Soil Islands Self-Conditioned by Welwitschia Mirabilis in the Namib Desert

Home , Acanthosicyos, Acrospermum, Namib

‘Allelofertile’ soil islands self-conditioned by Welwitschia mirabilis in the Namib Desert

Thesis by Dalia Hamzah Shabaan

In Partial Fulfillment of the Requirements For the Degree of Master of Science

King Abdullah University of Science and Technology Thuwal, Kingdom of Saudi Arabia

July, 2019

Shaaban D.H. 2 Master Thesis

EXAMINATION COMMITTEE PAGE

The thesis of Dalia Hamzah Shaaban is approved by the examination committee.

Committee Chairperson: Daniele Daffonchio Committee Members: Peiying Hong, Arnab Pain

Shaaban D.H. 3 Master Thesis

Shaaban D.H. 4 Master Thesis

ABSTRACT

Under the extreme arid conditions of deserts, long periods of drought, nutrient-poor soils and high temperatures severely challenge the primary productivity of the ecosystem. Desert plants have evolved morphological and physiological adaptations against abiotic stresses. Along with these adaptation strategies they can recondition their surrounding soil, which will result in the enrichment of nutrients and moisture in the soil surrounding the plant. Although such self-fertilization may support the growth of other sympatric plant species under the plant, competitive exclusion mechanisms (i.e., allelopathy) reduce this possibility. Consequently, this will affect the diversity and functionality of the edaphic microbial communities. I hypothesize that desert xerophytes recondition the soils surrounding their body along with combining the ‘fertility’ and ‘allelopathy’ mechanisms to create a favorable new niche in desert ecosystem. I tested this hypothesis on the soil reconditioned by Welwitschia mirabilis growing in its native environment, the Namib Desert, Namibia. The collected soils were first used to confirm that Welwitschia manipulates the surrounding soil creating a ‘fertile’ but ‘exclusive’ soil area around the plant. Along with evaluating the effect of the reconditioned soil on the germination and plant development under normal irrigation and controlled drought condition, using barley as phytometer. The physio-chemical (i.e., WHC and WP) and microbial community analyses demonstrate that W. mirabilis reconditions the surrounding soil creating an environmental gradient around itself, in which the fertility is increased, through the accumulation and incorporation of shed reproductive parts of the plants (i.e., cones) in the surrounding soil, that will stimulate the plant growth under drought stress. Along with the fertilization effect, soil reconditioning also favor the antagonist effect (i.e., allelopathy) against plant competitors (e.g., new germinating seeds) to protect its ecological niche. Furthermore, the microorganisms and/or soluble/thermolabile molecules contribute to the allelopathic effect activated by the soil-reconditioning around W. mirabilis. The

Shaaban D.H. 5 Master Thesis interactions among W. mirabilis, soil and microbes highlight an adaptive strategy that combines soil fertilization and allelopathy that I defined as “Alleolofertility” strategy. This allelofertility island surrounding the W. mirabilis may contributes to explain the evolutionary success of such a ‘living fossil’.

Shaaban D.H. 6 Master Thesis

ACKNOWLEDGEMENTS

I would like to express my sincere gratitude to my advisor professor Daniele Daffonchio for giving me the opportunity to work in his lab and for his guidance, immense knowledge, encouragement, and support throughout the course of this research. Besides my advisor, I would like to thank my thesis committee: Prof. Peiying Hong and prof. Arnab Pain for their insightful comments, and support. A very special gratitude goes out to Dr. Ramona Marasco, the research scientist. For being the BEST teacher and mentor, I have ever had, she has taught\helped me more than I could ever give her credit for here. Thanks for teaching me with all you heart, thanks for your encouragement, motivation, enthusiasm and patience with me throughout this journey. You have shown me, by example, what a good scientist should be. This work would not have been possible without you. Thanks to Gillian Maggs-Kölling and Eugene Marais from the Gobabeb Training and Research Center (Namibia), and Don A. Cowan and Jean-Baptiste Ramond from the University of Pretoria (South Africa) for their collaboration in this project. I am also grateful to all my fellow lab mates, who I had the pleasure to work with during my research. With a special mention to Sadaf Umer, Alan Barozzi and Gregoire Michoud for their help and assistance. Last but not the least, a heartful gratitude is extended to my family and friends, whom love and guidance are with me in whatever I pursue. They are the ultimate role models. who provide unending inspiration.

Shaaban D.H. 7 Master Thesis

TABLE OF CONTENTS

EXAMINATION COMMITTEE PAGE 2 COPYRIGHT PAGE 3 ABSTRACT 4 ACKNOWLEDGEMENTS 7 TABLE OF CONTENTS 8 LIST OF ABBREVIATIONS 10 LIST OF ILLUSTRATIONS 11 LIST OF TABLES 12 AIM OF THE WORK 13 Chapter 1. Ecology, adaptation and evolution of meta-organisms in desert ecosystem 17

1.1. Introduction 17 1.2 Plant ecological adaptation and their role in desert soil reconditioning 20 1.2.1 Xerophytic and pheratophytic adaptation of plants to survive in 20 desert ecosystem.

1.2.2 Plant self-conditioning of soil in desert environment. 22 1.2.2.1 Establishment and stability of fertility islands. 22 1.2.2.2 Allelopathy mechanisms to favor plant establishment in desert fertility island. 24

1.3 Biodiversity of microbial communities in desert ecosystem: ecology and functional role of edaphic microorganisms. 26

1.3.1 Study of edaphic microorganisms in desert. 27 1.3.2 Edaphic microbial diversity harbored in desert. 28 1.3.3 Functional services of desert microorganisms and their role in soil fertilization and plant adaptation. 32

1.4 Conclusion 34 Bibliography 35

Shaaban D.H. 8 Master Thesis

Chapter 2. ‘Allelofertile’ soil islands self-conditioned by Welwitschia mirabilis in the Namib Desert 51

Abstract 51 2.1 Introduction 52 2.2. Results and Discussion 54 2.2.1 W. mirabilis behaviours and ‘wash’ site ecosystem 54 2.2.2 Microbial communities’ structure and diversity across W. mirabilis allelofertility island gradient 58

2.2.3 Microbial communities’ taxonomic composition and predicted 62 functions across the W. mirabilis allelofertility island gradient

2.2.4 Effects of soils from ‘allelofertility island’ on germination and development of phytometer 67

2.3 Conclusion 70 2.4 Material and Methods 71 2.4.1 Site description and sampling 71

2.4.2 Water holding capacity and wilting point measurements 72

2.4.3 Total DNA extraction from soil 72 2.4.4 Metaphylogenomic analysis of edaphic microbial (bacteria, archaea and fungi) communities 72

2.4.5 Beta- and alpha-diversity of microbial communities 74

2.4.6 In vivo experiment to evaluate the effect of Welwitschia- allelofertility island soils on seed germination and plant development 74 under watered and controlled-drought conditions.

Bibliography 75 Supplementary material 87

Shaaban D.H. 9 Master Thesis

LIST OF ABBREVIATIONS

BSCs Biological Soil Crust C Carbon Ca Calcium K Potassium N Nitrogen NGS Next Generation Sequencing OUT Operational Taxonomic Unit P Phosphorus PCoA Principal Cordinate Analysis PET Potential Evapotranspiration PGP Plant Growth Promoting qPCR Quantitative Polymerase Chain Reaction S Sulfate SOM Soil Organic Matter UNEP United Nations Environment Program WHC Water holding Capacity WP Wilting Pint

Shaaban D.H. 10 Master Thesis

LIST OF ILLUSTRATIONS

Chapter 1. Ecology, adaptation and evolution of meta-organisms in desert ecosystems Figure 1. Image and distribution of arid land ...... 19 Figure 2. Plant adaptation strategies (xerophytes and phreatophytes)...... 21 Figure 3. The biotic and abiotic processess driving the formation of fertility island ...... 23 Figure 4. Pathways and dynamics for release of allelochemicals into the environments ...... 25 Figure 5. Microbial community shifts from deserts soil to the same soil managed by desert-farming ...... 29 Figure 6. Phylogenetic affiliation of microbial community recovered from Thar Desert, India ...... 31

Chapter 2. ‘Allelofertile’ soil islands self-conditioned by Welwitschia mirabilis in the Namib Desert. Figure 1. Distribution and habitat of W. mirabilis ...... 56 Figure 2. Humidity and temparture range in Welwitschia wash ...... 57 Figure 3. Alpha diversity estimates of the microbial communities ...... 60 Figure 4. Welwitschia allelofertility island recruitment process ...... 61 Figure 5. Betadiversity of microbial communities associated with Welwitschia allelofertility island ...... 62 Figure 6. Microbial community composition along Welwitschia allelofertility gradient . 63 Figure 7. Effect of Welwitschia soil on seed germination and development ...... 68 Figure 8. Plant biomass under normal and drought conditions ...... 69

Shaaban D.H. 11 Master Thesis

LIST OF TABLES

Chapter 2. ‘Allelofertile’ soil islands self-conditioned by Welwitschia mirabilis in the Namib Desert. Table 1. Measurments of Water holding capacity (WHC) and wilting point of soil (WP) along the Welwitschia allelofertility gradient ...... 58 Table 2. Quality metrics of high-througput sequenncing (Miseq Illumina) analysis ...... 59 Table 3. Functional predictions of the bacterial communities along the Welwitschia allelofertility gradient ...... 64 Table 4. Functional predictions of archaeal communities along the Welwitschia allelofertility gradient ...... 66 Table 5. Functional predictions of the trophic categories of fungi along the Welwitschia allelofertility gradient ...... 67 Table 6. PCR primers and protocol for thermal cycling ...... 73

Shaaban D.H. 12 Master Thesis

AIM OF THE WORK

The polyextreme environmental conditions of desert (i.e., water limitation, high temperature level, lack of nutrients, and soil salinity) limit plant establishment, growth, and reproduction (Day & Ludeke, 1993; Noy-meir, 1974). Certain plants, known as xerophytes, have developed specific morphological and physiological adaptive strategies to survive in the harsh conditions of the desert (Fahn & Cutler, 1992; Wickens, 2013). Beside these adaptations, the physical and biotic interactions driven by the plant can positively recondition their local environment (Finzi et al. 1998; Lovett et al., 2004). The modification of the surrounding soil often results in the creation of ‘fertility islands’, in which organic matter, nutrients and moisture are enriched in the soil (Camargo-Ricalde and Dhillion, 2003; Ridolfi, 2008). To ensure their development, plants can also activate competitive exclusion mechanisms (i.e., allelopathy) to disfavor the possibility that other sympatric plants proliferate in their niche this competing for resources (Molisch 1937; Rice 2012). The net effect of these positive (fertility) and negative (allelopathy) adaptative mechanisms of desert plants is the creation of unique specific ecological- niches that have important effects on the overall dynamics of desert vegetation (diversity, productivity, and distribution of the plant communities; Chen & Zeng, 2003), as well as on the assembly and functionality of the edaphic microbial communities (Bonkowski et al., 2009; Muller et al., 2016). Microorganisms (i.e., bacteria, archaea and fungi) are among the first and main colonizers of desert habitats, largely contributing in determining soil structure and composition (Bonkowski et al., 2009). Soil- and plant-associated microorganism play important roles in many aspects of desert ecosystem, among other they may help in preventing soil erosion and improve growth of desert plants (Eida, 2018; Soussi, 2016). In this scenario, understanding how plant, soil and microbes interact together and benefit each other, can provide new information regarding the strategies and mechanism that may enhance desert plant establishment and resistance to stresses.

Shaaban D.H. 13 Master Thesis

My work is part of an international project in collaboration with the Gobabeb Training and Research Center and the University of Pretoria that has as main purpose the study of the desert microbiome. Particularly, my Master thesis is aimed to verify if deserts plants can combine the ‘fertility’ and ‘allelopathy’ mechanisms previously described in literature (Camargo-Ricalde, 2003; Hierro, 2003; Ridolfi, 2008; Rice 2012) to create a more favorable niche in desert ecosystems. First, I intended to evaluate the reconditioning-process driven by desert plants using two main indicators of soil modification: i) the physio-chemical properties of soil (i.e., organic matter and water capacity) and ii) the associated edaphic microbial community (i.e., diversity, composition and functionality). Furthermore, I assessed the effect of the reconditioned soil on the growth and development of surrounding plants to prove the capacity of desert plant to combine fertilization and competitive mechanisms during stressful condition (i.e., drought).The extreme arid environment, such as the gravel plain of Namib Desert, has been selected to study the formation of ‘allelofertility’ island mediated by the “living fossil” plant Welwitschia mirabilis. I tested the fertility and allelopathy of soil surrounding W. mirabilis in microcosms assays by using a bioassay based on the germination of allochthonous seed (e.g., barley) and their development under different levels of stress. I also extracted the total DNA from the soil reconditioned by W. mirabilis and performed high-throughput metagenomic sequencing of bacterial, archaeal and fungal communities. The overall aims of my study were to: i) reveal the environmental conditions of the soil reconditioned by W. mirabilis (i.e., physio-chemistry and associated microbiome), ii) understand the ecological importance of the plant-soil-associated microbial community, and iii) test the hypothesis that W. mirabilis create an ‘self-conditioned allelofertility island’ to increase its fitness in the harsh conditions of the Namib Desert.

Shaaban D.H. 14 Master Thesis

Chapter 1: Ecology, adaptation and evolution of meta-organisms in desert ecosystems.

Abstract. Plants inhabiting desert ecosystem evolved specific morphological and physiological adaptations that allow them to survive the polyextreme conditions imposed by arid environment, such as low nutrients and moisture level. Beside these adaptations, plants are able to self-condition the surrounding microenvironment creating more favorable conditions for their survival by improving soil fertilization and their potential competitiveness. The modification of plant-microenvironment is a fundamental ecological adaptation for the plant itself, as well as for the overall ecosystem that benefit of it (i.e., desert organisms), including the microbial communities associated with such soils. In this review, I provided a critical overview on the edaphic microbial communities (i.e., bacteria, archaea and fungi) associated with desert soils and their role in the soil surrounding desert plants. The composition, structure and role of the edaphic microorganisms affected by plant soil-conditioning are discussed. A deeper understanding of the factors affecting microbial communities, population dynamics and interaction in the complex mechanisms of soil reconditioning processes by desert plants would help to understand the direct and indirect strategies evolved by xerophytic metaorganisms (plant and microorganisms) to ameliorate the overall plant fitness and resilience in deserts.

Keywords. Deserts, Plant xerophytic adaptation, Soil reconditioning, Microbiome, Fertility Island, Fertilization, Competition, Allelopathy

1.1. Introduction

Deserts (hot and cold) are dry-barren regions of sand or rocks/gravel (Figure 1.1A and B) having a moisture deficit over the course of the year (i.e., <200 mm of rain per year;

Shaaban D.H. 15 Master Thesis

Sandquist 2014, contrasted by a much higher water evapotranspiration). They cover about 35% of the Earth’s land area (Laity, 2009; Figure 1.1C), representing the most extensive terrestrial biome (Thomas, 1997; Laity, 2008). In desert areas, besides limited water availability, the elevated temperature and sun irradiation, high salinity and low nutrients further contribute to create a hostile environment for life (Chamizo et al., 2012; Lester, Satomi and Ponce et al., 2007; Sandquist, 2014; Stomeo et al., 2013). According to the UNEP (1992), deserts are defined as those regions with an aridity index (ratio among annual precipitation and potential evapotranspiration, P/PET) of less than one (Whitford, 2002; Tsakiris & Vangelis, 2005; Stomeo et al., 2013), indicating how losses of moisture through evaporation are more frequent than the annual precipitation received (Deserts, 2018). The polyextreme conditions of deserts (i.e., low resource availability, extreme temperatures and water scarcity) are the environmental selective forces driving the distribution and diversity of organisms (Noy-meir, 1974; Gutterman, 2002; Makhalanyane et al., 2015). Within these organisms, vegetation is an important component defining the desert biome. The first most important challenges for desert plants are water scarcity and low nutrient content of arid soils, in which organic matter is low (Aroca et al., 2012, Wallace, 1980). In these conditions, only specialized plants can survive and exhibit specific xerophytic and phreatophytic adaptive traits, including morphological and physiological changes of both above- and below-ground compartments (Sandquist 2014; Wickens, 2013; Danin, 1996, Fahn & Cutler, 1992). Such adaptations optimize water management by reducing the amount of water lost by evapotraspiration (i.e., xerophytes; Fahn & Cutler, 1992) and favoring moisture uptake at or near the water table (i.e., phreatophytes; Wickens, 2013). Besides these adaptative strategies, plants recondition the surrounding soil creating favorable niches for their establishment and growth. During this process, the soil surrounding the plant is enriched in nutrient and organic matter, with a consequent improvement of its water holding capacity (fertility island effect; Ridolfi, 2008). At the same time, strategies to enhance competition and exclusion of other plants (i.e.,

Shaaban D.H. 16 Master Thesis allelophaty) are also implemented, in order to limit the competition for the limited resources by other plants (Molisch 1937; Rice 2012). Such plant-mediated soil reconditioning strategies can remarkably affect the overall soil chemistry, as well as on the edaphic microbial community (Bonkowski et al., 2009; Muller et al., 2016).

Figure 1.1. Image and distribution of arid land. (A and B) Pictures of sandy dune and gravel plain areas of Namib Desert, Namibia, respectively. (C) Distribution map of non-polar arid land; map available online at https://pubs.usgs.gov/gip/deserts.

Microorganisms (i.e., bacteria, archaea, and fungi) are considered, together with the plants, one of the most important components of desert ecosystems (Bonkowski et al., 2009). They are defined as the ‘biological engine of the earth’ (Ritz et al. 2004) and are the first to colonize arid and poor soils (Mapelli et al. 2012; Borin et al. 2010).

Shaaban D.H. 17 Master Thesis

Microorganisms maintain and manage the soil structure and composition, modulating several processes within soil, including those associated with fertility (e.g., organic matter decomposition; Gregorich & Carter 1997). In arid environments, beneficial edaphic microorganisms are essential not only for soil but also for plant-metaorganisms; they interact with soil, and (directly or indirectly; Spaepen et al., 2009) with plants, providing valuable ecological services to promoting its growth, protection, and productivity (Bashan & Holguin, 2002; Marasco et al., 2012; Rodriguez et al. 2008) In this context, understanding the ecology of the edaphic microbiome harbored by the soil reconditioned by desert plants may improve the efforts to comprehend the overall ecological processes taking place during plant-metaorganisms adaptation in desert ecosystems. Thus, here I assess the available knowledge on the composition, structure and functionality of the microbial communities (i.e., bacterial, archaeal and fungal) associated with soil reconditioned by plants in hot deserts, categorizing the plants self- reconditioning of soil in two main categories: (i) fertilization (Ridolfi, 2008) and (ii) competition/exclusion (Rice, 2012) strategies.

1.2. Plant ecological adaptation and their role in desert soil reconditioning

1.2.1. Xerophytic and phreatophytic adaptation of plants to survive in desert ecosystem. Desert is an extreme environment in which only adapted organisms can withstand (Wickens, 2013; Danin, 1996, Fahn & Cutler, 1992). Although the abiotic stresses of desert (i.e., water limitation) create various challenges to its’ flora (Thorup, 1969; Aroca et al., 2012), desert plants evolved numerous mechanisms (i.e., morphological, physiological and ecological) dedicated to improve their water management by i) reducing transpiration, ii) increase water absorption and iii) meliorate water storage (Sandquist, 2014; Smith, 1998). Drought tolerance and avoidance are the main strategies used by desert plants for optimal fitness and success (Hesp, 1991; Schuster, 2016). Drought tolerant plants include xerophytes, halophytes, phreatophytes,

Shaaban D.H. 18 Master Thesis and succulents, while drought avoidant plants include annuals and ephemerals (Singh, 2015; Vishwakarma, 2017).

Figure 1.2. Plant adaptation strategies (xerophytes and phreatophytes). (A) Distinction between xerophytes and phreatophytes plants adaptation strategies; (image taken from www.slideshare.net/spottyzebra/desert-biomes. (B) Pencil cholla inhabit in Joshua Tree National Park; it is a xerophytic plant has evolved its leaves in spines for avoiding water evapotranspiration. (C) Acanthosicyos horridus occupy the dunes of Namib Desert, Namibia; it is a phreatophytic plant with long taproot involved in the uptake of deep soil water from the ephemeral rivers of this desert (e.g., Kuiseb).

In sandy and gravel hot desert, xerophytic and phreatophytic plants (Figure 1.2A) are the most abundant. The first category, the xerophytic plants, evolved specific adaptative traits to store as much of the water available and overcome water loss altering stomata characteristics and the photosynthetic physiology, developing small leaves or even none at all (e.g., cacti; Figure 1.2B; Mares, 2017). On the contrary, phreatophytes plants have long taproots (up to 50 mt) able to reach the water table in order to access the tap ground water supplies and survive during long periods of drought (e.g., Acanthosicyos horridus, Figure 2C; Klopatek, 1994). Due to this adaptation, phreatophytes have a relatively constant supply of water and therefore do not need to store it (Robinson, 1958).

Shaaban D.H. 19 Master Thesis

1.2.2 Plant self-conditioning of surrounding soil in desert environment

1.2.2.1 Establishment and stability of fertility islands. Depict the complex processes activated by the interaction among plant and soil is a key point to understand the stability and functionality of vegetation in response to the polyextreme conditions of deserts (Ehrenfeld et al. 2005). In arid soil, nutrients are concentrated in the surface because nutrients input mainly occur via dust, weathering and deposition of plant litter (Jobbágy, 2001). The reduced amount of rainfall further reduces the downward leaching of nutrient into soils (Lehmann, 2003). Desert plants (i.e., shrubs and trees) play an important role in the vertical and horizontal redistribution of soil nutrients (i.e., C, N and P; Jobbágy, 2001; Yao, 2017). They evolved the capacity to self-reconditioning the surrounding soil, modifying and redistributing nutrients and organic matter (Finzi et al. 1998; Lovett et al., 2004). This process results in a spatial heterogeneity of soil sources (Garcia-Moya and McKell et al., 1970; Schlesinger et al., 1990, 1996), in which a nutrient-rich zone capable of improved water retention, defined as ‘fertile islands’ (or ‘resource islands’) is surrounded by low-nutrient soils (i.e., inter-plant area; Schlesinger et al., 1996; Camargo-Ricalde and Dhillion, 2003). Such nutrient-rich zones originate from the complex interaction of biotic and abiotic processes that start with the accumulation of plant litter (i.e., dry plant material) under plant canopy and the surrounding area (Figure 1.3). At the same time, the transport of organic matter and biological material mediated by animal, wind and water (Chew and Whitford et al., 1992; Garner and Steinberger et al., 1989), along with the overgrazing disturbance (Kieft et al., 1998), contribute to the establishment of soil niche in which fertility is enhanced. Examples of desert plants able to manipulate the composition and properties of the surrounding soil are the spiny trees and shrubs belonging to the Prosopis genus (Fabaceae, Mimosoideae; Rossi & Villagra, 2003; Nobel, 1988; Schade & Hobbie, 2005). The autogenic fertilization of these plants significantly increases the concentration of organic matter, phosphorus, potassium and nitrogen under their canopies. In the study of Saixiyala et al., 2017, organic matter, total nitrogen and water content were

Shaaban D.H. 20 Master Thesis significantly higher in the soil beneath the Caragana intermedia shrub canopies than in the soil outside the canopies. A similar strategy is adopted by Welwitschia mirabilis, a ‘living fossil’ endemic to Namib Desert. Abrams and colleagues (1997) demonstrated that soils surrounding W. mirabilis plants significantly increase nutrients accumulation (i.e., organic carbon, nitrogen, phosphate and electrical conductivity) than the soils between plants. The majority of litter is composed by shedding of reproductive parts (i.e., flowers and cones) of the plants that are slowly incorporated into the soil (Figure 1.3C). The leaves are leathery and massive and not represent the main source of biological material (litter), but are fundamental to create a microenvironment with relatively moderate thermal and moisture regimes that facilitate an increased resilience to drought (Schulze et al., 1980; March, 1990). As measured by March (1990), the soil below W. mirabilis reaches 40°C and a moisture content of 0.5%, while the non- colonized soil exposed to sun was over 70°C with a significantly lower content of moisture (0.2%).

Figure 1.3. The biotic and abiotic processes driving the formation of fertility island. (A) Scheme showing the abiotic and biotic processes driving the development of fertility island under shrubs (modified from Li et al., 2017; Abrams et al., 1997). These processes may include (1) interception of windblown material, (2) root uptake, (3) enhanced infiltration, (4) litter deposition, (5) stemflow, and (6) throughfall. (B) Larrea tridentate, a creosote bush found in Western North America that can induce the formation of fertility island accumulating and entrapping litter under its canopy (Mabry, 1977). (C) Welwitschia mirabilis, endemic plant of Namib Desert, Namibia, able to enrich the nutrients and carbon contents in the surrounding soil (picture credit to Ramona Marasco).

Shaaban D.H. 21 Master Thesis

The abiotic (i.e., trapping of windblown materials) and biotic processes (i.e., deposition of litter) are not the only ones involved in the development of ‘fertile islands’ in dryland. In fact, the microorganisms inhabiting the soil (and their interactions) play an important role (Bashan & de Bashan, 2010; Makhalanyane et al., 2015; Kuske et al., 2002). They can modulate the nutrients availability, directly through the enrichment of organic matter and influencing the solubilization of several nutrients (i.e., P, K, Ca, S; Jones, 2011; Bolan, 2011); indirectly, by stimulating the growth of plants through mutualistic symbiosis’s with plants roots (i.e., rhizosphere and mycorrhizae; Cross and Schlesinger, 1999; Wardle et al. 2004; Whitford, 2002). The overall beneficial effect of nutrient accumulation and microenvironment formation is often coupled with an enrichment of beneficial microorganisms (e.g., Streptomyces spp. and N-fixing bacteria; Kuske et al., 2002) and an increment of the soil enzymatic activities (Thompson et al., 2006), such as microbial β‐glucosidase and polyphenol oxidase (Yao et al., 2019). These extracellular enzymes mediate the degradation, transformation and mineralization of soil organic matter starting from the cellulose and lignin components of litter, respectively (Thomas, 2012; Sinsabaugh 2010). Soil texture and soil organic matter (SOM) are key factors determining soil water holding capacity (Carter, 2002). Soil texture determines the surface to volume interaction with water, so the larger the surface area (i.e., smaller particle sizes of silt and clay), the easier it is for the soil to hold water (Hillel, 2012). Soil organic matter (SOM) from decayed material originated from living organism (i.e., plant litter) favors water retention and increments in soil SOM always positively correlate with gains in water holding capacity (Abrams et al 1997; Brady, 2008; Kögel-Knabner, 2002).

1.2.2.2 Allelopathy mechanisms to favor plant establishment in desert fertility island. During the creation of self-advantageous soil environments, plants can either positively (favoring; Figure 1.3B) or negatively (excluding; Figure 1.3C) affect the establishment of other neighboring grasses beneath their canopies (Saixiyala et al., 2017; Rossi & Villagra, 2003), strongly influencing the overall plant community structure and composition

Shaaban D.H. 22 Master Thesis

(Berg, 2009). Plants developing exclusion mechanisms such as allelopathy preserve soil fertility for their growth. These ‘allelopathic plants’ activate biological mechanisms in order to reduced seed germination and seedling growth of other plant species in their proximity (Molisch 1937; Rice, 2012; Cipollini, 2012). This process starts with the release in the surrounding soil of secondary metabolites, known as allelochemicals, from different plant tissues of the root system or the canopy and/or by decomposition, leaching and volatilization of organic matter residues (Cipollini, 2012; Figure 1.4A). Allelopathic inhibition mediated by plants is complex and can involve the interaction of different classes of chemicals, such as phenolic compounds, flavonoids, terpenoids, alkaloids, steroids (Li, 2010; Figure 1.4B). Furthermore, physiological and environmental stresses (i.e., limited nutrient and moisture) can also enhance the allelopathic suppression of competitor plants (Pedrol, 2006).

Figure 1.4. Pathways and dynamics for release of allelochemicals into the environment. (A) Schematic representation of possible pathways of release into the environment: (1) exudation and deposition on the leaf surface with subsequent washing off by rainfall; (2) exudation of volatile compounds from living green parts of the plant; (3) decay of plant residues (e.g., litter and dead roots); and (4) root exudation. (B) Input and output dynamics of allelochemicals in soil. In soil, diverse factors affect the availability of allelochemicals, and consequently their effective influence on target plants. Leaching, physiochemical processes, microbial breakdown and uptake by plants are factors that can reduce the soil concentration of allelochemicals, while incorporation/ absorption in/ by soil organic matter can favor their activity (from de Albuquerque et al., 2011).

Shaaban D.H. 23 Master Thesis

The increment and reduction of the allelopathic effect can be also modulated by the microbial degradation of natural product and allelochemicals (Cipollini, 2012). Soil microorganisms, in particular, Fusarium, Pseudomonas and Thielaviopsis can influence the bioavailability of allelochemicals (e.g., phenolic acids and alkaloids) by the decomposition of plant residues in soil (Cipollini, 2012). At the same time, allelopathic molecules can be not only converted, but also directly released by soil-borne microorganisms (first study of Kaminsky, 1981). Kaminsky (1981), with soil transfer experiments, demonstrated the inhibition of plant growth close to Adenostoma fasciculatum (a shrub of Rosaceae family) was due to soil-borne microorganisms and not to shoots leachates; when soil under canopy was transferred in an open field, phytotoxicity persists, while when soil from control sites was placed under the shrub, plants can colonize the area. Fumigation of soil under the canopy also reduced the phytotoxicity. Similar results were obtained in Israel, studying the competitive mechanisms of Coridothymus capitatus (Labiatae; plant density 16-fold less under the plant compared to intra-shrub soil; Katz et al., 1987). Microcosms experiments showed that annuals (e.g., Plantago psyllium and Erucaria hispanica) had reduced seed germination of 45% when grown in shrub-affected soil (Katz et al., 1987). Notably, treatment of soil with allelopathic plant litter increased bacterial populations (Vokou et al., 1984), with an enrichment of actinomycetes (36.2-fold compared to control soil among shrubs; Katz et al., 1987) possibly involved in the transformation/secretion of allelochemicals. Nevertheless, microorganisms may suppress plants also through competition, for instance by immobilizing nutrients (Jenkinson, 1998; Wardale and Nilsson, 1997). Details on the direct and indirect allelopathic mechanisms driven by microorganisms are discussed in the next section.

1.3 Biodiversity of microbial communities in desert ecosystem: ecology and functional role of edaphic microorganisms

Shaaban D.H. 24 Master Thesis

Microorganisms constitute the dominant life form on earth, and are the basis for the evolution of more complex life (i.e., animal and plant; Zavarzin, 2003; Madigan and Martinko 2006). Microorganisms (i.e., fungi, archaea and bacteria) have colonized extensively almost all Earth’s biospheres, including extreme environments (Rampelotto, 2013). The most important factor shaping ecosystems and regulating microbial communities is climate (Breshears et al., 2008; Kelly and Goulden, 2008; Noy-Meir 1974), and deserts, having limited water and nutrient availability, extreme temperatures, high radiation and limited availability of organic carbon (Sterflinger et al. 2012), provide one of the most extreme environmental conditions for microbial life (Direito et al. 2011).

1.3.1 Study of edaphic microorganisms in deserts. Deserts host a complex and biodiverse ecosystem containing billions of microbes that live, reproduce and interact each other’s in a single gram of sand/soil (Raynaud, 2014; Vollmer, 1997) and include bacteria, archaea and fungi (Bonkowski et al., 2009; Muller et al., 2016). Each community has its unique characteristics that correspond to the different functions they play in the ecosystem (Rousk et al., 2010). Moreover, the structure and abundance of such communities are mainly influenced by the range and severity of several factors (i.e., abiotic stresses) affecting the inhabited environment (Lauber et al., 2009; Angel et al., 2010; Rousk et al., 2010; Stomeo et al., 2012). Microorganisms play important roles in soil, such as chemical transformations and fertilization processes (Jacoby, 2017). While early studies supported the concept of desert ‘sterility’ by very low levels of viable/cultivable microorganisms (due to the limited portion [<1%] of environmental microorganisms cultivable; Riesenfeld et al. 2004), the applications of DNA/RNA-based molecular methods showed a contrasting picture (among others, Fierer and Jackson, 2006). The development of molecular culture-independent methods made possible to study the total microbial communities, leading to new insight and deepened our understanding of microbial life in desert ecosystem (Daffonchio et al., 2015; Riesenfeld et al. 2004). Metagenomic and PCR–

Shaaban D.H. 25 Master Thesis based methods including next generation sequencing [NGS, Tringe, 2005] and fingerprinting techniques [denaturing gradient gel electrophoresis (Muyzer et al. 1993), terminal-restriction fragment length polymorphism (Osborn et al. 2000)] have been developed to study the total microbial DNA extracted from environmental samples (Rastogi and Sani, 2011). These approaches do not rely on any form of cultivation, nor on any prior knowledge about the microbial communities present in the samples (Riesenfeld et al. 2004). These approaches revealed large portions of previously undetected microbial groups and highlight the potential importance of these previously cryptic taxa in extreme environments (de Los Ríos et al. 2010; Andrew et al 2012).

1.3.2 Edaphic microbial diversity harbored by desert. In arid and semiarid soils, the composition of soil microorganisms is significantly different form the one hosted my other soil biomes (e.g., forest, temperate soils; Fierer et al., 2012). Several studied revealed that desert arid and semi-arid soils host a reduced number of microbes, both in term of cell abundance and diversity, in comparison with desert soils managed with desert-farming techniques (Köberl et al., 2011, 2013, 2016; Marasco et al 2018; Makhalanyane et al., 2015). For instance, it has been observed that the desert-farming soils host approx. 1500- and 1000-times more gene copies of bacteria and fungi, respectively, than the bulk sand with the same origin but not managed and such communities were characterized by a higher phylogenetic diversity (richness) than the non-managed desert soil (Figure 1.5). The harsh environmental conditions largely common to all deserts, along with the microbial cell inputs determined by the circulation of dust associated to sand storms, determine common diversity traits in the microbial communities of different deserts (Nagy et al., 2005).

Bacteria. Despite the different geographic location of arid lands and deserts across the world (Figure 1.1C), the arid and semi-arid soils are consistently dominated by i)

Shaaban D.H. 26 Master Thesis heterotrophic bacteria belonging to Actinobacteria such as Rubrobacter, Arthrobacter and Thermopolyspora (Makhalanyane, 2015)) and Alphaproteobacteria such as Ochrobactrum (Koberl et al., 2011) and Firmicutes (Li and Kong, 2018); ii) photoautotrophic cyanobacteria, such as Pseudanabaenaceae, Chroococcidiopsis, Phromidium, Microcoleus (Rao and chan, 2016); iii) ascomycete fungi including Cladophialophora, Cladosporium, Leptosphaerulina, Alternaria; (Conley et al., 2006); iv) Euryarchaeota archaea such as Methanosarcina and Thermotoplasmata (Angel and Claus, 2012).

Figure 1.5. Microbial community shifts from desert soil to the same soil managed by desert-farming techniques. Quantitative and compositional analyses of microbial communities associated with desert and desert-farming soil in Egypt. Chemistry of soil are also reported (right and left bottom) to underline the enrichment of nutrients and organic carbon in desert-farming soil compared to barren desert sand. Figure from Köberl et al., 2011.

Molecular analysis (i.e., DNA extraction, pyrosequencing and real-time quantitative PCR approach) conducted on sandy deserts (e.g., Taklamakan desert, China; Gobi Desert,

Shaaban D.H. 27 Master Thesis

Mongolia [An et al. 2013]; Thar Desert, India [Rao et al., 2016; see Figure 1.6) confirm the dominance of heterotrophic bacteria of the Proteobacteria (12-25%), Firmicutes (25- 70%) and Actinobacteria (9-15%). Contrary, in the oldest and driest Atacama Desert (South America) and in the world-largest Sahara Desert (North Africa), Actinobacteria and Chloroﬂexi were the most abundant bacteria, with a low abundance of Acidobacteria and Proteobacteria (Neilson et al., 2012; Chanal et al., 2006). All the bacterial taxa ubiquitous of desert ecosystems have in common the evolution of multiple genetic and physiological mechanisms of resistance to arid and oligotrophic desert conditions; for instance, the Chloroflexi have a protective layer that envelope the cell (Tocheva, 2016), while Actinobacteria can create vegetative forms (e.g., spores) able to resist to long period of drought and low nutrient availability (Griffiths, 2017). Notably, Cyanobacteria are an important component of microbial communities both in hot and cold deserts (e.g., Sonoran and Taklimakan; Lacap-Bugler et al., 2017). Members of this phylum are photosynthetic or nitrogen-fixers, both fundamental functions in oligotrophic arid environments (Chan et al., 2013). They also contribute to stabilize the general conditions of desert soils, by improving soil stability, moisture retention and fertility (Belnap and Gardner 1993). The ability of cyanobacteria to withstand the stressful desert conditions (high UV irradiation, desiccation and water stress) provides a significant competitive advantage (Cockell and Knowland 1999; Starkenburg et al., 2011; Rajeev et al., 2013). Despite these advantages, cyanobacteria in the most extreme hyperarid deserts are generally restricted to protected sub-lithic niches and biological soil crusts (BSCs), with only limited cell numbers in surface soils (Makhalanyane, 2015). The stable associations between cyanobacteria and other organisms like fungi, lichens and mosses have a crucial role in stabilizing soil against erosion (de-Bashan et al., 2010, Bashan et al., 2012, Xu et al., 2013) and constitute a favorable niche for the germination of plant seeds (Belnap, 2013).

Shaaban D.H. 28 Master Thesis

Figure 1.6. Phylogenetic affiliation of microbial community recovered from the Thar Desert, India (modified from Rao et al., 2016). (A-C) Diversity of microbial community associated with arid soil (Jaisalmer), arid sand dune (Jaisalmer), semi-arid soil (Ajmer) and semi-arid endolithic (Ajmer) of the Thar Desert. Phylogenetic position is reported for bacteria (A) fungi (B) and archaea (C). Taxonomic affiliations are reported by color scale for each kingdom, while the soil origin is indicated by a letters-code: JB, arid soil (Jaisalmer); JSD, arid sand dune (Jaisalmer); LL, semi-arid soil (Ajmer); A, semi-arid endolith (Ajmer).

Fungi. Together with bacteria, also fungi have an important ecological role in desert ecosystems (Makhalanyane, 2015). Studies using both cultivation-dependent and - independent approaches revealed high level of fungal diversity in arid environments

Shaaban D.H. 29 Master Thesis

(e.g., Negev and Sonoran deserts Grishkan, 2010; Carrillo‐Garcia, 2010), according with their ability to tolerate stressful conditions (Waller et al., 2005; Sterflinger et al., 2012). The majority of fungi detected in the desert belong to the Basidiomycota and Ascomycota phyla and includes among others Cladosporium, Alternaria, Aspergillus, Penicillium, Periconia and Giberella (Conley et al., 2006). Examination of the phylotypes recovered from desert sand revealed the abundance of members capable of intense melanisation (e.g., Chaetothyriales; Sterflinger et al., 2012), which protects them from UV and solar irradiation.

Archaea. Finally, although they are relatively rare across many biomes, archaea are particularly abundant in desert soils (Fierer et al., 2012). The thermophilic and chemolithoautotrophic ammonia-oxidizers of the Thaumarchaeota phylum are the most abundant in arid and semi-arid soils (Treusch et al. 2005; Brochier-Armanet et al. 2008). Their ability to oxidize ammonia at low concentration play important roles in biogeochemical cycling of nitrogen in oligotrophic environments (Hatzenpichler et al., 2008). Recent metagenome sequencing of Indian desert soils found that stress-tolerant anaerobic Euryarchaeota phylotypes such as Methanosarcina were an important component of the desert soil prokaryotic community (Pandit et al., 2014; Rao et al 2016).

1.3.3 Functional services of desert microorganisms and their role in soil fertilization and plant adaptation. As consequence of the evolutionary pressure of moisture- and thermal-stress events, microorganisms have evolved special adaptation mechanisms for which only those genes required to survive and respond appropriately to the physical and chemical composition of these particular habitats are expressed and regulated (Bohnert et al., 1995; Boor 2006, Colica et al., 2014). Arid soil metagenomes revealed a higher abundance of genes associated with dormancy and stress response than in non- arid biomes (Fierer et al., 2012). Although microorganisms are the first soil colonizers (Mapelli et al., 2012) and have an important role in the stability and productivity of

Shaaban D.H. 30 Master Thesis deserts ecosystems (Belnap and Gardner 1993), their nutrient cycling rates are lower compare to more mesic habitats (Fierer et al., 2007a, 2012). The enhancement of soil fertility under plant canopies (i.e., fertility islands) promote microbial heterogeneity and consequently the functionality of the entire soil-microbiome and bio-geochemical cycles (Herman et al., 1995; Smith, 2015; Kuske et al., 2002). Soil beneficial bacteria and fungi are key drivers of litter and organic matters decomposition, and nutrient mineralization (Aguilera et al.,1999). Bacteria ameliorate plant nutrient status by providing nutrients that are largely unavailable to plants because they are stored in insoluble forms (Raddadi et al., 2007). For example, organic acids and extracellular enzymes produced by phospahte-solubilizing bacteria (i.e., Bacillus and Paenibacillus; Jorquera et al., 2008) increase the solubilization of soil phosphate, making it available for plant uptake (Puente et al., 2004). Similarly, the ferric iron (Fe2+) that can be easily taken up by plants often results from bacteria reduction of the low soluble Fe (OH)3 and from the chelating activity of microbial siderophores produced, among others, by Streptomyces, Bacillus, Pseudomonas and Fusarium; Egamberdiveya 2007). In desert soil, low nitrogen mineralization rates occur, especially in non-vegetated areas between shrubs (Austin et al., 2004; Schade and Hobbie 2005). This might induce the selection for diazotrophs and oligotrophic ammonia oxidizers microorganisms (e.g., cyanobacteria, green sulfur bacteria, rhizobia and

Thaumarchaeota; Marques, 2019) that fixing the atmospheric nitrogen in NH3 and

+ - oxidizing NH4 to nitrite (NO2 ), respectively, favor the nitrogen bioavailability and cycling (Makhalanyane, 2015). In the fertility island, plants develop their root system in the reconditioned soil (Bonanomi, 2008). Throughout the secretion of root exudates in the rhizosphere (defined as the thin layer of soil surrounding the root), plants establish an intimate chemical interaction that could result in a positive association with beneficial soil microbes (PGP, plant growth promoting), (Lugtenberg et al., 2009; Vessey et al., 2003). PGP microorganisms stimulate plant growth by both direct and indirect mechanisms, including (i) biofertilization (previously listed), (ii) biopromotion (mainly with the

Shaaban D.H. 31 Master Thesis synthesis of phytohormones such as auxins, cytokinins, gibberellins; Egamberdieva, 2017), and (iii) biocontrol of phytopathogen (activation of induced systemic resistance; Yang et al., 2009). PGP bacteria may regulate plant growth and development by using one or more of these mechanisms (Glick et al., 2007), as well as by activating allelopathic mechanisms (Cipollini, 2012; Figure 1.4). Soil toxicity (allelophaty) has been partly related to microbial activity that during alternating periods of drought and humidity accumulate toxins (Kaminsky, 1981). Friedman et al., (1989) demonstrated that in areas exposed to intermittent drought, drought-tolerant actinomycetes proliferate. Thus, water limitation alters microbial populations and the relative concentration of microbial allelochemicals in the soil resulting in different response/susceptibility of the plant to these molecules (Kennedy et al., 1991; Friedman et al., 1989). For instance, allelochemicals released by microorganisms may be specific (Scacchi et al., 1992; Hoagland, 1990) and act in different ways depending on the nutritional conditions of the soil (Barazani and Friedman 2001).

1.4 Conclusion

Plants have developed special adaptive strategies to overcome the polyextreme environmental conditions of deserts. The biotic and abiotic processes driven by desert plants (i.e., shrubs and trees) aimed to recondition their surrounding soil favor the accumulation of resources (i.e., nutrient and organic carbon). This process, create a favorable ecological niche for both plant and microorganisms, defined as “fertility island”. Microorganism colonizing this soil niche can provide essential ecological services such as bio-fertilization that can favor plant establishment (Li et al 2017). While microorganisms favor the availability of water and nutrients for the plant, in return the plant supplies carbon and nutrient resources for the microbial growth (Kuske et al., 2002). The increase in microbial activity will lead to enhanced accumulation of

Shaaban D.H. 32 Master Thesis resources (i.e., decomposition of organic matters, Jones, 2011), stabilization of soil structure and increased water retention (Carter, 2002). The processes of plant-soil-microbe interaction and habitat engineering in turn modify the diversity, assembly and distribution of soil microbial communities. The advances in molecular based approaches (DNA\RNA-based) can be applied to study the involvement of microorganisms in the formation and persistence of fertility islands and their role in sustaining ecosystem services for plant establishment and development in arid soils. Although there is an increasing number of studies describing the functional capacity of microbial communities, the link between community structure and function remains a central challenge in microbial ecology, especially in desert ecosystems.

Shaaban D.H. 33 Master Thesis

1.5 Bibliography

Abrams MM, Jacobson PJ, Jacobson KM, Seely MK. Survey of soil chemical properties across a landscape in the Namib Desert. Journal of Arid Environments. 1997 Jan 1;35(1):29-38. Aguilera JM, Stanley DW. Microstructural principles of food processing and engineering. Springer Science & Business Media; 1999 Sep 30. An S, Couteau C, Luo F, Neveu J, DuBow MS. Bacterial diversity of surface sand samples from the Gobi and Taklamaken deserts. Microbial ecology. 2013 Nov 1;66(4):850-60. Andrew DR, Fitak RR, Munguia-Vega A, Racolta A, Martinson VG, Dontsova K. Abiotic factors shape microbial diversity in Sonoran Desert soils. Appl. Environ. Microbiol.. 2012 Nov 1;78(21):7527-37. Angel R, Claus P, Conrad R. Methanogenic archaea are globally ubiquitous in aerated soils and become active under wet anoxic conditions. The ISME journal. 2012 Apr;6(4):847. Angel R, Soares MI, Ungar ED, Gillor O. Biogeography of soil archaea and bacteria along a steep precipitation gradient. The ISME journal. 2010 Apr;4(4):553. Aroca R, Porcel R, Ruiz-Lozano JM. Regulation of root water uptake under abiotic stress conditions. Journal of experimental botany. 2011 Sep 13;63(1):43-57. 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 Oct 1;141(2):221-35. Bashan Y, de-Bashan LE. Microbial populations of arid lands and their potential for restoration of deserts. InSoil biology and agriculture in the tropics 2010 (pp. 109-137). Springer, Berlin, Heidelberg. Bashan Y, Holguin G. Plant growth-promoting bacteria: a potential tool for arid mangrove reforestation. Trees. 2002 Mar 1;16(2-3):159-66.

Shaaban D.H. 34 Master Thesis

Belnap J, Gardner JS. Soil microstructure in soils of the Colorado Plateau: the role of the cyanobacterium Microcoleus vaginatus. Great Basin Naturalist. 1993;53(1):6. Belnap J, Gardner JS. Soil microstructure in soils of the Colorado Plateau: the role of the cyanobacterium Microcoleus vaginatus. Great Basin Naturalist. 1993;53(1):6. Belnap J, Lange OL, editors. Biological soil crusts: structure, function, and management. Springer Science & Business Media; 2013 Dec 1. Berg G, Smalla K. Plant species and soil type cooperatively shape the structure and function of microbial communities in the rhizosphere. FEMS microbiology ecology. 2009 Apr 1;68(1):1-3. Bohnert HJ, Nelson DE, Jensen RG. Adaptations to environmental stresses. The plant cell. 1995 Jul;7(7):1099. Bolan NS, Adriano DC, Kunhikrishnan A, James T, McDowell R, Senesi N. Dissolved organic matter: biogeochemistry, dynamics, and environmental significance in soils. InAdvances in agronomy 2011 Jan 1 (Vol. 110, pp. 1-75). Academic Press. Bonanomi G, Rietkerk M, Dekker SC, Mazzoleni S. Islands of fertility induce co-occurring negative and positive plant-soil feedbacks promoting coexistence. Plant Ecology. 2008 Aug 1;197(2):207-18. Bonkowski M, Villenave C, Griffiths B. Rhizosphere fauna: the functional and structural diversity of intimate interactions of soil fauna with plant roots. Plant and Soil. 2009 Aug 1;321(1-2):213-33. Boor KJ. Bacterial stress responses: what doesn't kill them can make them stronger. PLoS biology. 2006 Jan 17;4(1):e23. Borin S, Ventura S, Tambone F, Mapelli F, Schubotz F, Brusetti L, Scaglia B, D'Acqui LP, Solheim B, Turicchia S, Marasco R. Rock weathering creates oases of life in a High Arctic desert. Environmental microbiology. 2010 Feb;12(2):293-303. Brady NC, Weil RR, Weil RR. The nature and properties of soils. Upper Saddle River, NJ: Prentice Hall; 2008.

Shaaban D.H. 35 Master Thesis

Breshears DD, Huxman TE, Adams HD, Zou CB, Davison JE. Vegetation synchronously leans upslope as climate warms. Proceedings of the National Academy of Sciences. 2008 Aug 19;105(33):11591-2. Brochier-Armanet C, Boussau B, Gribaldo S, Forterre P. Mesophilic Crenarchaeota: proposal for a third archaeal phylum, the Thaumarchaeota. Nature Reviews Microbiology. 2008 Mar;6(3):245. Camargo-Ricalde SL, Dhillion SS. Endemic Mimosa species can serve as mycorrhizal" resource islands" within semiarid communities of the Tehuacán-Cuicatlán Valley, Mexico. Mycorrhiza. 2003 Jun 1;13(3):129-36. Carrillo‐Garcia Á, De La Luz JL, Bashan Y, Bethlenfalvay GJ. Nurse plants, mycorrhizae, and plant establishment in a disturbed area of the Sonoran Desert. Restoration Ecology. 1999 Dec;7(4):321-35. Carter MR. Soil quality for sustainable land management. Agronomy journal. 2002 Jan 1;94(1):38-47. Chamizo S, Cantón Y, Miralles I, Domingo F. Biological soil crust development affects physicochemical characteristics of soil surface in semiarid ecosystems. Soil Biology and Biochemistry. 2012 Jun 1;49:96-105. Chan Y, Van Nostrand JD, Zhou J, Pointing SB, Farrell RL. Functional ecology of an Antarctic dry valley. Proceedings of the National Academy of Sciences. 2013 May 28;110(22):8990-5. Chanal A, Chapon V, Benzerara K, Barakat M, Christen R, Achouak W, Barras F, Heulin T. The desert of Tataouine: an extreme environment that hosts a wide diversity of microorganisms and radiotolerant bacteria. Environmental Microbiology. 2006 Mar;8(3):514-25. Chew RM, Whitford WG. A long-term positive effect of kangaroo rats (Dipodomys spectabilis) on creosotebushes (Larrea tridentata). Journal of Arid Environments. 1992 May;22:375-86. Cipollini, D., Rigsby, C.M. & Barto, E.K. J Chem Ecol (2012) 38: 714. https://doi.org/10.1007/s10886-012-0133-7

Shaaban D.H. 36 Master Thesis

Cockell CS, Knowland J. Ultraviolet radiation screening compounds. Biological Reviews. 1999 Aug;74(3):311-45. Colica G, Li H, Rossi F, Li D, Liu Y, De Philippis R. Microbial secreted exopolysaccharides affect the hydrological behavior of induced biological soil crusts in desert sandy soils. Soil Biology and Biochemistry. 2014 Jan 1;68:62-70. Conley CA, Ishkhanova G, Mckay CP, Cullings K. A preliminary survey of non-lichenized fungi cultured from the hyperarid Atacama Desert of Chile. Astrobiology. 2006 Aug 1;6(4):521-6. Cross AF, Schlesinger WH. Plant regulation of soil nutrient distribution in the northern Chihuahuan Desert. Plant Ecology. 1999 Nov 1;145(1):11-25. Daffonchio, Hirt, & Berg. (2015). Plant-microbe interactions and water management in arid and saline soils. In Principles of Plant-Microbe Interactions (pp. 265–276). Springer International Publishing. Danin A. Plants of desert dunes. Springer Science & Business Media; 2012 Dec 6. de Albuquerque MB, dos Santos RC, Lima LM, de Albuquerque Melo Filho P, Nogueira RJ, Da Câmara CA, de Rezende Ramos A. Allelopathy, an alternative tool to improve cropping systems. A review. Agronomy for Sustainable Development. 2011 Apr 1;31(2):379-95. de los Ríos A, Valea S, Ascaso C, Davila A, Kastovsky J, McKay CP, Gómez-Silva B, Wierzchos J. Comparative analysis of the microbial communities inhabiting halite evaporites of the Atacama Desert. International Microbiology. 2010 Jun 1;13(2):78-89. De-Bashan LE, Bashan Y. Immobilized microalgae for removing pollutants: review of practical aspects. Bioresource technology. 2010 Mar 1;101(6):1611-27. De-Bashan LE, Hernandez JP, Bashan Y. The potential contribution of plant growth- promoting bacteria to reduce environmental degradation–A comprehensive evaluation. Applied Soil Ecology. 2012 Oct 1;61:171-89.

Shaaban D.H. 37 Master Thesis

Direito SO, Marees A, Röling WF. Sensitive life detection strategies for low-biomass environments: optimizing extraction of nucleic acids adsorbing to terrestrial and Mars analogue minerals. FEMS microbiology ecology. 2012 Jul 1;81(1):111-23. Egamberdieva D, Wirth SJ, Alqarawi AA, Abd_Allah EF, Hashem A. Phytohormones and beneficial microbes: essential components for plants to balance stress and fitness. Frontiers in microbiology. 2017 Oct 31;8:2104. Egamberdiyeva D. The effect of plant growth promoting bacteria on growth and nutrient uptake of maize in two different soils. Applied Soil Ecology. 2007 Jun 1;36(2- 3):184-9. Ehrenfeld JG, Ravit B, Elgersma K. Feedback in the plant-soil system. Annu. Rev. Environ. Resour.. 2005 Nov 21;30:75-115. Fahn A, Cutler DF. Xerophytes. Gebrüder Borntraeger; 1992. Fierer N, Bradford MA, Jackson RB. Toward an ecological classification of soil bacteria. Ecology. 2007 Jun;88(6):1354-64. Fierer N, Jackson RB. The diversity and biogeography of soil bacterial communities. Proceedings of the National Academy of Sciences. 2006 Jan 17;103(3):626-31. Fierer N, Leff JW, Adams BJ, Nielsen UN, Bates ST, Lauber CL, Owens S, Gilbert JA, Wall DH, Caporaso JG. Cross-biome metagenomic analyses of soil microbial communities and their functional attributes. Proceedings of the National Academy of Sciences. 2012 Dec 26;109(52):21390-5. Finzi AC, Van Breemen N, Canham CD. Canopy tree–soil interactions within temperate forests: species effects on soil carbon and nitrogen. Ecological applications. 1998 May;8(2):440-6. Friedman J, Hutchins A, Li CY, Perry DA. Actinomycetes inducing phytotoxic or fungistatic activity in a Douglas-fir forest and in an adjacent area of repeated regeneration failure in southwestern Oregon. Biologia Plantarum. 1989 Nov 1;31(6):487-95. Garcia-Moya E, McKell CM. Contribution of shrubs to the nitrogen economy of a desert‐ wash plant community. Ecology. 1970 Jan;51(1):81-8.

Shaaban D.H. 38 Master Thesis

Garner W, Steinberger Y. A proposed mechanism for the formation of ‘fertile islands’ in the desert ecosystem. Journal of arid Environments. 1989 May 1;16(3):257-62. Glick BR, Todorovic B, Czarny J, Cheng Z, Duan J, McConkey B. Promotion of plant growth by bacterial ACC deaminase. Critical Reviews in Plant Sciences. 2007 Oct 23;26(5-6):227-42. Gregorich EG, Carter MR, Doran JW, Pankhurst CE, Dwyer LM. Biological attributes of soil quality. InDevelopments in soil science 1997 Jan 1 (Vol. 25, pp. 81-113). Elsevier. Grishkan I, Nevo E. Spatiotemporal distribution of soil microfungi in the Makhtesh Ramon area, central Negev desert, Israel. fungal ecology. 2010 Nov 1;3(4):326- 37. Gutterman Y. Survival strategies of annual desert plants. Springer Science & Business Media; 2012 Dec 6. Hatzenpichler R, Lebedeva EV, Spieck E, Stoecker K, Richter A, Daims H, Wagner M. A moderately thermophilic ammonia-oxidizing crenarchaeote from a hot spring. Proceedings of the National Academy of Sciences. 2008 Feb 12;105(6):2134-9. Herman RP, Provencio KR, Herrera-Matos J, Torrez RJ. Resource islands predict the distribution of heterotrophic bacteria in Chihuahuan Desert soils. Appl. Environ. Microbiol.. 1995 May 1;61(5):1816-21. Hesp PA. Ecological processes and plant adaptations on coastal dunes. Journal of arid environments. 1991 Sep 1;21(2):165-91. Hillel D. Soil and water: physical principles and processes. Elsevier; 2012 Dec 2. Hoagland RE. Microbes and microbial products as herbicides: an overview. Jacoby R, Peukert M, Succurro A, Koprivova A, Kopriva S. The role of soil microorganisms in plant mineral nutrition—current knowledge and future directions. Frontiers in plant science. 2017 Sep 19;8:1617. Jenkinson, D. S. 1988. Determination of microbial biomass carbon and nitrogen in soils. In: Advances in Nitrogen Cycling in Agricultural Ecosystems. pp. 368–386. Wil- son, J. R., Ed., International, Walling-Ford.

Shaaban D.H. 39 Master Thesis

Jobbágy EG, Jackson RB. The distribution of soil nutrients with depth: global patterns and the imprint of plants. Biogeochemistry. 2001 Mar 1;53(1):51-77. Jones DL, Oburger E. Solubilization of phosphorus by soil microorganisms. InPhosphorus in action 2011 (pp. 169-198). Springer, Berlin, Heidelberg. Jorquera MA, Hernández MT, Rengel Z, Marschner P, de la Luz Mora M. Isolation of culturable phosphobacteria with both phytate-mineralization and phosphate- solubilization activity from the rhizosphere of plants grown in a volcanic soil. Biology and Fertility of soils. 2008 Sep 1;44(8):1025. Kaminsky R. The microbial origin of the allelopathic potential of Adenostoma fasciculatum H & A. Ecological Monographs. 1981 Feb;51(3):365-82. Katz ME, Ferguson J, Reed SI. Temperature-sensitive lethal pseudorevertants of ste mutations in Saccharomyces cerevisiae. Genetics. 1987 Apr 1;115(4):627-36. Kelly AE, Goulden ML. Rapid shifts in plant distribution with recent climate change. Proceedings of the National Academy of Sciences. 2008 Aug 19;105(33):11823- 6. Kennedy AC, Young FL, Elliott LF, Douglas CL. Rhizobacteria suppressive to the weed downy brome. Soil Science Society of America Journal. 1991;55(3):722-7. Kieft TL, White CS, Loftin SR, Aguilar R, Craig JA, Skaar DA. Temporal dynamics in soil carbon and nitrogen resources at a grassland–shrubland ecotone. Ecology. 1998 Mar;79(2):671-83. Klopatek JM, Stock WD. Partitioning of nutrients in Acanthosicyos horridus, a keystone endemic species in the Namib Desert. journal of Arid Environments. 1994 Mar 1;26(3):233-40. Köberl J, Prettenthaler F, Bird DN. Modelling climate change impacts on tourism demand: A comparative study from Sardinia (Italy) and Cap Bon (Tunisia). Science of the Total Environment. 2016 Feb 1;543:1039-53. Köberl M, Müller H, Ramadan EM, Berg G. Desert farming benefits from microbial potential in arid soils and promotes diversity and plant health. PLoS One. 2011 Sep 2;6(9):e24452.

Shaaban D.H. 40 Master Thesis

Köberl M, Schmidt R, Ramadan EM, Bauer R, Berg G. The microbiome of medicinal plants: diversity and importance for plant growth, quality and health. Frontiers in microbiology. 2013 Dec 20;4:400. Kögel-Knabner I. The macromolecular organic composition of plant and microbial residues as inputs to soil organic matter. Soil Biology and Biochemistry. 2002 Feb 1;34(2):139-62. Kuske CR, Ticknor LO, Miller ME, Dunbar JM, Davis JA, Barns SM, Belnap J. Comparison of soil bacterial communities in rhizospheres of three plant species and the interspaces in an arid grassland. Appl. Environ. Microbiol.. 2002 Apr 1;68(4):1854-63. Lacap-Bugler DC, Lee KK, Archer S, Gillman LN, Lau MC, Leuzinger S, Lee CK, Maki T, McKay CP, Perrott JK, de Los Rios-Murillo A. Global diversity of desert hypolithic cyanobacteria. Frontiers in microbiology. 2017 May 16;8:867. Laity JJ. Deserts and desert environments. John Wiley & Sons; 2009 Jan 28. Lauber CL, Hamady M, Knight R, Fierer N. Pyrosequencing-based assessment of soil pH as a predictor of soil bacterial community structure at the continental scale. Appl. Environ. Microbiol.. 2009 Aug 1;75(15):5111-20. Lehmann J, Schroth G. Nutrient leaching. Trees, Crops and Soil Fertility, CABI Publishing, Wallingford. 2003:151-66. Lester ED, Satomi M, Ponce A. Microflora of extreme arid Atacama Desert soils. Soil Biology and Biochemistry. 2007 Feb 1;39(2):704-8. Li J, Gilhooly III WP, Okin GS, Blackwell III J. Abiotic processes are insufficient for fertile island development: A 10‐year artificial shrub experiment in a desert grassland. Geophysical Research Letters. 2017 Mar 16;44(5):2245-53. Li Y, Kong Y, Teng D, Zhang X, He X, Zhang Y, Lv G. Rhizobacterial communities of five co- occurring desert halophytes. PeerJ. 2018 Aug 30;6:e5508. Li ZH, Wang Q, Ruan X, Pan CD, Jiang DA. Phenolics and plant allelopathy. Molecules. 2010;15(12):8933-52.

Shaaban D.H. 41 Master Thesis

Lovett GM, Weathers KC, Arthur MA, Schultz JC. Nitrogen cycling in a northern hardwood forest: do species matter?. Biogeochemistry. 2004 Feb 1;67(3):289- 308. Lugtenberg B, Kamilova F. Plant-growth-promoting rhizobacteria. Annual review of microbiology. 2009 Oct 13;63:541-56. Madigan MT, Martinko JM. Microorganisms and microbiology. Brock biology of microorganisms. 11th ed. Upper Saddle River, New Jersey (NJ): Pearson Prentice Hall. 2006:1-20. Makhalanyane TP, Valverde A, Gunnigle E, Frossard A, Ramond JB, Cowan DA. Microbial ecology of hot desert edaphic systems. FEMS microbiology reviews. 2015 Feb 25;39(2):203-21. Mapelli F, Marasco R, Balloi A, Rolli E, Cappitelli F, Daffonchio D, Borin S. Mineral– microbe interactions: biotechnological potential of bioweathering. Journal of Biotechnology. 2012 Feb 20;157(4):473-81. Marasco R, Rolli E, Ettoumi B, Vigani G, Mapelli F, Borin S, Abou-Hadid AF, El-Behairy UA, Sorlini C, Cherif A, Zocchi G. A drought resistance-promoting microbiome is selected by root system under desert farming. PloS one. 2012 Oct 31;7(10):e48479. Marasco R, Rolli E, Fusi M, Michoud G, Daffonchio D. Grapevine rootstocks shape underground bacterial microbiome and networking but not potential functionality. Microbiome. 2018 Dec;6(1):3. Mares MA. Encyclopedia of deserts. University of Oklahoma Press; 2017 Jan 19. Marques EL, Dias JC, Gross E, Silva AB, de Moura SR, Rezende RP. Purple Sulfur Bacteria Dominate Microbial Community in Brazilian Limestone Cave. Microorganisms. 2019 Feb;7(2):29. Marsh BA. The microenvironment associated with Welwitschia mirabilis in the Namib desert. Namib ecology. 1990;25:149-53.

Shaaban D.H. 42 Master Thesis

Mendes R, Garbeva P, Raaijmakers JM. The rhizosphere microbiome: significance of plant beneficial, plant pathogenic, and human pathogenic microorganisms. FEMS microbiology reviews. 2013 Sep 1;37(5):634-63. Molisch H. Der Einfluss einer Pflanze auf die andere, Allelopathie. Fischer Jena. 1937. Müller DB, Vogel C, Bai Y, Vorholt JA. The plant microbiota: systems-level insights and perspectives. Annual review of genetics. 2016 Nov 23;50:211-34. Muyzer G, De Waal EC, Uitterlinden AG. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction- amplified genes coding for 16S rRNA. Appl. Environ. Microbiol.. 1993 Mar 1;59(3):695-700. 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 Microbiology Ecology. 2005 Oct 1;54(2):233-45. Neilson JW, Quade J, Ortiz M, Nelson WM, Legatzki A, Tian F, LaComb M, Betancourt JL, Wing RA, Soderlund CA, Maier RM. Life at the hyperarid margin: novel bacterial diversity in arid soils of the Atacama Desert, Chile. Extremophiles. 2012 May 1;16(3):553-66. Nobel PS, McDaniel RG. Low temperature tolerances, nocturnal acid accumulation, and biomass increases for seven species of agave. Journal of Arid Environments. 1988 Nov 1;15(2):147-55. Noy-Meir I. Desert ecosystems: higher trophic levels. Annual Review of Ecology and systematics. 1974 Nov;5(1):195-214. Osborn AM, Moore ER, Timmis KN. An evaluation of terminal‐restriction fragment length polymorphism (T‐RFLP) analysis for the study of microbial community structure and dynamics. Environmental microbiology. 2000 Feb;2(1):39-50. Pandit AS, Joshi MN, Bhargava P, Ayachit GN, Shaikh IM, Saiyed ZM, Saxena AK, Bagatharia SB. Metagenomes from the saline desert of Kutch. Genome Announc.. 2014 Jun 26;2(3):e00439-14.

Shaaban D.H. 43 Master Thesis

Pedrol N, González L, Reigosa MJ. Allelopathy and abiotic stress. InAllelopathy 2006 (pp. 171-209). Springer, Dordrecht. Pettit RE. Organic matter, humus, humate, humic acid, fulvic acid and humin: their importance in soil fertility and plant health. CTI Research. 2004:1-7. Puente ME, Li CY, Bashan Y. Microbial populations and activities in the rhizoplane of rock-weathering desert plants. II. Growth promotion of cactus seedlings. Plant Biology. 2004 Sep;6(05):643-50. Raddadi N, Cherif A, Ouzari H, Marzorati M, Brusetti L, Boudabous A, Daffonchio D. Bacillus thuringiensis beyond insect biocontrol: plant growth promotion and biosafety of polyvalent strains. Annals of microbiology. 2007 Dec 1;57(4):481- 94. Rajeev L, Da Rocha UN, Klitgord N, Luning EG, Fortney J, Axen SD, Shih PM, Bouskill NJ, Bowen BP, Kerfeld CA, Garcia-Pichel F. Dynamic cyanobacterial response to hydration and dehydration in a desert biological soil crust. The ISME journal. 2013 Nov;7(11):2178. Rampelotto, Pabulo. "Extremophiles and extreme environments." (2013): 482-485. Rao S, Chan Y, Bugler-Lacap DC, Bhatnagar A, Bhatnagar M, Pointing SB. Microbial diversity in soil, sand dune and rock substrates of the Thar Monsoon Desert, India. Indian journal of microbiology. 2016 Mar 1;56(1):35-45. Rastogi G, Sani RK. Molecular techniques to assess microbial community structure, function, and dynamics in the environment. InMicrobes and microbial technology 2011 (pp. 29-57). Springer, New York, NY. Raynaud X, Nunan N. Spatial ecology of bacteria at the microscale in soil. PLoS One. 2014 Jan 28;9(1):e87217. Rice EL. Allelopathy. Academic press; 2012 Dec 2. Ridolfi L, Laio F, D’Odorico P. Fertility island formation and evolution in dryland ecosystems. Ecology and Society. 2008 Jun 1;13(1). Riesenfeld CS, Schloss PD, Handelsman J. Metagenomics: genomic analysis of microbial communities. Annu. Rev. Genet.. 2004 Dec 15;38:525-52.

Shaaban D.H. 44 Master Thesis

Ritz K, McHugh M, Harris J. Biological diversity and function in soils: contemporary perspectives and implications in relation to the formulation of effective indicators. InAgricultural Impacts on Soil Erosion and Soil Biodiversity: Developing Indicators for Policy Analysi s-OECD Expert Meeting Rome, Italy 2003 Mar (pp. 563-573). Robinson TW. Phreatophytes. US Government Printing Office; 1958. Rodriguez R, Redman R. More than 400 million years of evolution and some plants still can't make it on their own: plant stress tolerance via fungal symbiosis. Journal of experimental botany. 2008 Feb 10;59(5):1109-14. Rossi BE, Villagra PE. Effects of Prosopis flexuosa on soil properties and the spatial pattern of understorey species in arid Argentina. Journal of Vegetation Science. 2003 Aug;14(4):543-50. Rousk J, Bååth E. Fungal biomass production and turnover in soil estimated using the acetate-in-ergosterol technique. Soil Biology and Biochemistry. 2007 Aug 1;39(8):2173-7. Sandquist DR. Plants in deserts. Ecology and the Environment. 2014:297-326. Scacchi, A., Bortolo, R., Cassani, G. et al. J Plant Growth Regul (1992) 11: 39. https://doi.org/10.1007/BF00193842 Schade JD, Hobbie SE. Spatial and temporal variation in islands of fertility in the Sonoran Desert. Biogeochemistry. 2005 Apr 1;73(3):541-53. Schlesinger WH, Raikes JA, Hartley AE, Cross AF. On the Spatial Pattern of Soil Nutrients in Desert Ecosystems: Ecological Archives E077-002. Ecology. 1996 Mar;77(2):364-74. Schlesinger WH, Reynolds JF, Cunningham GL, Huenneke LF, Jarrell WM, Virginia RA, Whitford WG. Biological feedbacks in global desertification. Science. 1990 Mar 2;247(4946):1043-8. Schulze ED, Robichaux RH, Grace J, Rundel PW, Ehleringer JR. Plant water balance. BioScience. 1987 Jan 1;37(1):30-7.

Shaaban D.H. 45 Master Thesis

Schuster AC, Burghardt M, Alfarhan A, Bueno A, Hedrich R, Leide J, Thomas J, Riederer M. Effectiveness of cuticular transpiration barriers in a desert plant at controlling water loss at high temperatures. AoB Plants. 2016;8. Singh M, Kumar J, Singh S, Singh VP, Prasad SM. Roles of osmoprotectants in improving salinity and drought tolerance in plants: a review. Reviews in Environmental Science and Bio/Technology. 2015 Sep 1;14(3):407-26. Sinsabaugh RL. Phenol oxidase, peroxidase and organic matter dynamics of soil. Soil Biology and Biochemistry. 2010 Mar 1;42(3):391-404. Smith, S.D., Devitt, D.A., Sala, A. et al. Wetlands (1998) 18: 687. https://doi.org/10.1007/BF03161683 Spaepen S, Vanderleyden J, Okon Y. Plant growth-promoting actions of rhizobacteria. Advances in botanical research. 2009 Jan 1;51:283-320. Starkenburg, S.R., Reitenga, K.G., Freitas, T., Johnson, S., Chain, P.S., Garcia-Pichel, F. and Kuske, C.R., 2011. Genome of the cyanobacterium Microcoleus vaginatusFGP-2, a photosynthetic ecosystem engineer of arid land soil biocrusts worldwide. Sterflinger K, Tesei D, Zakharova K. Fungi in hot and cold deserts with particular reference to microcolonial fungi. Fungal ecology. 2012 Aug 1;5(4):453-62. Stomeo F, Valverde A, Pointing SB, McKay CP, Warren-Rhodes KA, Tuffin MI, Seely M, Cowan DA. Hypolithic and soil microbial community assembly along an aridity gradient in the Namib Desert. Extremophiles. 2013 Mar 1;17(2):329-37. Thomas DS. Sand seas and aeolian bedforms. Arid Zone Geomorphology: Process, Form and Change in Drylands. 1997:373-412. Thomas, A., Hoon, S. R., Mairs, H., Dougill, A. J. (2012). Soil Organic Carbon and Soil Respiration in Deserts: Examples from the Kalahari. In L. Mol, & T. Sternberg (Eds.), Changing Deserts: Integrating People and their Environment. (pp. 40-60). The White Horse Press.

Shaaban D.H. 46 Master Thesis

Thompson TL, Zaady E, Huancheng P, Wilson TB, Martens DA. Soil C and N pools in patchy shrublands of the Negev and Chihuahuan Deserts. Soil Biology and Biochemistry. 2006 Jul 1;38(7):1943-55. Thorup RM. Root development and phosphorus uptake by tomato plants under controlled soil moisture conditions. Agronomy Journal. 1969;61(5):808-11. Tocheva EI, Ortega DR, Jensen GJ. Sporulation, bacterial cell envelopes and the origin of life. Nature Reviews Microbiology. 2016 Aug;14(8):535. Treusch AH, Leininger S, Kletzin A, Schuster SC, Klenk HP, Schleper C. Novel genes for nitrite reductase and Amo‐related proteins indicate a role of uncultivated mesophilic crenarchaeota in nitrogen cycling. Environmental microbiology. 2005 Dec;7(12):1985-95. Tringe SG, Rubin EM. Metagenomics: DNA sequencing of environmental samples. Nature reviews genetics. 2005 Nov;6(11):805. Tsakiris G, Vangelis HJ. Establishing a drought index incorporating evapotranspiration. European water. 2005 Sep 10;9(10):3-11. Vessey JK. Plant growth promoting rhizobacteria as biofertilizers. Plant and soil. 2003 Aug 1;255(2):571-86. Vishwakarma K, Upadhyay N, Kumar N, Yadav G, Singh J, Mishra RK, Kumar V, Verma R, Upadhyay RG, Pandey M, Sharma S. Abscisic acid signaling and abiotic stress tolerance in plants: a review on current knowledge and future prospects. Frontiers in plant science. 2017 Feb 20;8:161. Vokou D, Margaris NS, Lynch JM. Effects of volatile oils from aromatic shrubs on soil microorganisms. Soil Biology and Biochemistry. 1984 Jan 1;16(5):509-13. Vollmer AT, Au F, Bamberg SA. Observations on the distribution of microorganisms in desert soil. The Great Basin Naturalist. 1977 Mar 31:81-6. Wallace A, Romney EM, Hunter RB. The challenge of a desert: revegetation of disturbed desert lands. Great Basin Naturalist Memoirs. 1980 Jan 1:216-25. Waller F, Achatz B, Baltruschat H, Fodor J, Becker K, Fischer M, Heier T, Hückelhoven R, Neumann C, Von Wettstein D, Franken P. The endophytic fungus Piriformospora

Shaaban D.H. 47 Master Thesis

indica reprograms barley to salt-stress tolerance, disease resistance, and higher yield. Proceedings of the National Academy of Sciences. 2005 Sep 20;102(38):13386-91. Wardle DA, Bardgett RD, Klironomos JN, Setälä H, Van Der Putten WH, Wall DH. Ecological linkages between aboveground and belowground biota. Science. 2004 Jun 11;304(5677):1629-33. Wardle DA, Nilsson MC. Microbe-plant competition, allelopathy and arctic plants. Oecologia. 1997 Jan 1;109(2):291-3. Whitford WG, Anderson J, Rice PM. Stemflow contribution to the ‘fertile island’effect in creosotebush, Larrea tridentata. Journal of Arid Environments. 1997 Mar 1;35(3):451-7. Whitford WG. Ecology of desert systems. Elsevier; 2002 Mar 25. Wickens GE. Ecophysiology of economic plants in arid and semi-arid lands. Springer Science & Business Media; 2013 Apr 17. WRI I. UNEP. 1992. Global Biodiversity Strategy. 1995. WWF. 2018. Deserts. Retrieved from https://www.worldwildlife.org/habitats/deserts. Xu X, Cao X, Zhao L. Comparison of rice husk-and dairy manure-derived biochars for simultaneously removing heavy metals from aqueous solutions: role of mineral components in biochars. Chemosphere. 2013 Aug 1;92(8):955-61. Yang D, Zhang S, Liu G, Yang X, Huang Z, Ye X. Facilitation by a spiny shrub on a rhizomatous clonal herbaceous in thicketization-grassland in Northern China: Increased soil resources or shelter from herbivores. Frontiers in plant science. 2017 May 16;8:809. Yang J, Kloepper JW, Ryu CM. Rhizosphere bacteria help plants tolerate abiotic stress. Trends in plant science. 2009 Jan 1;14(1):1-4. Yao Y, Shao MA, Jia Y, Li T. Distribution of soil nutrients under and outside tree and shrub canopies on a revegetated loessial slope. Canadian journal of soil science. 2017 Jun 29;97(4):637-49. Zavarzin GA, Kolotilova NN. Lectures on Natural History Microbiology.

Shaaban D.H. 48 Master Thesis

Shaaban D.H. 49 Master Thesis

Chapter 2: ‘Allelofertile’ soil islands self-conditioned by Welwitschia mirabilis in the Namib Desert

Abstract. In response to nutrients limitation and water scarcity, deserts plants have the capacity to reconditioning the surrounding environment to create favorable ecological- niches defined ‘fertility island’. To reduce the competition for nutrient and water, desert plant activates also exclusion mechanism (e.g. allelopathy) aimed to inhibit and reduce the establishment of other plants. These positive (fertilization) and negative (exclusion) processes driven by the plant significantly recondition the composition (i.e., nutrient and water content) of the desert soil, thus affecting the edaphic microbial communities associated. In this study, I aimed to assess the effect of Welwitschia mirabilis in the formation of ‘allelofertility island’, in which fertilization and exclusion mechanisms are combined to create a favorable microenvironment for the plant survival. Results showed how W. mirabilis significantly affected the composition and structure of microbial communities (bacteria, archaea and fungi). Microorganisms typical of arid environment (Actinobacteria, Alphaproteobacteria, Thaumarchaeota and Ascomycota) and able to carry important ecological service to the plant throughout fertilization process (i.e., decomposition of litter organic matter) were enriched in the soil of the allelofertility island. I explored if microorganisms/metabolites were involved in the exclusion process mediated by W. mirabilis in the allelofertile soil. Barley seeds incubated in the soil sampled around W. mirabilis plants showed a delayed germination, followed by a reduction biomass of adult plants. Sterilization and water-washing of this soil allowed a normal germination rate of the seeds, suggesting that either thermolabile water-soluble allelochemicals or antagonistic microorganisms can be involved in the allelopathic mechanisms activated by W. mirabilis. The overall results support the concept that desert plants, such as W. mirabilis, create ‘allelofertility island’ to modulate in their favor the diversity and functionality of the

Shaaban D.H. 50 Master Thesis surrounding edaphic microbial communities. This could be an important adaptative mechanisms in the evolutionary success of the ‘living fossil’ W. mirabilis.

Keywords. Welwitschia mirabilis, Allelofertile soil islands, Plant-soil-microbe interaction, Namib desert, Microbiome, Polyextreme conditions.

2.1. Introduction

Land heterogeneity and environmental severity make deserts one of the most hostile places for the establishment, development and distribution of plant communities (Anderson et al., 2004; Mitchell et al. 2007; Mou et al., 1995; Laity, 2009; Makhalanyane, 2015). Although, living in such environment may be challenging or in some cases even impossible, a wide number of plants have evolved morphological, physiological and behavioral adaptations to optimize water management and nutrient uptake (Michael, 1999; Wickens, 2013 Fahn and Cutler, 1992). Both biotic (i.e., deposition of litter and microbial activity) and abiotic processes (i.e., trapping of biological materials) are involved in the self-conditioning and redistribution of water and nutrients (i.e., organic carbon, nitrogen and phosphate) in the soil surrounding shrubs and trees inhabiting arid land (Coppinger et al., 1991; Schlesinger et al., 1996). The enrichment of nutrients, organic matter and water retention create the so called ‘fertility island’, an important adaptative strategy of desert plants to improve and meliorate their microenvironment (Rossi and Villagra, 2003; Schade and Hobbie, 2005). The process of soil reconditioning around the plant is facilitated by the activity of edaphic microorganisms (i.e., Streptomyces and Rhizobium sp; Bashan & de Bashan, 2010; Makhalanyane et al., 2015) that significantly contribute to the recycling of organic matter (Powlson, 2001) and nutrients solubilization (Alori, 2017). To protect the developed ecological niche, desert plants can activate competitive/exclusion mechanisms, including allelopathy. This phenomenon consists in the generation of detrimental effects of one plant on the germination, growth, survival,

Shaaban D.H. 51 Master Thesis and reproduction of another by releasing allelochemical metabolites. These allelopathic molecule can diffuse in the surrounding soil by leaching, root exudation, volatilization, residue decomposition, and other processes (Cipollini, 2012; Reigosa, 2006). The success of this strategy may be partly contributed by the interference and interactions of such allelochemicals with soil/plant associated microorganisms (Lou, 2016). The latest can affect bioavailability of allelochemicals by decomposing litter (e.g., Pseudomonas and Thielaviopsis, Cipollini, 2012), or by converting harmless plant-derived compounds to more toxic forms (e.g., Acrospermum viticola; Gagliardo and chilton, 1992), as well as by directly releasing allelopathic molecules (e.g., Streptomyces hygroscopicus; Barazani, 2001). Auto-fertilization and allelopathic interactions are important strategies in determining species distribution and abundance within plant communities in arid environments. One successful example is given by the iconic plant Welwitschia mirabilis (plant sub-division Gnetophytina) which can live and survive for thousands of years under the polyextreme conditions of the hyperarid Namib Desert (Cooper, 2000; Henschel and Seely, 2000; Chaw, 2000). The accumulation and incorporation of litter modify the soil surrounding W. mirabilis by accumulating nutrients including nitrogen (Abrams et al., 1997). Moreover, the shelter provided by the continuous growth of the evergreen broadleaves creates a microclimate that alleviate the thermal and moisture regime respect to the unvegetated barren soil (Schulze et al., 1980). Due to the constant selective pressure of desert (i.e., water and nutrient limitation; Aroca et al., 2012; Wallace, 1980), plants tend to protect their niche by competing/excluding the vegetation in their proximity through allelopathic strategies (Rice 2012). Possibly, leached phenolic compounds from the litter (e.g. the fruiting cones) diffuse in the soil, or root and leaves exudates and volatile compounds can weaken or inhibit the growth of native plants (Tedersoo, 2017). The development of the W. mirabilis environmental-niche, in which fertility is increased and allelophatic mechanisms implemented, guarantees to the plant a successful strategy to persist in the hyperarid Namib Desert (Rossi & Villagra, 2003; Nobel, 1988; Schade & Hobbie, 2005).

Shaaban D.H. 52 Master Thesis

The self-recondition of the surrounding soil further provides a favorable microenvironment for microorganisms, particularly of nitrogen-fixing and heterotrophic microorganisms (Kuske et al., 2002), which directly contribute to the biological fertility and nutrient cycling in the soil (Bolan, 2011). Even though the importance of microorganisms in fertility and allelofertility mechanisms has been postulated and recognized (Makhalanyane et al., 2015; Cipollini, 2012) as well as their ecological and protective services for desert plants (Jones, 2011; Jenkinson, 1998), no studies were conducted on the microbial ecology of the ‘allelofertility island’ that combine both the inhibiting and promoting effects. The assembly of the microbial communities in the soil re-conditioned by desert plans is driven by the modification of the soil composition and structure, as well as by the released/accumulated allelochemicals (Schlesinger et al., 1996; Rice, 2012; Cipollini, 2012). Here, I evaluate the self-conditioning effect (i.e., the creation of allelofertility islands) of W. mirabilis on the soil it colonizes and, on the assembly, and selection of the edaphic microbial community. I used the Namib Desert soil surrounding W. mirabilis plants, the litter produced by the plant and and the barren soil not influenced by the plant to measure in vivo - under both watered and controlled-drought conditions - the induced fertility and allelopathy effects. Additionally, by using high-throughput metagenomic sequencing of bacterial, archaeal and fungal communities I disentangled the effect of W. mirabilis on the diversity and functionality of the edaphic microorganisms associated with the ‘allelofertility island’.

2.2 Results and Discussion

2.2.1 W. mirabilis behaviours and ‘wash’ site ecosystem. W. mirabilis is a gymnosperm endemic of the arid and semiarid ecosystems of Namibia and Angola. The area colonized by this ‘living fossil’ is limited to 150,000 km2, from the Bentiaba River in southern Angola to the Kuiseb River in Namibia and up to 100 km inland of the coast (Figure 2.1A; Kers, 1967). The hyperarid central Namib Desert (Eckardt, 1999) is characterized by two

Shaaban D.H. 53 Master Thesis principal sources of water, fog and rainfall (Bornman, 1973; Henschel, 2019). Fog coming from the Atlantic Ocean and influencing up to 60 km2 inland (Eckardt, 2013; Frossard, 2015) provides an important source of water for those animals and plants able to collect and absorb it (Ebner, 2011; Henschel, 2008), while the sporadic rainfall events guarantee a limited water source to desert organisms (around 100 mm per year; Eckardt, 2013). W. mirabilis populations of Namib Desert tends to occur mainly along ephemeral watercourses and shallow plains (Jacobson and Lester, 1993; e.g., Kruiseb river, Figure 2.1A), indicating a possible dependence on ground water, in addition to precipitation and fog (Bornman, 1973; Van Jaarsveld and Pond, 2013). The idea that W. mirabilis can act as phreatophyte (absorbing water from ground water; Van Jaarsveld and Pond, 2013) is not based on evidence (Henschel et al., 2019), but only based on their distribution and similarity with other plants in the same environment (among others, Acanthosicyos horridus [!Nara melon] and Acacia greggii; Klopatek, 1994; Van Jaarsveld & Pond, 2013). In 2019, after the excavation of six W. mirabilis plants, Henschel and coauthors (2019) map the root system of the plants. The results showed that W. mirabilis has an elongated shallow root system penetrating up to 2 m deep in the soil but covering extended surface (up to 9 m) around the plant (Henschel et al., 2019). The roots create a complex network of both major and fine spongy roots to increment absorption of water from fog during the night (Bornman et al., 1972; Henschel et al., 2019; Henschel et al., 2019; Fan et al., 2017). W. mirabilis individuals studied in this work were located in the ‘W. mirabilis wash’ (Figure 2.1A and 2.1B). This site is a wide ravine characterized by a sandy channel in which the predominant plant species is W. mirabilis (Figure 2.1.B; Abrams et al., 1997). Plants had a diameter ranging from 95 to 190 cm (Table 2.1); according to their size, the plants are about 500-1000 years-old (Lukaszkiewicz, 2008). These plants have unique leaves: each plant has two leaves for its entire life, which grow outwards from the meristem like fingernail (Butler, 1973). As the plants age the leaves splits and give the appearance of multiple leaves (Figure 2.1C). Around the leaves, plants are surrounded

Shaaban D.H. 54 Master Thesis by a dark-soil zone (hereafter ‘allelofertility island’) created by the accumulation and incorporation of the litter in the soil (Abrams et al 1997, Figure 2.1C). The majority of litter appears to be due to shedding of the reproductive parts of the plants (the male reproductive cones; Supplementary Figure 2.1A and 2.1B) because the leaves are leathery and massive (Abrams et al 1997). The males’ plants produce relatively small cones (Supplementary Figure 2.1C) that presumably are rapidly decomposed by soil microorganisms (Abrams et al 1997; Kögel-Knabner, 2002). As previously observed by Abrams and collegues (1997) in the same site, soil surrounding W. mirabilis male plants has significantly higher levels of nutrients (Mg, Na, K, Fe, P, N), organic carbon and soil electrical conductivity compared to the soil between plants. The increment of these components around the plant underlines the capacity of W. mirabilis to self-fertilize its surrounding soil.

Figure 2.1. Distribution and habitat of W. mirabilis. (A) Distribution map of W. mirabilis along the arid and semiarid regions of Namibia and Angola (modified from Pekarek et al., 2016). Red circle indicated the ‘W. mirabilis wash’ site investigated in this study. (B) Photograph of the ‘W. mirabilis wash’ site inhabited by W. mirabilis plants. (C) Photographs of one of the W. mirabilis plants sampled; the black ‘allelofertility’ halo around the plant is visually distinguishable (darker soil) from the whitish barren soil.

The dark halo surrounding the selected plants extend over to 150-255 cm away from plant, covering a portion equal to the size of the plant that originated it (in average ratio

Shaaban D.H. 55 Master Thesis

1:1; Supplementary Table 2.1). As it possible observe in the Figure 2.1C, the intensity of the black color gradually decreases moving away from the plant. In the environmentally stressful habitat in which the W. mirabilis plants occurs, temperature and relative humidity of air and soil were very variable along the year, following both seasonality (winter vs summer) and day/night excursion (Figure 2.2). In the ‘W. mirabilis wash’, temperatures of superficial barren soil can be up to 50°C (Figure 2.2A). Below the plants this maximum decrease of few degree (~46°C), possibly due to the shelter provided by the plant leaves. In January 1982, despite higher temperatures were measured below the unvegetated soil surface (maximum, 60°C in one day measurement), a substantial reduction of temperatures were observed below the plant litter (maximum, 40°C; March, 1990).

Figure 2.2. Humidity and temperature range in the ‘W. mirabilis wash’. (A) Relative humidity (%) and (B) temperature experienced by air (blue), unvegetated barren soil (grey) and soil under W. mirabilis plants (brown) along the year (from April 2018 to May 2019).

In the case of relative humidity, plants reduce the variability observed in the bulk soil not influenced by the plant and maintain a more stable environment: for the 50% of the year soil under the plant had 40-60% of relative humidity, while these values were encountered by barren soil and air only for 28% of the year time (Figure 2.2B). Additionally, measures of the water holding capacity (WHC) conducted on our samples revealed that WHC of the litter (Z0) and the initial part of the allelofertility island (Z1) are significantly higher than those of barren unvegetated soil (Table 2.1). Soil WHC is a

Shaaban D.H. 56 Master Thesis very important characteristic for sustaining plant growth and it is primarily controlled by soil texture and organic matter content (Carter, 2002; Brady, 2008). A higher concentration of organic matter originated from the plant litter in the soil (Novara, 2015) increases the retention of water (Pettit, 2004). Such water retention significantly influences the wilting point of the soil (WP; Table 2.1) and consequently the water available for plant (WHC - WP; Brady, 2008; Kögel-Knabner, 2002). It is defined as the minimal amount of water in the soil that the plant requires not to wilt. Below this point a wilted plant can no longer recover its turgidity (for example in the in vivo experiment, see Figure 8). Notably, litter and the allelofertility island soil showed the higher WP values (Table 1), confirming the capacity of the W. mirabilis litter to positively increase the fertility of soil.

Table 2.1. Measurements of water holding capacity (WHC) and wilting point of soil (WP) along the W. mirabilis allelofertility gradient, from zone Z0 to zone Z5.

Category Zone Water Holding Capacity Wilting point Litter Z0 30.05 ± 14.85 (a) 15.88 ± 3.13 (a) Allelofertility start Z1 17.24 ± 6.57 (ab) 9.33 ± 4.71 (ab) Allelofertility middle Z2 12.87 ± 1.36 (b) 5.69 ± 0.84 (b) Allelofertility border Z3 10.31 ± 2.45 (bc) 4.25 ± 1.15 (bc) Barren Z4+Z5 6.61 ± 1.92 (c) 3.16 ± 0.19 (c)

These observations confirmed the results reported by March (1990), where significantly higher values of moisture were measured below W. mirabilis plants compared to nearby exposed areas. Modifications of both fertility and microclimate (thermal/moisture regimes) create favorable microhabitats for W. mirabilis. The microhabitats below W. mirabilis attract numerous animals and a variety of insects (Bornman, 1971), as well as enrich the presence and activity of microorganisms (Bonkowski et al., 2009; Muller et al., 2016).

2.2.2 Microbial communities’ structure and diversity across W. mirabilis allelofertility island gradient. After quality trimming, exclusion of non-target co-amplicons (i.e.,

Shaaban D.H. 57 Master Thesis chloroplast and mitochondria) and unclassified reads, a total of 10,642,750, 12,313,856 and 13,140,865 bacterial, archaeal and fungal reads, respectively, were generated (Table 2.2). The recovered sequences were clustered into 2,334 unique operational taxonomic units (OTUs; 97% nucleotide identity) for bacteria, 117 for archaea and 410 for fungi (Table 2.2).

Table 2.2. Quality metrics of high-throughput sequencing (MiSeq Illumina) analysis.

Kingdom Used Unassigned Non-target <0.001% N. OTU Bacteria 10,642,750 106,674 3,185 108,789 2,334 Archaea 12,313,856 733,101 51,220 6,915 117 Fungi 13,140,865 1,046,282 n.d. 20,140 410

Rarefaction curves indicating the OTU richness per sample generally approached saturation (Supplementary Figure 2.3). Litter microbial communities were much less diverse than those in the allelofertility island and the barren soils (shape of rarefaction curves in Supplementary Figure 2.3). The microbial diversity within each sample (alphadiversity) was analyzed based on the richness (number of OTUs) and evenness indices (Figure 2.3). The W. mirabilis allelofertility island gradient significantly affected the richness of bacterial (F5,113=180, p<0.0001), archaeal (F5,110=18.7, p<0.0001) and fungal (F5,114=3.6, p=0.005) communities. Notably, the distribution of richness changed in function of the kingdom analyzed (Figure 3A-C). Not considering the litter samples, the richness significantly increased starting from allelofertility island to barren soil in bacterial communities (F=230, p<0.0001, R2=0.7; Figure 3A), while in the archaeal community an opposite trend was observed (F=40, p<0.0001, R2=0.3; Figure 3B). No significant correlation between location (sampling zone) and richness was found for fungi (F=0.5, p=0.47; Figure 3C). For evenness estimate, a clear effect of W. mirabilis soil-reconditioning was found for archaeal (F5,110=4.6, p=0.0006; Figure 3D) and fungal (F5,114=13.2, p<0.0001; Figure 3E) communities, but not for bacteria (F5,113=1, p=0.4; Figure 3F). Except the lower archaeal

Shaaban D.H. 58 Master Thesis evenness of litter, highly comparable measures were recorded for the allelofertility island and the barren soil (F=2, p=0.16; Figure 3E). In contrast, barren soils fungal communities showed significantly higher eveness than the fertility island where it progressively decreased (F=51, p<0.0001, R2=0.34; Figure 3F).

Figure 2.3. Alpha diversity estimates of the microbial communities. (A-C) OTUs richness (number of observed OTUs) and (D-F) evenness were calculated for bacterial, archaeal and fungal communities. Box plots display the first (25%) and third (75%) quartiles, the median and the maximum and minimum observed values within each data set.

Through a process mediated by the decomposition and incorporation into the soil matrix of the plant litter (Abrams et al 1997; Kögel-Knabner, 2002), the allelofertility island selected its communities from the microbial-pools present in the surrounding soil (Makhalanyane et al., 2015). As showed by the Venn diagrams, the allelofertility island communities were also enriched with all the microorganisms carried by the litter (Figure 2.4A-C). In fact, no OTUs were detected as litter-specific and 31%, 46% and 80% of bacteria, archaea and fungi, respectively, were ubiquitously detected, i.e., associated with all the categories (litter, allelofertility and barren; Figure 2.4A-C). However, these values increased up to 86%, 72% and 95%, respectively, when only allelofertility island

Shaaban D.H. 59 Master Thesis and barren soils were considered (Figure 2.4A-C). Interestingly, generalists (represented by the spheres around the center of the triangles) and soil specialists (spheres along the allelofertility-barren edges of the triangles) were the main categories detected by ternary plot in all the communities (Figure 2.4D-F), with more abundant specialist-OTUs than generalist ones.

Figure 2.4. W. mirabilis allelofertility island recruitment process. Venn diagram detecting (A) bacterial, (B) archaeal and (C) fungal specialist (only present in one category) and generalist (shared among soil categories) OTUs. Ternary plot revealing relative abundance (dot size) and generalist or category-specific behaviors of (D) bacterial, (E) archaeal and (F) fungal OTUs among W. mirabilis allelofertility island’ gradient.

The observed selective process is mainly driven by incorporation and degradation of litter into the soil (Kögel-Knabner, 2002). The enrichment of organic matter in the soil is an important driver of edaphic microbial communities’ recruitment and assembly in arid and semi-arid environments (Bashan & de Bashan, 2010; Kuske et al., 2002). The complex moisture/chemistry gradient along the W. mirabilis allelofertility island transition determines specific micro-niches that ultimately leads to a gradual

Shaaban D.H. 60 Master Thesis differentiation between edaphic microbial communities associated with litter, allelofertility island and barren soil (ANOVA, bacteria: F5,110=27.7, p=0.001; archaea:

F5,113=54.6, p=0.001; fungi: F5,514=16.6, p=0.001). Principal coordinate analysis (PCoA) of OTUs’ composition revealed a ‘horseshoe shape’ ordination of the samples, starting from the litter (Z0), moving to the allelofertility island (Z1-Z3) and ending with the barren soils (Z4 and Z5). The ordinations explained up to 68.1, 70 and 51.1% of the total compositional (Bray-Curtis) variation in bacteria, archaea and fungi, respectively (Figure 2.5). Notably, cross-validation analysis of the defined zones (from Z0 to Z5) revealed relatively low percentages of misclassification for all the communities (5, 22. 4 and 17.5% in bacteria, archaea and fungi, respectively) with a consistent overlap among samples belonging to successive, zones while samples belonging to litter constitutes a clearly separate cluster (Supplementary Table 2.2).

Figure 2.5. Beta diversity of microbial communities associated with W. mirabilis allelofertility island. Principal coordinate analysis (PCoA) of (A) bacterial, (B) archaeal and (C) fungal communities based on Bray-Curtis similarity matrices.

2.2.3. Microbial communities’ taxonomic composition and predicted functions across the W. mirabilis allelofertility island gradient. Microbial communities associated with the allelofertility island created by W. mirabilis were comprised of the same dominant phyla/class with different overall contributions (Figure 2.6).

Shaaban D.H. 61 Master Thesis

Figure 2.6. Microbial community composition along the W. mirabilis allelofertility gradient. Average relative abundance of (A) bacterial, (B) archaeal and (C) fungal communities at different taxonomic level. Microbial communities’ composition at phylum, family and class levels, respectively.

Bacteria. Bacterial communities comprised a total of 19 bacterial pyla, 45 classes, 79 orders (95% classified), 104 families and 176 genera. In particular, samples were dominated by Proteobacteria (Alphaproteobacteria, 35%), followed by Actinobacteria (22%), Bacteroidetes (19%), Firmicutes (13%) and Chloroflexi (5%). Along the fertility gradient (from litter to barren soil), Firmicutes and Bacteroidetes showed a significant drop passing from 63% to 13% of the sequences’ relative abundance (Figure 2.6A). Opposite trend was detected for Actinobacteria and Chloroflexi that together passed from 4% in the litter and 17-21% in the allelofertility island, up to 37-40% in the barren unvegetated soils (Figure 2.6A). It was expected to observe that Alphaproteobacteria, Actinobacteria, Bacteroidetes, Firmicutes and Chloroflexi dominated the gravel plain of Namib Desert communities (Ronca, 2015). These ubiquitous phyla have been detected in arid and semi-arid soil worldwide scale (Delgado-Baquerizo et al., 2018; Makhalanyane et al., 2013; Ronca et al., 2015; Rao et al., 2016; Schulze-Makuch et al., 2018). Furthermore, members from these phyla are well-known for their diverse genetic

Shaaban D.H. 62 Master Thesis

(e.g. multi-stress related genes; Undabarrena et al., 2017) and physiological (e.g., Chloroflexi’s protective layered cell envelope structure, Overmann, 2008; Lacap et al., 2011; sporulation of Actinobacteria; Kroos and Maddock, 2003; Gao and Garcia-Pichel, 2011) resistance mechanisms to the arid and oligotrophic desert conditions

Table 2.3. Functional prediction of the bacterial communities along the W. mirabilis allelofertility gradient. Values indicate percentage of metabolic pathway categories (relative abundance) possibly involved in biogeochemical cycling of carbon and nitrogen in each zone of the gradient (from Z0 to Z5).

Location along allelofertility island gradient Predicted function Z0 Z1 Z2 Z3 Z4 Z5 Chemoheterotrophy 47.28 38.29 38.75 38.12 37.56 36.48 Aerobic chemoheterotrophy 47.13 37.75 38.09 37.27 36.35 35.63 Manganese oxidation 0.34 2.78 6.15 8.26 7.69 8.07 Nitrate reduction 0.32 1.48 1.09 0.97 3.85 4.19 Fermentation 0.43 1.30 2.01 1.89 2.09 1.58 Aromatic compound degradation 0.21 0.76 1.08 1.93 1.70 1.56 Methylotrophy 0.32 1.49 1.08 0.60 0.02 0.06 Methanol oxidation 0.32 1.46 1.06 0.58 0.02 0.06 Nitrate denitrification 0.32 1.46 1.06 0.58 0.02 0.06 Nitrite denitrification 0.32 1.46 1.06 0.58 0.02 0.06 Nitrous oxide denitrification 0.32 1.46 1.06 0.58 0.02 0.06 Denitrification 0.32 1.46 1.06 0.58 0.02 0.06 Dark hydrogen oxidation 0.32 1.46 1.06 0.58 0.02 0.06 Nitrite respiration 0.32 1.46 1.06 0.58 0.02 0.06 Nitrate respiration 0.32 1.46 1.06 0.58 0.02 0.06 Nitrogen respiration 0.32 1.46 1.06 0.58 0.02 0.06 Cellulolysis 0.05 0.14 0.24 0.96 1.88 1.42 Animal parasites or symbionts 0.34 0.50 0.52 1.01 0.56 0.57 Ureolysis 0.34 0.50 0.52 1.01 0.56 0.57

The 16S rRNA gene data analysis using Faprotax package (Aßhauer et al., 2015) showed that the majority of bacterial community’ members were involved in chemoheterotrophy and aerobic chemoheterotrophy pathways, showing a consistent distribution along the allelofertility gradient (36-38%; Table 2.3). Contrary, manganese oxidation and nitrate reduction showed increasing trends from litter to barren soil (Z0- Z5), as well as fermentation metabolisms, cellulolysis and aromatic compound degradation (Table 2.3), while methylotrophy and nitrogen cycle metabolic pathways were more in the allelofertility island soils (Z1 and Z2 zones; Table 2.3).

Shaaban D.H. 63 Master Thesis

Archaea. In archaeal communities two phyla have been detected (Euryarchaeota and Thaumarchaeota), distributed in 3 classes, 3 orders, 6 families and 12 genera. Differences in archaeal community composition between ecosystem types could be explained by the combined effects of precipitation gradient and vegetation cover (Angel et al., 2010). In the selected site, the Nitrososphaeraceae family dominated the soils (barren and allelofertility island) and the W. mirabilis litter (81% of relative abundance; Figure 2.6B). Only in the soil strongly reconditioned by the W. mirabilis (Z1 and Z2), Haloferacaceae, Haloadaptaceae and Halomicrobiaceae classes, along with unclassified Halobacteriales were enriched (up to 15% of relative abundance; Figure 2.6B). Generally, species of halophylic archaea (e.g., Halobacteriaceae, family of Halobacteriales), contains unique ether lipids that cannot be easily degraded, and are temperature resistant and highly salt-tolerant (Ruppel, 2013). They can exhibit phosphorus-solubilizing activities that contribute to phosphorus cycling in hypersaline environments (Yadav, 2015). They can also play an important role in nitrogen cycle under extreme conditions (e.g., extreme temperatures and pH values, salinity and water stress (Stres, 2007). Despite archaeal taxa are relatively rare across many biomes, they represent an important component of desert soils’ microbiome (Fierer et al., 2012), with Thaumarchaeota being the principal representatives. All known organisms of this phylum are chemolithoautotrophic ammonia-oxidizers and may play important roles in biogeochemical cycling in oligotrophic environments such as deserts (Brochier-Armanet et al., 2008). These observations were confirmed also by the functional prediction for archaeal community in our studied sites of the Namib Desert. A total of 22 of 117 OTUs (19%) were assigned to at least one functional group. The majority of archaeal community members were involved in the aerobic ammonia oxidation (98-99%), with a reduction of this activity in the allelofertility island Z1 and Z2 zones (Table 2.4). Notably, nitrate and nitrogen species respirations, also in limited percentage, were mainly predicted in the allelofertility island (Z1-Z3) rather than in the barren soil (Z4 and Z5) or

Shaaban D.H. 64 Master Thesis litter (Z0). Predicted aerobic chemoheterotrophic pathway were decreasing and randomly distributed along the allelofertility island (Table 2.4).

Table 2.4. Functional predictions of the archaeal communities along the W. mirabilis allelofertility gradient. Values indicate percentage of metabolic pathway categories (relative abundance) possibly involved in biogeochemical cycling of carbon and nitrogen in each zone of the gradient (from Z0 to Z5); n.d., not detected.

Location along allelofertility island gradient Predicted function Z0 Z1 Z2 Z3 Z4 Z5 Aerobic ammonia oxidation 98.20 28.59 20.29 95.16 99.94 99.92 Nitrification 98.20 28.59 20.29 95.16 99.94 99.92 Nitrate respiration 0.01 0.77 0.12 0.01 0.001 0.0003 Nitrate reduction 0.01 0.77 0.12 0.01 0.001 0.0003 Nitrogen respiration 0.01 0.77 0.12 0.01 0.001 0.0003 Aerobic chemoheterotrophy n.d. 0.01 n.d. 0.001 0.0003 n.d. Chemoheterotrophy n.d. 0.01 n.d. 0.001 0.0003 n.d.

Fungi. Fungal communities were composed by a total of four phyla (Ascomycota, Basidiomycota, Chytridiomycota and Zygomycota), along with 19% of unclassified SVs. Fungal SVs were distributed in 12 classes, 24 orders, 41 families and 75 genera. The classes Dothideomycetes, Sordariomycetes and Eurotiomycetes (all Ascomycota) are stress-resistant organisms adapted to desert environments (Sterflinger et al., 2012) and are equally widespread in the studied ecosystem (Figure 2.6C). The predicted feeding strategies (trophic categories, Nguyen et al., 2016; Table 2.5) showed that 62% of the fungal OTUs (49% of reads) were assigned to specific trophic categories: 27% to pathotroph-saprotroph-symbiotroph, 16% to pathotroph-saprotroph, 3.5% to saprotrophs and 2% to saprotroph-symbiotroph (Table 2.5). Notably the ywo most important categories were present in litter and allelofertility soils than in the barren one (Table 2.5). In fact, fungi deliver important ecological services to desert ecosystem, for instance decomposition of organic matter into small molecules (e.g., organic acids) easily recycled in C and N sources (Carlie, 2001). Fungi can also act as mutualists involved in the solubilization of phosphorous for plant uptake (Ritz, 2005).

Shaaban D.H. 65 Master Thesis

Moreover, fungal hyphae physically bind soil particles all together, creating stable aggregates that can increase water-holding capacity (Carlie, 2001).

Table 2.5. Functional predictions of the trophic categories of fungi along the W. mirabilis allelofertility gradient. Values indicate percentage of members (relative abundane) in each zone of the gradient (from Z0 to Z5).

Location along allelofertility island gradient Predicted Z0 Z1 Z2 Z3 Z4 Z5 Pathogen Saprotroph Symbiotroph 0.02 0.0003 0.0002 Nd 0.00 nd Pathotroph 0.05 0.13 0.22 0.63 1.62 1.71 Pathotroph Saprotroph 28.11 14.78 18.75 18.27 6.85 4.04 Pathotroph Saprotroph Symbiotroph 34.71 37.90 23.45 22.86 17.47 19.39 Pathotroph Symbiotroph 0.73 6.95 0.39 0.76 2.25 1.23 Saprotroph 0.44 2.79 3.19 3.68 4.26 7.20 Saprotroph Symbiotroph 0.01 0.003 0.0001 0.0001 0.0002 0.01 Symbiotroph 0.01 0.01 0.003 0.004 0.06 0.02 Not detected 35.92 37.44 53.99 53.80 67.49 66.41

2.2.4. Effects of soils from ‘allelofertility island’ on germination and development of phytometer. Barley was used as model phytometer to measure the physiological response of seeds and plants to the soil from the allelofertility island and to the barren soil and W. mirabilis litter. The germination process was strongly delayed by the allelofertility island soil: at 5 days, when the germination rate on litter and barren soil reached 75% and 60%, respectively, less than 10% of the seeds was germinated in the allelofertility island soil (Figure 2.7). At the end of the 10 days, the percentage of seed germination in allelofertility soil was comparable with the one on the other substrates (Figure 7A). The delay in the germination process due to allelopathic effect of soil reconditioned by W. mirabilis can reduce the probability of surrounding plants to establish under field circumstances. In fact, prolonged exposure of seeds to natural enemies, such as soil fungi and vertebrates and invertebrates’ predators, may increase the chances of mortality (Pereira et al., 2013b). To assess which soil components was responsible for the allelophaty observed (i.e., delay in the germination of allochthonous seeds), soils were tindalized to inactivate the

Shaaban D.H. 66 Master Thesis biotic part including microorganisms and thermal-sensitive biomolecules (Mahmood, 2014). After the tyndallization, seeds planted in the soil from the allelofertility island showed a normal germination rate, similar to the one of barren soil (97±4%), indicating that the allelopathic molecules/microorganisms were not anymore active. Moreover, in this case, the biomass of plants grown in barren soil was lower than the ones in allelofertility island’ soil, presumably due to the low nutrient content of the barren soil that is insufficient to sustain a regular growth of plant (Figure 2.8A). In fact, when the allelopathic effect was reduced/suppressed, the fertilization effect of the litter mixed with the soil around W. mirabilis allowed a better plant development.

Figure 2.7. Effect of W. mirabilis soil on seed germination and development. (A) Germination rate of barley seeds (n. of germinated seeds per day) planted in different soils. (B and C) Representative picture of germinated seeds at 4 and 7 days, respectively. From the left to the right; Agricultural soil, litter mix soil, allelofertility soil, barren soil.

Similar results were obtained using water-washed soils, in which water-soluble components and microbial cells were removed. Plants grown in the water-washed allelofertily island soil showed germination rate and plant development similar to those in barren soil, indicating that water-soluble molecules and/or microorganisms responsible of the allelopathic effect measured in stock soils were no longer present or were significantly reduced in the soil below a toxicity threshold (Figure 2.8B).

Shaaban D.H. 67 Master Thesis

Figure 2.8. Plant biomass under normal and drought conditions. (A) Plant biomass expressed as average ± standard deviation of 30 plants. (B and C) Picture of plant growth for 21 days in pot with different soils under (B) normal and (C) drought stress conditions. Pots follow the order of soil treatments reported in panel A (Agriculture, litter, allelofertility and barren soils). (D and E) Shoot and root of plants after 21 days of incubation in different soil. Order of treatments: agriculture, litter, allelofertility and barren soils.

Assays of 21 days duration were performed to assess the allelofertility effect of soils under drought conditions compared with normal irrigation. Plants recovered from the

Shaaban D.H. 68 Master Thesis allelofertility soil had a significantly lower biomass compared to those growing in the other substrates (Figure 2.8A and B). In particular, plants showed a smaller and less developed root system (Figure 2.8D), confirming that the delay in seed germination significantly affect the further development of plants. On the contrary, under controlled water stress condition (i.e., interruption of irrigation), plants grown in barren soil were severely affected after 5 days without watering, whereas the plants in litter and allelofertility soils showed turgid tissues and better development (Figure 2.8A, C and E). Compared to barren soil, litter and allelofertility soils significantly enhanced the plant biomass, respectively showing overall biomass accumulations of about 3-fold higher than in the barren soil and similar to the one of a commercial rich agriculture soil (Figure 2.8A). These results support the capacity of the litter-artificial soil mix and the allelofertility island soil to retain more water (improved WHC, Table 2.1) and consequently to have more water available for plant growth (WHC - WP). The capacity to provide an ongoing supply of water during periods of replenishment guarantees growth and survival for longer periods of drought (Hudson, 1994). Through the overall measurements of barley growth and development in W. mirabilis reconditioned soil, we confirmed the allelofertility mechanisms of W. mirabilis that I have hypothesized in the ‘Wash site’ (Figure 2.1C). In fact, soil reconditioned by W. mirabilis demonstrated a combination of (i) antagonism activity against plant competitors (e.g., new germinating seeds) but at the same time (ii) stimulation of plant growth especially under water stress, both of them possibly with the contribution of the soil microbiome. Such ecological and functional combination may contribute to explain the evolutionary success of the ‘living fossil’ W. mirabilis in the extreme conditions of the Namib Desert.

2.3. Conclusion

Here, I demonstrated that W. mirabilis apply a consistent radially-distributed selective pressure, that I defined as an “allelofertility-island effect”, under contrasting physico-

Shaaban D.H. 69 Master Thesis chemical conditions such as those of the gravel plain of the Namib Desert. Through soil self-reconditioning, mediated by the plant litter and possibly the root exudates, W. mirabilis modifies soil chemistry (i.e., organic carbon and WHC; March, 1990; Abrams, 1997) and increases habitat heterogeneity favoring fertility and allelophaty process. Such soil self-reconditioning process activated by W. mirabilis affected the diversity, interactions and function of the edaphic microbial communities (bacteria, archaea and fungi). The selective pressure of the allelofertility island consistently determined the recruitment and assembly of different microbial communities across the allelofertility gradient, that can boost and modulate ecological services of pivotal importance for W. mirabilis survival. The overall results elucidate the dynamic and interactions among plant, soil and microbes in the development of adaptive strategies involved in the enhancement of desert plant establishment and resistance to stresses.

2.4. Material and Methods

2.4.1. Site description and sampling. The study site ‘Welwitschia Wash’ was located in the gravel plains of the Namib Desert, approximately 14 km east of Gobabeb (23°36’44.01”S 15°10’09.01”E; Figure 1A). The area includes a 3-20 m wide ravine with a sandy channel, bordered by stony ledges where the predominant plant species was W. mirabilis (Herre, 1961; Jacobson et al., 1993; Figure 1B). Temperature (°C) and relative humidity (RH%) of air, barren soil and soil under plant were measured along one year (from April 2018 to May 2019) using HOBO sensors. In April 2018, five different male individuals of W. mirabilis growing in the wash were randomly selected. Plant had a diameter ranging from 95 to 190 cm (Supplementary Table 1). Each plant was surrounded by a darker-soil zone (hereafter, ‘Allelofertile island’) created by the accumulation/incorporation of the shed reproductive parts of the plants (i.e., male cone) in the soil (Abrams et al 1997, Figure 1C). A concentric-radial sampling design was adopted to collect soil samples along the allelofertility island gradient (from dark to white soil, Figure 1C and schematic representation in

Shaaban D.H. 70 Master Thesis

Supplementary Figure 2). Soil was sampled at five distances away from the W. mirabilis stock (Supplementary Figure 2 and Supplementary Table 1), passing from the dark-soil close to the plant (zone Z1, starting point of allelofertile island), to the brownish ones in the middle and border of allelofertile island (zones Z2 and Z3, respectively), and to the whitish barren soils (zones Z4 and Z5, plant-interspace soil). From each radial-zone, four replicates were collected for each plant, for a total of 100 samples of soils (60 fertility island, 40 barren). Samples from the central wooden body of the plant composed by litter (dry shedding male cone; Supplementary Figure 1) were also collected (Z0, n=20). Samples were stored at 4°C for soil chemistry and microcosm experiments and at -20°C for molecular analyses. All the samples were collected under the research/collecting permit number ES32861, which was delivered by the Namibian Ministry of Environment and Tourism.

2.4.2. Water holding capacity and wilting point measurements. Collected soils were saturated with water and incubated for 24 h under a pressure of 1/3 of bar. The soils were weighed and dried at 105°C for 24 h. The water hold by soils (WHC, water holding capacity) was calculated and expressed as percentage. To calculate wilting point (WP), soils were saturated with water, incubated for 48 h under a pressure of 15 bars and finally dried at 105°C for 24 h. Difference of weight between the dried soil and the wet soil express the WP of soil.

2.4.3. Total DNA extraction from soil. Total DNA was extracted from a 0.5 ± 0.1 g sub- sample of soils using the PowerSoil® DNA Isolation Kit (MoBio Inc., USA) following the manufacturer’s instructions. In the case of the litter (i.e., mix of dry flower and sand) collected form the center of the plant, samples were grinded in liquid nitrogen with sterile mortar and pestle before extracting the DNA. 5 l of the extracted DNAs were visualized by electrophoresis in 0.8% agarose gels add with SYBR Safe at the Molecular

Imager Gel DocTM XR+ Imaging System (Biorad). DNA was quantified using Qubit 3.0 fluorometer dsDNA BR Assay Kit (Thermo Fisher Scientific Waltham,MA, USA), following

Shaaban D.H. 71 Master Thesis the manufacturer’s instructions. Extracted DNA was stored at -20°C until further processing.

2.4.4. Metaphylogenomic analysis of edaphic microbial (bacteria, archaea and fung) communities. To identify bacterial and archaeal communities’ composition the V3-V4 hypervariable region of the 16S rRNA gene was targeted using the primer’ pairs 341F- 785R and A2F-534R respectively (Baker et al. 2003; Mapelli et al. 2018). To identify fungal community composition the region 2 of the internal transcribed spacer region (ITS2) was amplified using the primer pair ITS3F-ITS4F (Tedersoo et al. 2015). The reaction mixtures contained buffer 1 X, 1.5 mM MgCl2, 0.3 mM of dNTPs, 0.3 mM of each primer, 1 U HiFi Platinum Taq (Invitrogen), and 3-5 µl of DNA. If necessary, DNA was properly diluted. Cycling conditions used to amplify the bacterial and archaeal 16S rRNA and fungal ITS2 gene fragments were reported in Table 2.6.

Table 2.6. PCR primers and protocol for thermal cycling.

Target kingdom Primer Thermal cycling 94°C for 1 min Bacterial 16S rRNA I785R 94 °C for 30 sec, 55°C for 30 sec, 68°C for 30 sec (X 25) Mapelli et al., 2018 I341F 68°C for 5 mins 95°C for 10 min Archaeal 16S rRNA I534R 95 °C for 30 sec, 60°C for 15 sec, 68°C for 50 sec (X 25) Baker et al., 2003 I-A2F 68°C for 5 mins 96°C for 5min Fungal ITS2 ITS4RI 95°C for 1 min, 64°C for 1 min, 72°C for 45sec (X 30) Tedersoo et al., 2015 ITS3FI 72°C for 5 mins

The 16S rRNA and ITS2 metagenomic sequencing libraries were constructed using the 96 Nextera XT Index Kit (Illumina) following the manufacturer’s instructions. Libraries sequencing was done using the Illumina MiSeq platform with pair-end sequencing at the Bioscience Core Lab, King Abdullah University of Science and Technology. Using fastq- join algorithm (https://expressionanalysis.github.io/ea-utils/). Raw reads of forward and reverse were assembled for each sample into paired-end reads with a minimum overlap of 50 nucleotides and maximum of one mismatch within the region. The obtained pair-

Shaaban D.H. 72 Master Thesis end reads were analyzed using a combination of the UPARSE v8 (Edgar et al. 2013) and the QIIME v1.8 (Caporaso et al. 2010) software to form the Operational Taxonomic Units (OTUs) of 97% sequence similarity. Taxonomy was assigned to the representative sequences of the OTUs searching against the latest version of the Greengenes (DeSantis et al. 2006) and SILVA 132 databases for bacteria and archaea and the UNITE database for fungi (DeSantis et al., 2006; Koljalg et al., 2014). Only samples presenting a suitable sequencing depth (rarefaction curve in Supplementary Figure 2.3) and good’s coverage values >98% were used for the further analyses.

2.4.5. Beta- and alpha-diversity of microbial communities. Shared and exclusive OTUs (and their relative distribution) across among the different zones were calculated as described in Marasco et al 2018, using Venn-diagram and ternary plots using R package (ggplot and ggtern). Compositional (Bray-Curtis of the log-transformed OTUs table) similarity matrices were calculated and Principal Coordinates Analysis (PCoA) were performed in PRIMER v. 6.1 (Ramette, 2007; Clarke and Gorley, 2015). Permutational multivariate analyses of variance (PERMANOVA, main and multiple comparison tests) were also performed on the compositional distance matrices using the same software (Anderson, 2008). The considered explanatory variables were ‘Plant replicate’ (5 levels: from A to E) and ‘Allelofertility island gradient’ (6 levels: from Z0 to Z5). Alpha diversity indices (richness and evenness) were calculated using the PAST software.

2.4.6. In vivo experiment to evaluate the effect of Welwitschia-allelofertility island soils on seed germination and plant development under watered and controlled- drought conditions. Seed germination rate (n. germinated seed/day) and plant development (biomass) of barley were used as phyto-indicators to evaluate the effect of W. mirabilis soil-reconditioning (i.e., allelofertility island). The soil used were: i) the white barren soil not influenced by W. mirabilis plant, ii) the dark-soil close to W. mirabilis stock (affected by both litter and root exudates; Henschel et al. 2019; Valverde,

Shaaban D.H. 73 Master Thesis

2016), and iii) the artificial ‘dark’ soil created in laboratory mixing in equal part the barren sand (treatment i) and the litter collected from the center of the plant (treatment ii) to eliminate the contribute of plant exudates (Supplementary Figure 2.4A). A positive control composed by one part of agricultural soil and one parts of sand was also added. Plastic pots (10 cm diameter) were filled with the soils of the four different treatments; three pots for each treatment were prepared. The pots were watered with 50 ml of milliq water and 10 barley seeds added for each one. The pots were incubated in a growth chamber at day and night temperatures of 25±1°C, with approximately 100 μmol photons m−2s−1 of light supplied for 12 h during the daytime and 50% of relative humidity and were properly irrigated for all the duration of the experiment (21 days; Supplementary Figure 2.4A). Contemporary, a drought stress experiment was conducted; in this case, after 3 days of watering, irrigation was alternated by two drought periods of 5 days. (Supplementary Figure 2.4B) Seed germination and plant development tests (10 days incubation) were also performed with sterilized soils (Z1 vs Z5). Soils were sterilized using the tyndallization process (Bashan et al., 2014) to evaluate if the observed effects were mediated by living microorganisms or thermos-sensible bio-molecules present in the soil. Soil extracts were also tested; 200 g of each soil were mixed with 400 ml of distilled sterile water and shacked for 36 h (100 rpm). The solutions obtained were filtered to remove soil particles and used to water seeds (1:2 vol:vol) in sterile agriculture perlite. All the treatment was incubated following the condition previously reported. For all the assay the germination of seeds was evaluated every day (i.e., number of shoots coming out from the soil) and at the end of incubation time plants were harvested for biomass measurements (i.e., fresh and dry weight). Statistical analysis (one-way ANOVA) was used to compare treatments using the Kruskal-Wallis and Dunn’s multiple comparison tests in Graph-pad software.

Shaaban D.H. 74 Master Thesis

BIBLIOGRAPHY

Abrams MM, Jacobson PJ, Jacobson KM, Seely MK. Survey of soil chemical properties across a landscape in the Namib Desert. Journal of Arid Environments. 1997 Jan 1;35(1):29-38. Abrams, M., Jacobson, P., Jacobson, K., Seely, M. Survey of soil chemical properties across a landscape in the Namib Desert. Journal of Arid Environments (1997) 35: 29– 38 Alori ET, Glick BR, Babalola OO. Microbial phosphorus solubilization and its potential for use in sustainable agriculture. Frontiers in microbiology. 2017 Jun 2;8:971. Anderson, T.M., McNaughton, S.J., Ritchie, M.E., 2004. Scale-dependent relationships between the spatial distribution of a limiting resource and plant species diversity in an African grassland ecosystem. Oecologia 139, 277–287. Aroca R, Porcel R, Ruiz-Lozano JM. Regulation of root water uptake under abiotic stress conditions. Journal of experimental botany. 2011 Sep 13;63(1):43-57. Aßhauer KP, Wemheuer B, Daniel R, Meinicke P. Tax4Fun: predicting functional profiles from metagenomic 16S rRNA data. Bioinformatics. 2015 May 7;31(17):2882-4. Baker, G., Smith, J., Cowan, D. Review and re-analysis of domain-specific 16S primers. Jornal of microbiological methods. 55, 541-555 (2003) Barazani O, Friedman J. Allelopathic bacteria and their impact on higher plants. Critical reviews in microbiology. 2001 Jan 1;27(1):41-55. Barazani O. & Friedman J. 1999. Allelopathic bacteria and their impact on higher plants. Crit. Rev. Microbiol. 27: 741–755. Barto EK, Hilker M, Müller F, Mohney BK, Weidenhamer JD, Rillig MC. The fungal fast lane: common mycorrhizal networks extend bioactive zones of allelochemicals in soils. PLoS One. 2011 Nov 14;6(11):e27195. Bashan, Y., & de Bashan, L. E. (2010). Microbial populations of arid lands and their potential for restoration of deserts. In P. Dion (Ed.), Soil biology and agriculture in

Shaaban D.H. 75 Master Thesis

the tropics ()Soil Biology Series 21. Chapter 6 (pp. 109– 137). Berlin, Heidelberg, Germany: Springer. Bolan NS, Adriano DC, Kunhikrishnan A, James T, McDowell R, Senesi N. Dissolved organic matter: biogeochemistry, dynamics, and environmental significance in soils. InAdvances in agronomy 2011 Jan 1 (Vol. 110, pp. 1-75). Academic Press. Bonkowski M., Villenave C., Griffiths B. (2009). Rhizosphere fauna: the functional and structural diversity of intimate interactions of soil fauna with plant roots. Plant Soil 321 213–233. 10.1007/s11104-009-0013-2 Bornman CH, Elsworthy JA, Butler V, Botha CEJ. Welwitschia mirabilis: Observation on general habit, seed, seedling, and leaf characteristics. Madoqua 1972; (II)1: 53-6. Bornman LJ, Chris H, Botha CE. Welwitschia mirabilis: observations on movement of water and assimilates under fohn and fog conditions. Madoqua. 1973 Jan 1;1973(Series 2 Issue 2):25-31. Bornman, C. H. 1972a. Welwitschia mirabilis: paradox of the Namib Desert. Endeavour (English edition) 31: 95–99. Bornman, C. H. 1972b. Welwitschia mirabilis: das widersprüchlichste Gewächs der Namib. Endeavour (German edition) 31: 95–99. Bornman, C. H. 1977. Welwitschia mirabilis: structural and functional anomalies. Madoqua (II) 10: 21–31. Brady NC, Weil RR, Weil RR. The nature and properties of soils. Upper Saddle River, NJ: Prentice Hall; 2008. Brady NC, Weil RR, Weil RR. The nature and properties of soils. Upper Saddle River, NJ: Prentice Hall; 2008. Butler V, Bornman CH, Evert RF. Welwitschia mirabilis: morphology of the seedling. Botanical Gazette. 1973 Mar 1;134(1):52-9. Caporaso JG, Kuczynski J, Stombaugh J, Bittinger K, Bushman FD, Costello EK, et al. QIIME allows analysis of high-throughput community sequencing data. Nat Methods. 2010;7:335–6

Shaaban D.H. 76 Master Thesis

Carlile MJ, Watkinson SC, Gooday GW. The fungi. Gulf Professional Publishing; 2001 Jan 23. Chaw SM, Parkinson CL, Cheng Y, Vincent TM, Palmer JD. Seed plant phylogeny inferred from all three plant genomes: Monophyly of extant gymnosperms and origin of Gnetales from conifers. Proc Natl Acad Sci USA. 2000; 97: 4086–91. PMID: 10760277 Chew RM, Whitford WG. A long-term positive effect of kangaroo rats (Dipodomys spectabilis) on creosotebushes (Larrea tridentata). Journal of Arid Environments. 1992 May;22:375-86. Cooper-Driver GA, Wagner C, Kolberg H. Patterns of Aspergillus niger var. phoenicis (Corda) Al-Musallam infection in Namibian population of Welwitschia mirabilis Hook. f. J Arid Environ 2000; 46: 181-98. Coppinger, K.D., Reiners, W.A., Burke, I.C., Olson, R.K., 1991. Net erosion on a sagebrush steppe landscape as determined by cesium-137 distribution. Soil Science Society of America Journal 55, 254–258. Coppinger, K.D., Reiners, W.A., Burke, I.C., Olson, R.K., 1991. Net erosion on a sagebrush steppe landscape as determined by cesium-137 distribution. Soil Science Society of America Journal 55, 254–258. Cross, A. F., & Schlesinger, W. H. (1999). Plant regulation of soil nutrient distribution in the northern Chihuahuan Desert. Plant Ecology, 145, 11–25. https://doi.org/10.1023/A:1009865020145 Danin A. Plant adaptations to environmental stresses in desert dunes. In: Cloudsley- Thompson J, Punzo F, editors. Adapt. Desert Org; 1996. DeSantis, T. Z. et al. Greengenes, a chimera-checked 16S rRNA gene database and workbench compatible with ARB. Appl. Environ. Microbiol. 72, 5069–5072 (2006). Desert Ecological Research Unit, Desert Research Foundation of Namibia, Gobabeb Training and Research Centre, P.O. Box 953, Walvis Bay, Namibia (E-mail: [email protected])

Shaaban D.H. 77 Master Thesis

Ebner M, Miranda T, Roth-Nebelsick A. Efficient fog harvesting by Stipagrostis sabulicola (Namib dune bushman grass). Journal of arid environments. 2011 Jun 1;75(6):524- 31. Eckardt, F. D. & Spiro, B. The origin of sulphur in gypsum and dissolved sulphate in the Central Namib Desert, Namibia. Sedimentary Geol 123, 255–273 (1999). Eckardt, F. D. et al. The nature of moisture at Gobabeb, in the central Namib Desert. J Arid Environ 93, 7–19 (2013). Edgar RC. UPARSE: highly accurate OTU sequences from microbial amplicon reads. Nat Methods. 2013;10:996–8. Eller BM, von Willert DJ, Brinckmann E, Baasch R. Ecophysi- ological studies on Welwitschia mirabilis in the Namib desert. S Afr J Bot 1983; 2: 209-23. Fahn, A., & Cutler, D. F. (1992). Xeromorphytes. Gebrüder Borntraeger. Fan Y, Miguez-Macho G, Jobbágy EG, Jackson RB, Otero-Casal C. Hydrologic regulation of plant rooting depth. Proceedings of the National Academy of Sciences. 2017 Oct 3;114(40):10572-7. Fan, Y., Miguez‐Macho, G., Jobbágy, E. G., Jackson, R. B., & Otero‐Casal, C. (2017). Hydrologic regulation of plant rooting depth. Proceedings of the National Academy of Sciences, 114, 10572–10577. Frossard A, Ramond JB, Seely M, Cowan DA. Water regime history drives responses of soil Namib Desert microbial communities to wetting events. Scientific reports. 2015 Jul 21;5:12263. Gagliardo RW, Chilton WS. Soil transformation of 2 (3H)-benzoxazolone of rye into phytotoxic 2-amino-3H-phenoxazin-3-one. Journal of chemical ecology. 1992 Oct 1;18(10):1683-91. Garner, W., Steinberger, Y., 1989. A proposed mechanism for the formation of ‘fertile islands’ in the desert ecosystem. Journal of Arid Environments 16, 257–262. Giess, W. 1969. Welwitschia mirabilis Hook. fil. Dinteria 3: 3–55. Glick BR. Plant Growth-Promoting Bacteria: Mechanisms and Applications. Scientifica. 2012; 2012:15.

Shaaban D.H. 78 Master Thesis

Gomes, M., Garcia, Q., Barreto, L., Pimenta, L., Matheus, M., Figueredo, C. 2017. Allelopathy: An overview from micro- to macroscopic organisms, from cells to environments, and the perspectives in a climate-changing world. Academic Press, New York. 72/2: 113—129 González‐Ruiz, T., Rodríguez‐Zaragoza, S., & Ferrera‐Cerrato, R. (2008). Fertility islands around Prosopis laevigata and Pachycereus hollianus in the drylands of Zapotitlán Salinas, México. Journal of Arid Environments, 72, 1202–1212. https://doi.org/10.1016/j.jaridenv.2007.12.008 Gupta S, Larson WE. Estimating soil water retention characteristics from particle size distribution, organic matter percent, and bulk density. Water resources research. 1979 Dec;15(6):1633-5. Hanson, A. D. 1980. Interpreting the metabolic response of plants to water stress. HortScience 15:623-29. Henschel JR, Seely MK. Ecophysiology of atmospheric moisture in the Namib Desert. Atmospheric Research. 2008 Mar 1;87(3-4):362-8. Henschel JR, Wassenaar TD, Kanandjembo A, Louw MK, Neef G, Shuuya T, Soderberg K. Roots point to water sources of Welwitschia mirabilis in a hyperarid desert. Ecohydrology. 2019 Jan; 12(1):e2039. Henschel, J., Wassenaar, R., Kanandjembo, A., Louw, M., Neef, G., Shuuya, T., Soderberg, K et al. (2019). Roots point to water sources of Welwitschia mirabilis in a hyperarid desert. RESEARCH ARTICLE. DOI: 10.1002/eco.2039 Henschel, J.R., Seely, M.K., 2000. Long-term growth of Welwitschia mirabilis, a long lived plant of the Namib desert (including a bibliography). Plant Ecol. 150, 7e26. Henschel, J.R., Seely, M.K., 2008. Ecophysiology of atmospheric moisture in the Henschel, JR, Wassenaar, TD, Kanandjembo, A, et al. Roots point to water sources of Welwitschia mirabilisin a hyperarid desert. Ecohydrology. 2019; 12:e2039 Hook, P.B., Burke, I.C., Lauenroth, W.K., 1991. Heterogeneity of soil and plant N and C associated with individual plants and openings in North American shortgrass steppe. Plant and Soil 138, 247–256.

Shaaban D.H. 79 Master Thesis

Hu, R., Wang, X., Zhang, Y., Shi, W., Jin, Y., Chen, N., 2016. Insight into the influence of Hudson BD. Soil organic matter and available water capacity. Journal of Soil and Water Conservation. 1994 Mar 1;49(2):189-94. Jacobson, K. M., & Lester, E. (2003). A first assessment of genetic variation in Welwitschia mirabilis Hook. Journal of Heredity, 94(3), 212-217. Jenkinson, D. S. 1988. Determination of microbial biomass carbon and nitrogen in soils. In: Advances in Nitrogen Cycling in Agricultural Ecosystems. pp. 368–386. Wil- son, J. R., Ed., International, Walling-Ford. Joh R. Henschel & Mary K. Seely Jones DL, Oburger E. Solubilization of phosphorus by soil microorganisms. InPhosphorus in action 2011 (pp. 169-198). Springer, Berlin, Heidelberg. Kaminsky R. The microbial origin of the allelopathic potential of Adenostoma fasciculatum H & A. Ecological Monographs. 1981 Feb;51(3):365-82. Kers, L. E. (1967). The distribution of Welwitschia mirabilis Hook. f. Svensk Botanisk Tidskrift, 61(1), 97-125. Klopatek JM, Stock WD. Partitioning of nutrients in Acanthosicyos horridus, a keystone endemic species in the Namib Desert. journal of Arid Environments. 1994 Mar 1;26(3):233-40. Kögel-Knabner I. The macromolecular organic composition of plant and microbial residues as inputs to soil organic matter. Soil Biology and Biochemistry. 2002 Feb 1;34(2):139-62. Kögel-Knabner I. The macromolecular organic composition of plant and microbial residues as inputs to soil organic matter. Soil Biology and Biochemistry. 2002 Feb 1;34(2):139-62. Kuske CR, Ticknor LO, Miller ME, Dunbar JM, Davis JA, Barns SM, Belnap J. Comparison of soil bacterial communities in rhizospheres of three plant species and the interspaces in an arid grassland. Appl. Environ. Microbiol.. 2002 Apr 1;68(4):1854-63. Kutschera, L., Lichtenegger, E., Sobotik, M. & Haas, D. 1997a. Die Wurzel, das neue Organ, ihre Bedeutung für das Leben von Welwitschia mirabilis und anderer Arten

Shaaban D.H. 80 Master Thesis

der Namib sowie von Arten angrenzender Gebiete, mit Erklärungen des geotropen Wachstums der Pflanzen. Pflanzensoziologisches Institut, Klagenfurt, Austria. 94 pp. Laity JJ. Deserts and desert environments (Vol. 3). Wiley; 2009. Lodewyckx, C., Vangronsveld, J., Porteous, F., Moore, E. R. B., Taghavi, S., Mezgeay, M., et al. (2002). Endophytic bacteria and their potential applications. Crit. Rev. Plant Sci. 21, 583–606. doi: 10.1080/0735-260291044377 Lou, Y., Davis, A.S. & Yannarell, A.C. Plant Soil (2016) 399: 357. https://doi.org/10.1007/s11104-015-2698-8 Lukaszkiewicz J, Kosmala M. Determining the age of streetside trees with diameter at breast height-based multifactorial model. Arboriculture and Urban Forestry. 2008;34(3):137. Lukaszkiewicz, J., Kosmala, M et al. Determining the age of streetside trees with diameter at breast hight-based multifactorial model. Arboriculture & Urban Forestry. 2008; 34(3): 137-143 Mahmood T, Mehnaz S, Fleischmann F, Ali R, Hashmi ZH, Iqbal Z. Soil sterilization effects on root growth and formation of rhizosheaths in wheat seedlings. Pedobiologia. 2014 May 1;57(3):123-30. Makhalanyane TP, Valverde A, Gunnigle E, Frossard A, Ramond J-BJB, Cowan DA. Microbial ecology of hot desert edaphic systems. FEMS Microbiol Rev. 2015;39:203– 21. Makhalanyane, T. P., Valverde, A., Gunnigle, E., Frossard, A., Ramond, J. B., & Cowan, D. A. (2015). Microbial ecology of hot desert edaphic systems. FEMS Microbiology Reviews, 39(2), 203–221. Marsh, B., (1990). The Microenvironment Associated with Welwitschia mirabilis in the Namib Desert Desert Ecological Research Unit of Namibia, P. 0. Box 1592, Swakopmund, 9000 Namibia & Zoology Department, University of Cape Town, Rondebosch, Cape Town, 8000 South Africa

Shaaban D.H. 81 Master Thesis

Martinez-Meza E, Whitford WG. Stemflow, throughfall and channelization of stemflow by roots in three Chihuahuan desert shrubs. Journal of Arid Environments. 1996 Mar 1;32(3):271-87. Mitchell, R.J., Campbell, C.D., Chapman, S.J., Osler, G.H.R., Vanbergen, A.J., Ross, L.C., Cameron, C.M. & Cole, L. 2007. The cascading effects of birch on heather moorland: a test for the top-down control of an ecosystem engineer. Journal of Ecology 93: 540–554. Molisch, H. 1937. Der Einfluss einer Pflanze auf die andere-Allelopathie. Fischer, Jena. Mou, P., Jones, R.H., Mitchell, R.J., Zutter, B., 1995. Spatial distribution of roots in Swetgum and Lobolly Pine monocultures and relations with above-ground biomass and soil nutrients. Functional Ecology 9, 689–699. Muller D.B., Vogel C., Bai Y., Vorholt J.A. (2016). “The plant microbiota: systems-level insights and perspectives,” in Annual Review of Genetics Vol. 50 ed. Bonini N. M., editor. (Palo Alto, CA: Annual Reviews; ) 211–234 Nilsen ET, Sharifi MR, Rundel PW. Comparative water relations of phreatophytes in the Sonoran Desert of California. Ecology. 1984 Jun;65(3):767-78. Nobel PS, McDaniel RG. Low temperature tolerances, nocturnal acid accumulation, and biomass increases for seven species of agave. Journal of Arid Environments. 1988 Nov 1;15(2):147-55. Novara A, Rühl J, La Mantia T, Gristina L, La Bella S, Tuttolomondo T. Litter contribution to soil organic carbon in the processes of agriculture abandon. Solid Earth. 2015 Apr 24;6(2):425-32. NOY-MEIR. I.. 1979/80. Structure and function of a desert ecosystem. Israel Journal of Botany 28: 1-19. Pekarek, E., Jacobson, K., & Donovan, A. (2006). High levels of genetic variation exist in Aspergillus niger populations infecting Welwitschia mirabilis Hook. Journal of Heredity, 97(3), 270-278.

Shaaban D.H. 82 Master Thesis

Pereira SR, Laura VA, Souza AL. Seed dormancy overcoming as a strategy for forest restoration in tropical pasture. Pesquisa Agropecuária Brasileira. 2013 Feb;48(2):148-56. Pettit RE. Organic matter, humus, humate, humic acid, fulvic acid and humin: their importance in soil fertility and plant health. CTI Research. 2004:1-7. Powlson, D.S., Hirsch, P.R. & Brookes, P.C. Nutrient Cycling in Agroecosystems (2001) 61: 41. https://doi.org/10.1023/A:1013338028454 Reigosa MJ, Pedrol N, González L, editors. Allelopathy: a physiological process with ecological implications. Springer Science & Business Media; 2006 Feb 28. Rice E. L. (1984). Allelopathy, 2nd ed. New York: Academic Press Rice KJ, Dyer AR. Seed aging, delayed germination and reduced competitive ability in Bromus tectorum. Plant Ecology. 2001 Aug 1;155(2):237-43. Ritz K. Fungi. Encydopedia of soil in the environment; 2005, 111-119 Ronca S, Ramond JB, Jones BE, Seely M, Cowan DA. Namib Desert dune/interdune transects exhibit habitat-specific edaphic bacterial communities. Frontiers in microbiology. 2015 Sep 4;6:845. Rossi BE, Villagra PE. Effects of Prosopis flexuosa on soil properties and the spatial pattern of understorey species in arid Argentina. Journal of Vegetation Science. 2003 Aug;14(4):543-50. Rossi, B. E., & Villagra, P. E. (2003). Effects of Prosopis flexuosa on soil properties and the spatial pattern of understorey species in arid Argentina. Journal of Vegetation Science, 14, 543–550. https://doi.org/10.1658/ 1100- 9233(2003)014%5B0543:EOPFOS%5D2.0.CO;2 Roth-Nebelsick A, Ebner M, Miranda T, Gottschalk V, Voigt D, Gorb S, et al. Leaf surface structures enable the endemic Namib desert grass Stipagrostis sabulicola to irrigate itself with fog water. J R Soc Interface. 2012;9:1965–74. Ruiz TG, Zaragoza SR, Cerrato RF. Fertility islands around Prosopis laevigata and Pachycereus hollianus in the drylands of Zapotitlán Salinas, México. Journal of Arid Environments. 2008 Jul 1;72(7):1202-12.

Shaaban D.H. 83 Master Thesis

Ruppel S, Franken P, Witzel K. Properties of the halophyte microbiome and their implications for plant salt tolerance. Functional Plant Biology. 2013 Aug 23;40(9):940-51. Salvatore, M., Carafa, A., Carratu’, G., et al (2013). Growth and reproductive phenology of Welwitschia mirabilis hook. F. The open plant science journal. 7, 39-46. Schade JD, Hobbie SE. Spatial and temporal variation in islands of fertility in the Sonoran Desert. Biogeochemistry. 2005 Apr 1;73(3):541-53. Schlesinger, W. H., Raikes, J. A., Hartley, A. E. & Cross, A. F. 1996. On the spatial pattern of soil nutrients in desert ecosystems. Ecology 77: 364–374. Schlesinger, W. H., Raikes, J. A., Hartley, A. E. & Cross, A. F. 1996. On the spatial pattern of soil nutrients in desert ecosystems. Ecology 77: 364–374. SCHULZE, E. D., ELLER. B, M., THOMAS, D. A., WILLERT, von D. J. and BRINCKMANN, E., 1980. Leaf temperatures and energy balance of Welwitschia mirabilis in its natural habitat. Oecologia 44: 258-262. Stres B, Bonete MJ, Martínez-Espinosa RM, Mahne I, Bothe H. Organisms of the Nitrogen Cycle under extreme conditions: low temperature, salinity, pH value and water stress. InBiology of the Nitrogen Cycle 2007 Jan 1 (pp. 369-379). Elsevier. Sun, Y., Zhang, Y., Feng, W., Qin, S., Liu, Z., Bai, Y., Yan, R., Fa, K., 2017. Effects of xeric Taylor-George S, Palmer F, Staley J, et al. Fungi and bacteria involved in desert varnish formation. Microb Ecol 1983;9:227–45. Tedersoo L, Anslan S, Bahram M, Põlme S, Riit T, Liiv I, Kõljalg U, Kisand V, Nilsson RH, Hildebrand F, Bork P. Shotgun metagenomes and multiple primer pair-barcode combinations of amplicons reveal biases in metabarcoding analyses of fungi. MycoKeys. 2015(10):1-43. Tedersoo L. (2017) Global Biogeography and Invasions of Ectomycorrhizal Plants: Past, Present and Future. In: Tedersoo L. (eds) Biogeography of Mycorrhizal Symbiosis. Ecological Studies (Analysis and Synthesis), vol 230. Springer, Cham

Shaaban D.H. 84 Master Thesis

Thompson, T.L., Zaady, E., Huancheng, P., Wilson, T.B., Martens, D.A., 2006. Soil C and N pools in patchy shrublands of the Negev and Chihuahuan Deserts. Soil Biol. Biochem. 38, 1943–1955. Valverde A, De Maayer P, Oberholster T, Henschel J, Louw MK, Cowan D (2016) Specific Microbial Communities Associate with the Rhizosphere of Welwitschia mirabilis, a Living Fossil. PLoS ONE 11(4): e0153353. doi:10.1371/journal. pone.0153353 Valverde A, De Maayer P, Oberholster T, Henschel J, Louw MK, Cowan D (2016) Specific Microbial Communities Associate with the Rhizosphere of Welwitschia mirabilis, a Living Fossil. PLoS ONE 11(4): e0153353. doi:10.1371/journal. pone.0153353 Van Jaarsveld EJ, Pond U. Uncrowned Monarch of the Namib: Welwitschia Mirabilis. Penrock publications; 2013. von Willert DJ, Eller BM, Brinckmann E, Baasch R. CO2 Gas Exchange and traspiration of Welwitschia mirabilis Hook. F. Cen- tral Namib Desert 1982; Oecologia 55: 21-9. Wallace A, Romney EM, Hunter RB. The challenge of a desert: revegetation of disturbed desert lands. Great Basin Naturalist Memoirs. 1980 Jan 1:216-25. Williamson GB, Obee EM, Weidenhamer JD. Inhibition ofSchizachyrium scoparium (poaceae) by the allelochemical hydrocinnamic acid. Journal of Chemical Ecology. 1992 Nov 1;18(11):2095-105. Yadav AN, Sharma D, Gulati S, Singh S, Dey R, Pal KK, Kaushik R, Saxena AK. Haloarchaea endowed with phosphorus solubilization attribute implicated in phosphorus cycle. Scientific reports. 2015 Jul 28;5:12293. Yao, Y., ShaO, M., Fu, X., Wang, X., Wei, X et al. Effects of shrubs on soil nutrients and enzymatic activities over a 0–100 cm soil profile in the desert-loess transition zone. Catena 174 (2019) 362–370.

Shaaban D.H. 85 Master Thesis

Supplementary Material

Supplementary Figure 2.1. Litter produced by W. mirabilis male plants. (A) W. mirabilis growing in the Wash site and relative allelofertility island created by the litter (i.e., male cone). (B) Detail of dry male cone fall down and accumulate as litter in the center of the W. mirabilis stock (C) W. mirabilis male cones in flower.

Supplementary Figure 2.2. W. mirabilis allelofertility island. Sampling design adopted to characterize the microbial community associated with different zone of W. mirabilis allelofertility island. Samples are collected at increasing distance from the plant in the allofertility island (Z1, Z2 and Z3; white labels in the scheme) and 3-5 mt away from the plant (Z4 and Z5; red labels in the scheme) in an unvegetated area between W. mirabilis plants. Samples of litter (Z0; blue label in the scheme) are also collected from the center of each plant.

Shaaban D.H. 86 Master Thesis

Supplementary Figure 2.3. Rarefaction curves for bacterial, archaeal and fungal OTUs, clustering at 97% sequence similarity. Curves represent sequences for multiple samples of litter (Z0; black line), allelofertility island (Z1, Z2 and Z3; brown-shade lines) and unvegetated barren soil (Z4 and Z5; grey-shade lines).

Shaaban D.H. 87 Master Thesis

Supplemenatry Figure 2.4 In vivo experiment to evaluate the effect of W. mirabilis- allelofertility island soils on seed germination under watered and controlled-drought conditions. Three types of soil were used: litter mix soil (black color); allelofertility soil (brown color); barren soil (gray color). Plastic pots (10 cm diameter) were filled with the soils of the four different treatments; three pots for each treatment were prepared, and 10 barley seeds added for each one. (A) Under normal irrigation conditions, pots were watered with 50 ml of milliq water, pots were properly irrigated for all the duration of the experiment. (B) Under controlled-drought conditions, after 3 days of watering, irrigation was alternated by two drought periods of 5 days.

Shaaban D.H. 88 Master Thesis

Supplementary Table 2.1. List of all soil samples collected from the five W. mirabilis plants Plant diameter, sample code, location along the fertility island gradient and distance from the center of the plant are reported for each set of samples. Size of allelofertility island is indicated by the distance of border samples for each sampled plant. Diameter and distance are expressed in cm. W, W. mirabilis; AFI, allelofertility island. (*) Distance value measured from the center of the plant.

Plant W diameter Sample code AFI gradient Distance* AFI Zone A 190 cm A01- A04 AFI Start 95 Z1 A05- A08 AFI Middle 175 Z2 A09- A12 AFI Border 255 Z3 A13- A16 Bulk 1 445 Z4 A17- A20 Bulk 2 595 Z5 A21- A24 Litter 0 Z0 B 140 cm B01- B04 AFI Start 70 Z1 B05- B08 AFI Middle 120 Z2 B09- B12 AFI Border 200 Z3 B13- B16 Bulk 1 370 Z4 B17- B20 Bulk 2 570 Z5 B21- B24 Litter 0 Z0 C 130 cm C01- C04 AFI Start 65 Z1 C05- C08 AFI Middle 135 Z2 C09- C12 AFI Border 205 Z3 C13- C16 Bulk 1 365 Z4 C17- C20 Bulk 2 565 Z5 C21- C24 Litter 0 Z0 D 120 cm D01- D04 AFI Start 60 Z1 D05- D08 AFI Middle 110 Z2 D09- D12 AFI Border 240 Z3 D13- D16 Bulk 1 360 Z4 D17- D20 Bulk 2 560 Z5 D21- D24 Litter 0 Z0 E 95 cm E01- E04 AFI Start 47.5 Z1 E05- E08 AFI Middle 87.5 Z2 E09- E12 AFI Border 147.5 Z3 E13- E16 Bulk 1 347.5 Z4 E17- E20 Bulk 2 547.5 Z5 E21- E24 Litter 0 Z0

Shaaban D.H. 89 Master Thesis

Supplementary Table 2.2. Cross validation of microbial community associated with the samples collected at increasing distance from the plant along the allelofertility island (from Z0 to Z5).

A) Bacterial communities (correct, 94.9%) Group Z0 Z1 Z2 Z3 Z4 Z5 Total %correct Z0 0 0 0 0 18 0 18 100 Z1 0 0 0 0 0 20 20 100 Z2 20 0 0 0 0 0 20 100 Z3 0 19 1 0 0 0 20 95 Z4 0 1 17 2 0 0 20 85 Z5 0 0 2 18 0 0 20 90

B) Archaeal communities (correct, 77.6%) Group Z0 Z1 Z2 Z3 Z4 Z5 Total %correct Z0 16 1 0 0 0 0 17 94.1 Z1 2 17 1 0 0 0 20 85 Z2 0 1 18 1 0 0 20 90 Z3 0 0 3 13 2 1 19 68.4 Z4 0 0 0 1 13 6 20 65 Z5 0 0 0 0 7 13 20 65

C) Fungal communities (correct, 82.5%) Group Z0 Z1 Z2 Z3 Z4 Z5 Total %correct Z0 20 0 0 0 0 0 20 100 Z1 0 14 4 2 0 0 20 70 Z2 0 0 17 3 0 0 20 85 Z3 0 0 2 16 2 0 20 80 Z4 0 0 0 1 16 3 20 80 Z5 0 0 0 2 2 16 20 80