1. 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
2. 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, Chair
Department of Biology
Tom Zink
Department of Biology
John O’Leary
Department of Geology
Approval Date
4. iv
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
5. v
ABSTRACT OF THE THESIS
Improving Ecosystem Function:
Facilitating Restoration of Degraded Biocrusts
Using Mixed Culture Inoculation
by
Sharon Reeve
Master 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.
6. TABLE OF CONTENTS
PAGE
ABSTRACT................................................................................................................................
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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7. TABLE OF CONTENTS
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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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8. LIST OF TABLES
PAGE
Table 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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9. LIST OF FIGURES
PAGE
Figure 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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10. 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.
1
11. 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).
2
12. 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
3
13. 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
4
14. 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).
5
15. 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
6
16. 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).
7
17. 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
8
18. 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
9
19. 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: derived
from Terrestrial Vegetation of California by Michael G.
Barbour et al. 2007
10
20. 11
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.).
21. 12
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
22. 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?
13
23. 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
14
Figure 2. Coastal sage scrub location study sites. Credit: derived from Google Maps
24. 15
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
25. 16
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
*
K2
HPO4
MgSO4
7H2
O
CaCl2
2H2
O
Citric Acid H2
O
Ferric Ammonium Citrate*
Na2
EDTA H2
O
Na2
CO3
Trace Metal Mix
Distilled Water
Agar
*KNO3
substituted for NaNO3
*Ferric Citrate substituted for
Ammonium Citrate
*pH should be 7.1
.
.
.
.
1.5 g
0.04 g
0.075 g
0.036 g
0.006 g
0.006 g
0.001 g
0.02 g
1.0 ml
1.0 L
10.0 g
Growth Media Weight/Volume
26. 17
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 (20.3μmol 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 (20.3μmol
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
27. 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
18
Figure 5. Flasks of inoculant from left to right: MT A , MT A -2, MT B & SMER A.
Credit: Sharon Reeve
28. 19
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 H2
SO4
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
29. (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
20
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% of soil remains on sieve after five dipping cycles
4 10-25% of soil remains on sieve after five dipping cycles
5 25-75% of soil remains on sieve after five dipping cycles
6 75-100% of soil remains on sieve after five dipping cycles
Stability
Class
1-6
Criteria for Assignment to Stability Class
Time Dipped into Water
Figure 6. Soil sieve apparatus. Figure 7. Reference CSS biocrusts.
Credit: Sharon Reeve
30. 21
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 cmsurface 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.
31. 22
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
32. 23
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
33. 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.
24
34. 25
RESULTS
Culturing 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
(20.3μmol 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
35. 26
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
36. 27
Figure 11. Chlorophyll a concentrations of five treatments. (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
chlorophyll
a
(ug/Chla/g)
Inocula-on
Treatments
Chlorophyll
a
per
Treatment
a
a,b
b
c
c
Control Autoclaved
Inoculated
InoculatedGround
Crust
Inoculated
Autoclaved
Control
(μgChla/gsoil)
Chlorophyll a per Treatment
Inoculation Treatments
chlorophylla
Variable
log (Chla)
Stability
EPS
EPS
R2
0.733
0.579
0.129
0.125
df (model)
4
4
4
1
df (error)
45
45
45
48
F
30.9
15.441
1.673
6.858
P
<0.001
<0.001
0.173
0.012
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).
37. 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)
Figure 12. Extractable polysaccharide concentrations in five treatments. (Columns with
non-repetitive letters are significantly different. One-way ANOVA, n = 50, α = 0.05. Means
and standard errors of 10 replicates).
Control Autoclaved
Inoculated
InoculatedGround Crust
Inoculated
Autoclaved
Control
Extractable Polysaccharides(μgC/gsoil)
a a
a a
a
38. 29
Figure 13. Stability index of soils from the five treatments (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
Stability
Index
Treatment
Stability
a
a,b
b,c
c,d
d
Control Autoclaved
Inoculated
Inoculated
Inoculated
GroundAutoclaved
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).
39. 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.)
30
Variable
log (Chla)
Stability
EPS
R2
0.757
0.593
0.123
df
(mdl)
3
3
3
df
(err)
36
36
36
F
107.249
39.493
4.897
F
5.057
11.877
0.06
F
0
1.027
0.102
P
<0.001
<0.001
0.033
P
0.031
0.001
0.809
P
0.989
0.318
0.751
Table 5. Statistical Values for Two-way Analysis of Variance
Inoculated Autoclaved Interaction
40. 31
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 A
KC311913 Atacama Desert isolate
42. 33
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).
43. 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
44. 35
DISCUSSION
Leptolyngbya 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
45. 36
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
46. 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
47. 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
48. 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
