IMPROVING ECOSYSTEM FUNCTION:

FACILITATING RESTORATION OF DEGRADED BIOCRUSTS

USING MIXED CULTURE INOCULATION

A Thesis

Presented to the

Faculty of

San Diego State University

In Partial Fulfillment

of the Requirements for the Degree

Master of Science

in

Biology

by

Sharon Reeve

Fall 2014

SAN DIEGO STATE UNIVERSITY

The Undersigned Faculty Committee Approves the

Thesis of Sharon Muczynski:

Improving Ecosystem Function:

Facilitating Restoration of Degraded Biocrusts

Using Mixed Culture Inoculation

David Lipson, ChairDepartment of Biology

Tom ZinkDepartment of Biology

John O’LearyDepartment of Geology

Approval Date

Copyright © 2014

by

Sharon Reeve

All Rights Reserved

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Action is the antidote to despair. — Joan Baez

I believe our biggest issue is the same biggest issue that the whole world is facing, and that is habitat destruction.

—Steve Irwin

viv

ABSTRACT OF THE THESIS

Improving Ecosystem Function:Facilitating Restoration of Degraded Biocrusts

Using Mixed Culture Inoculationby

Sharon ReeveMaster of Science in Biology

San Diego State University, 2014

In arid and semiarid ecosystems where physiological constraints prevent most vascular plant establishment, the biological soil crust (biocrust) community is ecologically critical. The key to the survival of biocrust is the versatility and adaptability of cyanobacteria and green algae. The organismal construct of the biocrust community is especially vulnerable to compressional forces, and is slow to recover without assistance. Given the importance of biocrusts to so many aspects of healthy ecosytem function, it would be advantageous if land managers prioritized restoring damaged biocrusts. Coastal sage scrub (CSS) is a unique and imperiled valuable habitat and is nearly unmatched in the biodiversity of unique plants and animals. Recent work in biocrust restoration finds that assisted restoration speeds recovery of functionality in biocrusts. At present, studies of biocrust restoration in CSS habitat do not exist. This study examines the feasibility of isolating and culturing a mix of endemic CSS cyanobacteria and green algae to inoculate native CSS soil, thereby facilitating recovery of disturbed biological soil crusts. It further looks at markers for culture growth, chlorophyll a, extractable polysaccharides, and stability, to gauge whether inoculation and growth of the culture have increased soil function. Growth of the mixed culture and increases in functionality are compared between autoclaved soil inoculations and native soil inoculations to determine the extent that native crust organisms can regrow without inoculation, and how the inoculum interacts with the native microbial community. A putative novel genus and species of cyanobacteria related to Leptolyngbya was isolated and tentatively included in the genus, Trichotorquatus. The mixed culture included a green algae, possibly a species of Trebouxia. Mixed inoculum added to native soil significantly increased chorophyll a levels and soil stability, and increased extractable polysaccharides after just two months, demonstrating recovery of function. Autoclaving soil reduced increases in functionality indicating the importance of the intact soil community for growth. It may be possible in the future to restore biocrust in CSS using mixed culture inoculation.

TABLE OF CONTENTS

PAGEABSTRACT................................................................................................................................ LIST OF TABLES.................................................................................................................... LIST OF FIGURES....................................................................................................................CHAPTER 1 INTRODUCTION.......................................................................................................... 2 LITERATURE REVIEW................................................................................................ Characterization of Biological Soil Crust....................................................................... Biocrust Ecosystem Services.......................................................................................... Biocrust Damage by Compression................................................................................. Biocrust Carbon and Nitrogen Storage........................................................................... Restoration of Biocrust................................................................................................... Coastal Sage Scrub—Imperiled Habitat......................................................................... Biocrust Restoration of Coastal Sage Scrub................................................................. Research Questions....................................................................................................... 3 MATERIALS AND METHODS.................................................................................. Study Sites.................................................................................................................... Field Sampling.............................................................................................................. Growth Medium............................................................................................................ Isolating Strains of Cyanobacteria and Green Algae.................................................... Chlorophyll a Measurements........................................................................................ Reference Measurements for Characteristics and Chlorophyll a................................. Reference Measurements for Extractable Polysaccharides.......................................... Reference Measurements for Soil Stability.................................................................. DNA Extraction, Purification, and Cloning.................................................................. Inoculation Experiment................................................................................................. Final Analysis of Inoculation Experiment.................................................................... Experimental Design and Statistical Analysis.............................................................. 4 RESULTS..................................................................................................................... Culturing Experiments.................................................................................................. Chlorophyll a Results...................................................................................................

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TABLE OF CONTENTS

PAGE Extractable Polysaccharides Results............................................................................. Stability Test Results.................................................................................................... Correlation Test Results................................................................................................ PCR Test Results.......................................................................................................... 5 DISCUSSION............................................................................................................... Leptolyngbya foveolarum/ Trichotorquatus.................................................................. Green Algae Trebouxia sp. ........................................................................................... Chlorophyll a................................................................................................................ Extractable Polysaccharides......................................................................................... Stability......................................................................................................................... Conclusion.................................................................................................................... 6 ACKNOWLEDGEMENTS.......................................................................................... 7 REFERENCES.............................................................................................................

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LIST OF TABLES

PAGETable 1. Growth Media Ingredients BG-11 ..............................................................................Table 2. Stability Class Table: Criteria for the Assignment of Crust to Stability Classes........Table 3. Baseline Values for Reference Biological Soil Crusts................................................Table 4. Statistical Values for One-way Analysis of Variance..................................................Table 5. Statistical Values for Two-way Analysis of Variance..................................................Table 6. Correlations Between Treatments................................................................................

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LIST OF FIGURES

PAGEFigure 1. Coastal sage scrub locations.....................................................................................Figure 2. Coastal sage scrub location study sites..................................................................... Figure 3. Coastal sage scrub biological soil crust A. Lichenized crust at SMER B. Visible cyanobacterial crust from Mission Trails.........Figure 4. Collected biocrust.....................................................................................................Figure 5. Flasks of inoculant MT A, MT A-2, MT B, & SMER A..........................................Figure 6. Soil sieve apparatus...................................................................................................Figure 7. Reference CSS biocrusts...........................................................................................Figure 8. Before inoculation.....................................................................................................Figure 9. Representative photographs of each treatment.........................................................Figure 10. Single strand cyanobacteria from isolation............................................................. Figure 11. Chlorophyll a concentrations in five treatments.....................................................Figure 12. Extractable polysaccharide concentrations in the five treatments..........................Figure 13. Stability index of soils from five treatments...........................................................Figure 14. Maximum likelihood phylogenetic tree..................................................................Figure 15. Neighbor-joining phylogenetic tree showing placement of SMER-A cyanobacterial isolate in novel genus, Trichotorquatus ..........................................................Figure 16. Green algae Trebouxia sp. from Mission Trails isolation culture...........................Figure 17. Mix of green algae, Trebouxia sp. and Trichotorquatus sp.....................................Figure 18. Unknown green algae recovered from ground crust inoculation............................

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INTRODUCTION

Intact biological soil crusts or biocrusts play a pivotal role in the development and function of biodiverse semi-arid and arid landscapes (Garcia-Pichel 2003; Maestre et al. 2005). In arid ecosystems where physiological constraints prevent most vascular plant establishment, the biological soil crust community is ecologically critical (Belnap and Lange 2003). These thin veneers of biological activity transform harsh desert landscapes, making them conducive to life and enabling complex ecosystems to form. Multi-functional biological soil crusts can be found on much of the Earth’s surface and perform many vital ecosystem services that are valuable to the optimal health of aridlands (Belnap and Lange 2003). Disturbance, to the detriment of the system, simplifies the biodiversity of biocrusts. Even small, seemingly non-visible damage to soil crusts can lead to long-lasting ecosystem function losses (Bowker et al. 2011; Tongway 1994). Further damage to biocrusts can accelerate loss of biological function leading to irreversible desertification (Belnap and Eldridge 2001). Loss of biocrust to desertification is devastating to ecosystems and costly in the loss of environmental services (Pimental et al. 1995). Given the importance of biocrusts, it is unfortunate that restoration efforts of arid and semi-arid lands rarely include plans for remediation of biological soil crusts (Bowker et al. 2008). For degraded ecosystems, it is necessary to assist in rehabilitation in order to prevent the negative feedback of desertification (Bowker 2007). This is especially timely now as San Diego’s climate is getting warmer and drier (Messner et al. 2011). Research on biological soil crust restoration is lacking, but what is available strongly supports inclusion of biocrust remediation into site restoration plans (Bowker 2007). Given the importance of biocrust in a dual role as ecosystem engineer and as modulator of the soil-atmosphere interface, it is imperative that land owners protect and attempt to restore these ecosystems (Bowker et al. 2011; Eldridge et al. 2010). In spite of the overwhelming evidence of many important ecosystem functions provided by biocrusts, no functional restoration techniques have been developed in Coastal Sage Scrub (CSS) habitats (Bowker et al. 2006). Some research has shown success in using ground biocrust to restore damaged areas, but that approach is destructive (Cole et al. 2010). New research indicates that assisted recolonization of biological soil crusts through inoculation of cyanobacteria can accelerate restoration of compromised areas (Acea et al. 2003; Zheng et al. 2011; Xu et al. 2012). Restoration inoculation has not been attempted in the unique ecosystem of San Diego CSS. This study looks at whether it is possible to isolate and grow cyanobacteria and green algae from CSS biocrust, and whether inoculation of native soil with a mixed culture from the isolation results in improved function.

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LITERATURE REVIEW

CharaCterization of BiologiCal Soil CruSt Occurring in arid and semi-arid environments, biological soil crusts (biocrusts) are only millimeters thick, yet are inordinately important for terrestrial ecosystem function (Xiao et al. 2011). In ecosystems where lack of water limits vascular plants, the biocrust community figures prominently in the landscape and covers large areas of dry land environments (Gar-cia-Pichel et al. 2001). Biocrusts are environmentally influential, globally distributed, and comprise a large biomass (Castillo-Monroy et al. 2011). Biocrusts contribute globally with total biomass estimated in the range of 109 metric tons (Garcia-Pichel 2003). Biocrusts are photosynthetically active, major microbial communities occurring on up to 70% of semi-arid and aridland, which make up 40% of Earth’s surface (Bowker 2007; Gilad et al. 2004). The key to survival of biocrust is the versatility and adaptability of cyanobacteria in the face of inhospitable environments (Jia et al. 2008). This group of organisms can tolerate extreme fluctuations in light, salinity, UV radiation, and moisture (Potts 1999; Belnap and Lange 2003). Cyanobacteria are ubiquitous prokaryotes, and are the photosynthetic primary producers and founding organisms of biocrust communities (Bates and Garcia-Pichel 2009; Rosentreter et al. 2007). Originally from aqueous environments, cyanobacteria are thought to be the oldest oxygen-producing organisms (Belnap et al. 2001). Their photosynthesis profoundly changed Earth’s atmosphere and allowed for evolution of organisms that can use oxygen for respiration (Berman-Frank et al. 2003). They are found in nearly every habitat on Earth, aqueous or land, thermophilic or in symbiosis with other organisms. Filamentous cyanobacteria generally dominate biocrusts with 70-98.6% of the biovolume, and contribute a large proportion of the ecosystem services provided by biocrust communities (Malam Issa et al. 2007; Řeháková et al. 2011). Filamentous cyanobacteria are aerobic photoautotrophs, and are mobile in moist soils (Belnap et al. 2001; Garcia-Pichel and Pringault 2001; Bowling et al. 2011). The filaments of cyanobacteria move through soil, maximizing photosynthesis, minimizing harmful levels of solar radiation, and colonizing new arid habitat (Evans and Johansen 1999; Belnap and Lange 2003). Some species of cyanobacteria can grow under particularly harsh levels of genetically damaging radiation because of protective pigments (Rosentreter et al. 2007; Bates and Garcia-Pichel 2009). The dark, protective, UVR-absorbing pigments scytonemin and gloecapsin are produced by two widespread genera of cyanobacteria: Nostoc and Scytonema (Yeager et al. 2004; Sinha et al. 2008; Peli et al. 2011). Scytonemin even protects desiccated cyanobacteria (Bowker et al. 2002; Tamaru et al. 2005).

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Cyanobacteria are photo- and hydrotactic, so are able to track light and water, two crucial functions optimizing survival and dispersal (Garcia-Pichel and Pringault 2001). Cyanobacteria can also survive the severe drought conditions that are often present in arid ecosystems (Gray et al. 2007; Peli et al. 2011). The sheath material of cyanobacteria, particularly Microcoleus, is a gelatinous matrix of extracellular polysaccharides (EPS) that adhere to soil particles (Tirkey and Adhikary 2005; Bates and Garcia-Pichel 2009). The polysaccharides work by cementing soil particles together, and by having a macromolecular structure that is easily adsorbed on clay particles (Maestre et al. 2011). Hyphal density is strongly correlated with soil stability, and nutrient levels. As the cyanobacteria grow they form a network of soil aggregates that stabilize soil, prevent erosion, enhance plant germination, and encourage further establishment of biocrust genera (Hawkes 2003; Li et al 2005). The network of soil aggregates and polysaccharides can absorb up to ten times their volume in water, slowly releasing it after precipitation (Belnap et al. 2006). Cyanobacteria are not the only biological soil crust organisms that generate sticky polysaccharides. The outer sheath of green algae and cell wall components are constructed of polysaccharides (König, J. and Peveling 1984) The slow moisture release increases the time that biocrust organisms, can be metabolically active, which is crucial in arid environments (Veste et al. 2001; Bowker 2007). Slow moisture release is also key for creating soil conditions that foster vascular plant growth and establishment. Biocrusts are complex microbial communities made of varying proportions of cyanobacteria, lichens, algae, bacteria, fungi, and bryophytes (Belnap and Lange 2003; Belnap et al. 2006). The assemblage of organisms in a particular biocrust depends on many factors. The local climate, aspect, slope, elevation, pH, presence of plants, and particularly the soil chemistry from parent material all factor into which biological soil crust organisms occur and in what proportions (Garcia-Pichel et al. 2003; Rosentreter et al. 2007; Bowker and Belnap 2008; Ochoa-Hueso et al. 2011; Pietrasiak et al. 2011; Hernandez and Knudsen 2012). The makeup of the biocrust varies greatly with locality and with time (Rosentreter et al. 2007). Younger, cyanobacterially dominated crusts are organismally simple, smooth, light-colored, and generally shallow at 1-2 mm depth (Belnap and Lange 2003). Later in biocrust development, smaller cyanobacteria, green algae, liverworts, fungi, lichens and bryophytes, all colonize on the soil surface and give crusts their distinctive coloration and patterning (Rosentreter et al. 2007). Older crusts become more complex in number of genera and topography and are often dark-colored, at least 3-5 mm but up to 14 mm deep (Belnap and Eldridge 2003; Langhans et al. 2009). Site conditions influence the organisms; for example, the biological soil crust of the Southern California coast has quite a different species makeup and appearance from the biocrust in the Colorado Plateau (Belnap and Lange 2003). Biocrusts are an important interface between atmosphere and the terrestrial environment, mediating energy, precipitation influx, and chemical

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exchange (Belnap et al. 2003; Castillo-Monroy et al. 2011). The biocrust community acts as an ecosystem engineer, able to alter, modify and control the physical state and chemical reactivity of the immediate habitat (Gilad et al. 2004; Bowker et al. 2005; Wright and Jones 2006; Eldridge et al. 2010). Indeed, the influence of biocrust on the local environment is considerable.

BioCruSt eCoSyStem ServiCeS

Biocrusts perform a number of valuable ecosystem services, including nitrogen fixation, carbon sequestration, soil stabilization, run-off reduction, precipitation mediation, dust nutrient collection, soil surface albedo moderation, microclimate amelioration, and nutrient chelation (Belnap et al. 2001; Lal 2004; Belnap et al. 2006; Bowker et al. 2008; Bowker et al. 2008a; Chaudhary et al. 2009; Bowker et al. 2010; Darby et al. 2010). Biocrusts also form the base of a biodiverse set of organisms and macroinvertebrates that contribute to ecosystem function and nutrient deposition through their individual characteristics, and through species interactions (Bowker et al. 2010; Darby et al. 2010). A study of the complexity of fauna from a Chihuahuan desert biocrust found an average of 44 genera of micro- and macroarthropods reflecting a rich biodiversity at multiple trophic levels occurring at an almost microscopic scale (Shepherd et al. 2002). In older crusts that have developed a higher proportion of lichens, mosses, and vascular plant roots, the resulting biocrust ecosystem forms a rolling micro-topography (Belnap and Lange 2003). Biocrust community-created variation in land relief influences species distribution, and to the extent it is maximized, reflects the health of the system (Belnap et al. 2001; Bowker et al. 2011; Castillo-Monroy et al. 2011). The micro-topography of biocrusts provides a complex landscape in which microfauna to hunt, hide, and procreate (Darby et al. 2010). The development of complexity in the miniature landscape stimulates the species richness that is so important to ecosystem function in biocrusts (Bowker et al. 2011; Maestre et al. 2012). The physical modifications and changes in chemical processes brought about by biocrust activity are considerable. Multiple species assimilating and breaking down organic matter create diverse chemical compounds and cycle nutrients (Bowker et al. 2011). Biocrust organisms even affect the assemblage of below-ground bacterial communities (Castillo-Monroy et al. 2011). Finally, biocrust organisms induce changes that make the area habitable by vascular plants (Schlesinger 1997), and promote germination (Belnap and Lange 2003; Godínez-Alvarez et al. 2012). A partial reason is that the pinnacled, discontinuous area between the sparse plants of aridlands serves as nutrient collection sites. Biocrusts collect nutrient-containing dust, and the chemical processes of the organisms supply a widespread

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variety of nutrients, including N, Ca²+, Co, P, K+, Mg 2+, Mn, and Zn that are often limited under vascular plants (Leonard et al. 1995; Darby et al. 2010). Phosphorus is often the limiting nutrient for plants, and biocrust organisms secrete substances increasing the bioavailability of phosphorus (Rosentreter et al. 2007). Metabolic processes in biocrust contribute to better fertility levels, and levels are also enhanced by the negatively charged, polysaccharide-based, sheath material of cyanobacteria, which binds positively charged macronutrients and prevents them from leaching (Leonard et al. 1995; Hawkes 2003; Pintado et al. 2005; Collins et al. 2008). Biocrusts deter soil erosion and desertification (Belnap et al. 2001). Not only do arid and semi-arid lands cover a large percentage, approximately 40%, of the Earth’s surface, they are also home to 38% of the human population (Wright and Jones 2006). With this sector of the population growing, the amount of arable land they live on is being lost to erosion and finally to desertification, decreasing the amount of food that can be grown (Pimentel 2006). Soil erosion is a huge worldwide environmental problem, second only to population growth (Pimentel 2006). Desertification is broadly defined as reduced land productivity and loss of ecosystem function, or loss of biological potential of an ecosystem (Pimentel 1995, Asner et al. 2003). Desertification is costly—roughly $23 billion a year is lost from mismanagement of arable land—and negatively impacts at least a billion people (Arnalds and Archer 2000). Worldwide, desertification affects 3.6 billion hectares, or about one-quarter of Earth’s surface (Wiegand and Jeltsch 2000). Much of the process is anthropogenic in origin: land-clearing, overgrazing, urbanization, air pollution, and excessive cultivation are all examples (Lovich and Bainbridge 1999; Asner et al. 2003; Gomez et al. 2012). The biocrust community stabilizes the soil and maintains its structural integrity (Bowker et al. 2008). The presence of well-developed biocrust decreases soil erodibility dramatically (Belnap and Eldridge 2001; Xiao 2011). Disturbance simplifies the biodiversity of biological soil crusts and can lead to water and wind erosion, loss of hard-won nutrient stores, and desertification (Belnap and Eldridge 2001; Bowling et al. 2011). The negative effects of degradation lead to loss of structure, loss of species, and finally, loss of function (Bowker 2007; Bowker et al. 2010). Destruction of biocrust destroys the critical aridland food web leaving the soil unprotected from abiotic forces (Belnap and Eldridge 2001). Sediment loss also means losing valuable soil nutrients, which significantly affects nutrient cycling (Barger et al. 2006). Furthermore, damage to biotic processes intensifies the physical forces of degradation in a negative feedback loop that reinforces the process of desertification (Gilad et al. 2004; Bowker 2007). Once a system has reached the endpoint of desertification, most biological crusts do not recover (Belnap and Eldridge 2001; Gleason and Freid 2006).

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BioCruSt Damage By CompreSSion Over 30 studies have confirmed the damage that the compressional forces of livestock graz-ing, vehicular and human traffic have caused to biocrusts, and to their capacity to store car-bon (Green et al. 2008; Grote et al. 2010). The organismal construct of the biocrust commu-nity is especially vulnerable to compressional forces, in part because cyanobacteria evolved before terrestrial organisms (Potts 1999). Off-road vehicles, long term grazing pressures, and trampling by humans all damage biocrust (Belnap and Lange 2003). Southern California has a long history of ranching from the mid-1800s (Lovitch and Bainbridge 1999). The destruc-tive impact from grazing animals eliminates the ecosystem function of soil crusts (Rosen-treter et al. 2007). Estimates place soil erosion rates from over-grazed pastures at over 100 tons ha-1 year-1 (Pimentel et al. 1995). Extensive land degradation is already altering Earth’s climate (Asner et al. 2003; Gleason and Freid 2006). This has intensified the need for restor-ing or protecting biocrusts. Until recently, biocrusts were not included in land management decisions, but that is changing as we learn about the value of protection (Hernandez and Sandquist 2011). Given the importance of biocrusts to so many aspects of healthy ecosytem function, it would be advantageous if land managers attempted to restore these ecosystems (Bowker et al. 2010b; Eldridge et al. 2010; Bowker et al. 2011). One study showed that global warming is likely to reduce cover, to simplify species richness, and ultimately, to reduce ecosystem function of biocrusts (Maestre et al. 2010). Another study posited that warmer temperatures together with grazing pressure will lead to greater C losses to the atmo-sphere (Thomas et al. 2011). Better conservation and restoration of damaged biocrusts will help offset this loss (Bowker 2007). Recent studies on functionality find that biocrust species are not redundant, and that each species contributes uniquely to healthy biocrust function, making restoration an even more desirable goal (Bowker et al. 2010b; Castillo-Monroy et al. 2011). Additionally, Maestre et al. (2012) found that ecosystem functionality increases with a greater number of species. Another compelling reason for restoration is that many biocrust species manufacture novel secondary metabolic compounds that assist in pathogen defense and nutrient uptake (Maestre et al. 2011). In a community situation, these metabolites could be mutually beneficial with other organisms and possibly synergistic in effect (Belnap and Lange 2003). Land degradation and desertification contribute substantially to the rise in atmospheric CO2 (Solomon et al. 2007; Singh et al. 2011). Biocrusts have the capacity to store large amounts of carbon and nitrogen. Most climate models fail to include the carbon fixed by soil organisms (Solomon et al. 2007). A possible strategy to offset the atmospheric rise in CO2 is to foster carbon sequestration in biocrusts (Lal 2004b). Biocrusts in aridlands have high soil C sequestration potential (Lal 2004). Fostering terrestrial carbon sequestration

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accomplishes two things: reduction of atmospheric CO2, and improvement to ecosystem function of the biocrusts. Since 1850, land use changes and cultivation have added 136 +/- 55 pentagrams of CO2 to the atmosphere (Lal 2004). Protection or restoration of degraded biocrusts is one way to sequester more carbon in the biosphere. One report suggests that restoration of 1.1 billion hectares of aridlands could sequester 0.2 to 0.4 gigatons of carbon per year (Lal 2004).

BioCruSt CarBon anD nitrogen Storage In biocrusts, the actions of cyanobacteria, lichens, green algae, and heterocystic bacteria (Hawkes 2003; Collins et al. 2008) add significant nitrogen to impoverished soils worldwide, releasing 5-70% fixed N, making them habitable for higher plant forms (Belnap and Lange 2003). In arid-land ecosystems, bacteria are important for nitrification, and fungi are important for translocating and mineralizing nitrogen (Collins et al. 2008). Arid soils colonized by biocrust are generally nitrogen limited, and in that way, biocrust acts as an eco-system engineer (Belnap and Lange 2003; Gilad et al. 2004; Bowker et al. 2006). Estimates of annual fixation range from 0.7 to 100 kg N ha-1 year -1 (Bates and Pichel 2009). In areas of high biocrust cover, dissolved organic N is the major form of nitrogen (Maestre et al. 2011). Cyanobacterial and green algae photosynthate products fuel the mobilization and mineral-ization of N (Green et al. 2008; Wang et al. 2008; Zhang and Feng 2008). Some genera of cyanobacteria are important nitrogen fixers of atmospheric N² in specialized structures called “heterocysts” and change N² into a form that is usable for vascular plants (Belnap et al. 2001; Starkenburg et al. 2011). When crusts are degraded or lost, soil N content decreases by 25-75% (Green et al. 2008). Southern California has drier conditions which makes restoration with cyanobacteria and green algae particularly advisable and viable (Barbour et al. 2007). Cyanobacteria can function in extreme drought conditions, unlike many vascular plants (Patzelt et al. 2014). Some green algae species can withstand even drier conditions than cyanobacteria (König, J. and Peveling 1984). Cyanobacteria and green algae are poikilohydric, so can survive cycles of dessication and rehydration (Belnap and Lange 2003). Water potential is the energy state of water, with pure water having a water potential of zero MPa. Most vascular plants have a limit of around -1.5 MPa, below this limit, death occurs because the plant cells do not have enough water to function (Raven et al. 2005). Nostoc, a common cyanobacteria genera can survive -100 MPa by producing extracellular polysaccharides (Potts 1999). This adaptation to environmental stress allows cyanobacteria and green algae to withstand long periods of drought (anhydrobiosis), sometimes even years of dormancy, and to become metabolically active within hours of precipitation (Belnap and Lange 2003; König, J. and Peveling 1984).

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Studies have shown that, because of their environment, biocrusts in California are naturally rich in cyanobacterial species and dominate the open spaces between vascular plants. (Pietrasiak et al. 2011). Additionally, other studies of cyanobacterially dominated biocrusts show increased germination, seedling survival, and growth of an endemic plant species (Pendleton et al. 2004; Godínez-Alvarez et al. 2012).

reStoration of BioCruSt

Biocrusts are slow to recover from disturbance. Cyanobacteria are the dominant pioneer species of biocrusts and are tolerant and resilient in the face of environmental extremes, which makes them good candidates for reintroduction into degraded biocrust areas (Belnap et al. 2003; Bowker 2007; Xu et al. 2012). The problem is, once biocrusts are impaired or destroyed, it takes, by some estimates, 14-100 years or more to recover, depending on conditions (Lovich and Bainbridge 1999; Belnap and Eldridge 2001). This factor alone has discouraged conservation and restoration. Recovery times vary with environmental conditions (Belknap et al. 2001). In arid landscapes with discontinuous plant cover, disturbance-induced declines are supplemented by accelerated soil erosion, loss of vascular plants, and persistent alteration of the biogeochemical characteristics of the intact system (Belnap and Eldridge 2001; Gilad et al. 2004; Bowker et al. 2005, Barger et al. 2006). That said, recent work in restoration finds that assisted restoration speeds recovery of functionality (Buttars et al. 1998; Belnap and Eldridge 2001; Malam Issa et al. 2007; Maqubela et al. 2009, Xiao et al. 2011; Zheng et al. 2011). A recent study in China found the presence of cyanobacterial polysaccharides increased seed germination and metabolic activity of adjacent vascular plants (Xu et al. 2012). Restoration techniques for biocrust are in their infancy. Typically, when restoration of a landscape is attempted, biocrusts are not included (Bowker et al. 2008). We know now that this incomplete approach negates the wealth of ecosystem services provided by biocrusts, and delivers a poor facsimile of the original rich ecosystem (Bowker 2007). Available data suggest that it is possible and advisable to restore biocrusts. Restoring biocrusts restores ecosystem function, which ultimately benefits humans (Bowker 2007). It makes sense to start restoration work with pioneer species of cyanobacteria. Unfortunately, there is a paucity of studies on establishing cyanobacterial populations as a tool for restoration. Most existing literature on biocrust restoration originates in other countries, and most have only a laboratory component (Pendleton et al. 2004; Malam Issa et al. 2007; Prabu and Udayasoorian 2007; Maqubela et al. 2009; Zheng et al. 2011). One laboratory study of cyanobacterial inoculation of poorly aggregated soil resulted in the growth of a dense superficial network of cyanobacterial filaments and

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extracellular polysaccharides after 6 weeks (Malam Issa et al. 2007). Further testing revealed that inoculated soil was resistant to soil aggregate breakdown, demonstrating gain of some biocrust function in a short period of time (Malam Issa et al. 2007). There was also an increase of soil N up to 40% and an increase of soil C (Malam Issa et al. 2007). Other studies have shown that inoculation with cyanobacteria Microcoleus vaginatus can speed soil crust recovery and increase species diversity (Buttars et al. 1998; Belnap and Eldridge 2001). After trampling soil disturbance, Buttars et al. (1998) used pelletized cyanobacteria as an inoculate, which increased organic matter, nutrient content, and soil stability. Unfortunately, this success was not seen in further field experiments (Buttars et al. 1998). Subsequent research using live inocula showed greater success (Pendleton et al. 2004; Prabu and Udayasoorian 2007; Maqubela et al. 2009; Zheng et al. 2011). Laboratory research by Pendleton et al. (2004)using live cultures of Microcoleus and Nostoc in pots of soil resulted in increased plant survival at low soil fertility. Another laboratory study in India using the native cyanobacteria genus, Westiellopsis, found a five-fold increase in numbers of cyanobacteria after 90 days (Prabu et al. 2007). Another study in Africa using a species of cyanobacteria, Nostoc, for inoculation in hoop houses found increases in soil nitrogen (17- 40%), carbon, soil aggregation, and Maize crop yield (Maqubela et al. 2009). Another six-month experiment used a slurry of cyanobacteria and amendments in a growth chamber and resulted in visible aggregate and improved rates of C and N fixation (Zheng et al. 2011). One study, (Cole et al. 2010) looked at the success rate of transplanting crusts as an alternative to inoculation. The experiment had regrowth success, but it necessitated destruction of donor biocrust sites, so is not a viable sustainable method. One of the only studies using fresh inoculum in situ was a recent study in China using two strains of fresh cyanobacterial inoculum on degraded aridland (Wang et al. 2009). After three years algal cover had increased 48.5%, with an additional 14 cyanobacterial species, and new moss species establishing on the biocrust (Wang et al. 2009). Additionally, biocrust thickness, compressional strength, organic C, total N, and chlorophyll a content had all increased (Wang et al. 2008). Considering the past success of cyanobacterial inoculation, especially the success of fresh inoculum in the laboratory, it makes sense to try live inoculation of cyanobacteria on coastal sage scrub soils of Southern California. Success of experimental inoculation using a non-destructive method could be a first step in developing a new restoration technique.

CoaStal Sage SCruB—imperileD haBitat

Coastal sage scrub (CSS) is an imperiled valuable habitat unique to coastal and inland California (O’Leary 1989; Davis et al. 1994; Figure 1). CSS in San Diego is home to an

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unmatched and exceedingly rich population of flora and fauna (Dobson et al. 1997). In fact, San Diego County is the most biodiverse county in the continental United States (Dobson et al. 1997). There are more species of living things here than anywhere else in the country (Rebman and Simpson 2006). Nearly 2,000 individual species of plants are endemic to the county, and of those, plants occurring near the coast of San Diego in coastal sage scrub locations are unique to just a small geographic area (Rebman and Simpson 2006;

Barbour et al. 2007). CSS supports many species; over 200 of the animal and plant species have low numbers that warrant protection, including the cactus wren (Campylorhynchus brunneicapillus), California gnatcatcher (Polioptila californica), and San Diego desert woodrat (Neotoma lepida intermedia) (Schoenherr 1992). These plants and animals only exist here, and have evolved to live in the specific microclimates of Southern California and Northern Baja California (Schoenherr 1992). This geographic region is listed in the Top Ten Biodiversity Hot Spots in the world (Dobson et al.1997). Protecting and restoring the biodiversity of the biocrusts makes good economic sense because these actions will protect the valuable ecosystem functions provided by intact crusts (Garcia-Pichel 2003). Ecosystem function, arable land, and biodiversity are all at risk when biological soil crusts are lost. Biocrust restoration and reestablishment is an essential step in regaining the health of degraded terrestrial environments (Bowker 2007). Measures must be taken to preserve the species richness of this precious and unique endemic community in San Diego. Development is not the only force threatening CSS and biocrust communities; invasive plants cost the state of California over 82 million dollars a year in control measures alone (Pimental et al. 2000). Invasive plants are dangerous because they change the nutrient balance (Belnap et al. 2001; Evans et al. 2001; Eviner et al. 2010) and mycorrhizal

Figure 1. Coastal sage scrub locations. Credit: derivedfrom Terrestrial Vegetation of California by Michael G. Barbour et al. 2007

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composition of the soil (Hawkes et al. 2005). Invasive plants also increase fuel load and fire frequency (Lovitch and Bainbridge 1999, Talluto and Suding 2008). The introduction of invasive exotic plants initiates a negative feedback loop that increases subsequent invasion (Yelenik et al. 2004; Eviner et al. 2010). These changes in the ecosystem create conditions unsuitable for endemic plants. Additionally, increased fire frequency puts human settlement at risk. In San Diego County this is certainly an issue. Studies have shown that intact biocrust hinders establishment of invasive plants by inhibiting germination and root penetration (Deines et al. 2007; Hernandez and Sandquist. 2011). Several studies, worldwide, show that intact biocrust inhibits the germination and establishment of exotic grasses (Belnap and Lange 2003; Hernandez and Sandquist. 2011). Germination of native grasses was not deterred by intact biocrust (Belnap and Lange 2003). Additionally, loss of intact crust decreases available plant nutrients and favors establishment of invasive exotics that can survive in impoverished soils (Belnap and Lange 2003). Since natural recovery is typically slow, if it occurs, assisted recovery could propel the system forward benefiting the ecology of the biocrust, and ultimately, of the whole ecosystem (Bowker 2007). Restoring biocrust could slow exotic grass establishment and decrease the heightened fire risk associated with the increased fire load of invasion (Eviner et al. 2010). Techniques for biological soil crust reestablishment are lacking and there is a need for them, now that we know how valuable they are for ecosystem function, to develop techniques to heal damage and to encourage biocrust growth (Bowker et al. 2007).

BioCruSt reStoration of CoaStal Sage SCruB Studies of biological soil crusts of CSS habitat are few; studies involving restoration of CSS biocrust do not exist at all. Given that biocrust organisms vary with environmen-tal factors and across ecosystems, results of CSS biocrust restoration research could differ from the restoration research carried out on other parts of the globe (Malam Issa et al. 2007; Maqubela et al. 2009; Prabu and Udayasoorian 2007; Wang et al. 2008; Zheng et al. 2011). Some CSS types, especially in semi-arid Southern California, do not receive sufficient pre-cipitation to support further appreciable colonization by lichens and mosses, and remain in a cyanobacterially dominant state (Belnap and Lange 2003; Hernandez and Knudsen 2012). In Southern California, algal crusts dominated by cyanobacteria are cryptic, existing beneath and dispersed through gravel (Pietrasiak et al. 2011). It is important to note that the assemblage of biocrust organisms is not static and can change incrementally or substantially, given the local soils and conditions, revealing a rich mosaic of microhabitats or environmental niches with biodiversity potential (Belnap and Lange 2003; Rosentreter et al. 2007; Bowker et al. 2010b.).

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This community-created variation influences species distribution and, to the amount it is maximized, reflects the health of the system (Belnap et al. 2001; Castillo-Monroy et al. 2011). Because develop-ment and urbanization have reduced coastal sage scrub habitat to 10-15% of the original land area (Figure 1), it has become imperative that we restore damaged areas or we will lose them forever (Lo-vitch and Bainbridge 1999). Given the value of this rapidly dwindling valuable habitat, further study of this habitat, including ways to preserve it, seems timely and appropriate (Westman 1981). Developing new techniques to restore the fragile and important biocrust in sage scrub could assist in reducing fire risk and exotic grass invasion while preserving soil structure and arresting erosion. Globally, restoration techniques could help preserve arable land and increase carbon and nitrogen sinks (Domingo et al. 2011). This study examines the feasibility of isolating and culturing an endemic mixture of CSS cyanobacteria and green algae to inoculate native CSS soil, thereby facilitating recovery of disturbed biological soil crusts. Given the importance of biocrust and the length of time for recovery, land managers should attempt to restore these ecosystems (Eldridge et al. 2010; Bowker et al. 2011). Additionally, invasive plants also threaten to degrade the CSS ecology (Westman 1981). A stumbling block exists, as not much is known about California biological crusts and even less about crusts in CSS habitats (Belnap et al. 2001; Hernandez and Sandquist 2011). Analyzing the health and components of the existing biocrust is overlooked in most habitat restoration (Bowker et al. 2008). If biocrusts are not included in the restoration, degraded areas of coastal sage scrub are only partially rehabilitated by eradicating invasives and replacing them with native vascular plants (Bowker et al. 2008; Godefroid et al. 2011). Indeed, even the traditional process of habitat restoration, as currently practiced, tends to ignore the soil crust organisms, and instead, concentrates on killing invasive plants with herbicide formulations and planting vascular plants (Godefroid et al. 2011). Although not much research has been done on the negative affects of pesticides and herbicides on biocrust organisms, what has been done finds that, minimally, they kill soil bacteria, and decrease the amount of nitrogen fixation by heterocystic cyanobacteria (Bhunia et al. 1991; Kremer and Means 2009; Megharaj et al. 1989). The only required tests for pesticide toxicity are run on what are considered the active ingredients only (Cox and Surgan 2006). Inert ingredients are not required to be listed, let alone tested, so knowledge gaps exist on how negatively the complete formulations affect the soil community (Cox and Surgan 2006). Studies are finding the inert ingredients often increase the toxicity of the pesticides (Cox and Surgan 2006; Mesnage et al. 2013). For example, recent work on the additive, POE-15 (polyethoxylated tallowamine), a common additive to the active ingredient glyphosate herbicide, finds POE-15 causes negative cell effects at 1 and 3 ppm making the POE-15 the more toxic ingredient (Mesnage et al. 2013). When inert and active ingredients are combined in formulations, the synergistic toxicology is greater then the ingredients alone (Cox and Surgan 2006). Clearly, restoration involving the use of pesticides retards the progress

of a restoration, and their use may be reconsidered if the health of biocrust organisms were included in the plans. We are in danger of losing this valuable habitat unless restoration begins to include the health of the whole system, including status of the biocrust. In order to preserve a resource, one must have knowledge of that resource. Since so little is known about biocrust in CSS habitats, it makes an ideal study subject. Because cyanobacteria do so much to improve soil conditions for other plants (Xu et al. 2012), successful inoculation with native strains could be used to restore ecosystem function in degraded areas. This study will increase our general knowledge on the composition of biological soil crusts in coastal California. Part of the study will use pioneer genera of cyanobacteria and green algae from CSS. Isolates will be grown and used to inoculate damaged crusts. Additionally, ground native biocrust will be used as a comparison because using the whole biocrust profile may increase success (Irvine et al. 2013). Autoclaving some treatments will provide a proxy for pesticide use by killing soil organisms. A comparison will be made of autoclaved treatments and nonautoclaved treatments. Looking more closely at biocrust locally and at the role of cyanobacteria in establishment of new biocrust will undoubtably reveal some new findings that will assist in CSS preservation. For example, mycorrhizae, little known a few years ago, are now commonly used as an inoculate for growing native plants because it was found that all but a few natives benefit from mycorrhizal association (Lovitch and Bainbridge 1999). Until recently, we did not know of the importance of mycorrhizae, and how necessary they are for better habitat restoration and now they have become integral for some native plant growers. Perhaps, in the future, inoculating with a native culture of cyanobacteria, or a mixed culture with green algae will be an integral part of habitat restoration.

reSearCh QueStionS

There are three research questions for this study. 1) Can endemic biocrust organisms be isolated and cultured? 2) Will an inoculum of native cyanobacteria and green algae grow on CSS soil? 3) Will inoculum growth increase biological function as measured by EPS, stability, and chlorophyll a in CSS soil?

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MATERIALS AND METHODS

StuDy SiteS

The SDSU Santa Margarita Ecological Reserve (SMER) is located in north San Diego County near the city of Temecula on 1,790 ha of coastal sage scrub countryside at 33°26’ N, 117°9’ W (Figure 2). The reserve has a substantial amount of undisturbed and disturbed CSS along the Santa Margarita River at an elevation of approximately 393 m. This area has a Mediterranean climate with average annual precipitation of 280 mm (Zink et al. 1995). Approximately 93% of the average annual precipitation occurs from October through April. Marine fog supplies a portion of the moisture. Temperatures are moderated by the Pacific Ocean 29 km west of the site. The site has a mild average mean temperature

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Figure 2. Coastal sage scrub location study sites. Credit: derived from Google Maps

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of 16.4 ° C. Observed area has not burned in the last 50 years. The reserve is representative of disturbed Diegan coastal sage scrub habitat (Holland 1986; Oberbauer 1996). The soil is Las Posas rocky loam derived from igneous rock parent material and is classified as a well-drained Alfisol (NRCS Web Soil Survey). Dominant plant species on the site are Artemisia californica (typical mean cover of 46%), Salvia mellifera, Malosma laurina, Acmispon glabra, Salvia apiana, and invasive non-native forbs Bromus hordeaceus sp. hordeaceus, Bromus madritensis, and Brassic nigra (NRCS Web Soil Survey). Typical height for CSS shrubs is 1-1.5 m. There is a mix of drought deciduous shrubs and evergreen shrubs with lesser amounts of herbaceous perennials and annual plants. Coastal Sage Scrub vegetation type is also called “soft chaparral” (O’Leary 1989).

Biological soil crust in coastal sage scrub can be cryptic when dormant (Figure 3), and recognizing its appearance when not actively growing is difficult (Belnap and Lange 2003; Rosentreter et al. 2007). Mission Trails Regional Park is located 13 km northeast of San Diego (Figure 2) and has 2,340 ha of dedicated open space at 32°48’N 117°01’W. The site was used by the military for more than 40 years, and as a park since 1960. This large park has 81 km of hiking trails all through CSS, grassland, and chaparral habitats. San Diego River bisects the park. Topography is rolling with elevations ranging from sea level to maximum peak of 485 meters. Biological soil crust was sampled near the Visitor Center in disturbed Diegan coastal sage scrub as evidenced by non-native exotic plant species (Holland 1986; Oberbauer 1996). The vegetation is a mix of aromatic, soft-leaved, drought deciduous subshrubs and sclerophyllous evergreen shrubs with some perennials and annuals (O’Leary 1989). Areas above 333 meters

Figure 3. Coastal Sage Scrub Biological Soil Crust. A. Lichenized crust at SMER B. Visible cyanobacterial crust from Mission Trails Credit: Sharon Reeve

A B

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Table 1. Growth Media Ingredients BG-11. are classified as Inland Diegan coastal sage scrub (Oberbauer 1996). The area sampled has well-drained Friant rocky fine sandy loam soils from weathered metasedimentary rock parent material (NRCS Web Soil Survey).

fielD Sampling

Several approximately 490 mm2

roughly circular pieces of actively growing biological soil crust were collected from Santa Margarita Ecological Reserve and from Mission Trails Regional Park. A variety of biocrust types were collected—those dominated by bryophytes as well as those dominated by cyanobacteria—so a good cross-section of age and development was obtained for CSS habitat. Also, biocrusts were collected from the gaps

between plants and did not contain any observable plant material. Biocrusts were stored under dry conditions at room temperature (20-25° C) in covered petri dishes (Figure 4).

Biocrusts were air-dried for a week, and ground with a sterile mortar and pestle and used for a dilution series in an attempt to isolate individual cyanobacterial species. All the biocrusts were crushed and mixed to produce a homogeneous sample, one sample for each location.

growth meDium

The growth media BG-11 (Table 1) was developed to optimize the growth of cyanobacteria and is a standard support for use in culturing cyanobacteria in the laboratory (Stanier et al. 1971). Nitrogen

Figure 4. Collected biocrust.Credit: Sharon Reeve

Source: Stanier et al. 1971

NaNO3*K2HPO4

MgSO4 7H2OCaCl2 2H2OCitric Acid H2OFerric Ammonium Citrate*Na2EDTA H2ONa2CO3

Trace Metal MixDistilled WaterAgar

*KNO3 substituted for NaNO3*Ferric Citrate substituted for Ammonium Citrate*pH should be 7.1

..

.

.

1.5 g0.04 g0.075 g0.036 g0.006 g0.006 g0.001 g0.02 g1.0 ml1.0 L10.0 g

Growth Media Weight/Volume

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was included in case non-heterocystic cyanobacteria was isolated. One liter of sterile BG-11 growth medium was prepared in the laboratory with slight modifications (Table 1) (Stanier et al. 1971).

iSolating StrainS of CyanoBaCteria anD green algae

A limiting dilution series for each sample was prepared in each of two 24-well plates. The biocrust samples from the two locations were each diluted with 1 ml of distilled water, and then thoroughly swirled and mixed. A separate plate was used for the biocrust from Mission Trails and one for the biocrust from Santa Margarita. The 24-well plate contained 1.8 mL of BG-11 in each well. The inoculum (0.2 mL) was pipetted in each well of the first column, mixed and serially diluted through the remaining columns (Rippka 1988; Lee et al. 2014). Plates were sealed with parafilm, incubated at the constant temperature of 30˚C, and kept under a compact fluorescent light with the irradiance of 20.3μmol m-2 s-1. After a month, sufficient growth had occurred to further isolate the cyanobacteria or green algae on agar and BG-11 plates. For this phase of the experiment, BG-11 media was prepared as before, except agar was added. Sterile petri dishes (Figure 4) were filled halfway with the mixture and streaked with culture. Each petri dish was fitted with a lid and sealed in parafilm. All 20 of the plates were set in the incubator at 30˚C (Mazor et al. 1996) and kept under a compact fluorescent light, operating 24 hours a day, with the radiance of 20.3μmol m-2 s-1. Growth was observed and samples of the colonies were observed under the microscope after a couple of weeks. After the initial experimentation, fresh samples of biological soil crust were collected from Santa Margarita Ecological Reserve and from Mission Trails Regional Park and isolation and culturing procedure was repeated. In preparation to scale up the experiment and to maximize growth, streaked petri dishes of BG-11 and agar were exposed to different light levels in the 30°C incubator. The lights operated 24 hours a day. Mesh cloth was used to alter the light irradiance levels to four levels: low (4.5μmol m-2 s-1) (Mazor et al. 1996), low/med (6.6μmol m-2 s-1), med (μmol m-2 s-1), and high (20.3 μmol m-2 s-1). For comparison, some dishes were also grown in the window at a noon reading of 388 μmol m-2 s-1. After initial experimentation with light levels, the cultures were moved to the high light setting (20.3 μmol m-2 s-1) in the incubator. To determine if adding a solid matrix would increase growth relative to liquid culture, visible isolated colonies were transferred with a sterile loop into four 150 mm petri dishes for each site. Isolated colonies that demonstrated a high growth rate were used preferentially. Two of the five petri dishes contained four rounded spoons of autoclaved sand, and 60 ml of BG-11 in each, two more contained 90 ml of BG-11 and no sand, and the fifth was an uninoculated control with just BG-11. The set of 10 large prepared petri dishes

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were placed in the large incubator with four warm white fluorescent tubes in the four corners of the unit. The irradiance was measured at 17.6 μmol m-2 s-1 and the temperature was set at 30˚C with continuous light. While no specific measurements were taken, observationally, all treatments grew well, and with or without sand. After two months steps were taken to scale up the amount of inoculum.

Chlorophyll a meaSurementS

To scale up quantities of cyanobacteria and green algae to use for experimental inoculation, each culture from the four liquid petri dishes was added to one of four sterilized Erlenmeyer flasks with 250 ml of BG-11 and grown in the incubator with continuous light irradiance of 17.6 μmol m-2 s-1 and temperature of 30˚C. There were three flasks with isolates from Mission Trails, MT A, MT A-2, and MT B (Figure 5). There was one flask with isolate from Santa Margarita, SMER A (Figure 5). The cultures grew to high density in less than a month. For analysis of the chlorophyll a in the flasks, flask contents were blended thoroughly and 1 ml of contents was centrifuged, the pellet extracted and added to 1 ml DMSO in sealed test tubes (Castle et al. 2011). DMSO was used as a solvent for extracting chlorophyll a

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Figure 5. Flasks of inoculant from left to right: MT A , MT A -2, MT B & SMER A.Credit: Sharon Reeve

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because it was found to be the most effective for extracting chlorophyll a by Pompelli et al. 2012 (Castle et al. 2011; Pompelli et al. 2012). Agitation was found to increase the efficiency of chlorophyll a extraction, so all the samples were vortexed twice for 20 minutes before incubation at 65°C for 45 minutes, and again after incubation (Castle et al. 2011). Tubes were centrifuged for 5 minutes at high speed and 200μl of supernatant from each tube was extracted and pipetted into a 96-well standard microplate with DMSO blanks. Measurements for chlorophyll a were taken in the spectrophotometer reading absorbance at 665 nm., Spectra MAX 190 by Molecular Devices used in conjunction with the software SOFTmax PRO 4.0 Life Sciences Edition by Molecular Devices Corp. To determine chlorophyll a, the equation from Ritchie 2008 was used (11.4062 x (6650 nm) x V)/(g soil-1) x L (where V is the extraction volume in ml, L is the pathlength in cm, units are ug/g Ritchie 2008; Castle et al. 2011).

referenCe meaSurementS for CharaCteriStiCS anD Chlorophyll a Reference measurements were taken using six freshly collected biological soil crust samples from Mission Trails (Figure 7) that contained no observable moss or lichens. Reference statistics of depth, weight, bulk density, chlorophyll a, DOC, and stability for local CSS crust were measured and recorded. The average depth of the soil crust and the average density of 1 cm square of biocrust was measured for use as guidelines and reference in the inoculation experiment. Crust thickness was gauged by slicing with a razor blade and observing where the soil fell away from the aggregated biocrust. Reference chlorophyll a was determined using DMSO extraction (Castle et al. 2011; Pompelli et al. 2012). After collection of the crusts, they were placed in petri dishes and air dried for several days (Mazor et al. 1996). A sterilized mortar and pestle were used to grind them into a fine dust and 3 ml DMSO was added to 1.5 gm of ground crust in sealed test tubes, and chlorophyll a was then extracted and analyzed in the spectrophotometer as before (Pompelli et al. 2012; Castle et al. 2011; Ritchie 2008). Crust morphology was observed under a light microscope.

referenCe meaSurementS for extraCtaBle polySaCChariDeS

The weak acid extraction method for extractable polysaccharides from Redmile-Gordon et al. 2014 was modified and used to yield EPS from air dried biocrust (Redmile-Gordon et al. 2014). A mixture of 25 ml of 0.5M H2SO4 and 0.5 g of biocrust was autoclaved for an hour, the extract was centrifuged for 20 minutes and the supernatant was extracted (Redmile-Gordon et al. 2014) To indirectly measure polysaccharides from cyanobacterial growth, dissolved organic carbon was measured using a Mn(III) reduction assay (Barlett and Ross 1988). After a 24-hour dark reaction time, the results were compared to a set of incremental glucose standards

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(ranging from 0-700 μM glucose) in the spectrophotometer set at A490nm, Spectra MAX 190 by Molecular Devices used in conjunction with the software SOFTmax PRO 4.0 Life Sciences Edition by Molecular Devices Corp.

referenCe meaSurementS for Soil StaBility

Soil stability was measured (Figure 6) using an adaptation of standardized field method developed by Herrick et al. 2001. A Stability Class Table was constructed using

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Table 2. Stability Class Table: Criteria for the Assignment of Crust to Stability Classes Source Herrick et al. 2001

0 Soil too unstable to sample (falls through sieve) 1 50% of structural integrity lost within 5s of insertion 2 50% of structural integrity lost 5-30s after insertion 3 50% of structural integrity lost 30-300s after insertion OR <10%ofsoilremainsonsieveafterfivedippingcycles 4 10-25%ofsoilremainsonsieveafterfivedippingcycles 5 25-75%ofsoilremainsonsieveafterfivedippingcycles 6 75-100%ofsoilremainsonsieveafterfivedippingcycles

StabilityClass1-6

Criteria for Assignment to Stability ClassTime Dipped into Water

Figure 6. Soil sieve apparatus. Figure 7. Reference CSS biocrusts.Credit: Sharon Reeve

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data and criteria from past soil stability studies (Tongway 1994). Values in the table gave consistent and verifiable results, and were comparable to laboratory-based testing (Herrick et al. 2001). Biological soil crust fragments approximately 6-8 mm in diameter were used as they produced truer results than larger or smaller fragments (Herrick et al. 2001). A five-compartment sieve was constructed to allow testing of multiple fragments at once (Figures 6 and 7). All fragments were uniformly air dried before testing. Five biological soil crust cubes were placed in the sieve and dipped according to the chart and results were recorded.

Dna extraCtion, purifiCation, anD Cloning DNA analysis using polymerase chain reaction (PCR) was performed on DNA that was extracted from the flasks of isolated material using the PowerMax Soil® DNA Isolation kit and standard kit protocol (MoBio Laboratories Inc. Carlsbad, CA USA). Cyanobacterial 16S rRNA genes were amplified by PCR using the universal bacterial primers, F8 (50-AGAGTTTGATCCTGGCTCAG) and R1111 (5′-TTGCGCTCGTTGCGGGACT-3′) (Nubel et al. 1997; Ouellette et al. 2006; Lipson et al. 2009). The PCR mixture for each sample contained one unit taq polymerase (Fisher BioReagents™ Taq DNA Polymerase), “Buffer A” (Fisher Biosciences), 3.0 mM MgCl2, 1.25 μM of each primer, 200 μM of nucleotide triphosphate, and 20 mg 1-1 bovine serum albumin. After an initial 2 minute denaturation step at 94°C, the PCR reaction was continued by 32 cycles of 94°C for 1 minute, 56°C for 1 minute, and 72°C for 1 minute, and finally, an extension phase at 72°C for 10 minutes. Sequencing of PCR products was done by Eton Biosciences. The resulting nucleotide sequences were submitted to NCBI Basic Local Alignment Search Tool (BLAST) for analysis of homologous sequences in nucleotide and protein databases. Sequences were compared to the BLAST clone library (McGinnis and Madden 2004) for the most closely aligned sequence matches and a phylogenetic tree was constructed using the FastDNAml function of BioEdit (Hall 1999).

inoCulation experiment

A randomized complete block design was used for the inoculation experiment. A large rack of 50 small identical pots with a 5 cm x 5 cm surface area was used. There were 10 replicates of 5 treatments: 1) control (C) 2) autoclaved control (AC) 3) ground crust inoculation (GI) 4) autoclaved soil with live inoculum (AI), and 5) soil with live inoculum (I). To prepare for the inoculation experiment, native soil from SMER had been stockpiled from previous work and salvaged for use in this project. A bucket of 5 kg of soil, was mixed well and homogenized. A portion, 2 kg of the soil was autoclaved for 40 minutes at 120°C in a covered stainless steel container. Sterilizing the soil was a surrogate for pesticide application.

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For the experiment, the pots were washed in detergent, and soaked in 1% bleach to clean, and then rinsed in distilled water. Seventy grams of sieved soil were added to each pot. To 30 pots, 70 g of native soil were added, and to 20 pots, 70 g of sterile soil were added. The soil was tamped down to make all the surfaces flat and uniform (Figure 8). The first 10 pots were controls, 10 pots were autoclaved controls, 10 pots had a ground crust inoculation, 10 pots contained autoclaved soil and were treated with liquid inoculum, and the last 10 pots were untreated native soil with liquid inoculum. Two of the five treatments involved BG-11 mixed with inoculum, so to equalize the experiment, the other three treatments received an equal volume of BG-11. The third set of pots was treated with ground up crust. The weight of ground up crust applied to each was figured using the average of the chlorophyll a values for the reference crusts by surface area and applying a tenth of the reference amount of chlorophyll a for the proportional surface area of the pot. In other words, all GI pots started with 0.1 the amount of chlorophyll a, as a mature coastal sage scrub biological soil crust to allow for potential growth. A cm3 of reference CSS biocrust weighed 0.7 gm. In both the liquid inoculation and the ground up crust treatments the amount of chlorophyll a in the reference crusts was used as a guide to the application rates. Like the ground crust treatment, the liquid inoculum was prepared using the reference crust measurements for chlorophyll a and applying 0.1 of that amount to the same size surface area of the pot. The four flasks of cultured cyanobacteria and green algae were measured for chlorophyll a and a ratio of the contents was prepared based on this measurement, so all strains were equally represented. All 50 pots were labeled and randomly distributed on a rack placed in the incubator set at 30°C (Mazor et al. 1996) and kept under compact fluorescent lights with a constant irradiance of 20.3 μmol m-2 s-1. The pots were covered in plastic wrap and watered evenly every week with 15 ml of distilled water per pot and the order rerandomized. The inoculation experiment was started on 27 June 2014 (Figure 8) and ended on 22 August 2014 (Figure 9).

final analySiS of inoCulation experiment At the conclusion of the experiment all 50 pots were transferred to the lab where they were air-dried for 10 days. The contents of each pot was removed as carefully as possible, and the top 1 cm was sliced off of each sample and divided into quarters for analysis. Four of the sliced quarters were tested, one quarter of the sample was analyzed for extractable polysaccharides, another quarter was analyzed for chlorophyll a, another quarter was tested for stability, and the fourth quarter was observed under the microscope. Extractable polysaccharide values for the pots were analyzed using the weak acid extraction method described previously for the reference crust (Redmile-Gordon et al. 2014) and read in the spectrophotometer. Chlorophyll a values were measured using the previously described

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DMSO extraction method (Ritchie 2008; Castle et al. 2011) and the spectrophotometer. Soil aggregate stability was tested using a soil sieve with samples of experimental pot crusts (Figure 6.) and the scale developed by Herrick et al. (2001) as previously described in baseline measurements in (Table 2). At the conclusion of the experiment photographs of all of the pots were taken against a white background (Figures 9A-H).

Fig. 8. Before inoculation.

Fig. 9C. Ground inoculated.

Fig. 9F. Auto. inoculated.

Fig. 9A. control.

Fig. 9D. Ground inoculated.

Fig. 9G. Inoculated.

Fig. 9B. Autoclaved control.

Fig. 9E. Auto. inoculated.

Fig. 9H. Inoculated.

Figure 9. Photographic results of inoculation experiment Credit: All photos S. Reeve

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experimental DeSign anD StatiStiCal analySiS

The inoculation experiment had 5 treatments. There was a control and an autoclaved control, inoculation with ground crust on native soil, and liquid culture inoculation on native soil and on autoclaved soil. This was a balanced experiment with 10 replicates per treatment and a single factor ANOVA with five levels. An additional single factor ANOVA was performed comparing all inoculated treatments versus non-inoculated treatments for extractable polysaccharides. Two-way analysis of variance, removing the ground crust treatment, and comparing the controls and the liquid inoculated treatments was carried out for both chlorophyll a and stability. The soil stability test (Table 2), gave quantifiable nonparametric results (Table 2 and Figure 13) (Herrick et al. 2001). The rating scale used in this test was a ranking not based on any strict value or strict numerical interpretation—the data resulting were ordinal data and analyzed using a non-parametric Kruskal-Wallis test for significance.

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RESULTSCulturing experimentS

After the first series of isolation of cyanobacterial and algal cultures using dilution series, streaking, microscopy and keying confirmed that all of the cyanobacteria observed belonged to one species, what we initially identified morphologically as Leptolyngbya foveolarum (Gomont) Anagnostidis & Komárek 1988 (Figure 10) (Stancheva et al. 2014; Komárek and Anagnostidis 2005). During the duplication of the isolation steps it was observed from the growth that the cyanobacteria cultures grew better in the high light of the incubator (20.3 μmol m-2 s-1) and were photo inhibited in direct sunlight. In the environment, filamentous cyanobacteria move in

Figure 10. Single strand cyanobacteria from isolation resembling Leptolyngbya foveolarum (Gomont) Anagnostidis & Komárek 1988 Credit: Sharon Reeve

response to levels of intense solar radiation by moving deeper into the substrate (Belnap et al. 2001). The petri dishes had no substrate for the cyanobacterial filaments to move behind to alter the intensity of the solar radiation. Further experimentation using petri dishes with sand and petri dishes without sand found (based on visual observation) that the culture of cyanobacteria grew well with or without substrate. The goal for the next phase was to maximize growing conditions to grow larger amounts of cyanobacteria for use in inoculation. Liquid BG-11 was chosen for ease of use in large 500 ml Erlenmeyer flasks. After the initial set up of the four flasks of the isolated cyanobacteria cultures that grew best, the cultures grew quickly. Of the four cultures that grew best, three were from Mission Trails, and one was from Santa Margarita. Levels of growth in chlorophyll a, extractable polysaccharides, and stability were used as surrogates for measuring increases of biological function. Chlorophyll a levels indicated growth of cyanobacterial filaments and green algae populations. Extractable

20 µm

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polysaccharides indicated the presence of extracellular polysaccharides, a component of filaments of cyanobacteria and a structural component of green algae. The measure of stability indicated how successful the growth of cyanobacterial filaments had knit together soil particles. A set of reference biological soil crusts from Mission Trails and Santa Margarita were measured (Table 3) for these parameters and compared with the growth of the 50 pots in the inoculation experiment. The four flasks of cyanobacteria and green algae in BG-11 were also measured for chlorophyll a and proportionate amounts were mixed for the cyanobacterial inoculum. SMER A had the most chlorophyll a and was the only mixed culture of cyanobacteria and green algae. Notice the different hue of lighter green on the ground crust inoculated pots in Figures 9C and 9D. The pots inoculated with the liquid culture of cyanobacteria and algae are darker green in Figures 9E-H.

Chlorophyll a reSultS

Comparisons of the data were done with one-way analysis of variance (Table 4). There was a difference between most treatments in the amount of chlorophyll a that the pots contained (Figure 11). A posthoc test, Tukey’s Honest Significant Difference (HSD), was performed to point to the specific groups that were significantly different. The test revealed that all of the inoculation treatments were significantly higher in chlorophyll a than the controls (Figure 11). Both of the liquid culture inoculated treatments were significantly higher in chlorophyll a than the ground crust inoculated treatment. A two-way ANOVA (Table 5) that removed the ground crust treatment found that the autoclaved treatments had significantly

Table 3. Baseline Values for Reference Biological Soil Crusts

Reference Crusts

Mean Value Standard Error +/-

Chlorophyll a (ug/g soil)

Extractable Polysaccharides(ug C/g)

Aggregate Stability

2.59

696.0

0.167

15.75

4397.0

5.833

27

Figure 11. Chlorophyll aconcentrationsoffivetreatments.(Columns with non-repetitive letters are significantly different. One-way ANOVA , n = 50, α = 0.05, means and standard errors of 10 replicates).

0  1  2  3  4  5  6  

Control   Sterilized  Control  

Ground  Crust   Autoclaved  Treatment  

Liquid  Inoculum  

chloroph

yll  a    

(ug/Ch

la/g)  

Inocula-on  Treatments  

Chlorophyll  a  per  Treatment  

         a                  a,b          b  

     c      c  

Control AutoclavedInoculated

InoculatedGroundCrust

Inoculated

Autoclaved Control

(μg

Chl

a/ g

soi

l)

Chlorophyll a per Treatment

Inoculation Treatments

chlo

roph

yll a

Variable

log (Chla)StabilityEPSEPS

R2

0.7330.5790.1290.125

df (model)

4441

df (error)

45454548

F

30.915.441

1.6736.858

P

<0.001<0.001

0.1730.012

Table 4. Statistical Values for One-way Analysis of Variance

less chlorophyll a than non-sterilized soil (n = 40, α = 0.05, p = 0.031). The two-way ANOVA (Table 5) also found that the chlorophyll a levels of liquid inoculated treatments were significantly greater than non-inoculated treatments (n = 40, α = 0.05, p < 0.001).

27 28

extraCtaBle polySaCChariDeS reSultS

The initial one-way ANOVA indicated no statistical differences in the levels of EPS between the treatments (n = 50, α = 0.05) (Table 4 and Figure 12). Further exploration with a two-way ANOVA that compared all inoculated treatments with non-inoculated treatments, the inoculated treatments had significantly more EPS (Table 5) (n = 50, α = 0.05, p = 0.033).

0  

1000  

2000  

3000  

4000  

5000  

6000  

con   st  con   ground  crust   auto  inoc   inoc  

extractable  C  (ug  g-­‐1)  

Dissoved  Organic  Carbon  (ug  C/g  soil)  

Figure12.Extractablepolysaccharideconcentrationsinfivetreatments.(Columns with non-repetitive letters are significantly different. One-way ANOVA, n = 50, α = 0.05. Means and standard errors of 10 replicates).

Control AutoclavedInoculated

InoculatedGround CrustInoculated

Autoclaved Control

Extractable Polysaccharides(μ

g C

/ g s

oil) a a

a aa

29

Figure13.Stabilityindexofsoilsfromthefivetreatments(Columns with non-repetitive letters are significantly different. Kruskal-Wallis one-way ANOVA by ranks, a non-parametric test, n = 50, α = 0.05)

0  

0.5  

1  

1.5  

2  

2.5  

3  

3.5  

4  

C   SC   GC   AT   LI  

Stab

ility  In

dex  

Treatment  

Stability  

 a  

   a,b  

b,c  

     c,d  

d  

Control AutoclavedInoculated

InoculatedInoculated

Ground Autoclaved Control

Stability

StaBility teSt reSultS

The Kruskal-Wallis one-way ANOVA by ranks, non-parametric test showed significant differences among the treatments (Table 4 and Figure 13). Further testing with a Post Hoc test was performed using Tukey’s HSD to analyze individual contributions to significance. The inoculated treatments were significantly higher in stability than the controls. Inoculated soil was significantly more stable than the Autoclaved Inoculated soil (Figure 13). Analysis using a two-way ANOVA and removing the ground crust treatment showed that autoclaving significantly reduced stability (Table 5) (p = 0.001, n = 40, α = 0.05). Also, inoculation significantly improved stability of those treatments over non-inoculated treatments in a two-way ANOVA (Table 5) (p < 0.001, n = 40, α = 0.05). Although most of the experimental treatments gained stability and function, they still lagged behind the reference crusts that easily rated 6 with only one 5 rating (Table 3 and Figure 13).

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Correlation teSt reSultS

Further statistical analysis found that both chlorophyll a and extractable polysaccharides were significantly correlated with stability (Table 6). Correlation between chlorophyll a and stability was significant at (p = 0.0001) and significant between extractable polysaccharides and stability at (p = 0.0021). Statistically, this means that both autoclaved and native soil significantly increased in chlorophyll a and stability after the inoculation treatment.

Variable

Chla

Stability

EPS

Chla

—

0.0001

0.0581

Stability

0.573

—

0.0021

EPS

0.3021

0.471

—

Table 6. Correlations Between Treatments (The upper diagonal shows the Pearson correlation coefficients among variables and the lower diagonal shows the p-values for each correlation.)

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Variable

log (Chla)StabilityEPS

R2

0.7570.5930.123

df (mdl)

333

df (err)

363636

F

107.24939.493

4.897

F

5.05711.877

0.06

F

01.0270.102

P

<0.001<0.001

0.033

P

0.0310.0010.809

P

0.9890.3180.751

Table 5. Statistical Values for Two-way Analysis of Variance

Inoculated Autoclaved Interaction

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pCr teSt reSultS The particular CSS crust isolate cloning sequence placed in the rough phylogenetic tree was collected at Santa Margarita Ecological Reserve and called SMER A (Figure 14).Phylogenetic analysis placed the sequence in a group with an uncultured cyanobacterium from a Mongolian BSC (Kemmling et al. 2012) and an isolate from the Atacama desert (Patzelt et al. 2014), and this group appeared to be separate from

Figure 14. Maximum likelihood phylogenetic tree, based on partial 16S rRNA sequences, showing relationship of cyanobacterial isolate SMER-A to other cultured and uncultured cyanobacteria.

NR074309 Synechococcus elongatus

GQ167549 Nostoc calcicola

AF284803 Microcoleus vaginatus

FM210757 Leptolyngbya laminosa

KC463201 South African BSC

AY493574 Leptolyngbya frigida

KM020005 Pseudanabaena galeata

KM020005 Pseudanabaena catenata

FR798945 Leptolyngbya foveolarum

HF678483 Leptolyngbya boryana

FJ230796 hypolithic biofilm

JX255129 Mongolian BSC clone

SMER AKC311913 Atacama Desert isolate

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157Oculatella_VRUC135_X8-4809

158Oculatella_VRCU192_DO295208_1

159Oculatella_VRCU198_DO295207

160Oculatella_subterranea_AD501_Zam_HQ917688

161Oculatella_subterranea_SP301_Zam_HQ917689

162Oculatella_subterranea_SP1401_1Zam_HQ917690

163Oculatella_subterranea_SP1401_2Zam_HQ917691

164Oculatella_subterranea_SP1402_Zam_HQ917692

155Oculatella_GSE_PSE_49_070_2cons

175Oculatella_HA4348_LM1cons

176Oculatella_HA4348_LM3cons

173Oculatella_LLi18_DQ786166

174Oculatella_CR_3_EF545622

275Trichotorquatus_WJT9NPBG15_P16A

275Trichotorquatus_WJT54NPBG7_P43C

275Trichotorquatus_WJT54NPBG7_P43D

275Trichotorquatus_WJT55NPBG7_P44C

275Trichotorquatus_WJT40NPBG3_P38H

275Trichotorquatus_WJT36NPBG11_P31B

275Trichotorquatus_WJT19NPBG5_P19A

281Trichotorquatus_WJT66NPBG9_P49E

035Pseudoanabaena_HA_4215_MV1_DK_cons

340Neosynechococcus

259Trichotorquatus_ATA_2_1_CV25c

341San Diego_Leptolyngbyaceae_sp

242Trichotorquatus_TAA2_2HA1_27_2_HMO19680

271Trichotorquatus_WJT32NPBGA

301Trichotorquatus_F12_1_HA2_PF11_1_RT

156Oculatella_GSE_PSE_52_07L_5b

245Leptolyngbya_LSP_X8-4809

227Leptolyngbya_PCC7375_A8039011

Figure 15. Neighbor-joining phylogenetic tree showing placement of SMER-A cyanobacterial isolate in novel genus, Trichotorquatus (Patzelt et al. 2014, courtesy of Nicole Petrasiak).

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Figure 16. Green Algae Trebouxia sp. from Mission Trails isolation culture used in inoculations Credit: Sharon Reeve

20 μm

Leptolyngbya sequences in the database (Figure 14). The sequence from the SMER A isolate was phylogenetically closely related to a new genera recently described (Patzelt et al. 2014). To further explore the possibility that the SMER A isolate may fall within this new classification, the sequence was sent to one of the researchers for the Atacama Desert cyanobacterial sequences to see if it matched with the new genera of cyanobacteria they were describing (Patzelt et al. 2014). Awaiting further investigation, Dr. Nicole Petrasiak has placed the sequence in the genus Trichotorquatus (Figure 15). The green of the growth from the liquid inoculum (Figures 9E-H) was a darker green than the growth from the ground crust inoculation (Figures 9C-D). The liquid culture inoculated native soil revealed a mix of the green algae, possibly Trebouxia sp., and the Trichotorquatus sp. (Figure 17). In Figure 16, the green algae, appears possibly to be a Trebouxia species from MissionTrails. Observations under light microscope magnification of 20x revealed a different green algae in the ground crust treatment (Figure 18).

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Figure 17. Mix of Green Algae, Trebouxia sp. and Trichotorquatus sp. recovered from soil surface of inoculated, nonautoclaved (native) treatment Credit: Sharon Reeve

10 μm

34

Figure 18. Unknown Green Algae from Ground Crust Inoculation recovered from soil surface of ground crust inoculation treatment Credit: Sharon Reeve

.03 mm

35

DISCUSSIONLeptoLyngbya foveoLarum/ trichotorquatus

A research goal for this thesis was to determine if isolating and culturing local crust organisms was possible. Furthermore, if so, could CSS soil be inoculated with the culture and would there be an increase in soil function as a result? The first phase of experiment—collecting, isolating, and culturing— took place twice because the results were unexpected. I did not find the expected species of cyanobacteria. All existing literature pointed to the dominance of Microcoleus vaginatus in the majority of arid and semi-arid land systems. Community structure of biological soil crust is different depending on the properties of the site (Belnap and Lange 2003). Many crusts are cyanobacterially dominated in coastal sage scrub and in harsh environments like the Mojave Desert (Yeager et al. 2004; Wang et al. 2008; Pietrasiak et al. 2011; Pietrasiak et al. 2013). As expected, CSS habitat is certainly cyanobacterially dominated both in SMER and MT, but not by Microcoleus, at least not where we looked. To be clear, lack of Microcoleus vaginatus in specimens collected and in culture does not mean lack in soil. It could be that collection didn’t take place in the right area. We expected to see Microcoleus vaginatus because it is dominant in many similar ecosystems; and it is commonly studied, as this species is plentiful and morphologically easily identified under the light microscope because of the multiple strands of densely interwoven trichomes in one sheath (Leonard et al. 1995; Garcia-Pichel 2001; Rosentreter et al. 2007). Also, phylogenetically, M. vaginatus is recognizable in a PCR because of a distinctive 11-base pair insert within the 16S rRNA that makes it unmistakable (Dvorak et al. 2012). Microcoleus vaginatus has also been cultured for use as an experimental cyanobacterial inoculant (Wang et al. 2008; Zheng et al. 2011). Initially in this experiment, after reading copious research reports about how ubiquitous Microcoleus vaginatus is in many Western environments, it was expected to be in many of the slides of organisms from CSS crusts. It was surprising, therefore, that none of the slides had recognizable Microcoleus vaginatus. Conversely, many slides had another cyanobacteria, and it was commonly seen, even after repeating the isolation culture. Morphologically, it was keyed out as Leptolyngbya foveolarum (Figure 10), (Gomont) (Anagnostidis and Komárek 1988). After looking at rRNA evidence, this researcher will refer to this cyanobacteria as a species of Trichotorquatus sp. Recent 16S rRNA sequence data is more reliable than morphology, and the evidence suggests this is more correct for this species (Patzelt et al. 2014). There could be a number of reasons for not finding Microcoleus. It could be that conditions were provided in the isolation and culturing that allowed Trichotorquatus sp. to outcompete Microcoleus vaginatus. This is unlikely for two reasons: first, Microcoleus was

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never seen even in samples from fresh crust, and second, conditions for growth are similar for both species (Mazor et al. 1996; van der Grinten et al. 2005). Unlike other species of cyanobacteria, both Microcoleus vaginatus and Trichotorquatus sp. tolerate high light levels and higher pH soils. Many research papers have chosen M. vaginatus for ease of culture, so it would have been cultured if it were present (Wang et al. 2008; Zheng et al. 2010). The answer to this might be found by looking at other ecosystems where Trichotorquatus sp./Leptolyngbya foveolarum is found in abundance. The genus Trichotorquatus has only recently been described by Patzelt et al. (2014). Research was carried out using the conventional name of Leptolyngbya foveolarum. The two species in the different genera, Leptolyngbya and Trichotorquatus, are closely related and share similar simple, non-distinct morphologies, and share similar habitat preferences. Until future research sorts out the phylogenetic differences, it may be that not every article I referenced referring to Leptolyngbya foveolarum is actually Trichotorquatus sp. but they are analogous for the purposes of this research. Řeháková et al. (2011) found both species, Microcoleus vaginatus and Trichotorquatus sp. in abundance in the dry mountain region of Ladakh, NW Himalaya, but Trichotorquatus sp. thrived at higher elevations and in newer soils with less organic matter while Microcoleus thrived at lower elevations with higher organic matter (Řeháková et al. 2011). In Svalbard, an archipelago in the Arctic Ocean, Trichotorquatus sp. was found in abundance, and it was hypothesized that it was a robust generalist and colonizer of new environments (Stibal et al. 2006). Another look at this region found more Trichotorquatus sp. colonizing newly deglaciated barren soil with high pH and low organic matter; however, Microcoleus was also found there, but to a much lesser extent (Kastovska et al. 2005). Another study found Trichotorquatus sp. tolerated high pH and high light levels of up to 200 μmol m-2 s-1 (van der Grinten et al. 2005). Unlike previous reports, done in the 1960s, of Microcoleus prevalence in the Mojave Desert, a recent study found Trichotorquatus species were the most abundant cyanobacterial genera at Fort Irwin in the Mojave Desert (Alwathnani and Johansen 2011). Trichotorquatus sp. was considered common and found at all six sampling sites (Alwathnani and Johansen 2011). An earlier study found Trichotorquatus sp. at San Nicolas Island in the Channel Islands in a high pH sandy loam soil (Fletchtner et al. 2008). Microcoleus vaginatus was present at higher levels at a different site with a higher sand component and high pH, and also present at lower levels at all six sites (Fletchtner et al. 2008). A very recently published paper from research in the Atacama Desert suggests that because of 16S rRNA gene cloning/sequencing Trichotorquatus and Microcoleus species are in great need of phylogenetic restructuring (Patzelt et al. 2014). Members in the genus of Leptolyngbya are very small and difficult to differentiate under the

37

light microscope. Many species that resembled the small single stranded cyanobacteria were placed in this genus and are now just getting sorted out with DNA testing. Perhaps in the past, morphological interpretation underestimated species diversity, and also assumptions may have been made for similar, but unexamined ecosystems. With new phylogenetic methods and new environments being explored, changes will continue to take place in classifications. What we thought was Leptolyngbya foveolarum may be classified as a species of Trichotorquatus in the near future (Figures 10, 14, & 15). Interestingly, recent unpublished soil metagenomic analysis found in our lab’s preliminary work did not find any sequences for Microcoleus vaginatus from their sampling, in the same location where the Trichotorquatus sp. was isolated at Santa Margarita Ecological Reserve (Sherlynette Castro, unpublished data). Other cyanobacteria of related genera were found our lab’s preliminary work, but not Microcoleus vaginatus. So it could be that Microcoleus vaginatus, so common in many desert soils, is not as ubiquitous in coastal sage scrub as previously conjectured. It could be that coastal sage scrub is unique, and while somewhat similar to desert environments, has different dominant cyanobacteria and different proportions of these organisms. More study is needed to fully understand the cyanobacterial makeup of coastal sage scrub. It may be that the soil type, pH, or some other factor that we have not yet identified, increases the likelihood of seeing one type of cyanobacteria over another, even though we now speculate that they require the same growing conditions. What does matter for the goals of this study, is demonstrate that the cultured species of Trichotorquatus is a hardy organism because it was isolated and cultured twice. The fact that it is easily grown could make it a good subject for large scale culture and growth on damaged CSS biocrust sites.

green algae trebouxia sp. Mixed with the cyanobacteria in some of the MT flasks was the green algae thought to be a Trebouxia sp., but awaiting confirmation from ITS gene sequencing results. The culture used for inoculation was a mixed culture of Trichotorquatus and Trebouxia species. Initially, the experiment was planned to inoculate with only cyanobacteria, but there are benefits to having a mixed culture of eukaryotic Trebouxia species and prokaryotic Trichotorquatus species. Both organisms are drought tolerant and grow rapidly, and could be beneficial to use for inoculation in semi-arid coastal sage scrub; and used together, hostile conditions may force one to perish while the other survives. Using both effectively increases the chances of survival and benefit to biological soil crust restoration. Although commonly thought of as an aquatic organism, green algae is a common component of desert biological soil crusts, both as a free-living organism and as a symbiont

37 38

in lichens (Belnap and Lange 2003; Gray et al. 2007; Holzinger and Karsten 2013; Patzelt et al. 2014). In a recent paper on the hyper-arid Atacama Desert, species of green algae were able to survive even drier conditions than cyanobacteria could tolerate (Patzelt et a. 2014). Green algae, like cyanobacteria, endure drought by entering a state of anhydrobiosis, where cells cease to function, but remain living (Gray et al. 2007). Furthermore, once moisture is detected, algae can become metabolically active within an hour of rehydration (Gray et al. 2007). Specifically, Trebouxia sp. reduce water loss by accumulating organic osmolytes such as polyols (Holzinger and Karsten 2013). Given the protracted drought California is experiencing this ability to survive without metabolizing, differentiates and gives inoculation, as a restoration technique the advantage over vascular plant restoration as a more viable technique. Vascular plants die without water, unlike cyanobacteria and green algae, they have no protective mechanism to manage drought. Also like cyanobacteria, green algae is constructed of significant amounts of polysaccharides in the cell walls and outer sheath (Konig and Peveling 1984). Polysaccharides stabilize and aggregate the soil even if the organism dies—another advantage to using a mixed culture to inoculate damaged biocrust in restoration (Garcia-Pichel 2003). Soils aggregated by polysaccharides are protected from the forces of wind and water erosion (Belnap and Lange 2003). Using both organisms in a mixed culture confers advantages to the inoculation.

Chlorophyll a Another goal for this study was to find a non-destructive way to begin to rehabilitate the biological potential of the CSS biocrust. There was a question of whether cultured local cyanobacteria and green algae would grow sufficiently on native CSS soils. They did grow, and they grew fast. There was luxuriant growth on the soils in Figures 9G and 9H. Compare that to actively growing biocrust at Mission Trails in Figure 3B. This growth occurred in less than two months. It was interesting that even autoclaved soil grew cyanobacteria to the extent that it was not statistically different than the inoculated native soil in the one-way ANOVA (n = 50, α = 0.05). Looking at the two-way ANOVA comparing liquid inoculated treatments versus controls, we see autoclaving significantly reduces chlorophyll a (n = 40, p = 0.031). This is interesting as the process of autoclaving was used in this experiment as a surrogate for pesticide application. Removing the suite of metabolically active organisms in the soil has negative ramifications for biological activity. It suggests that the health of the soil community is important for the growth of cyanobacteria and green algae. The photographic evidence also shows superior growth on the native soil. Compare Figures 9E and 9F with the native soil growth in Figures 9G and 9H. Native soil has an assemblage of organisms that may

synergistically work together to foster growth of cyanobacteria and green algae, the two-way ANOVA certainly bears this out. Autoclaving heats up the soil and kills all organisms. Nonautoclaved treatments had significantly more chlorophyll a than the autoclaved treatments indicating that soil organism-harming practices like the use of pesticides sets back the growth of biocrust (Irvine 2011, Megharaj et al. 1989). Uncompromised native soil is the better substrate to grow a mixed culture inoculum. Statistical results signify that interaction of the intact microbial community is important for the growth of photosynthesizing cyanobacteria and green algae. The two-way ANOVA also found that inoculation significantly boosted the amount of chlorophyll a in the liquid inoculated treatments (n = 40, p < 0.001). Statistical results show inoculation with a mixed culture of cyanobacteria and green algae made an impressive difference in the chlorophyll a content of the treated soils. For these distinct differences to occur, inoculation worked because the mixed culture grew measurably in a short period of time. The statistical difference is emblematic of the impressive growth of green algae and cyanobacteria in a brief time span. The results of the one-way ANOVA found equal significance in growth of chlorophyll a levels in both native and autoclaved soils means good things for the potential to remediate even damaged native CSS soils not containing a full complement of organisms with a mixed culture inoculation, because, according to this experiment, even sterile soils grew cyanobacteria and green algae. Since the focus of this experiment was to find a non-destructive way to remediate biocrust, it was not disappointing to find the destructive ground crust treatment not statistically different than the growth of chlorophyll a on the control treatments. Biocrust organisms grew in non-inoculated native soil, but the difference is not apparent in the photo for the Control in Figure 9A. It was, however, seen in the statistical results from chlorophyll a levels for the Ground Crust treatment and the Control. The growth in the Ground Crust treatment was statistically different than the Autoclaved Control, so some growth had taken place in the Ground Crust treatment—just not to the impressive degree as the liquid mixed culture inoculated soils. Compare the photos in Figures 9C-H to see the green growth of the Ground Crust Inoculated. Once again, had the experiment run longer the apparent growth of green algae and other organisms (Figure 18) from the Ground Crust may have become statistically different than the Control, but would probably never catch up with the growth of the inoculated crusts, at least not for a long while. So in the short time that this experiment ran, inoculating soil with a liquid culture of cyanobacteria is preferable to destroying existing biocrust for use in rehabilitating damaged biocrust. The liquid inoculated soil had an average of 4.42 μg chla/g soil, compared to 15.75 μg chla/g soil for the reference

39

39 40

crust and less than 1 μg chla/g soil for autoclaved control. Inoculation of soil and growth of the culture increased the amount of chla to almost a third of the reference crust amount in just a few weeks.

extraCtaBle polySaCChariDeS

None of the treatments was statistically different from the others in the one-way ANOVA. A two-way ANOVA looking at all inoculated treatments versus non-inoculated found a significant difference (n = 50, α = 0.05, p = 0.012). Native soil inoculated with liquid mixed culture samples had the highest values for EPS (Figure 12), and it is possible had the experiment run longer that the difference may have become more pronounced as the cyanobacteria and green algae continued to grow. The polysaccharide extraction measurement was a proxy for the growth of extracellular polysaccharides contained in the filaments of cyanobacteria, cell walls, and sheaths of green algae. Longer experimentation time almost certainly would have resulted in large significant differences, at least between inoculated and non-inoculated treatments, and possibly between other treatment types. Weak acid extraction was the technique used for the separation of carbon from the sample. This method is not precise for EPS and captures a large background of general soil organic matter in addition to EPS. This may account for the differences in the one-way ANOVA not being significant. If a more precise method had been used it is foreseeable that significance would have been found with the one-way ANOVA. More importantly, extractable polysaccharides did increase, meaning that there was growth of EPS because of cyanobacterial and green algal growth and inoculated treatments were significantly different. Inoculated soils showed an increase in function, an increase in aggregation, and signs of carbon sequestration.

StaBility

Even though EPS levels were not found to be significant with the one-way ANOVA, stability measurements showed that EPS was at work binding soil particles together. The results also demonstrated the role of other organisms at work when the autoclaved, otherwise known as sterile soil control fell apart and had no stability compared to the other soil treatments. It was significantly less stable than the Ground Crust Inoculated and both of the Liquid culture inoculated soils. The growth of EPS in the Autoclaved Inoculated soil made it significantly more stable than the Autoclaved Control soil. Significantly, in a little less than two months enough cyanobacterial filaments had knit the soil particles together well enough that the stability increased substantially, even in sterilized soil. Both of the Liquid Inoculated treatments were significantly more stable than all of the other treatments. Liquid inoculated soil averaged 4.42 compared to 1.1 for the autoclaved control and 5.83 for the reference

41

crust. A large increase in soil function for the inoculated soil. This bodes well for using liquid mixed culture inoculum for increasing soil stability in restoration work and proves that improvement can be seen in soil structure in a short period of time with measurable and quantifiable results.

ConCluSion

This experiment demonstrated that remediation of damaged biological soil crusts in coastal sage scrub is possible and desirable in terms of increased function and stability in a short period of time. A developed biocrust can make a difference in whether a vascular plant can germinate and whether there will be sufficient resources in the lean and dry soils of coastal sage scrub habitat. If restoration philosophy shifts to make restoration of biological soil crusts the focus of initial restoration efforts, there may be a greater degree of success when planting vascular plants as the logical second step. Clearly, further research needs to be done, but the evidence is clear that inoculation does work and that it increases function at least in lab soils. The most successful growth was seen on nonautoclaved soil with an intact biome. Logically it follows that more emphasis should be placed on preserving the ecology of the biocrust organisms because they may be working synergistically to promote growth in ways we don’t fully understand yet. One way to do this is to decrease the application of pesticides knowing that they will negatively affect the growth of biocrust organisms. Given how precious and endangered coastal sage scrub habitat is, and how climate change is decreasing precipitation and raising temperatures, it makes sense to prioritize the preservation and restoration of biological soil crust. Biological soil crusts are perhaps the most important biotic component for maintaining arid and semi-arid land systems where conditions limit the growth of vascular plants (Bowker et al. 2005). Certainly biocrusts play many ecosystem roles and bring about changes in hostile environments that tip the balance to allow vascular plants to germinate and establish (Chaudhary et al. 2009; Eldridge et al. 2010). Notable ways they do this are through soil building and soil-moisture retention. As San Diego is expected to become warmer and drier in the future (Messner et al. 2011), developing techniques to increase soil-moisture retention could be key to conserving and rehabilitating our local unique and valuable ecosystems (Bowker 2007) and decreasing desertification (Pimental et al. 1995). Mixed culture inoculation could help prevent erosion, because even when dormant, extracellular polysaccharides of cell walls, sheaths, and filaments continue to bind soil particles. Unlike the plants installed in a restoration effort that require costly inputs of hydration, once inoculated, cyanobacteria can simply go dormant when they lack moisture and resume growing when the conditions are appropriate. Given the nature of biological soil crust to positively affect the hydrological cycle, more research into developing the technique of liquid mixed culture inoculation should be pursued.

ACKNOWLEDGEMENTS

My most sincere thanks and eternal gratitude go to my advisor, David Lipson, for encouraging me to pursue what must have initially seemed a crazy idea, and for helping me to see it to fruition. Without his unbridled enthusiasm and thorough knowledge, this thesis might well have died on the vine. I am also indebted to John O’Leary, my committee member, who shares my passion for wild California and the processes that shape the landscape and plants that live there. Thank you, John, for painstakingly proofreading my thesis and giving me excellent feedback. It was kind of committee member, Matthew Edwards, to take the time to teach me statistics so pa-tiently. Thank you Matthew! I also owe a debt of gratitude to Tom Zink for agreeing to serve as my committee member when Matthew was teaching at another location. I am grateful for the communications with forthcoming scientists who freely gave their opinions and shared their knowledge. Thank you Jayne Belnap, Nicole Pietrasiak, Robert Sheath, and Rosalina Stancheva for your help in identification. Thank you to the fabulous group of kind-hearted lab mates Archana Srinivas, Jaime Zlamal, Melanie Merkley, Lisa Ordinez, Robert Wagner, and Sherly Castro. They were quick with support, and happily accessible with good advice and helpful laboratory skills. I wish you all continued success and the best that life has to offer. Also figuring as an integral person in this process, (and it is a process!), is my friend Brenda Helt. A friend–sure–that word gets tossed around easily, but she is the personification of loving support and well-timed swift kicks. It doesn’t hurt that she is also a respected aca-demic with really great writing advice, which in no way, compares with her kind heart, sense of justice, and compassionate soul. Her husband, Jon, was supportive in an important albeit quieter way. It has been a tough year for personal reasons, and I was not sure I could keep on the road of this graduate degree, except for all of your help. It fills my heart to know that people like you walk this planet and I am so grateful for your help! More grateful than you will ever, or, could possibly, know. Warm gratitude to my yoga shala. Your daily classes, insights into Buddhist teachings and into the magic that is yoga, transformed me every day into someone better able to take on life’s challenges as Sharon Reeve.

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