scale. PloS One 2012, 7:e48565. Chamizo S, Cantón Y, Lazaro R, Sole-Benet A, Domingo F. Crust composition and disturbance drive infiltration through biological soil crusts in semiarid systems. Ecosystems 2012, 15:148– 61. Michel P, Payton IJ, Lee WG, During HJ. Impact of disturbance on above-ground water storage capacity of bryophytes in New Zealand indigenous tussock grassland ecosystems. N Zeal J Ecol 2013, 37:114–36. Belnap J. The potential roles of biological soil crusts in dryland hydrologic cycles. Hydrol Process, 2006, 20:3159–78. Chamizo S, Cantón Y, Rodríguez-Caballero E, Domingo F. Biocrusts positively affect the soil water balance in semiarid ecosystems. Ecohydrology. 2016, 9:1208–21. Kidron GJ, Monger HC, Vonshak A, Conrad W. Contrasting effects of microbiotic crusts on runoff of desert surfaces. Geomorphology 2012, 139:484–94. Darby BJ, Neher DA, Belnap J. Impact of biological soil crusts and desert plants on soil microfaunal community composition. Plant Soil, 2010, 328:421–31. Baran R, Brodie EL, Mayberry-Lewis J, Hummel E, Da Rocha UN, Chakraborty R, Bowen BP, Karaoz U, Cadillo-Quiroz H, Garcia-Pichel F, Northen TR. Exometabolite niche partitioning among sympatric soil bacteria. Nat Comm 2015, 6:doi:10.1038/ncomms9289. Xie CF, Lou HX. Secondary metabolites in bryophytes: An ecological aspect. Chem Biodiv 2009, 6:303–12. Solhaug KA, Gauslaa Y, Nybakken L, Bilger W. UV-induction of sunscreen pigments in lichens. New Phytol, 2003, 158:91–100. Büdel B, Karsten U, Garcia-Pichel F. Ultraviolet-absorbing scytonemin and mycosporine-like amino acid derivates in exposed, rock-inhabiting cyanobacterial lichens. Oecologia 1997, 112:165–72. Hooper DU, Chapin FSI, Ewel JJ, Hector A, Inchausti P, Lavorel S, Lawton JH, Lodge DM, Loreau M, Naeem S, Schmid B, Setala H, Symstad AJ, Vandermeer J, Wardle DA. Effects of biodiversity on ecosystem functioning: a consensus of current knowledge. Ecol Monogr 2005, 75:3–35. Cardinale BJ, Duffy JE, Gonzalez A, Hooper DU, Perrings C, Venail P, Narwani A, Mace GM, Tilman D, Wardle DA, Kinzig AP, Daily GC, Loreau M, Grace JB, Larigauderie A, Srivastava DS, Naeem S. Biodiversity loss and its impact on humanity. Nature 2012, 486:59–67. Cardinale BJ, Matulich KL, Hooper DU Byrnes JE, Duffy E, Gamfeldt L, Balvanera P, O’Connor MI, Gongalez A. The functional role of producer diversity in ecosystems. Am J Bot 2011, 98:572– 92. Lefcheck JS, Byrnes JE, Isbell F, Gamfeldt L, Griffin JN, Eisenhauer N, Hensel MJS, Hector A, Cardinale BJ, Duffy JE. Biodiversity enhances ecosystem multifunctionality across trophic levels and habitats. Nat Commun 2015, 6:6936. Bowker MA, Maestre FT, Escolar C. Biological crusts as a model system for examining the biodiversity-function relationship in soils. Soil Biol Biochem 2010, 42:405–17.

96 | 5 Bryophyte and Lichen Diversity on Arid Soils: Determinants and Consequences

[116] Bowker MA, Maestre FT, Mau RL Diversity and patch-size distributions of biological soil crusts regulate dryland ecosystem multifunctionality. Ecosystems 2013, 16:923–33. [117] Maestre FT, Escudero A, Martínez I, Guerrero C, Rubio R. Does spatial pattern matter to ecosystem functioning? Insights from biological soil crusts. Func Ecol 2005, 19:566–73. [118] Mulder CP, Uliassi DD, Doak DF. Physical stress and diversity-productivity relationships: the role of positive interactions. Proc Natl Acad Sci 2001, 98:6704–8. [119] Rixen C, Mulder CPH. Improved water retention links high species richness with increased productivity in arctic tundra moss communities. Oecologia 2005, 146:287–99. [120] Hu C, Liu Y, Song L, Zhang D. Effect of desert soil algae on the stabilization of fine sands. J Appl Phycol 2002, 14:281–92. [121] Castillo-Monroy AP, Bowker MA, Maestre FT, Rodríguez-Echeverría S, Martinez I, BarrazaZepeda CE, Escolar C. Relationships between biological soil crust, bacterial diversity and abundance and ecosystem functioning: Insights from a semi-arid Mediterranean environment. J Veg Sci 2011, 1:165–74. [122] Pasari JR, Levi T, Zavaleta ES, Tilman D. Several scales of biodiversity affect ecosystem multifunctionality. Proc Nat Acad Sci 2013, 110:10219–22. [123] Tilman D, Isbell F, Cowles JM Biodiversity and ecosystem functioning. Annu Rev Ecol Evol Syst 2014, 45:471–93. [124] Venail P, Gross K, Oakley TH, Narwani A, Allan E, Flombaum P, Isbell F, Joshi J, Reich PB, Tilman D, van Ruijven J, Cardinale BJ. Species richness, but not phylogenetic diversity, influences community biomass production and temporal stability in a re-examination of 16 grassland biodiversity studies. Funct Ecol 2015, 29:615–26. [125] Wilsey BJ, Polley HW. Realistically low species evenness does not alter grassland speciesrichness–productivity relationship. Ecology 2004, 85:2693–700. [126] Maestre FT, Castillo AP, Bowker MA, Ochoa-Hueso R. Species richness and composition are more important than spatial pattern and evenness as drivers of ecosystem multifunctionality. J Ecol 2012, 100:317–30. [127] Castillo-Monroy AP, Bowker MA, García-Palacios P, Maestre FT. Aspects of soil lichen biodiversity and aggregation interact to influence subsurface microbial function. Plant Soil 2015, 386:303–16. [128] Bowker MA, Mau RL, Maestre FT, Escolar C, Castillo AP. Functional profiles reveal unique ecological roles of various biological soil crust organisms. Funct Ecol 2011, 25:787–95. [129] Büdel B, Vivas M, Lange OL. Lichen species dominance and the resulting photosynthetic behaviors of Sonoran Desert soil crust types (Baja California, Mexico) Eco Proc 2012, 2:6. [130] Walker BH. Biodiversity and functional redundancy. Cons Bio 1992, 6:18–23. [131] Naeem S. Species redundancy and ecosystem reliability. Cons Bio 1998, 12:39–45. [132] Bowker MA, Maestre FT, Eldridge DJ, Belnap J, Castillo-Monroy AP, Escolar C, Soliveres S. Biological soil crusts (biocrusts) as a model system in community, landscape, and ecosystem ecology. Biodivers Conserv 2014, 23:1619–37. [133] Gotelli NJ, Ulrich W, Maestre FT. Randomization tests for quantifying species importance to ecosystem function. Methods Ecol Evol 2011, 2:634–642. [134] Cornelissen JHC, Lang SI, Soudzilovskaia NA, During HJ. Comparative cryptogam ecology: a review of bryophyte and lichen traits that drive biogeochemistry. Ann Bot-London 2007, 99:987–1001. [135] Castillo-Monroy AP, Bowker MA, García-Palacios P, Maestre FT. Aspects of lichen biodiversity and aggregation interact to influence subsurface microbial function. Plant Soil 2014, 386:303–16.

Andrea Porras-Alfaro, Cedric Ndinga Muniania, Paris S. Hamm, Terry J. Torres-Cruz, and Cheryl R. Kuske

6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils Desert ecosystems represent a rich reservoir of unexplored fungal diversity with complex assemblages of microbial communities. Deserts are considered one of the most hostile habitats for life on Earth [1, 2]. They encompass extreme conditions for life, including drastic changes in temperature, high ultra violet and infrared radiation, low moisture availability, long periods of dryness, low nutrient availability, and osmotic stress [3, 4]. All these characteristics require organisms with specific adaptations to survive in this intense and variable environment [5–7]. Fungi in these areas include a high number of taxa with hyaline and melanized hyphae that inhabit rock surfaces, biocrusts, rhizosphere soils, and plant tissues (󳶳 Fig. 6.1) [3, 6, 8, 9]. Taxa with melanized hyphae are known as dark septate fungi (DSF) (󳶳 Fig. 6.2a,b). Dark septate fungi (DSF) are a nonmonophyletic group of fungi that includes a diverse taxonomic assemblage within Ascomycota. Orders such as Pleosporales, Sordariales, Capnodiales, Xylariales, Helotiales, and Hypocreales include a number of DSF commonly isolated from multiple substrates in deserts including soils and plants [10]. Dark septate fungi are dominant inside plant tissue as endophytes, on the surface of rocks, and in biocrusts, a microbial community composed of algae, cyanobacteria or moss together with fungi, bacteria, and archaea [3, 11]. They are also considered as being of special interest in the medical field because they are allergens and cause pulmonary and skin diseases in immunocompromised and healthy individuals [12]. A majority of fungi in arid lands grow as asexual forms (mitosporic) or as sterile mycelia (󳶳 Fig. 6.2) and are thus difficult to characterize, but advances in molecular techniques and the low cost of sequencing have recently allowed large surveys in these areas, showing important potential for the description of novel taxa [8, 9, 13–16]. This chapter focuses on the description of fungal diversity in the different microenvironments characteristic of arid lands. We will discuss their roles as plant and biocrust symbionts, their function in nutrient cycling, their responses to climate and land use changes, and their potential as pathogens in humans.

6.1 Spatial Heterogeneity of Fungal Communities in Arid Lands The sparse distribution of plants and biocrusts in arid ecosystems creates a series of microenvironments in which fungi can be supported by the photosynthetic products and organic matter in zones where primary producers are present (i.e., islands of DOI 10.1515/9783110419047-006

98 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

(a)

(b)

(c)

(d)

(e)

(f)

(g)

6.1 Spatial Heterogeneity of Fungal Communities in Arid Lands |

99

󳶣 Fig. 6.1: Diverse microenvironments for fungal communities in desert ecosystems. (a) Coleogyne ramosissima (blackbrush) in a lichen dominated biocrust, (b) grasses and cyanobacteria dominated biocrust, (c) lichen dominated biocrust in gypsum soils, (d) desert varnish, (e) patchy distribution of plant communities, (f) lichen dominated biocrust, (g) moss dominated biocrust.

(a)

(b)

(c)

(d)

(e)

(f)

Fig. 6.2: Common fungi in arid systems. (a) Dark septate endophyte colonizing a grass root, (b) dark septate endophyte on root surface, (c) ectomycorrhizal fungi in piñon pine roots, (d) arbuscular mycorrhizal fungus, (e) microcolonial fungi inside pits on rock surface, scale bar 200 μm [5], (f) keratinophilic bait from soil using sterile snake skin.

100 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

fertility)(󳶳 Fig. 6.1) [17]. Biocrusts and rhizosphere zones account for the highest diversity of fungi in arid lands [8, 9, 15, 18, 19], but other communities are found in more extreme conditions such as desert varnish and gypsum deposits [5, 20, 21]. Distinct fungal communities in deserts are supported by the high heterogeneity created by the combination of seasonal climate, variable distribution of nutrients and water, and a mosaic of microenvironments [8, 17, 22].

6.1.1 Biocrusts Biocrusts, also known as biological soil crusts or microbiotic crusts, are prominent features of desert ecosystems (󳶳 Fig. 6.1). Biocrusts can cover up to 70% of the ground in some deserts [23]. This common arid microenvironment supports large microbial communities that involve a photosynthetic component (algae, cyanobacteria, or moss) combined with a microbial mat of fungi, archaea, and other bacteria; in which the bacterial biomass is 50–500 fold higher than the biomass of surrounding noncrusted soils [24, 25]. Biocrusts are classified by their color and texture or by the communities of microorganisms found in them [24, 26]. The darker crusts are dominated by cyanolichens and mosses (󳶳 Fig. 6.1a, c,f-g), and light crusts include cyanobacteria such as Microcoleus vaginatus (󳶳 Fig. 6.1b). The structure of microfungal communities in biocrusts is influenced by the photosynthetic partner and has shown large spatial heterogeneity from small areas to large regional scales (󳶳 Fig. 6.3a) [19, 25, 27]. Fungi show very patchy distributions even at the millimeter scale with high hyphal density areas while other areas lack hyphal components [24]. The patchy distribution has been confirmed using molecular methods in which comparison of biocrusts in close proximity show high variation and little overlap in terms of their fungal community composition (󳶳 Fig. 6.3a) [16]. Diversity studies on biocrusts reveal abundance of different fungi that rank from 40–106 species using a combination of cultured based techniques and molecular markers (mainly based on Sanger sequencing and DGGE bands). The most abundant genera within Ascomycota, the dominant phylum, include taxa such as Alternaria, Acremonium, Chaetomium, Phoma, Preussia, Stachybotrys, and Ulocladium [15, 18, 24, 27]. Many species within these genera are considered pathogens and decomposers that likely benefit from the carbon and nitrogen fixed by the photosynthetic partners. Steven et al. [15] reported at least 78 unique OTUs (operational taxonomic units) using cloning and sequencing of the LSU (large subunit) in biocrusts from Utah, USA. Culture based studies have reported 71 species and 48 genera in the western Negev Desert in Israel [27]. A recent study using 454 Titanium sequencing of biocrusts showed a slightly larger diversity than previously reported for biocrusts (140–228 OTUs for the LSU rRNA region) [16]. Next generation sequencing techniques facilitate the detection of larger numbers of taxa, the comparison of studies, and the determination of potential culture based bias toward fast growing fungi.

6.1 Spatial Heterogeneity of Fungal Communities in Arid Lands

A. Sand crusts 108

B. Shale crusts 121

88

39 (36%) 109

83

40 50 (37%) 22 (41%) 25 21 24

101

C. Between sand and shale

79

41 46 (52%) 12 (52%) 18 12 8 45 (54%)

|

Sand

210 (66%)

317

Shale

107

136 (56%)

243

Percent of shared OTUs

D. Taxonomic composition of conserved OTUs 100 80 Unclassified Fungi

60

Chytriomycota Basidiomycota

40

Unclassified Ascomycota Rare Ascomycota

20

Dothideomycetes

0

Sand (25 OTUs)

Shale (18 OTUs)

Sand and Shale (107 OTUs)

E. Taxonomic distribution of root-associated fungi CL1–N–64 CL2–N–67 CL3–N–60 CL4–C–50 CL5–C–43 CL6–C–77 CL7–N–22 CL8–N–23 CL9–N–20 CL10–C–17 CL11–C–21 CL12–C–28 CL13–N–26 CL14–N–27 CL15–N–24 CL16–C–22 CL17–C–21 CL18–C–29 0%

20% Pleosporales Phallales Onygenales

40% Agaricales Hypocreales Pezizales

60% Xylariales Glomerales unknown

80% 100% Sordariales Halosphaseriales

Fig. 6.3: Fungal diversity in the biological soil crust of the Colorado Plateau. (a–c) Shared OTUs for different replicate samples showing little overlap among fungal communities and large spatial heterogeneity. (d) Taxonomic composition of shared OTUs showing dominance of Dothideomycetes and a large number of unclassified fungi at this site. (e) Dominance of Pleosporales (Dothideomycetes) is also observed in individual plants (each bar) of Bouteloua gracilis in a semiarid grassland. Modified from [9, 16].

102 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

Dominance by dark septate fungi ranges from 83–98%, including abundant taxa within the Dothideomycetes, Sordariomycetes, Eurotiomycetes, and the Pezizomycetes (󳶳 Fig. 6.3a) [14, 15, 18, 24, 27]. Dominant taxonomic groups are consistent across culture based and molecular studies using different techniques such as DGGE, Sanger sequencing, and 454-Titanium sequencing. Pleosporales is the dominant fungal order in arid land biocrusts, in some cases representing up to 92% of the sequences [16, 18, 19], making this order one of the most important groups in terms of abundance and diversity in biocrusts. Specific areas, such as the Chihuahuan desert, report larger numbers of undescribed taxa within this order with little similarity to known fungi, illustrating how incomplete the fungal diversity from these systems is represented in curated databases [14, 18]. The large number of undescribed taxa opens new opportunities for the description and characterization of new species. For example, Knapp et al. [13] recently described three new genera and five new species within the order Pleosporales from a semiarid region. Other fungal phyla such as Basidiomycota and lower lineages of fungi, including zygomycetes (mainly Mortierellales) and chytridiomycetes, are present in biocrusts in a smaller proportion (< 1−20%). Agaricomycetes are dominant within Basidiomycota represented by taxa in the orders Agaricales, Cantharellales, Corticales, Polyporales, and Tremellales, including several yeast species [19]. Many of these fungal orders include plant pathogens, decomposers, and important mycorrhizal fungi. Lichenized fungi are also common in arid soils, even in cases when lichens are hard to distinguish from cyanobacterial dominated biocrusts [14, 16, 28]. Lichens are discussed in detail in Chapter 5 in this book. Within the basal lineages of fungi, Mortierella alpina seems to be quite common across different types of biocrusts [14, 29], and reports of chytrids using molecular methods shows great potential for the description of new species [16, 18]. Dominant fungi in biocrusts have adapted to the harsh conditions on the surface soil, including high UV radiation, high temperatures during the summer, and extremely limited water. Their melanized hyphae not only protects them against these conditions, but likely provides protection to cyanobacteria, algae, and other microorganisms in the biocrust [3]. It is possible that hyphal mats may also play a role in stabilizing the soil surface and limiting erosion in arid lands [3]. Fungi associated with different types of biocrusts affect nutrient availability through decomposition and transfer of nutrients with nearby grasses [30]. Fungal hyphae have been observed in direct contact with clusters of Microcoleus vaginatus, the dominant cyanobacteria in biocrusts [24]. Rhizosphere soils and biocrusts share a great proportion of specific fungal taxa [15, 18], and the overlaping fungal communities in these different patches are relevant to the support of fungal networks (also referred to as fungal loops) [17] that facilitate the interchange of nutrients between the biocrusts and rhizosphere zones. Green et al. [30] showed that grasses and biocrusts transport N (and C) through fungal networks. In this trace element study, 15 N was translocated from biocrusts and grasses at rates of up to 100 cm/day [30].

6.1 Spatial Heterogeneity of Fungal Communities in Arid Lands |

103

Microbial communities in the biocrusts are highly sensitive to changes in precipitation regimes with dramatic reductions in biocrust cover with altered precipitation patterns [15, 31, 32], but additional data needs to be collected to determine potential effects of changing climate on the structure of their fungal communities. Biocrusts show great potential for conducting simple and low cost manipulations in the field [15, 33]. Their distribution and spatial heterogeneity facilitate the establishment of studies in microbial diversity, biogeography, and responses to climate change [31].

6.1.2 Plant Associated Fungi in Deserts In addition to biocrust fungi, plant associated fungal communities (rhizosphere, mycorrhizal fungi, and endophytes) represent very important habitats for fungal diversity in arid lands (󳶳 Fig. 6.2). Plant associated fungi include taxa in every fungal phylum and represent multiple ecological strategies varying from mutualists, commensalists, pathogens, and saprobes. The fungal colonizers inside roots, stems, leaves, and seeds include more specialized community of fungi [9, 18, 34, 35], such as mycorrhizal and nonmycorrhizal species with large colonization rates by endophytic dark septate fungi [9, 35, 36]. Biocrusts and rhizosphere soils share an important proportion of fungal taxa. The structure of their fungal communities differs but dominant colonizers are frequently detected in both microenvironments [15, 18]. As in biocrusts, rhizosphere fungal communities are influenced by the presence of organic matter, nutrients, season, precipitation, and levels of CO2 [15, 37–41]. Ascomycota fungi are dominant (68–88%) in rhizosphere soils with lower and variable proportions of Chytridiomycota, Blastomycotina, Mucoromycotina, and Mortierellomycotina (< 1–31%) [15, 18, 22, 37]. Dothideomycetes, Eurotiomycetes, Leotiomycetes, and Sordariomycetes, all classes within Ascomycota, are common [8, 15]. In the shrub Larrea tridentata (creosote) in the Mojave desert, Dothideomycetes within the order Pleosporales were abundant [15, 40]. Similar proportions of dominant taxa at the class and order levels are consistent in multiple studies including arid grasslands in New Mexico, USA [18, 42] and are associated with plants in the family Asteraceae in a semiarid grassland in Europe [43]. Hudson et al. [22], using a metagenomic approach for rhizosphere soils in a semiarid grassland in New Mexico, also detected high proportions of Ascomycota (65%) with important contributions of Basidiomycota (30.9%) and arbuscular mycorrhizal fungi (AMF, 5.4%), which are more difficult to detect using conventional PCR based approaches [22]. 6.1.2.1 Mycorrhizal Fungi Mycorrhizal colonization in arid lands is not as abundant in comparison to more mesic environments but is still an important component of arid land fungal diversity [42,

104 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

44, 45]. Mycorrhizal fungi have important roles in the acquisition of nutrients, such as nitrogen and phosphorus. They facilitate the attachment of plant roots to the soil, access to water, and other essential nutrients [46, 47]. The stressful conditions of arid ecosystems favor two main groups of mycorrhizal fungi: arbuscular mycorrhizal fungi (AMF) and ectomycorrhizal fungi (EMF)(󳶳 Fig. 6.2c,d). 6.1.2.2 Arbuscular Mycorrhizal Fungi Represented by species in the phylum Glomeromycota, AMF are the most common plant symbionts found in about 80% of vascular plants (󳶳 Fig. 6.2d) [48, 49]. AMF play major roles in the establishment of plant communities in low-nutrient arid land soils by facilitating nutrient absorption, water uptake, and soil stabilization [48, 50, 51]. Though not as diverse and abundant as in other ecosystems, such as temperate forests, AMF communities in arid ecosystems portray some level of species richness and varying levels of colonization on plants. For example, general estimates of AMF biomass abundance in plants range from 4 g m−2 in deserts in comparison to 44 g m−2 in temperate grasslands [52]. In terms of species diversity, AMF taxa defined based on SSU rRNA analyses revealed lower numbers of AMF (27 taxa) for desert environments in comparison to temperate broadleaf mixed forests (82 taxa), temperate seminatural grasslands (90 taxa), and subtropical savannas and grasslands (43 taxa). Diversity was comparable or higher in deserts with respect to boreal forests (12 taxa), subtropical dry broadleaf forests (18 taxa), and temperate coniferous forests (12 taxa) [53]. The differences in diversity may be a result of the low number studies available for deserts that are poorly represented in molecular curated databases and the techniques used to detect these fungi in the environment. For example, the use of next generation sequencing has helped reveal an abundance of AMF fungi in piñon pine, which was considered primarily colonized by ectomycorrhizal fungi in juniper-piñon woodland in New Mexico [54]. The order Glomerales with Glomus group A is the dominant cluster of species [44]. Other dominant genera include Claroideoglomus and Scutellospora [44, 51, 55]. The orders Archaeosporales and Diversisporales are represented by genera such as Archaeospora, Diversispora, and Acaulospora but colonization levels are low [51]. In arid lands, AMF colonization rates vary greatly for different sites. Some fungi unique to desert ecosystems have relatively high colonization rates varying from 37 to 95% depending on their location, nutrient availability, and environmental conditions [44, 51, 55], while some grasses showed very low colonization rates [35, 45, 56] AMF nutrient acquisition and survival is highly dictated by water availability at these sites. The diversity and rates of root colonization by AMF tend to decrease with dryness but hyphae can survive for long periods under dry conditions [55, 57]. For some AMF, such as Acacia laevis and Scutellospora calospora, infectivity during the dry season also depends on the time of sporulation. The hyphae of A. laevis have the

6.1 Spatial Heterogeneity of Fungal Communities in Arid Lands |

105

capacity to infect plants for 11 weeks in dry soils if they did not receive water before sporulation started [55]. In addition to season, plant diversity and plant ecophysiological adaptations to stressful conditions create abiotic constraints that dictate the composition and growth of AMF communities [58]. Plants such as Atriplex halimus, a common plant of arid and semiarid regions, excretes salt as an adaptation to this stressful environment [59]. Thus, salt tolerant fungi dominate the diversity of AMF in A. halimus. Also, particular vegetation in areas with a high level of gypsum (gypsophytes) tends to present unique AMF structures in Glomus species that are specific for these sites [20]. 6.1.2.3 Ectomycorrhizal Fungi (EMF) Represented by species in the phyla Basidiomycota and Ascomycota, EMF are essential for desert trees and flowering plants [60, 61]. Ectomycorrhizal fungi link plant roots to the soil and surrounding plant communities, increasing nutrient efficiency in an environment with low nutrient quality and in some areas with high soil toxicity [62]. The most common type of basidiomycetes collected in these areas include Amanita species such as A. rubescens, A. citrina, and A. muscaria; Hebeloma species such as H. sinapizans and H. crustuliniforme; Laccaria laccata; Paxillus involutus, and Russula vesca [62]. Using 454-Titanium sequencing, Dean et al. [54] also reported a diverse assemblage of genera in piñon-juniper woodlands in New Mexico, including Cenococcum, Inocybe, Tricholoma, Rhizopogon, and Geopora, showing the potential of next generation sequencing for the documentation of ectomycorrhizal fungi in these poorly studied sites (󳶳 Fig. 6.2c) [54]. Mycoheterotrophic plants such as desert orchids are nonphotosynthetic plants that obtain all their nutrients, including carbon, from fungi rather than photosynthesis [63], They are also dependent on ectomycorrhizal networks for their survival. Fungi associated with desert mycoheterotrophs belong to the class Agaricomycetes, with Russulales, Sebacinales, and Boletales being the most common orders and Rhizopogon and Sebacina being the most common genera [64, 65]. Other mycorrhizal communities include desert truffles. They constitute a diverse group of hypogeous ectomycorrhizal fungi also known as turma [60, 61] and play a major role in maintaining certain plant communities in arid lands [61]. Desert truffles include species in the genus Terfezia, Tirmania, Picoa, and Balsamia, and mainly colonize the roots of plants in the family Cistaceae, known as rockroses, such as Cistus, Tuberaria, and Helianthemum [66–68]. Because of their adaptations to stressful conditions in arid ecosystems, they are spread worldwide with a higher number of reports in well studied sites in the Middle East, the Mediterranean basin, the African Kalahari, and the Australian desert [7, 60]. In these regions truffles also have economic importance in the food industry, where they are used as an expensive seasoning. The most commonly found species are Terfezia leptoderma, T. boudieri, T. claveryi, and Picoa lefebvrei [60, 61].

106 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

6.1.2.4 Nonmycorrhizal Fungi (Endophytes) Fungal endophytes have been recovered from leaves, stems, roots, and seeds of many species of arid plants. The term endophyte refers to fungi that inhabit plant tissues without causing any damage to their hosts [69, 70]. Root endophytes do not form the characteristic structures for nutrient transfer commonly observed in mycorrhizal fungi (i.e., vesicles, arbuscules, Hartig net, mantle). These plant-fungal associations occur with diverse species across all fungal phyla and are found in every studied plant across the globe [10, 69, 71]. In arid ecosystems, endophytes are important for nutrient transfer and plant survival because they provide protection against stressful conditions, such as drought and heat, but also against biotic factors such as herbivory [47, 69, 72]. Compared to other ecosystems, the diversity of fungal endophytes in arid lands is relatively low, but the rate of plant colonization can vary greatly among plant species [72–75]. Endophytes are phylogenetically diverse, showing important levels of novel species even at low colonization rates. An analysis of 22,000 plant segments from desert trees and shrubs showed colonization rates of 1–3.5% on stems and leaves with more than 60% of the isolates likely representing novel species [34]. Large numbers of potential novel species have also been recovered from roots in piñon-juniper woodlands [54] and grasses [9, 21, 35, 42, 44]. Root colonization rates in grasses are high (60–90%) with variation among plant species and tissue types (aboveground vs. belowground communities) [9, 21, 35, 42]. Dominant taxa in roots are similar to those observed in rhizosphere and biocrust soils, including many Dothideomycetes, Eurotiomycetes, Sordariomycetes, and a proportion of Basidiomycota mainly within Agaricomycetes (󳶳 Fig. 6.3e). Species such as Alternaria, Fusarium, Aspergillus, Chaetomium, Preussia, Monosporascus, Darksidea, and Moniliophthora appeared to be generalists, isolated from diverse plant species and tissues [10, 13, 35]. Other species such as Phoma pomorum show higher levels of specificity for specific tissues such as stems and leaves [72], resulting in more selective endophytic communities [13, 34]. Unlike mycorrhizal fungi, the functions of nonmycorrhizal fungi (endophytes and other rhizosphere associated fungi) are not well defined. Their ecological roles likely vary based on tissue, environmental factors, and host; ranging from mutualists to plant pathogens to saprobes [69]. For example, species of the genera Olpidium, Monosporascus, and Moniliophthora are well known plant pathogens, but are usually abundant in association with healthy roots of desert plants, mainly from the family Poaceae (󳶳 Fig. 6.3e) [9, 35, 42, 66]. Coprophilous fungi traditionally found in animal dung have also been recovered from arid land grasses [9]. Herrera et al. [76] suggested a potential link between the endophytic and coprophilic life stages in which the fungi are ingested by animals as plant endophytes and they continue as coprophiles once excreted. Among the different types of endophytes in arid lands, dark septate fungi are considered to be the most dominant, in some cases exceeding the abundance of AMF

6.2 Roles in Nutrient Cycling and Effects of Climate Change on Fungal Communities | 107

(󳶳 Fig. 6.2a,b) [9, 10, 35, 44]. Melanized septate hyphae are normally observed inside root tissue with the formation of microsclerotia (󳶳 Fig. 6.2a) and intercellular and intracellular colonization (󳶳 Fig. 6.2b) [9, 42, 56, 77]. Colonization is more common in the root cortex with extraradical mycelium spreading from the intercellular spaces in the roots into the soil [56]. Functional roles for the majority of DSF are still unclear, but fungal inoculation experiments in several plant species reveal the potential to increase plant thermotolerance and survival under drought conditions. Some species of Curvularia have been reported to confer thermotolerance to plants [78, 79]. A Paraphaeosphaeria quadriseptata isolate from a Sonoran desert cactus provides protection to model plants such as Arabidopsis thaliana to lethal temperatures through regulation of heat shock proteins [47]. This genus is also one of the most common taxa recovered from grasses such as Bouteloua gracilis, B. eriopoda, among others [9, 74] More specialized communities of endophytes in desert ecosystems include fungi in gypsum deposits or very specialized environments, like the Caatinga deserts in Brazil. With a worldwide coverage over 100 million ha, gypsum soils represent another specialized ecosystem in arid and semiarid regions with low annual precipitation and large numbers of endemic plant species (󳶳 Fig. 6.1) [21, 80]. Gypsum soils are characterized by high concentrations of calcium sulfate (CaSO4 ), low nutrient content, and low porosity. Thus, gypsophiles and gypsovags, the most common type of plants found in gypsums, have unique mycorrhizal and endophytic communities [81, 82]. Colonization rates vary widely among different plant tissues and species endemic to gypsum soils [21, 80, 83]. The variation of endophytic and mycorrhizal communities is likely correlated with the physiological and ecological demands of the plants as a response to stressful conditions of this environment. Commonly isolated genera from healthy plant tissues include Alternaria, Sporormiella, Phoma, Fusarium, Rhizoctonia, Epicoccum, Pleospora, and Cladosporium [21, 82]. Other specialized endophytic communities have been identified in the Caatinga deserts in Brazil. The dominant type of desert vegetation in this area includes cacti, shrubs and thorny trees, as well as arid grasses [84]. Species of Penicillium and Aspergillus are, common and unique species for these areas have been described, including A. caatingaensis and A. pernambucoensis. Other unique Neosartorya species include N. indohii, N. paulistensis, N. takakii, N. tatenoi, N. tsurutae, and N. udagawae [84, 85].

6.2 Roles in Nutrient Cycling and Effects of Climate Change on Fungal Communities Arid lands are characterized by low soil N content and are more responsive to low N input as a result of anthropogenic deposition [86]. Fungal interactions and responses

108 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

to N and C additions are diverse and complex. Two decades ago the biotic component of the global N cycle was attributed only to bacterial metabolism. Today we know that fungi have a fundamental role in N transformations in arid soils. Fungi are capable of dissimilatory nitrate reduction with production of NO, N2 O, and N2 [87, 88]. In arid lands fungi are resilient to N deposition in short and long term N deposition experiments, where little changes in diversity, community structure, and fungal biomass have been observed with respect to bacterial communities [8, 9, 18, 86]. The main C source for soil fungi is supplied by plants and cyanobacterial crusts [17, 30] and by the rapid turnover of soil proteins in arid lands [89, 90]. During periods of active growth, plant photosynthate may be translocated to biocrusts, the center of N-fixation [17]. Fungi account for a substantial fraction, even the majority, of N2 O production in arid land soils, since they can operate at low water potentials, and N2 O is the principal product of fungal mineralization of amino acids through denitrification via heterotrophic nitrifiers [87, 90]. In addition to their roles in nutrient cycling, fungi play important roles in decomposition processes that are highly regulated by abiotic factors. Photochemical oxidation (photodegradation) plays a major role facilitating the enzymatic oxidation processes carried out by bacteria and fungi [4, 91, 92]. Fungal communities that can tolerate high UV radiation and low moisture can quickly respond to the small pulses of water characteristic of arid environments. Fungi associated with plant litter consist of filamentous dark septate ascomycetes and yeasts. Gallo et al. [91] reported dominant communities of Sporiobolales, Coniochaetales, Cystofilobasidiales, and Pleosporales in litter of juniper and piñon in arid woodlands of New Mexico. In deserts, small mammals contribute to the accumulation of plant litter, allowing fungal communities to actively grow in a more humid environment with increased amounts of organic carbon. This higher level and movement of organic matter directly impacts the dispersal and structure of fungal communities, including specialized coprophilous fungi [76, 93, 94].

6.3 Extremophiles in Deserts Extremophilic fungi are those that can survive in conditions that are considered stressful or lethal for other organisms. As previously mentioned, fungi in deserts show adaptations to high UV radiation and low moisture, but in the mosaic of microenvironments there are even more specialized fungal communities exposed to higher selective pressures such as very high temperatures (40–70°C), and extremely low organic matter. We focus on two fairly well studied groups: thermophilic fungi and microcolonial fungi in rock varnish.

6.3 Extremophiles in Deserts | 109

6.3.1 Thermophilic and Thermotolerant Fungi Thermophilic fungi can grow in a range of temperatures between 40–50°C [95], with optimal growth at 45°C. Thermotolerant fungi include representatives that can grow between 40–50°C but their optimal growth temperature is at 25°C instead of 45°C [96, 97]. Unlike bacteria, Eukaryotes experience irreversible membrane damage above 65°C [95]. In desert ecosystems, these fungi can encounter conditions favorable for growth during the monsoon season, in which high temperatures will hold for long periods of time [96]. Thermophilic fungi reported in deserts include taxa within two major groups, the Ascomycota and Zygomycota (Mucoromycotina). Common orders of thermophiles in deserts include fungi within Sordariales, Eurotiales, and Mucorales [96]. Mucor miehei, M. thermohyalospora, Rhizomucor tauricus, R. pusillus, Talaromyces, Remersonia thermophila, and Stilbella thermophila are frequently reported in arid grasslands, as well as in many microenvironments in hot deserts [96]. Thermophilic fungi have been isolated from different substrates, including bulk soil, litter, animal dung, biocrusts, and rhizosphere soils [7, 96]. In Saudi Arabia up to 48 species of thermophilic and thermotolerant fungi were isolated from different types of desert soils, with two thirds of the species being thermotolerant and one third recognized as thermophiles [98]. Thermophilic fungi have also been studied from desert soils in Egypt, dominated by taxa such as Chaetomium thermophilum, Malbranchea pulchella var. sulfurea, Rhizomucor pusillus, Myriococcum albomyces, Talaromyces thermophilus, and Torula thermophila [99]. Powell et al. [96] showed that thermophiles vary seasonally in an arid grassland in New Mexico, with the highest number of propagules in summer and spring during the highest precipitation period. The amount of records for thermophilic fungi in desert soils is relatively limited despite their ubiquitous distribution based on recent reports [96]. This is likely due to the bias on isolation temperatures in culture based studies and the notion that fungal diversity in deserts is low [7, 98].

6.3.2 Rock Varnish and Microcolonial Fungi in Deserts In deserts, several organisms, including cyanobacteria, chlorophytes, fungi, mosses, heterotrophic bacteria, and lichens, can produce rock surface communities that are biologically active, forming thin and complex layers on the top few centimeters of rock surfaces [3]. These microcolonies can be found in association with specific mineral deposits known as rock varnish (󳶳 Fig. 6.1d, 󳶳 Fig. 6.2e). Rock varnish are present on rock surfaces [5] and are coatings mainly made of clays, oxides, hydroxides, manganese, and iron. They are found in deserts and semiarid regions all over the world. These dark coatings are hard and have a unique chemistry; they are usually black when they are

110 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

rich in iron and manganese, dark brown or pigmented opaline silica when rich in iron oxides, and can be red when deficient in manganese [5, 6]. The origin of rock varnish is not completely understood; it could be the result of abiotic processes, but it has also been suggested that their formation could be mediated by microorganisms that are commonly observed on these surfaces [5, 6]. Microcolonial fungi are the predominant biological organisms on desert varnish rock coatings; this fact has led researchers to study them as one of the forming agents of desert varnish (󳶳 Fig. 6.1d, 󳶳 Fig. 6.2e) [5, 6]. 6.3.2.1 Characteristics of Microcolonial Fungi Microcolonial fungi (MCF) have the ability to survive where other organisms are rarely found. They were first described in the Sonoran Desert by Perry and Adams in 1977, using scanning electron microscopy and morphological analysis [6]. Microcolonial fungi are globally distributed and have been reported in the Sonoran, Mojave, Gobi, Namib, Great Victoria, Gibson, Simpson, Arabian, and Nubian deserts [1, 6, 100], and in semiarid areas of the Mediterranean and the USA [7, 101]. These fungi form clusters on desert rocks and rock coatings of approximately 100 μm in diameter and have spheroidal subunits of approximately 5 μm in diameter with black or dark brown pigmentation [1, 6, 100]. These fungi are part of epi (surface) and endolithic (inside rock or in pores of mineral grains) communities and they can penetrate sedimentary soft rocks such as limestone, sandstone, and marble, and hard rocks such as granite and basalt [7]. One of the first reports on microcolonial fungi in deserts was published by Staley et al. [7] in 1982 on rocks collected in the western United States and Australia. The microcolonial structures were grown in the laboratory, obtaining slow growing fungal colonies that were mainly composed of a single isolate. The fungi on these rocks are metabolically active and have been referred to as blackberries and black globular units due to their color and shape [6]. Even though very limited morphological diversity has been observed, studies using DNA sequencing have shown high genus and species diversity within several orders of ascomycetes [7]. 6.3.2.2 Adaptations of Microcolonial Fungi Microcolonial fungi are recognized as one of the most stress tolerant eukaryotic organisms [7, 102]. Their colony morphology is thought to be a response to the environmental stressful conditions allowing for an optimal surface–volume ratio, decreasing water loss and reducing the fungal surface exposed to sun radiation and different stressors [7, 102]. Other factors of stress adaptation include the melanization of multilayered cell walls and the generation of trehalose to stabilize enzymes under desiccation [7, 101, 102]. It has been suggested that these fungi are chemoorganotrophs, since they rely on nutrients and carbon from external sources brought to the rock surface by the wind, like small particles of organic matter (e.g., pollen grains) [1, 6]. Micro-

6.4 Human Pathogenic Fungi in Desert Ecosystems

|

111

colonial fungi do not actively grow during hot periods regardless of the humidity but can survive for long periods under the severe desert conditions [100]. Pigments such as melanin, mycosporines, and carotenoids protect them from UV light [6, 101, 103], and their vegetative cells are highly stress tolerant and long living [6]. Colonies of these fungi produce large amounts of extracellular polymeric substances (EPS), which might provide protection from the sun [6, 7, 103], and can absorb water and hold it against the rocks for longer periods [3]. 6.3.2.3 Importance of Microcolonial Fungi Black microcolonial fungi are responsible for biological deterioration of marble and limestone monuments and statues, growing as a dark brown or black crust on their surfaces. They are considered one of the most damaging microorganisms in terms of the deterioration of monumental stones in all cities worldwide, not just arid lands. For example, a study by Marvasi et al. [104] characterized Sarcinomyces petricola as the yeast responsible for the dark spots found on two valuable statues (“Ratto delle Sabine” and “Copia del David”) located in the Piazza della Signoria in Florence, Italy. The study of these fungi is important in order to decide on proper procedures to restore and conserve monuments. Microcolonial fungi allow us to study the limits of life on Earth, evolution, and adaptation to extreme environmental conditions by eukaryotic organisms [105]. It is suspected that rock varnish coatings exist on Mars, and our understanding of how microcolonial fungi have developed several adaptations against harsh environmental conditions can provide good models to study rock coatings that can facilitate detection of life on other planets [6]. Studies of stress resistance by these fungi have provided promising results on their ability to survive space and Martian conditions [7, 102]. Cryptomyces antarticus (a cold desert microcolonial fungus) has even been shown to survive simulated Martian conditions and real space exposure [101, 105].

6.4 Human Pathogenic Fungi in Desert Ecosystems Arid soils are not immune to the ubiquitous distribution of fungal pathogens. In desert ecosystems, fungi reproduce mainly through asexual reproduction, creating large amounts of propagules or drought resistant spores that can be easily dispersed by wind, even at transcontinental distances [3]. Changes in climate and extreme droughts followed by dust storms and the increase in the number of infectious lung diseases have brought attention to the study of pathogenic fungi in desert ecosystems [106]. Opportunistic infections may occur in immunocompromised individuals due to a decreased ability to fight infections, such as those with HIV/AIDS or leukemia, in organ transplant patients, children, or the elderly.

112 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

6.4.1 Coccidioides immitis and C. posadasii From the family Onygenaceae, containing true human pathogens, the genus Coccidioides is of particular interest in desert ecosystems. This soil borne fungus, which reproduces using arthrospores, is endemic to arid regions of Mexico, Central and South America, and the southwestern United States [107]. Coccidioidomycosis, better known as Valley Fever, starts as a lung infection that can evolve into pneumonia and even become systemic and spread to other organs such as the skin, brain, and bones, and particularly endangers immunocompromised populations [108]. Outbreaks often occur among farmers and construction workers after dust storms [109] or earthquakes and during other events when the soil is disturbed [110, 111]. The CDC reported one of the overall highest incidences in 2011 with 42.6 cases per 100,000 people, with the largest number of cases among 60–79 year olds (69/100,000) in states where Valley Fever is endemic and has been reported (Arizona, California, Nevada, New Mexico, and Utah). The number of cases from 1998 to 2014 ranged from 2,271 to 22,641 [112]. The San Joaquin Valley in southern California is one of the most important endemic areas in the United States for Coccidioides immitis. The more prevalent Coccidioides posadasii has been detected across the southwestern US and is endemic to Mexico and South America, predominantly Argentina, Venezuela, and Brazil [113]. Temperature and soil texture seem to be the only two factors that regulate the presence of Coccidioides based on a study of nine sites in California, Utah, and Arizona [114]. Coccidioides-bearing soils are characterized by very fine sand particles and silt, and its distribution seems to be limited to very specific areas of the planet [114]. Like in the case of other true human pathogens, the detection of Coccidioides in the environment is very difficult due to its sporadic distribution. Only 0.55% (4 out of 720) positive soil samples were obtained in California [115]. More sensitive detection is possible using BALB/c mice as biosensors with 8.9% positive detection in soils from the Tuscan area in Arizona, which is known for the presence of Coccidioides posadasii [116]. Intraperitoneal inoculation into mice was also successful in isolating C. posadasii from 6 out of 24 (25%) soil samples from Brazil [117]. This technique has facilitated the examination of Coccidioides spp. in endemic areas [117].

6.4.2 Dematiaceous and Keratinolytic Fungi in Deserts Fungi in the family Arthrodermataceae, as well as other taxa found in desert soils, are keratinolytic known for their ability to degrade keratin and grow on skin, hair, and nails of animals. The ability to break down keratin, a stable and resistant cytoskeletal filament in human and animal cells, is considered a virulence factor of those fungi known as dermatophytes [118]. Dermatophytes can cause a common skin infection in humans known as ringworm or tinea. These infections are confined to the dead

6.4 Human Pathogenic Fungi in Desert Ecosystems

|

113

Table 6.1: Percentage of Arthrodermataceae fungi isolated from desert soils. Bahrain Microsporum gypseum Trichophyton mentagrophytes Arthroderma curreyi T. terrestre Chrysosporium indicum C. pannicola Arthroderma cuniculi C. tropicum References

3.75 2.5

Israel

Kuwait

India

Iran

Tunisia

4.4

7.5

12.5 1.66

22.96

27.4 3.7

3.5 15.7

17.5 10

5.83 19.16 7.5

[121]

20 [122]

10 [123]

2.5

2.5 [120]

14.07

11

[125]

3.7 14 [141]

superficial regions of the skin and are highly contagious, but in the majority of the cases they can be treated with the application of antifungal creams [119]. The dermatophytic macroconidial species of Epidermophyton, Microsporum, and Trichophyton can be found ubiquitously in the environment, including deserts. The most common desert soil dermatophyte is Microsporum gypseum, isolated from several countries, including Bahrain, Israel, Kuwait, India, Egypt, and Iran [120–125] (󳶳 Tab. 6.1). In addition to true dermatophytes, other saprophytic fungi can also cause opportunistic infections in humans. In desert soils, keratinophiles can take advantage of keratin as a carbon source in a low nutrient environment. Alternaria, a robust keratinophile and a very abundant fungus in deserts, has been reported as the causing agent of phaeohyphomycotic cysts in immunosuppressed individuals [126]. Fusarium solani and Fusarium oxysporum, both reported keratinophiles and common in deserts (󳶳 Fig. 6.2f), are also considered the most common causative agents of Fusarium mycosis [127]. Paecilomyces, Geomyces, and Chaetomium keratinophiles and opportunistic pathogens are also common in arid soils [15, 18, 125].

6.4.3 Eumycetoma Eumycetoma is a fungal chronic pseudotumorous infection of the skin and subcutaneous tissue with high incidence in tropical, subtropical, and arid regions. The infection progresses with granulomatous lesions and discharge of grains with fungal particles that spread into adjacent tissue, bone, fascia, and ligaments [128, 129]. Males between 16–50 years old with agricultural occupations have the highest incidence of this infection [129, 130]. The most common infection site is the foot that has been exposed to soil or plant material containing a pathogenic fungus [131] after a traumatic injury. Diagnosis is often accomplished by a biopsy and examination of the grains produced by the fungus, culture based methods, or DNA sequencing from infected

114 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

tissue. Madurella mycetomatis is the usual etiological agent, but eumycetomas have also been reported for other common genera including Exophiala jeanselmei, Leptosphaeria senegalensis, Madurella grisea, Fusarium, Aspergillus, Curvularia, Acremonium, and Paecilomyces, among others [129–132], many of which are common taxa in deserts. The mycetoma belt includes South America, Sudan, Somalia, Senegal, and southern India [132]. Extensive reports from arid regions include the Republic of Niger, Mexico, Brazil, Iran, India, and Somalia [129, 131, 132]. Sudan shows the highest number of eumycetoma cases in the world (70% of cases) with Mexico second with an average of 70 cases per year [131, 132].

6.4.4 Mycotoxins Mycotoxins are a diverse group of toxic and carcinogenic compounds produced by fungi. In economically poor arid regions they are not very well documented, but represent a major problem for human and animal health. Many of the fungi responsible for the production of mycotoxins are xerophilic (i.e., they can grow in low humidity or low water content) and are abundant in desert soils. The most prominent species of fungi producing mycotoxins are Penicillium, Aspergillus, and Fusarium; with the production of significant toxins such as aflatoxin, fumonisins, ochratoxin A, trichothecenes, and zearalenone [133, 134]. Mycotoxins can cause adverse effects that result in illnesses of animals as well as serious problems for human health. For example, Fusarium moniliforme, colonizing maize is known to cause leukoencephalomalacia in horses and has cancer promoting activity due to fumonisins [135]. Ochratoxin A is the nephrotoxic responsible for human Balkan endemic nephropathy and other urinary tract tumors [136]. Aflatoxin contamination by Aspergillus is common in arid ecosystems such as the sub-Saharan Africa. This fungus benefits from high humidity and temperature, but drought conditions increase the risk of aflatoxin contamination [137]. Aflatoxin is the most potent naturally occurring carcinogenic substance and is likely responsible for the highest incidence of hepatocellular cancer in Africa [138]. Kenya reported an acute outbreak of aflatoxicosis with 317 cases in July 2004, with a fatality rate of 39% caused by A. flavus contamination and ingestion of contaminated maize [139]. The replacement of millets and sorghum for maize as the preferred cereal for food puts higher numbers of individuals at risk, since maize seems to have higher colonization rates by aflatoxin producing Aspergillus strains [137, 140].

References |

115

6.5 Importance of Fungal Biodiversity in Arid Lands Plant and biocrust associated fungi comprise a large untapped reservoir of fungal diversity. Most studies have focused on specific plant species or sites combining molecular and cultured based methods, but the advent of next generation molecular techniques (e.g., genomics, transcriptomics, metagenomics) is opening new opportunities to study fungi in arid lands and their response to climate and land use changes [16, 22, 32]. Challenges are still present with the low number of fungal genomes available and the low number of functional categories that are well annotated. Metagenomic studies have proved to be of great value even with the disproportionate number of bacteria (97–99%) vs. fungal (0.5–1.5%) metagenome reads in arid soils. The metabolic potential and diversity of specific taxa that are difficult to detect using regular PCR based or culture based techniques have been revealed in current studies [15, 22]. Arid lands in general are considered critical zones of biological interactions [2, 3]. These fragile ecosystems are threatened by environmental changes and their disturbance could result in large scale impact on other ecosystems, including marine environments, through dust deposition, increase of human infections, among others [2]. Fungi represent a key component of the dynamics of these ecosystems. A better understanding of the structure and function of fungal communities in deserts will facilitate the establishment of practices to ameliorate damage, improve preservation of arid sites, maximize their potential for discovery of new species, and generate applications in agriculture and the medical field. Acknowledgment: AP-A support was provided by National Science Foundation (award number 1457002) and the Sevilleta Long Term Ecological Research Site. Support for CRK is from the US Department of Energy, Biological and Environmental Research Division, through a science focus area grant.

References [1] [2] [3] [4] [5]

Staley JT, Palmer F, Adams JB. Micro colonial fungi: common inhabitants on desert rocks? Science 1982, 215:1093–5. Pointing SB, Belnap J. Disturbance to desert soil ecosystems contributes to dust-mediated impacts at regional scales. Biodivers Conserv 2014, 23:1659–67. Pointing SB, Belnap J. Microbial colonization and controls in drylands systems. Nat Rev Microbiol 2012, 10:551–62. Huxman T, Snyder K, Tissue D, et al. Precipitation pulses and carbon fluxes in semiarid and arid ecosystems. Oecologia 2004, 141:254–68. Parchert KJ, Spilde MN, Porras-Alfaro A, Nyberg AM, Northup DE. Fungal Communities Associated with Rock Varnish in Black Canyon, New Mexico: Casual Inhabitants or Essential Partners? Geomicrobiol J 2012, 29:752–66.

116 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

[6]

[7] [8] [9]

[10] [11] [12] [13] [14]

[15]

[16]

[17] [18] [19] [20] [21] [22] [23] [24]

[25]

Perry RS, Gorbushina A, Engel MH, Kolb VM, Krumbein WE, Staley JT. Accumulation and deposition of inorganic and organic compounds by microcolonial fungi. Proc Third Eur Workshop Exo-Astrobiol, 2004, 55–8. Sterflinger K, Tesei D, Zakharova K. Fungi in hot and cold deserts with particular reference to microcolonial fungi. Fungal Ecol 2012, 5:453–62. Mueller RC, Belnap J, Kuske CR. Soil bacterial and fungal community responses to nitrogen addition across soil depth and microhabitat in an arid shrubland. Front Microbiol 2015, 6:891. Porras-Alfaro A, Herrera J, Sinsabaugh RL, Odenbach KJ, Lowrey T, Natvig DO. Novel root fungal consortium associated with a dominant desert grass. Appl Environ Microbiol 2008, 74:2805– 13. Jumpponen A, Trappe JM. Dark septate endophytes: a review of facultative biotrophic rootcolonizing fungi. New Phytol 1998, 140:295–310. Belnap J, Lange OL. Biological Soil Crusts: Structure, Function, and Management. Berlin, Heidelberg, Springer, 2002. Barberán A, Ladau J, Leff JW, et al. Continental-scale distributions of dust-associated bacteria and fungi. P Nat Acad Sci 2015, 112:5756–61. Knapp DG, Kovács GM, Zajta E, Groenewald JZ, Crous PW. Dark septate endophytic pleosporalean genera from semiarid areas. Persoonia 2015, 35:87–100. Bates ST, Garcia-Pichel F, Nash III TH. Fungal components of biological soil crusts: insights from culture-dependent and culture-independent studies. In: Nash TH III, Geiser L, McCune B, Triebel D, Tomescu AMF, Sanders WB (eds). Biology of Lichens – Symbiosis, Ecology, Environm Monitoring, Systematics, Cyber Applications. Verlagsbuchhandlung, Stuttgart: J. Cramer in der Gebrüder Borntraeger 2010, 197–210. Steven B, Gallegos-Graves LV, Yeager C, Belnap J, Kuske CR. Common and distinguishing features of the bacterial and fungal communities in biological soil crusts and shrub root zone soils. Soil Biol Bioch 2014, 69:302–12. Steven B, Hesse C, Gallegos-Graves LV, Belnap J, Kuske CR. Fungal Diversity in Biological Soil Crusts of the Colorado Plateau. Proc 12th Biennial Conf Science Management Colorado Plateau 2014:in press. Collins SL, Sinsabaugh RL, Crenshaw C, et al. Pulse dynamics and microbial processes in aridland ecosystems. J Ecol 2008, 96:413–20. Porras-Alfaro A, Herrera J, Natvig DO, Lipinski K, Sinsabaugh RL. Diversity and distribution of soil fungal communities in a semiarid grassland. Mycologia 2011, 103:10–21. Bates ST, Nash III TH, Garcia-Pichel F. Patterns of diversity for fungal assemblages of biological soil crusts from the southwestern United States. Mycologia 2012, 104:353–61. Alguacil MM, Roldan A, Torres MP. Complexity of semiarid gypsophilous shrub communities mediates the AMF biodiversity at the plant species level. Microb Ecol 2009, 57:718–27. Porras-Alfaro A, Raghavan S, Garcia M, Sinsabaugh RL, Natvig DO, Lowrey TK. Endophytic fungal symbionts associated with gypsophilous plants. Botany 2014, 92:295–301. Hudson CM, Kirton E, Hutchinson MI, et al. Lignin-modifying processes in the rhizosphere of arid land grasses. Environ Microbiol 2015, 17:4965–78. Belnap J. Some Like It Hot, Some Not. Science 2013, 340:1533–4. Bates ST, Garcia-Pichel F. A culture-independent study of free-living fungi in biological soil crusts of the Colorado Plateau: their diversity and relative contribution to microbial biomass. Environ Microbiol 2009, 11:56–67. Steven B, Gallegos-Graves LV, Belnap J, Kuske CR. Dryland soil microbial communities display spatial biogeographic patterns associated with soil depth and soil parent material. FEMS Microbiol Ecol 2013, 86:101–13.

References | 117

[26] Pietrasiak N, Regus JU, Johansen JR, Lam D, Sachs JL, Santiago LS. Biological soil crust community types differ in key ecological functions. Soil Biol and Biochem 2013, 65:168–71. [27] Grishkan I, Kidron GJ. Biocrust-inhabiting cultured microfungi along a dune catena in the western Negev Desert, Israel. Eur J Soil Biol 2013, 56:107–14. [28] States JS, Christensen M. Fungi associated with biological soil crusts in desert grasslands of Utah and Wyoming. Mycologia 2001, 93:432–9. [29] Bates ST, Nash TH, Sweat KG, Garcia-Pichel F. Fungal communities of lichen-dominated biological soil crusts: Diversity, relative microbial biomass, and their relationship to disturbance and crust cover. J Arid Environ 2010, 74:1192–9. [30] Green LE, Porras-Alfaro A, Sinsabaugh RL. Translocation of nitrogen and carbon integrates biotic crust and grass production in desert grassland. J Ecol 2008, 96:1076–85. [31] Johnson SL, Kuske CR, Carney TD, Housman DC, Gallegos-Graves LV, Belnap J. Increased temperature and altered summer precipitation have differential effects on biological soil crusts in a dryland ecosystem. Glob Change Biol 2012, 18:2583–93. [32] Steven B, Kuske CR, Reed SC, Belnap J. Climate change and physical disturbance manipulations result in distinct biological soil crust communities. Appl Environ Microb 2015, 81:7448–59. [33] Bowker MA, Maestre FT, Eldridge D, et al. Biological soil crusts (biocrusts) as a model system in community, landscape and ecosystem ecology. Biodivers Conserv 2014, 23:1619–37. [34] Massimo NC, Nandi Devan MM, Arendt KR, et al. Fungal endophytes in aboveground tissues of desert plants: infrequent in culture, but highly diverse and distinctive symbionts. Microb Ecol 2015, 70:61–76. [35] Herrera J, Khidir HH, Eudy DM, Porras-Alfaro A, Natvig DO, Sinsabaugh RL. Shifting fungal endophyte communities colonize Bouteloua gracilis: effect of host tissue and geographical distribution. Mycologia 2010, 102:1012–26. [36] Mandyam K, Fox C, Jumpponen A. Septate endophyte colonization and host responses of grasses and forbs native to a tallgrass prairie. Mycorrhiza 2012, 22:109–19. [37] Lipson DA, Kuske CR, Gallegos-Graves LV, Oechel WC. Elevated atmospheric CO2 stimulates soil fungal diversity through increased fine root production in a semiarid shrubland ecosystem. Glob Chang Biol 2014, 20:2555–65. [38] Shamir I, Steinberger Y. Vertical distribution and activity of soil microbial population in a sandy desert ecosystem. Microb Ecol 2007, 53:340–7. [39] Bell C, McIntyre N, Cox S, Tissue D, Zak J. Soil microbial responses to temporal variations of moisture and temperature in a Chihuahuan desert grassland. Microb Ecol 2008, 56:153–67. [40] Nguyen LM, Buttner MP, Cruz P, Smith SD, Robleto EA. Effects of elevated atmospheric CO2 on rhizosphere soil microbial communities in a Mojave Desert ecosystem. J Arid Environ 2011, 75:917–25. [41] Lipson DA, Wilson RF, Oechel WC. Effects of elevated atmospheric CO2 on soil microbial biomass, activity, and diversity in a chaparral ecosystem. Appl Environ Microb 2005, 71:8573– 80. [42] Khidir HH, Eudy DM, Porras-Alfaro A, Herrera J, Natvig DO, Sinsabaugh RL. A general suite of fungal endophytes dominate the roots of two dominant grasses in a semiarid grassland. J Arid Environ 2010, 74:35–42. [43] Wehner J, Powell JR, Muller LAH, et al. Determinants of root-associated fungal communities within Asteraceae in a semi-arid grassland. J Ecol 2014, 102:425–36. [44] Porras-Alfaro A, Herrera J, Natvig DO, Sinsabaugh RL. Effect of long-term nitrogen fertilization on mycorrhizal fungi associated with a dominant grass in a semiarid grassland. Plant and Soil 2007, 296:65–75.

118 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

[45]

[46] [47] [48]

[49] [50] [51] [52] [53]

[54]

[55] [56] [57]

[58]

[59] [60]

[61]

[62]

[63] [64]

Johnson NC, Rowland DL, Corkidi L, Egerton-Warburton LM, Allen EB. Nitrogen enrichment alters mycorrhizal allocation at five mesic to semiarid grasslands. Ecology 2003, 84:1895–908. Tisdall JM, Oades JM. Organic matter and water-stable aggregates in soils. J Soil Science 1982, 33:141–63. McLellan CA, Turbyville TJ, Wijeratne EM, et al. A rhizosphere fungus enhances Arabidopsis thermotolerance through production of an HSP90 inhibitor. Plant Physiol 2007, 145:174–82. Brundrett MC. Mycorrhizal associations and other means of nutrition of vascular plants: understanding the global diversity of host plants by resolving conflicting information and developing reliable means of diagnosis. Plant Soil 2009, 320:37–77. Wu Y, Jiang J, Shen W, He X. Arbuscular mycorrhiza fungi as an ecology indicator for evaluating desert soil conditions. Front Agricul China 2010, 4:24–30. Johnson D, Leake JR, Read DJ. Novel in-growth core system enables functional studies of grassland mycorrhizal mycelial networks. New Phytol 2001, 152:555–62. Kruger M, Teste FP, Laliberte E, et al. The rise and fall of arbuscular mycorrhizal fungal diversity during ecosystem retrogression. Mol Ecol 2015, 24:4912–30. Treseder KK, Cross A. Global distributions of arbuscular mycorrhizal fungi. Ecosystems 2006, 9:305–16. Öpik M, Vanatoa A, Vanatoa E, et al. The online database MaarjAM reveals global and ecosystemic distribution patterns in arbuscular mycorrhizal fungi (Glomeromycota). New Phytol 2010, 188:223–41. Dean SL, Warnock DD, Litvak ME, Porras-Alfaro A, Sinsabaugh R. Root-associated fungal community response to drought-associated changes in vegetation community. Mycologia 2015, 107:1089–104. Jasper DA, Abbott LK, Robson AD. The survival of infective hyphae of vesicular-arbuscular mycorrhizal fungi in dry soil: an interaction with sporulation. New Phytol 1993, 124:473–9. Barrow JR. Atypical morphology of dark septate fungal root endophytes of Bouteloua in arid southwestern USA rangelands. Mycorrhiza 2003, 13:239–47. Symanczik S, Courty PE, Boller T, Wiemken A, Al-Yahya’ei MN. Impact of water regimes on an experimental community of four desert arbuscular mycorrhizal fungal (AMF) species, as affected by the introduction of a non-native AMF species. Mycorrhiza 2015, 25:639–47. Barness G, Rodriguez Zaragoza S, Shmueli I, Steinberger Y. Vertical distribution of a soil microbial community as affected by plant ecophysiological adaptation in a desert system. Microb Ecol 2009, 57:36–49. Walker DJ, Lutts S, Sánchez-García M, Correal E. Atriplex halimus L.: Its biology and uses. J Arid Environ 2014, 100–101:111–21. Gutierrez A, Morte A, Honrubia M. Morphological characterization of the mycorrhiza formed by Helianthemum almeriense Pau with Terfezia claveryi Chatin and Picoa lefebvrei (Pat.) Maire. Mycorrhiza 2003, 13:299–307. Zitouni-Haouar Fel H, Fortas Z, Chevalier G. Morphological characterization of mycorrhizae formed between three Terfezia species (desert truffles) and several Cistaceae and Aleppo pine. Mycorrhiza 2014, 24:397–403. Kozdroj J, Piotrowska-Seget Z, Krupa P. Mycorrhizal fungi and ectomycorrhiza associated bacteria isolated from an industrial desert soil protect pine seedlings against Cd(II) impact. Ecotoxicology 2007, 16:449–56. Leake JR. The biology of myco-heterotrophic (‘saprophytic’) plants. New Phytol 1994, 127:171–216. Bruns TD, Read DJ. In vitro germination of nonphotosynthetic, myco-heterotrophic plants stimulated by fungi isolated from the adult plants. New Phytol 2000, 148:335–42.

References |

[65] [66] [67] [68] [69] [70] [71] [72] [73] [74]

[75] [76]

[77] [78] [79] [80] [81] [82] [83] [84] [85]

[86] [87]

119

Taylor DL, Bruns TD, Leake JR, Read DJ. Mycorrhizal specificity and function in myco-heterotrophic plants. Mycorrhizal Ecol 2003, 157:375–413. Bhatnagar A, Bhatnagar M. Microbial diversity in desert ecosystems. Curr Sci 2005, 89:91–100. Loizides M, Hobart C, Konstandinides G, Yiangou Y. Desert Truffles: the mysterious jewels of antiquity. Field Mycol 2012, 13:17–21. Jamali S, Banihashemi Z. Hosts and distribution of desert truffles in Iran, based on morphological and molecular criteria. J Agric Sci Technol 2012, 14:1379–96. Porras-Alfaro A, Bayman P. Hidden fungi, emergent properties: endophytes and microbiomes. Annu Rev Phytopathol 2011, 49:291–315. Wilson D. Endophyte: the evolution of a term, and clarification of its use and definition. Oikos 1995, 73:274–6. Arnold AE, Maynard Z, Gilbert GS, Coley PD, Kursar TA. Are tropical fungal endophytes hyperdiverse? Ecol Lett 2000, 3:267–74. Sun Y, Wang Q, Lu X, Okane I, Kakishima M. Endophytic fungal community in stems and leaves of plants from desert areas in China. Mycol Prog 2011, 11:781–90. Arnold AE, Maynard Z, Gilbert GS. Fungal endophytes in dicotyledonous neotropical trees: patterns of abundance and diversity. Mycol Res 2001, 105:1502–7. Herrera J, Poudel R, Nebel KA, Collins SL. Precipitation increases the abundance of some groups of root-associated fungal endophytes in a semiarid grassland. Ecosphere 2011, 2:1–14. Loro M, Valero-Jiménez CA, Nozawa S, Márquez LM. Diversity and composition of fungal endophytes in semiarid Northwest Venezuela. J Arid Environ 2012, 85:46–55. Herrera J, Poudel R, Khidir H. Molecular Characterization of Coprophilous Fungal Communities Reveals Sequences Related to Root-Associated Fungal Endophytes. Microb Ecol 2011, 61:239–44. Wu Y, Liu T, He X. Mycorrhizal and dark septate endophytic fungi under the canopies of desert plants in Mu Us Sandy Land of China. Front Agr China 2009, 3:164–70. Rodriguez RJ, Henson J, Van Volkenburgh E, et al. Stress tolerance in plants via habitatadapted symbiosis. ISME J 2008, 2:404–16. Redman RS, Sheehan KB, Stout RG, Rodriguez RJ, Henson JM. Thermotolerance generated by plant/fungal symbiosis. Science 2002, 298:1581. Alguacil MM, Roldan A, Torres MP. Assessing the diversity of AM fungi in arid gypsophilous plant communities. Environ Microbiol 2009, 11:2649–59. Palacio S, Escudero A, Montserrat-Marti G, Maestro M, Milla R, Albert MJ. Plants living on gypsum: beyond the specialist model. Ann Bot 2007, 99:333–43. Peláez F, Collado J, Arenal F, et al. Endophytic fungi from plants living on gypsum soils as a source of secondary metabolites with antimicrobial activity. Mycol Res 1998, 102:755–61. Landwehr M, Hildebrandt U, Wilde P, et al. The arbuscular mycorrhizal fungus Glomus geosporum in European saline, sodic and gypsum soils. Mycorrhiza 2002, 12:199–211. Oliveira LG, Cavalcanti MAQ, Fernandes MJS, Lima DMM. Diversity of filamentous fungi isolated from the soil in the semiarid area, Pernambuco, Brazil. J Arid Environ 2013, 95:49–54. Matsuzawa T, Campos Takaki GM, Yaguchi T, Okada K, Gonoi T, Horie Y. Two new species of Aspergillus section Fumigati isolated from caatinga soil in the State of Pernambuco, Brazil. Mycoscience 2014, 55:79–88. Sinsabaugh RL, Belnap J, Rudgers J, Kuske CR, Martinez N, Sandquist D. Soil microbial responses to nitrogen addition in arid ecosystems. Front Microbiol 2015, 6:819. Crenshaw CL, Lauber C, Sinsabaugh RL, Stavely LK. Fungal control of nitrous oxide production in semiarid grassland. Biogeochemistry 2008, 87:17–27.

120 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

[88] Chen H, Mothapo NV, Shi W. Soil moisture and pH control relative contributions of fungi and bacteria to N2O production. Microb Ecol 2015, 69:180–91. [89] Stursova M, Crenshaw CL, Sinsabaugh RL. Microbial responses to long-term N deposition in a semiarid grassland. Microb Ecol 2006, 51:90–8. [90] McLain JET, Martens DA. N2O production by heterotrophic N transformations in a semiarid soil. Appl Soil Ecol 2006, 32:253–63. [91] Gallo ME, Porras-Alfaro A, Odenbach KJ, Sinsabaugh RL. Photoacceleration of plant litter decomposition in an arid environment. Soil Biology and Biochemistry 2009, 41:1433–41. [92] Day TA, Zhang ET, Ruhland CT. Exposure to solar UV-B radiation accelerates mass and lignin loss of Larrea tridentata litter in the Sonoran Desert. Plant Ecol 2007, 193:185–94. [93] Clarke LJ, Weyrich LS, Cooper A. Reintroduction of locally extinct vertebrates impacts arid soil fungal communities. Mol Ecol 2015, 24:3194–205. [94] Masunga GS, Andresen O, Taylor JE, Dhillion SS. Elephant dung decomposition and coprophilous fungi in two habitats of semi-arid Botswana. Mycol Res 2006, 110:1214–26. [95] Magan N. Fungi in extreme environments. In: Kubicek CP, Druzhinina IS (eds). Environmental and microbial relationships, 2nd edn. Springer-Verlag Berlin Heidelberg, 2007, 350. [96] Powell AJ, Parchert KJ, Bustamante JM, Ricken JB, Hutchinson MI, Natvig DO. Thermophilic fungi in an aridland ecosystem. Mycologia 2012, 104:813–25. [97] de Oliveira TB, Gomes E, Rodrigues A. Thermophilic fungi in the new age of fungal taxonomy. Extremophiles 2015, 19:31–7. [98] Abdel-Hafez SII. Thermophilic and thermotolerant fungi in the desert soils of Saudi Arabia. Mycopathologia 1982, 80:15–20. [99] Hemida SK. Thermophilic and thermotolerant fungi isolated from cultivated and desert soils, exposed continuously to cement dust particles in Egypt. Zentralblatt für Mikrobiologie 1992, 147:277–81. [100] Palmer FE, Emery DR, Stumbler J, Staley JT. Survival and growth of microcolonial rock fungi as affected by temperature and humidity. 1987, 107:155–62. [101] Marzban G, Tesei D, Sterflinger K. A review beyond the borders: Proteomics of microcolonial black fungi and black yeasts. Nat Sci 2013, 5:640–5. [102] Zakharova K, Tesei D, Marzban G, Dijksterhuis J, Wyatt T, Sterflinger K. Microcolonial fungi on rocks: a life in constant drought? Mycopathologia 2013, 175:537–47. [103] Gorbushina AA, Kotlova ER, Sherstneva OA. Cellular responses of microcolonial rock fungi to long-term desiccation and subsequent rehydration. Stud Mycol 2008, 61:91–7. [104] Marvasi M, Donnarumma F, Brandi A, et al. Black microcolonial fungi as deteriogens of two famous marble statues in Florence, Italy. I. Biodeterior Biodegrad 2012, 68:36–44. [105] Selbmann L, Zucconi L, Isola D, Onofri S. Rock black fungi: excellence in the extremes, from the Antarctic to space. Curr Genet 2015, 61:335–45. [106] Reid CE, Gamble JL. Aeroallergens, allergic disease, and climate change: impacts and adaptation. Ecohealth 2009, 6:458–70. [107] Galgiani JN, Ampel NM, Blair JE, et al. Coccidioidomycosis. Clin Infect Dis 2005, 41:1217–23. [108] Dixon DM. Coccidioides immitis as a select agent of bioterrorism. J Appl Microbiol 2001, 91:602–5. [109] Williams JH, Phillips TD, Jolly PE, Stiles JK, Jolly CM, Aggarwal D. Human aflatoxicosis in developing countries: a review of toxicology, exposure, potential health consequences, and interventions. Am J Cli Nutr 2004, 80:1106–22. [110] Schneider E, Hajjeh RA, Spiegel RA, et al. A coccidioidomycosis outbreak following the Northridge, Calif, earthquake. JAMA 1997, 277:904–8. [111] Petersen LR, Marshall SL, Barton-Dickson C, et al. Coccidioidomycosis among workers at an archeological site, northeastern Utah. Emerg Infect Dis 2004, 10:637–42.

References |

121

[112] Centers for Disease C, Prevention. Increase in reported coccidioidomycosis–United States, 1998–2011. MMWR Morbidity and mortality weekly report 2013, 62:217. [113] Baptista-Rosas RC, Catalán-Dibene J, Romero-Olivares AL, Hinojosa A, Cavazos T, Riquelme M. Molecular detection of Coccidioides spp. from environmental samples in Baja California: linking Valley Fever to soil and climate conditions. Fungal Ecol 2012, 5:177–90. [114] Fisher FS, Bultman MW, Johnson SM, Pappagianis D, Zaborsky E. Coccidioides niches and habitat parameters in the southwestern United States: a matter of scale. Ann N Y Acad Sci 2007, 1111:47–72. [115] Greene DR, Koenig G, Fisher MC, Taylor JW. Soil isolation and molecular identification of Coccidioides immitis. Mycologia 2000, 92:406–10. [116] Barker BM, Tabor JA, Shubitz LF, Perrill R, Orbach MJ. Detection and phylogenetic analysis of Coccidioides posadasii in Arizona soil samples. Fungal Ecol 2012, 5:163–76. [117] de Macêdo RCL, Rosado AS, da Mota FF, et al. Molecular identification of Coccidioides spp. in soil samples from Brazil. BMC Microbiol 2011, 11:108–16. [118] Scott JA, Untereiner WA. Determination of keratin degradation by fungi using keratin azure. Medical Mycology 2004, 42:239–46. [119] Weitzman I, Summerbell RC. The dermatophytes. Clin Microbiol Rev 1995, 8:240–59. [120] Deshmukh SK, Mandeel QA, Verekar SA. Keratinophilic fungi from selected soils of Bahrain. Mycopathol 2008, 165:143–7. [121] Feuerman E, Alteras I, Hönig E, Lehrer N. The isolation of keratinophilic fungi from soils in Israel. A preliminary report. Mycopathol 1975, 56:41–6. [122] Al-Musallam AA, Al-Zarban SS, Al-Sanè NA, Ahmed TM. A report on the predominant occurrence of a dermatophyte species in cultivated soil from Kuwait. Mycopathol 1995, 130:159–61. [123] Deshmukh SK, Verekar SA. Prevalence of keratinophilic fungi in usar soils of Uttar Pradesh, India. Microbiol Res 2011, 2:15. [124] Bagy MMK. Saprophytic and keratinophilic fungi isolated from desert and cultivated soils continuously exposed to cement dust particles in Egypt. ZBL Mikrobiol 1992, 147:418–26. [125] Malek E, Moosazadeh M, Hanafi P, et al. Isolation of Keratinophilic Fungi and Aerobic Actinomycetes From Park Soils in Gorgan, North of Iran. Jundishapur J Microbiol 2013, 6:1–5. [126] Boyce RD, Deziel PJ, Otley CC, et al. Phaeohyphomycosis due to Alternaria species in transplant recipients. Transpl Infect Dis 2010, 12:242–50. [127] O’Donnell K, Sutton DA, Fothergill A, et al. Molecular phylogenetic diversity, multilocus haplotype nomenclature, and in vitro antifungal resistance within the Fusarium solani species complex. J Clin Microbiol 2008, 46:2477–90. [128] Yera H, Bougnoux ME, Jeanrot C, Baixench MT, De Pinieux G, Dupouy-Camet J. Mycetoma of the Foot Caused by Fusarium solani: Identification of the Etiologic Agent by DNA Sequencing. J Clin Microbiol 2003, 41:1805–8. [129] Zarei Mahmoudabadi A, Zarrin M. Mycetomas in Iran: a review article. Mycopathologia 2008, 165:135–41. [130] López-Martínez R, Méndez-Tovar LJ, Bonifaz A, et al. Actualización de la epidemiología del micetoma en México. Revisión de 3,933 casos. Gac Med Mex 2013, 149:586–92. [131] Estrada R, Chávez-López G, Estrada-Chávez G, López-Martínez R, Welsh O. Eumycetoma. Clin Dermatol 2012, 30:389–96. [132] Fahal AH, Hassan MA. Mycetoma. British J Surgery 1992, 79:1138–41. [133] Bankole S, Schollenbeger M, Drochner W. Mycotoxin contamination in food systems in subSaharan Africa. Bydgoszcz: Soc Mycotox Res 2006, 22:163–9. [134] Fink-Grernmels J. Mycotoxins: their implications for human and animal health. Veterin Quart 1999, 21:115–20.

122 | 6 Fungal Diversity, Community Structure and Their Functional Roles in Desert Soils

[135] Gelderblom WC, Jaskiewicz K, Marasas WF, et al. Fumonisins–novel mycotoxins with cancer-promoting activity produced by Fusarium moniliforme. Appl Environ Microbiol 1988, 54:1806–11. [136] Pfohl-Leszkowicz A, Manderville RA. Ochratoxin A: An overview on toxicity and carcinogenicity in animals and humans. Mol Nutr Food Res 2007, 51:61–99. [137] Hell K, Mutegi C. Aflatoxin control and prevention strategies in key crops of Sub-Saharan Africa. Afri J Microbiol Res 2011, 5:459–66. [138] Strosnider H, Azziz-Baumgartner E, Banziger M, et al. Workgroup report: public health strategies for reducing aflatoxin exposure in developing countries. Environ Health Persp 2006, 114:1898–903. [139] Probst C, Njapau H, Cotty PJ. Outbreak of an acute aflatoxicosis in Kenya in 2004: identification of the causal agent. Appl Environ Microbiol 2007, 73:2762–4. [140] Bandyopadhyay R, Kumar M, Leslie JF. Relative severity of aflatoxin contamination of cereal crops in West Africa. Food Addit Contam 2007, 24:1109–14. [141] Anane S, Al-Yasiri MYH, Normand AC, Ranque S. Distribution of keratinophilic fungi in soil across Tunisia: a descriptive study and review of the literature. Mycopathologia 2015, 180:61–8.

T.G. Allan Green

7 Limits of Photosynthesis in Arid Environments Abstract: Soils in arid zones are often covered with biological soil crust (BSC) typically composed of bacteria, fungi, cyanobacteria, algae, lichens (lichenized fungi) and bryophytes (mosses and liverworts). BSC have major effects on the stability and functioning of the soils. All organisms in BSC are poikilohydric, meaning that they can desiccate and are only active when wet. Photosynthesis of BSC, therefore, shows response curves to incident light, temperature, CO2 concentration, and thallus water content (WC). Photosynthesis of BSC is typically optimal at high light, around 15 to 20°C and ambient CO2 above 1000 ppm. Response to WC can be complex, but photosynthesis is limited at low WC and often, due to diffusion limitations, at higher WC. BSC rarely carry out photosynthesis under optimal conditions. Environmental water status is the major limiter, and in arid areas BSC are active for around 30% of the total time. In addition, they are active at light intensities and temperatures that are lower than the habitat means. Further limitations occur from thallus water content effects, either from low WC when drying or partially hydrated by dew, but also because many BSC organisms show depressed photosynthesis at high WC. The latter effect can be so intense that the organisms make little carbon gain from heavy rainfalls. As a result, overall carbon fixation is probably only around 1% of the theoretical maximum. The ability of BSC organisms to acclimate to a changing environment has probably been greatly underestimated and may occur in a few days, so that it might even be fast enough to influence the results of laboratory studies.

7.1 Introduction Biological soil crusts (BSC) are a mixture of autotrophic and heterotrophic organisms that (i) live within or on top the uppermost millimeters of soil creating a consistent layer and (ii) aggregate soil particles due to their presence and activity [1]. BSC are composed of a wide range of organisms, typically including bacteria, fungi, cyanobacteria, algae, lichens (lichenized fungi) and bryophytes (mosses and liverworts) of which all except bacteria (excluding cyanobacteria) and fungi are photosynthetic. Although local conditions strongly affect the presence of the different organisms, successional stages are recognized for BSC with initial colonization by filamentous cyanobacteria followed by smaller green algae and cyanobacteria and, finally, when the surface has stabilized, lichens and mosses [1]. BSC organisms cannot be treated as small higher plants but show important differences in their physiology and ecology. Firstly, and a physiological trait that links all BSC organisms, is that they are poikilohydric, meaning that their water status tends to equilibrate with the surrounding environment; they are wet and active when the enviDOI 10.1515/9783110419047-007

124 | 7 Limits of Photosynthesis in Arid Environments

ronment is wet, and dry and dormant under dry conditions. When dry, BSC organisms can withstand extremes of light and temperature (both high and low). Poikilohydry, through water supply and support, also enforces a size limitation on organisms with the vast majority being less than a centimeter high [2]. This, in turn, means that they are confined to a two-dimensional habitat in which they are almost always within the atmospheric boundary layer, bringing important changes to the interactions with the environment such as in heat exchange [2]. BSC occur throughout the world but, because of competition for light, are best developed in habitats in which competition by phanerogamous plants is limited. Such environments are hot, cool and cold semiarid and arid areas, and also polar and alpine zones. Such habitats are not productive, however their large extent means that they are estimated to contribute around 1% of global net primary production [3]. Because of their marginal climates BSC in these areas are also suggested to be more susceptible to future climate changes [4], and this is one important reason to gain a better understanding of the limits to photosynthesis by BSC.

7.2 Photosynthetic Responses to Environmental Factors, a Background 7.2.1 Rates, Chlorophyll and Mass Lange [5] summarizes the then available maximal net photosynthetic rates under optimal conditions (NPmax ) for a wide variety of soil crusts, and these span over two orders of magnitude between around 0.1 and 11.5 μmol m−2 s−1 . The majority of NPmax for BSC lie between 2 and 5 μmol m−2 s−1 (󳶳 Tab. 7.1), which are high rates compared to the more typical 1 to 2 μmol m−2 s−1 for rain forest lichens [6].

Table 7.1: LMA (mass per unit area), CO2 exchange rates, quantum efficiency and chlorophyll content for seven BSC lichen species LMA Species

g m−2

Maximal net Dark Quantum photosynthetic rate respiration efficiency μmol m−2 s−1 nmol g−1 s−1 μmol m−2 s−1

Collema cristatuma 310 2.8 Fulgensia fulgensb 440 5.2 Lecanora muralisc 510 6.5 Cladonia convolutad 630 5.4 Squamarina lentigerae 684 4.0 Collema tenax f 1190 3.9 Diploschistes diacapsisg 2000 5.0

9.03 11.82 12.75 8.57 5.85 3.28 2.5

Source of data: a [7], b [8], c [9], d [10], e [11], f [12], g [13].

0.95 1.25 1.60 1.80 1.50 1.80 1.50

0.015 0.026 0.025 0.024 0.015 0.011

Chlorophyll mg m−2 43 450 564 280 227 170 1350

7.2 Photosynthetic Responses to Environmental Factors, a Background | 125

Chlorophyll contents of BSC span a large range and can be comparable with those of average C3 leaves, which require 500–700 mg chl m−2 to achieve maximal quantum yield of CO2 uptake [5]. The chlorophyll contents of BSC lichens span a wide range from a low 42.7 mg chl m−2 for Collema cristatum to an exceptional 1350 mg chl m−2 for D. diacapsis (󳶳 Tab. 7.1) [5]. There are differences between the various BSC types. Zhao et al. [14] report 20.7, 29.0 and 38.1 mg chl m−2 for algal, mixed and moss dominated BSC from Tengger Desert in China, and Kidron et al. [15] measured 16.7 to 43.4 mg chl m−2 for cyanobacterial BSC and 53.2 mg chl m−2 for moss dominated BSC in the Negev Desert. For the Qubqi Desert, Mongolia, Lan et al. [16] found a large increase in chlorophyll content with BSC development from 30 mg chl m−2 in cyanobacterial dominated early crusts to 210 mg chl m−2 for fully developed moss dominated crusts. There appears to be no significant link between BSC chlorophyll content (mg chl m−2 ) −1 and NPmax (μmol m−2 s ) (󳶳 Tab. 7.1). Although data are limited, lichens forming BSC appear to be “heavy” in comparison to those growing in forests, showing a wide range in leaf mass per area (LMA, g dry weight m−2 ) from 310 gdw m−2 for Collema cristatum, to 2000 gdw m−2 for Diploschistes diacapsis (󳶳 Tab. 7.1). This compares to mean values of 86 gdw m−2 and 97 gdw m−2 for Lobaria scrobicularia and Lobaria pulmonaria, and 73 gdw m−2 Pseudocyphellaria crocata (Merinero et al. 2014), and 59 to 91 gdw m−2 for Pseudocyphellaria dissimilis from inside a New Zealand rain forest [17]. Similar magnitudes of LMA are reported for a wide range of lichens summarized in [18]. Data for bryophytes are not as easy to interpret as for lichens. Lichens, albeit a symbiosis, are a discrete organism and relatively easy to separate from soil crusts. Bryophytes, and mosses in particular, are known for being intimately bound with the soil crusts and can contribute to the structural strength of the BSC. As well as not being easy to separate from the crust, mosses have substantial portions of the plant below ground, which are not photosynthetic and will always be respiring when active. Studying Grimmia laevigata Alpert and Oechel [19] found 85.5 gdw m−2 for green parts of the plant and 161.5 gdw m−2 for brown parts (total 247 gdw m−2 ). Longton [20] found 241–692 gdw m−2 (100% cover) for Bryum argenteum and 1012–1108 gdw m−2 for B. antarcticum (= Henediella heimii) with the former growing in sheets and the latter in clumps. In contrast, Wu et al. [21] report 26.5 gdw m−2 for the desert moss Syntrichia caninervis in the Gurbantünggüt Desert, China, and Green and Snelgar [22] showed the thalloid liverworts Monoclea forsteri and Marchantia foliacea, New Zealand rain forest, to have only 33 and 35 gdw m−2 but still achieve a maximal net photosynthetic rates of 0.81 and 0.99 μmol m−2 s−1 , respectively. There appears to be no relationship between NPmax (area basis) and LMA, but there is a significant negative relationship between NPmax (dry weight basis) and LMA [23].

126 | 7 Limits of Photosynthesis in Arid Environments 7.2.2 Response of Net Photosynthesis (NP) to Light (PPFD, μmol m−2 s−1 )

(a)

4.0

8.0 Quantum efficiency

15°C

3.0

10°C

2.0 5°C

1.0 0.0 Light compensation

–1.0

Dark respiration rate

Light saturation

CO2 exchange – μmol m–2 s–1

Net photosynthesis (μmol CO2 m–2 s–1)

󳶳 Fig. 7.1a shows the typical saturation response of net photosynthesis to light by a lichen or bryophyte. Marked on the response curve are the so-called cardinal points: light level or photosynthetic photon flux density (PPFD) required to achieve maximal NP (PPFDsat ), quantum efficiency of NP to light (QE), which is initial slope of the response curve at low light, light level to achieve compensation (i.e. zero NP, PPFDcomp ), and dark respiration rate (DR), which is NP at zero light. The PPFDsat is typically around 700 μmol m−2 s−1 for BSC, and as a result they are referred to as sun plants [5]. However, BSC do not achieve the same photosynthetic rates as higher plants, which have leaves with protected photosynthetic cells and are able to build canopies. The high PPFDsat of BSC can be interpreted as a protection against the occasional bursts of high light or maintenance of the ability to benefit from such conditions; these are not exclusive. The light compensation point is positively correlated with high PPFDsat [24] and BSC have relatively high values for PPFDcomp, often 60 to 100 μmol m−2 s−1 , which are also influenced by temperature, being lower at low temperatures. This has the effect of lowering carbon gain at low light levels, such as might be found after sunrise. BSC also have low quantum efficiencies, from 0.015 to 0.026 (󳶳 Fig. 7.1a), which are less than those found for shade lichens and higher plants – 0.05 and 0.06, respectively. It is not surprising that with their high saturation light level for NP, BSC organisms appear to be well protected against potential damage to photosystems from high light. The highest light levels for BSC when hydrated and active are found in continen-

–2.0

6.0 4.0 2.0 0.0 –2.0 Collema Diploschistes Psora

–4.0 –6.0

0

200 400 600 800 1000 1200 PPFD (μmol m–2 s–1)

0 (b)

10

20 30 40 Temperature – °C

50

Fig. 7.1: (a) Typical response curve of net photosynthesis (μmol CO2 m−2 s−1 ) to incident light (PPFD, μmol m−2 s−1 ) of a soil crust at three temperatures (5, 10 and 15°C), showing the main cardinal points: light required to obtain maximal NP (PPFDmax ), quantum efficiency, light level to give compensation (no net CO2 exchange, PPFDcomp ), and dark respiration rate (DR). (b) Response of photosynthesis to temperature for BSC lichens; the response curves are generated at saturating light and optimal thallus water content (modified from [12]). Color coding of symbols: black – Collema tenax, red – Diploschistes diacapsis, blue – Psora cerebriformis: symbol shapes: • – net photosynthesis, 󳵳 – dark respiration, 󳶃 and dashed lines – Gross photosynthesis (NP – DR).

7.2 Photosynthetic Responses to Environmental Factors, a Background | 127

tal Antarctica where mean PPFD when active can reach around 700 μmol m−2 s−1 [25], and mosses have constitutive protection against high light with the xanthophyll cycle components present in similar quantities in both light and shade adapted forms. This protection of the photosystems is complimented by UV absorbing compounds [26]. It is now also becoming clear that bryophytes and lichens employ other methods to handle excess light and are physiologically agile in this area. One example is that both CO2 and O2 can act as interchangeable electron sinks, and the nonsaturating component of electron flow is photoreduction of oxygen [27, 28]. Although nonphotochemical quenching (NPQ) is found in both algae and plants, these organisms rely on two different proteins for its activation, light harvesting complex stress-related protein and photosystem II subunit S, respectively. In the moss Physcomitrella patens, however, both proteins are present and active [29]. As a general rule, no negative effects of high light or UV would be expected for BSC unless levels are applied that have little ecological relevance, e.g., shade adapted forms being exposed to very high light levels.

7.2.3 Response of Net Photosynthesis to Temperature In contrast to the rather constant response of NP to PPFD for BSC there seems to be a wider range of adaptions to temperature. Examples of typical responses of net photosynthesis to temperature (measured at saturating light and optimal thallus water content) are shown in 󳶳 Fig. 7.1b, with all three species showing a similar form of response. Net photosynthesis has an optimum temperature that is over 30°C for Collema and lower, around 20°C but with a much broader range with little change in NP, for the other two species. The decline in NP at higher temperatures is driven by the increasing dark respiration (exponential increase with temperature) up to about 30°C and at higher temperatures by a fall in photosynthetic capacity (gross photosynthesis, GP), which reaches a maximum at just over 30°C for all three species. A maximal rate of gross photosynthesis at around 30°C seems to be relatively common in lichens and mosses and is even found in Antarctic species [30], indicating that the underlying photosynthetic mechanisms show little change with environment. Differences in optimal temperature for NP are also reported for different organisms in the same habitat. For example, 20–27°C, 15°C and 20°C for cyanobacteria, lichens, and mosses, respectively, in the Mu Us Desert, Ningxia, northwest China (󳶳 Tab. 7.2, from [31]).

7.2.4 Response of Net Photosynthesis to Thallus Water Content (WC) Thallus water content in BSC is usually expressed as mm rain equivalent (mm, equal to liters per m2 ) and not, as is routine for lichens and bryophytes as % dry weight, (% dw = [wet weight − dry weight] ⋅ 100/dry weight), because of the difficulty in sepa-

128 | 7 Limits of Photosynthesis in Arid Environments

Table 7.2: Comparison of photosynthetic rates and light response, and thallus water content (WC) for BSC dominated by cyanobacteria, lichens and mosses; data from [31]. BSC type

NP max

Optimal temperature

PPFD to saturate NP

(°C)

Cyanobacterial Lichen Moss

μmol CO2 m−2 s−1 2.67 3.06 6.02

μmol m−2 s−1 μmol m−2 s−1 mm rain equivalent 900 70 0.38 870 90 0.92 1200 50 2.10

20–27 15 20

PPFD compensation

Optimal WC for NP

Maximal WC mm rain equivalent 1.3 2.5 3.8

rating BSC organisms from their substrate. At very low thallus water content there is no CO2 exchange, but as WC rises, so does NP until a maximum is reached (󳶳 Fig. 7.2). At NPmax the organisms are at, or close to, full turgor (relative water content, RWC, = 1.0) and at the so-called optimal water content WCopt [2]. Homoiohydric plants do not exceed RWC of 1.0, but lichens and bryophytes can do this because of variable amounts of external water held in capillary spaces outside the cells. As a result, maximal RWC in BSC organisms can be much higher than 1.0, often up to 2.0 or 3.0 for lichens and substantially higher for bryophytes (see 󳶳 Tab. 7.2 for a comparison of cyanobacteria, mosses and lichens at a desert site). The change in NP at WC above WCopt is strongly 3

2 10

Number of events Diploschistes Psora Didymodon

1

5 0

0

Net photosynthesis ( μmol m–2 s–1)

Number of rainfall events

15

–1 0

1 2 3 Rainfall – mm (0.2 mm categories) or Thallus water content – mm rain equivalent

4

−1

Fig. 7.2: Line graph: Response of net photosynthetic rate (right hand axis, μmol m−2 s ) measured at saturating PFD and 15°C to thallus water content (mm precipitation equivalent) for two lichens, • – Diploschistes diacapsis and 󳶃 – Psora decipiens, and one moss, 󳵳 – Didymodon rigidulus from Tabernas Desert, Almeria, Spain. Bar graph: distribution of rainfall occurrence with each bar representing the number of occurrences of a rainfall event of a particular size; X axis is rainfall event size in 0.2 mm categories. Note the “plateau” of the moss (󳵳).

7.3 Optimal Versus Real Photosynthetic Rates | 129

species dependent and can vary from maintenance of NPmax to a strong decline in NP, sometimes to negative values. The decline in NP at high WC is due to increased CO2 diffusion resistances caused by blockage from capillary water and cell wall expansion [32]. Three examples are shown in 󳶳 Fig. 7.2 and also for two species in 󳶳 Tab. 7.1. Diploschistes diacapsis has a WCopt of 0.5 mm and a maximal WC of 1.2 mm, whereas for the second lichen, Psora decipiens, the equivalent values are 1.2 mm and 2.5 mm, respectively. Both species show a sharp maximum in NP. In contrast, the moss has a WCopt of 1.2 mm and a maximal WC of 3.9 mm. In addition it shows a relatively small decline in NP from WCopt to around 3.6 mm. This is a reasonably general difference with bryophytes having higher WCopt and maximal WC than lichens. Both lichens and bryophytes show a wide range in their response curves and these appear to be adaptive. For example, the very low WCopt and maximal WC values for D. diacapsis appear to allow the species to benefit from dew fall [23].

7.2.5 Response of Net Photosynthesis to CO2 Concentration Net photosynthesis typically shows a similar form of saturation response to CO2 concentration as shown for light (󳶳 Fig. 7.1a). Most lichens require around 1000 ppm CO2 to saturate NP while mosses and liverworts, despite normally having single-cell thick leaves require around 1500 ppm CO2 . There is little information available for BSC, but studies on cyanobacterial dominated BSC show a linear response of NP to 1000 ppm CO2 [33]. The actual CO2 concentration around and/or within BSC remains enigmatic. There is evidence from many ecosystems from Antarctic mosses to rain forests that actual CO2 levels close to the soil surface can be higher than global CO2 concentrations due to an efflux of CO2 from the soil [34]. CO2 concentrations within the soils covered with BSC can reach 1200 ppm and are almost always above the ambient atmospheric levels [33, 35]. Such concentrations indicate a continual efflux of CO2 from the soil and must include sources in addition to recycling of BSC fixed carbon. Possible major sources are higher plant roots and associated mycorrhizae. The latter can receive up to 20% of the carbon fixed by the host plant [36].

7.3 Optimal Versus Real Photosynthetic Rates According to the response curves presented in 󳶳 Fig. 7.1a,b, 󳶳 Tab. 7.2, BSC at optimal WC will reach NPmax at a light level ≥ 500 μmol m−2 s−1 and temperatures ≥ 15°C. Higher light levels will have no effect on NP as most BSC seem to be well protected against excess light. Higher temperatures will lead to lower NP but not in the underlying photosynthetic rate until GPmax is not reached at around 30°C. From these data it might be expected that the normal habitat of BSC in arid areas is one of high light and moderate to high temperatures.

130 | 7 Limits of Photosynthesis in Arid Environments

Active Inactive

10

(a)

Number of data points

1000 1000 Inactive Active

800

800

600

600

400 200

400

0

0

200 400 600 800 1000

200 0

0

1600 1400 1200 1000 800 600 400 200 0

20 30 40 Temperature

50

60

0 (b)

500 1000 1500 2000 2500 3000 PPFD (100 μmol m–2 s–1 bands)

1000

Lichen Psora decipiens

1000 800

800

600

600

400 200

400

0

0

200 400 600 800 1000

200 0

0 (c)

Moss Didymodon rigidulus

Number of data points

1600 1400 1200 1000 800 600 400 200 0

Number of data points

Number of data points

In reality, all BSC photosynthetic organisms are poikilohydric and will only be active when hydrated. It is, therefore, necessary to distinguish between conditions when the organisms are active and when they are inactive. In the latter case they are typically resistant to extremes of light, desiccation and temperature [23]. With the exception of the rare example where fruticose lichens become active solely following equilibration with humid air [37] BSCs in hot arid areas are hydrated either by rain or by dew [38, 39] and in the cold Antarctic desert by melt water [25]. Dew and rain produce different patterns of activation for mosses and lichens in BSC. Activation by dew starts for both mosses and lichens during the night and ends in the morning soon after sunrise as they desiccate. The net result is that the organisms are active at lower temperatures and light levels than the overall conditions for the habitat. In particular, dry lichens and mosses become very hot, reaching over 60°C, because they are good insulators when dry. In contrast, rain can activate the BSC at any time of day. Both lichens and mosses rapidly activate and can stay so for several

10

20

30

Temperature

40

50

60

0 (d)

500 1000 1500 2000 2500 3000 PPFD (100 μmol m–2 s–1 bands)

Fig. 7.3: Distribution of active and inactive times (number of data points in year) in relation to temperature (a,c, 5°C bands) and light (b,d, 100 μmol m−2 s−1 bands) for the moss Didymodon rigidulus (a,b) and the lichen Psora decipiens (c,d) forming BSC at Tabernas Desert, Spain. Left hand panels: activity (left hand black bars) and inactivity (right hand gray bars); right hand panel: activity (right hand red bars) and inactivity (left hand black bars). Note: active and inactive bars are reversed in left and right hand panels.

7.4 Limits to Photosynthesis in Arid Areas

|

131

days but, once again, both temperature and incident light are lower than optimal values because of the cloud cover. Net photosynthesis follows the same pattern with a so-called gulp in the early morning after dew activation [39]. The contrast between temperature and light levels when active and when inactive is shown in 󳶳 Fig. 7.3. The data are from continuous monitoring at Tabernas Desert, Almeria [38, 39] for the year 2013 and the lichen P. decipiens and the moss D. rigidulus. Both species behave very similarly to PPFD when active, concentrated below about 500 μmol m−2 s−1 , although when inactive levels can reach 2500 μmol m−2 s−1 . For temperature, activity is concentrated below 20°C, although both species can reach 60°C, and most activity is at around 7.5°C for the moss and 12.5°C for the lichen. From August to March the majority of the active time is at night, as one might expect from dew activation lichens and mosses, while in summer months activity is mainly in the daytime, reflecting rain activation [39]. The pattern of different, suboptimal conditions when active has also been well documented by continuous monitoring in Antarctica [25]. Schlensog et al. [40] showed that mean light levels when active increasingly differ from overall incident light as the proportion of the time that the organisms are active declines.

7.4 Limits to Photosynthesis in Arid Areas 7.4.1 Length of Active Time Because of their poikilohydric lifestyle it is no surprise that the greatest limiter of photosynthesis by BSC in arid zones is water availability. 󳶳 Fig. 7.4a shows the annual run of activity for BSC in the Tabernas Desert, Spain (the annual precipitation is 230 mm but variable) obtained by continuous chlorophyll fluorescence monitoring [39]. The mean monthly time active for three lichens and one moss over 1 year was 20.7% ± 3.6 with a low of 0.0% in June and high of 74.7% in November (󳶳 Fig. 7.4a). Activity in the dark typically exceeds that in the light, especially in the high activity months, so that BSC were active in the light only 8.3% of the total time (󳶳 Fig. 7.4a). However, carbon gain only occurs at light levels above the photosynthetic compensation point. Activity in the year 2013 and for the moss D. rigidulus and lichen P. decipiens were 10.3% and 11.4%, respectively, and applying compensation points of 70 and 80 μmol m−2 s−1 gives a carbon gain only for 2.8% and 4.0% of the year, respectively. Carbon loss through respiration occurs for about twice as long as positive NP, albeit mainly at lower temperatures at night. A similar pattern is summarized for six lichens by Evans and Lange [41] and is a further indication that low water availability severely limits photosynthetic carbon gain by BSC.

132 | 7 Limits of Photosynthesis in Arid Environments

1.0 60

0.5 0.0

40 20 0

Cumulative time active (%)

1.5

Light/dark ratio

80

Sep Oct Nov Dec Jan Feb Mar Apr May Jun Jul Aug Sep Oct

Proportion of time active (%)

Temperature (°C)