Survival of two introduced plant growth promoting
micro-organisms in green roof growing substrates in southern
Finland
Long Xie
Master’s thesis
University of Helsinki
Department of Agricultural Sciences
Horticulture
June, 2014
HELSINGIN YLIOPISTO  HELSINGFORS UNIVERSITET  UNIVERSITY OF HELSINKI
Tiedekunta/Osasto  Fakultet/Sektion  Faculty
Laitos  Institution  Department
Faculty of Agriculture and Forestry
Department of Agricultural Sciences
Tekijä  Författare  Author
Long Xie
Työn nimi  Arbetets titel  Title
Survival of two introduced plant growth promoting micro-organisms in green roof
soil in southern Finland
Oppiaine  Läroämne  Subject
Plant Production Science, horticulture
Työn laji  Arbetets art  Level
Aika  Datum  Month and year
Sivumäärä  Sidoantal  Number of pages
Master’s thesis
June, 2014
76
Tiivistelmä  Referat  Abstract
Glomus intraradices and Bacillus amyloliquefaciens are two commercially used
plant growth promoting micro-organisms. They associate with plant roots to
facilitate host plants to absorb nutrients, induce resistance against pathogens and
pests, and regulate growth through phytohormones. Growth conditions for plants
on green roofs are often unfavorable. In order to test whether growth and
development of green roof plants could be enhanced via improving the microbial
interface, G. intraradices and B. amyloliquefaciens were inoculated on
experimental plots on a green roof in the summer of 2012. The experimental plots
were marked as R (inoculated with B. amyloliquefaciens from Rhizocell), M
(inoculated with G. intraradices from MYC4000), and C (control). The green roof
was made of sedum-herb-grass mats. The plants included e.g. stonecrops,
bluegrasses, yellow rockets, white clover, mullein, pennycress, and moss. The
survival and development of G. intraradices and B. amyloliquefaciens were
studied respectively from Poa alpina roots and soils in the summers of 2012 and
2013. G. intraradices was not detected in P alpina roots according to root staining
and microscopy. Probable reasons for the lacking of G. intraradices include high
phosphorus content in the soils, high soil temperature, and low soil moisture. PCR
and qPCR were used to detect Bacillus content in green roof soils. The abundance
of B. amyloliquefaciens was related to soil water content and soil temperature.
During the last two measurements in 2012, 4 weeks of high moisture content in the
soil resulted in large increase of B. amyloliquefaciens content in both M and R
groups, but then decreased substantially due to drought and heat in 2013. In 2013,
Only R group increased from the third to the last measurement, indicating probable
resistance of the B. amyloliquefaciens strain from Rhizocell additive. The
synergistic effect of B. amyloliquefaciens and G. intraradices might be responsible
for the thousand-fold increase of Bacillus content in M group in 2012.
Avainsanat  Nyckelord  Keywords
B. amyloliquefaciens; G. intraradices; P. alpina; detection; microscopy, PCR;
qPCR;
Säilytyspaikka  Förvaringsställe  Where deposited
Department of Agricultural Sciences and Viikki Campus Library
Muita tietoja  Övriga uppgifter  Further information
Supervisor: Prof. Jari Valkonen
Table of Contents
Abbreviations ........................................................................................................................... 5
1 Introduction........................................................................................................................... 6
2 Literature review ................................................................................................................... 8
2.1 Green roofs serve for urban ecosystems ........................................................................ 8
2.2 Plant growth promoting micro-organisms ..................................................................... 9
2.3 Glomus intradices ........................................................................................................ 10
2.3.1 Infection process of Glomus intraradices ............................................................. 10
2.3.2 Glomus intraradices helps plants to uptake nutrients ........................................... 12
2.3.3 Glomus intraradices helps plants to survival under environmental stresses ........ 12
2.3.4 Glomus intraradices helps plants to survive under biotic stresses ....................... 13
2.4 Bacillus amyloliquefaciens .......................................................................................... 14
2.4.1 Life cycle of Bacillus amloliquefaciens ................................................................ 14
2.4.2 Bacillus amyloliquefaciens helps plants to survive under environmental stresses 16
2.4.3 Bacillus amyloliquefaciens helps plants to survive under biotic stresses ............. 17
2.5 AMF colonization of Poa alpina ................................................................................. 19
3 Research objectives............................................................................................................. 21
4 Material and methods.......................................................................................................... 22
4.1 Green roof installation and experimental design ......................................................... 22
4.2 Plant material and inoculants ....................................................................................... 23
4.3 Analysis of Glomus intraradices ................................................................................. 24
4.3.1 Root treatment and staining .................................................................................. 24
4.3.2 Microscopic examination of roots ........................................................................ 25
4.3.3 Available phosphorus content measurement in the soil ........................................ 26
4.4 Analysis of Bacillus amyloliquefaicens ....................................................................... 26
4.4.1 Extraction of total DNA samples .......................................................................... 26
4.4.2 Detection of Bacillus amyloliquefaciens............................................................... 27
4.4.3 Quantification of Bacillus amyloliquefaciens ....................................................... 27
4.4.4 Calculation of Bacillus amyloliquefaciens content in the soil .............................. 27
5 Results................................................................................................................................. 29
5.1 Soil temperature/moisture on green roof during summers........................................... 29
5.1.1 Soil temperature .................................................................................................... 29
5.1.2 Soil moisture ......................................................................................................... 31
5.2 Available phosphorus in the green roof soil ................................................................ 33
5.3 No mycorrhizal structures of Glomus intraradices were found .................................. 33
5.4 Content of Bacillus amyloliquefaciens in the soil........................................................ 34
5.3.1 Detection of Bacillus amyloliquefaciens............................................................... 34
5.3.2 Bacillus amyloliquefaciens content in soil samples .............................................. 35
5.3.3 Large increase of Bacillus amyloliquefaciens content in M group ....................... 38
6 Discussion ........................................................................................................................... 39
6.1 Hypothesis of absence of Glomus intraradices ........................................................... 39
6.1.1 High phosphorus content might inhibits AMF growth ......................................... 39
6.1.2 Soil temperature could influences AMF growth ................................................... 41
6.1.3 Soil moisture could influences AMF growth ........................................................ 42
6.1.4 Micro-organism competition might leads to AMF absence.................................. 43
6.1.5 AMF was not observed due to the unsuitable sampling time ............................... 44
6.2 Development pattern of Bacillus amyloliquefaciens content ....................................... 44
6.2.1 Existence of Bacillus amyloliquefaicens in non-inoculated soil ........................... 44
6.2.2 Soil temperature affects Bacillus amyloliquefaciens reproduction ....................... 45
6.2.3 Soil moisture affects Bacillus amyloliquefaciens reproduction ............................ 46
6.2.4 Resistant Bacillus amyloliqufaciens from Rhizocell additive survived through
winter ............................................................................................................................. 47
6.2.5 Synergetic effect between the two micro-organisms ............................................ 48
7 Conclusions and future perspectives ................................................................................... 49
8 Acknowledgements ............................................................................................................. 51
References.............................................................................................................................. 53
Appendices ............................................................................................................................ 69
Abbreviations
AC
Arbuscule colonization
AMF
Arbuscular mycorrhizal fungi
API
Analytical profile index
HC
Hyphae colonization
IAA
Indole-3-acetic acid
ISR
Induced systemic resistance
NFW
Nuclease free water
P
Phosphorous
PCR
Polymerase chain reaction
PSM
Phosphate solubilizing micro-organisms
qPCR
Quantitative polymerase chain reaction
UV
Ultraviolet
VC
Vesicle colonization
VOCs
Volatile organic compounds
6
1 Introduction
Accompanied by urbanization and increasing fossil fuel consumption, air pollution
has been recognized as one of the most crucial environmental issues in many cities
(Akimoto, 2003; Mage et al., 1996; Cole & Neumayer, 2004), causing millions of
premature deaths from respiratory diseases each year (Katsouyanni & Pershagen,
1997; Bernstein et al., 2004; Kim et al., 2004).
Scientific studies have revealed that trees and shrubs can effectively mitigate urban
air pollution by filtrating and removing air pollutants, such as O3, NO2, SO2, and CO
(Bolund & Hunhammar, 1999; Nowak et al., 2006). As complimentary to traditional
parks and gardens on the ground level, green roofs in urban areas have been more
and more employed in urban green space planning, and its superiorities have been
manifested. A green roof is a roof of a building covered with vegetation and growth
substrates. Green roofs can provide ecosystem services to ambient environment,
including mitigating air pollution, relieving urban heat island effect, conserving
energy, and retaining stormwater (Peck et al., 1999; DeNardo et al., 2005;
Oberndorfer et al., 2007; Takebayashi & Moriyama, 2007; Yang et al., 2008).
Growing on top of buildings, green roofs can provide great advantages in densely
populated urban areas, especially where parks and gardens are scarce. In addition,
green roof provides hedonic value (Morancho, 2003).
However, the application of green roofs has been restricted due to some challenges.
Henry and Frascaria-Lacoste (2012) pointed out three challenges, which are extreme
weather conditions on rooftop, choice of suitable plants, and high maintenance cost.
In our study, two plant-growth-promoting micro-organisms were inoculated in green
roof soil, namely Glomus intraradices and Bacillus amyloliquefaciens to facilitate
plants to maintain good physiological conditions.
7
G. intraradices and B. amyloliquefaciens have been repeatedly reported to facilitate
plant growth through symbiosis. The benefits are induced systemic resistance (ISR),
nutrient absorption, and resistance to environmental stresses. Nowadays, they are
produced as commercial agricultural inoculants and used in plant production under
natural environment. The question is whether they can promote plant growth on
rooftop, withstanding extreme weather conditions, forming symbiosis with green
roof plants and cutting down maintenance costs. Prior to answering these questions,
we need to find out whether they can survive in the green roof substrates, and what
their development patterns are. To our knowledge, this is the first time to study
micro-organism inoculation in a green roof environment.
8
2 Literature review
2.1 Green roofs serve for urban ecosystems
During the last century, billions of people have flooded into cities for a better living,
a process called urbanization. From the year 2003 to 2011, 4.4% of total population
moved to cities in developing countries, and 3.2% in developed ones (United Nation,
2004; 2012). Increasing population in cities inevitably results in more pronounced
urban environmental problems (Grimm, et al., 2008).
As the term suggests, a green roof is a rooftop covered with vegetation and growing
substrates for the plants. Green roofs have long been regarded as engineering or
horticultural challenges, and serving aesthetic purpose (reviewed by Oberndorfer et
al., 2007). However, the contemporary proliferation of green roofs is supported by
more and more scientific evidence of a number of ecosystem services to mitigate
urban environmental issues. In addition, as green roofs grow on top of buildings,
they do not take up lands in populated urban areas. Because of their advantages,
green roofs have been adopted in many developed countries in green space planning,
such as Germany, Singapore, and the United States (reviewed by Oberndorfer et al.,
2007).
Despite the beneficial services and functions, the application of green roofs has been
restricted by some challenges. Extreme weather conditions on rooftop, choice of
suitable plants, and high installation and maintenance costs are the three major
problems (Henry & Frascaria-Lacoste, 2012; Nurmi et al., 2013). If these challenges
could be met, green roofs could be even more widely used, not only in developed
countries, but also in developing ones, where urban environmental issues are severe
and urgent.
9
2.2 Plant growth promoting micro-organisms
In the rhizosphere where plant roots and soil meet, countless micro-organisms reside
and propagate at a much denser level than other regions. Some of them might be
neutral or lethal to the growth and survival of plants, but others could support their
host plants via various mechanisms (reviewed by Compant et al., 2010). Due to their
plant-growth-promoting characteristics, these micro-organisms have been used as
commercial inoculants. Glomus intraradices and Bacillus amyloliquefaicens are two
of them (Table 1).
Table 1. Commercially used microbial inoculants
Kingdom
Genus
Species
Benefits
Bacteria
Bacillus
amyloliquefaciens
Nutrient absorption, soil borne pathogens
lichenformis
resistance, environmental stress
megaterium
tolerance, phytohormones production
sbtilis
thuringlensis
Bradyrhizobium
japonicum
Nitrogen fixation
Delftia
acidovorans
Alkaline conditions tolerance
Pseudomonas
chlororaphis
Pathogen resistance, nutrient absorption
fluorescens
trivialis
Fungi
Serratia
plymuthica
Pathogen resistance
Streptomyces
griseoviridis
Pathogen resistance
Beauveria
bassiana
Pest and microbial control
Coniothyrium
minitans
Pathogen control
Glomus
intraradices
Nutrient absorption, pathogen resistance
Metarhizium
anisopliae
Pests control
Paecilomyces
lilacinus
Nematode control
Phlebiopsis
gigantea
Pathogen resistance
Trichoderma
asperellum
Pathogen resistance
harzianum
Virus
Verticillium
lecanii
Pest control
Baculoviridae.
Cydia pomonella
Pest control
granulovirus
Source from Fargro Ltd. http://www.fargro.co.uk/products/default.asp?dist=WORLD
10
2.3 Glomus intradices
Glomus intraradices is an endomycorrhizal fungus distributed in almost all soil
ecosystems and types (reviewed by Strack et al., 2003). As an arbuscular mycorrhizal
fungus (AMF), it benefits vascular plants by forming mutualistic symbiote called
mycorrhiza with plant roots, facilitating the growth and survival of plants (St-Arnaud
et al., 1996; reviewed by Strack et al., 2003).
Glomus intraradices can form tree-like structured organs named arbuscules in the
cell of plant roots (Fig. 1D). Arbuscule functions as a carbohydrate/mineral exchange
organ between host plants and G. intraradices (Strack et al., 2003). It also forms
bulb-like, storage organs called vesicles (Fig. 1E). Biermann (1983) found that
vesicles of two Glomus species could act as propagules, which are infective and
crucial to the effectiveness of root colonization process. After colonizing host plant
roots and forming internal organs, G. intraradices develops a network of external
mycelium, which is known as extraradical mycelium (Bago et al., 1998). This
extraradical mycelium not only enlarges the absorption surface areas of plant roots,
but also acts as an environmental sensor (Van Aarle et al., 2002; Bago et al., 2004).
2.3.1 Infection process of Glomus intraradices
The infection of G. intraradices was revealed by fungal structure staining with
Trypan Blue (Phillips & Hayman, 1970). The process starts with germination of G.
intraradices spore. After attached to the surface of plant root, appressoria are formed
to penetrate the root surface. The penetration process is facilitated by nonaggressive
cell wall lytic enzyme. Afterwards, the hyphae develop arbuscules and vesicles
within the cell. Finally the spores appear, indicating that G. intraradices can enter
another cycle of colonization (Fig. 1). A picture of G. intraradices structures in
tomato root cell stained with Trypan Blue is shown in Fig. 2 (Myllys et al., 2013). In
this picture, hyphae, arbuscules and vesicles are manifested.
11
A
Spore
B
Spore
C
Spore
Appressoria
Root cells
Hyphae
D
F
E
Vesicle
Spore
Arbuscule
Fig.1. The life cycle of G. intraradices in root cells adapted from Strack et al., 2003. The process
includes spore germination (A), appressorium formation (B), hyphae formation (C), arbuscule
formation (D), vesicle formation (E), and spore formation (F).
Fig.2 Vesicles and arbuscules of G. intraradices in tomato root stained with Trypan Blue. Asterisk
stands for vesicle and arrowhead points out arbuscule (Myllys et al., 2013).
Root colonization by AMF can affect morphology and function of both host plants
and the fungi (Bonfante & Perotto, 1995). For instance, arbuscule is a key feature of
AMF. They are short-lived as they finally senesce after 4-10 days of symbiosis
(Sanders et al., 1977). Arbuscule takes up a large volume of root cell in highly
colonized root system, but it is still separated from the cell protoplast by host plasma
membrane. Also the central major vacuole in the cell is fragmented, organelles
12
increase significantly in number, and nucleus moves to the center and increases in
volume (Balestrini et al., 1994). It has been demonstrated by the improved labelling
techniques combined with epifluorescence microscopy that plant cytoskeletal
components (microtubules and microfilaments) play central roles in the molecular
information exchange between host plants and fungi before the hyphae penetration
(Genre & Bonfante, 1997, 1998; Blancaflor et al., 2001; Timonen & Peterson, 2002).
Studies have suggested that phytohormones released by plants can regulate
colonization of AMF, including cytokinins (Allen et al., 1980), abscisic acid (Bothe
et al., 1994), and jasmonic acid (Regvar et al., 1996). It further substantiates the
reciprocal interaction between plants and AMF.
2.3.2 Glomus intraradices helps plants to uptake nutrients
As a member of AMF, G. intraradices can help host plants to uptake, transport and
transfer phosphorous especially when limited P is available to the roots, in exchange
of photosynthetically fixed carbohydrates provided by host plants (Koide & Kabir,
2000; Bücking & Shachar-Hill, 2005). In some soil types, organic P makes up a large
proportion of total P source (50-85%), but plants need inorganic P mostly.
Consequently, plants adopt strategies to utilize organic P (Dalal, 1977; Schachtman et
al., 1998). Koide and Kabir (2000) performed an experiment, which showed that
plants could hydrolyze organic P (5-bromo-4-chloro-3-indolyl phosphate and
phenolphthalein diphosphate) into inorganic form through production of an enzyme
called phytase. Moreover, AMF-infected plants can release more phytase than
non-infected ones. However, it is not established whether AMF release phytase or
they induce the plants to produce more phytase (Koide & Kabir, 2000; Feng et al.,
2003). Other nutrients have also been reported to be transferred from mycorrhiza to
host plants (Khalil, et al., 1994).
2.3.3 Glomus intraradices helps plants to survival under environmental stresses
Glomus intraradices can alleviate the stresses caused by unfavorable growth
13
conditions. In one experiment, kidney beans were inoculated with G. intraradices
and grown under salinity stress. It was found that AMF-inoculated plants gained
salinity tolerance by regulating hydraulic conductance (Aroca et al., 2007). Other
Glomus species have also been found to possess salinity tolerance. Lettuce
inoculated with G. fasciculatum exhibited less yield decline under high saline soil
than non-inoculated ones (Ruiz-Lozano et al., 1996). Researchers concluded that the
salt tolerance is not a nutrient-dependent mechanism but a physiological outcome
brought by AMF colonization, which is in accordance with Aroca’s finding.
Drought tolerance is another benefit gained through AMF colonization. Studies were
made on lettuce inoculated separately with seven Glomus species. All of the AMFs
exhibited enhanced drought tolerance, yet each showed diverse effectiveness. Among
the seven species, G. intraradices ranked the fifth best species in terms of inducing
drought tolerance (Ruiz-Lozano, et al., 1995).
Glomus intraradices has been found to help plants growing in heavy metal
contaminated environment to survive (Hildebrandt et al., 2007). González-Guerrero
et al. (2008) found out that the hyphae of G. intraradices bind heavy metals to their
cell wall and keep them from emitting to the plant tissues.
2.3.4 Glomus intraradices helps plants to survive under biotic stresses
Furthermore, G. intraradices can mitigate diseases caused by plant pathogens. Even
though Caron et al. (1986) observed a decrease of G. intraradices colonization at
high P concentration in soil and plant tissues, the presence of G. intraradices could
still suppress root necrosis caused by Fusarium oxysporum f. sp. radicis lycopersici.
Meanwhile, treatment with only high P concentration could not alter the severity of
the symptoms. In other words, it was G. intraradices, not the addition of P nutrients
that could mitigate root necrosis. Also St-Arnaud (1994) found that P concentration
did not affect the Pythium ultimum infection, but inoculation with G. intraradices
14
could reduce propagules population of P. ultimum as much as 90%.
2.4 Bacillus amyloliquefaciens
Bacillus amyloliquefaciens is a Gram-positive, spore-forming bacterium closely
related to B. subtilus (Priest et al., 1987). Bacillus group members can elicit the
defense and tolerance of host plants against biotic and environmental stresses. The
beneficial outcome is called induced systemic resistance (ISR) (Van Loon & Glick,
2004, Kloepper et al., 2004).
Bacillus amyloliquenfaciens was given the name for its production of α-amylase and
protease (Fukumoto, 1943). Because of its close relation to B. subtilis, B.
amyloliqueficens has long been recognized as a subspecies or a variant of B. subtilis
(Tsuru, 1962; Gordon et al., 1973). It was not until 1987 that Priest et al. established
B. amyloliquefacines as a separate species, even though it was difficult to distinguish
on the basis of classical phenotypic tests. Physiologically, B. subtilis and B.
amyloliequefaciens release different enzymes and use different metabolism pathways
(Priest, 1977). DNA of B. amyloliefaciens shares less than 25% homology with DNA
of B. subtilis, compared with at least 50 to 60% homology shared by the strains of
the same species (Welker & Campbell, 1967; Seki et al., 1975). Later, the
unrelatedness of B. amyloliquefacines and B. subtilis has been proven by new
techniques, such as API test (O’Donnell et al., 1980, Logan et al., 1984), gas-liquid
chromatography (O’Donnell et al., 1980), and pyrolysis mass spectrometry (Shute et
al., 1984).
2.4.1 Life cycle of Bacillus amloliquefaciens
Bacillus amloliquefacines is rod-shaped soil bacterium, 0.7 to 0.9 by 1.8 to 3.0 μm.
Bacterial cells form chains that are motile to facilitate their movement (Priest et al.,
1987). As a Bacillaceae member, B. amyloliquefaciens forms endospores under
adverse conditions. The endospores can be dispersed in the soil and germinate later,
15
infecting plants through root system and forming symbiosis (Nicholson et al., 2000).
Bacillus species are widely distributed in almost all soil types, but they are
commonly distributed in rhizosphere, where they form a close connection to plant
roots. It can be explained by the effect of root exudates (Maheshwari, 2011). Root
exudates are biochemical substances that released in the rhizosphere by plant roots,
affecting the dynamics and distribution of micro-organisms in the neighboring soil
bulks (Schöttelndreier & Falkengren-Greup, 1999). In need of nutrients, bacillus
move towards plant roots, a process called tropotaxis. When reaching the root surface,
efficient Bacillus can survive the plant immune system (Maheshwari, 2011) and form
structures to anchor on root surface (Reva et al., 2004). Successful colonization
occurs when biofilm is formed on the surface of seeds, root, or root hairs (Reva et al.,
2004; Maheshwari, 2011; Chen et al. 2012). This layer of biofilm not only facilitates
the colonization of Bacillus, but also prevents competition from other
micro-organisms (López et al., 2009). After colonization, Bacillus produce
antimicrobials and plant hormones to promote plant growth (Maget-Dana et al., 1992;
Idriss et al., 2002; Yu et al., 2002; Makarewicz et al., 2006; Idris et al., 2007; López
et al., 2009; Maheshwari, 2011).
When adequate nutrients are available to Bacillus, it undergoes vegetative cycle,
featured by splitting of the mother cell into two daughter cells. When nutrient
depletion occurs, it undergoes sporulation cycle, forming stress resistant endospores
(Fig. 3, reviewed by Errington, 2003). It is commonly accepted that endospores from
Bacillus are resistant to very adverse environments including extreme temperature
(Fox & Eder, 1969; Margosch & Ehrmann, 2006), desiccation (Koike et al., 1992),
UV/gamma radiation (Setlow, 1988), and high/low pressure (Margosch et al., 2004).
16
Coat
Germination
Growth
Cortex
Sporulation
Vegetative
cycle
Prespore
Septum
Polar
Division
division
Mother cell
Fig.3. Sporulation cycle of G. intraradices . Adopted from Errington (2003).
2.4.2 Bacillus amyloliquefaciens helps plants to survive under environmental stresses
Researchers from India published a paper about the effect of B. amyloliquefaciens on
rice plants under salt stress (Nautiyal et al., 2013). Their experiment shows that B.
amyloliquefaciens NBRISN13 (SN13) increased salt tolerance of rice plants under
100 mM concentration, according to various vegetation growth parameters. Increased
shoot length (27.43%), root length (73.71%), and dry weight (55.47%) were
observed. They further tested the gene expression, finding that B. amyloliquefaciens
exerted gene regulation on rice plant which involved 14 salt stress responsive genes.
Bacillus amyloliquefaciens-inoculated plants also show drought tolerance. Normal
wheat (Triticum aestivum L.) seedlings showed 96% survival after 4 days drought
period, 70% after 5 days, and 50% after 7 days, whereas seedlings inoculated with B.
amyloliquefaciens 5113 exhibited 100% survival after 4 days, 88% after 5 days, and
67% after 7 days (Kasim et al., 2013). Other growth parameters, such as water
content and dry/fresh weight, have also certified the drought resistance trait of B.
amyloliquefaciens-inoculated plants.
Bacillus amyloliquefaciens can promote absorptive ability of plants under nutrient
17
deficiency, especially phosphate (Rodríguez & Fraga, 1999; Idriss et al., 2002). B.
amloliquefaciens was classified as a phosphate solubilizing micro-organism (PSM)
to produce a series of phosphate solubilizers. Phosphate solubilizers including lactic,
isovaleric, and isobutyric acid can solubilize inorganic phosphate and mineralize
organic phosphate to make phosphorous available to the plants (Krasilnikov, 1962;
Gaur & Ostwal, 1972, Vazquez et al., 2000). Researchers have found that B.
amyloliqueficens possesses the ability to secret phytase as scavenger enzyme that
dephosphorylates phytate phosphorus (myo-inositol hexakisphosphate) and makes
phosphorus available to plant (Makarewicz et al., 2006). Idriss et al. (2002) found
that B. amyloliquefaciens could significantly increase phytase content in the culture
medium compared with control groups. Meanwhile other growth performance
indicators, including shoot weight, root weight, and root length, were increased by
87.7%, 101.8%, and 46.1% respectively (Idriss et al., 2002).
2.4.3 Bacillus amyloliquefaciens helps plants to survive under biotic stresses
Besides inducing environmental resistance, B. amyloliquefaciens has been widely
recognized to plant increase resistance toward biotic stresses caused by fungi,
bacteria, virus, and pests.
In a study about Tomato mottle virus (ToMoV), B. amyloliquefaciens had shown to
reduce the severity of ToMoV infection in tomato plants, substantiated by both visual
symptom severity rating and Southern dot blot viral DNA analysis. The mechanism
might include resistance towards ToMoV, whitefly (ToMoV vector), or the
transmission process (Murphy & Zehnder, 2000). B. amyloliquefaciens strain
EXTN-1 was found to enhance resistance against Pepper mild mottle virus (PMMoV)
by triggering the expression of defense associated gene(s) (Park et al., 2001). Other
viruses having been reported to be constrained by B. amyloliquefaciens includes
Cucumber mosaic cucumovirus (CMV) (Zehnder et al., 2000), Tobacco mosaic virus
(TMV) (Shen et al., 2010), and Beet necrotic yellow vein virus (BNYVV)
18
(Desoignies et al., 2011).
In terms of antifungal activity, B. amyloliquefaciens strain BO7 was found to
antagonize the vascular wilt fungus Fusarium oxysporum f. sp. lycopersici (Fol)
(Vitullo et al., 2012). Significant reduction of wilt symptom caused by Fol was
observed in experimental groups pre-treated with BO7. The research was carried
forward to find out that bacterial surfactins play the major roles by initiating the
formation of a biofilm layer on the surface of root system. It facilitates B.
amyloliquefaciens to develop exclusive interaction with plant roots, and reduce the
competition from other micro-organisms (López et al., 2009). Fengycins and iturins
are another two antimicrobials produced by B. amyloliquefaciens, and they exhibit a
synergistic effect (Maget-Dana et al., 1992; Yu et al., 2002).
The isolated compounds from B. amyloliquefaciens could restrict bacterial pathogens.
Ryu et al. (2004) exposed Arabidopsis plants to volatile organic compounds (VOCs)
extracted from B. amyloliquefaciens IN937a before the plants was infected by
Erwinia carotovora. Among 9 VOCs of plant-growth-promoting rhizobacteria, B.
amyloliquefaciens and B. subtilis showed greatest relief of Arabidopsis from E.
carotovora infection. They found that the longer Arabidopsis was exposed to VOCs,
the better resistance was manifested. It suggests that VOCs can trigger chemical
changes in plant metabolism, resulting in enhanced plant defense. They further
identified the bioactive substance in the VOCs as 2,3-butanediol. In addition to its
antibacterial feature, 2,3-butanediol is involved in growth prompting of Arabidopsis
(Ryu et al., 2004). Other research groups have found antibacterial features of B.
amyloliquefaciens against Ralstonia solanacearum (Hu et al., 2010), Xanthomonas
axonopodis pv. glycines (Li et al., 2008; Prathuangwong & Buensanteai, 2007), and
Paenibacillus larvae (Benitez et al., 2012).
Bacillus amyloliquefaciens can suppress plant pests such as nematodes (Lian et al.,
2007; Burkett-Cadena et al., 2008; Mendoza et al., 2008). It was found that protease
19
Apr219 was the active ingredient causing the nematicidal effect, along with subtilisin
(Huang et al., 2004). Liu et al. (2013) discovered that plantazolicin produced by B.
amyloliquefaciens strain FZB42 is another nematicidal biochemical.
2.5 AMF colonization of Poa alpina
Known as bluegrass, Poa alpine is a perennial meadow grass species that commonly
grows in North America and Europe. P. alpina is heavily leafy plant growing in
scattered, dense clumps. Leaves are mostly basal, thin, and narrow blades ranging
from 1-12 cm in length. The stem is 10-40 cm, terminated by pyramidal pinkish
inflorescences (Flora of North America Editorial Committee, 2007). P. alpina starts
to sprout at the early spring and stops growth in September in Finland. It is a
common fodder grass in meadow because of its drought/frost resistance, high protein
content, and grazing tolerance (Yang et al., 2010).
Poa alpina, together with some other Poa species, have been found susceptible to
AMF colonization worldwide (reviewed by Read & Haselwandter, 1981). Typical
AMF structures (hyphae, vesicles, and arbuscules) are formed, and they can form
double infection with Rhizoctonia in the roots system of P. alpina (Cripps &
Eddington, 2005). However, this symbiosis is highly dependent on geographical
conditions and climate. For instance, it is found that many Poa species are
nonmycorrhizal in Arctic and Antarctic regions, but the same Poa species can be
colonized by AMF in greenhouse condition (Bledsoe et al., 1990; DeMars & Boerner,
1995).
There are a number of methods that can be used to determine AMF colonization level
with advantages and disadvantages. These methods include quantitative real-time
PCR (qPCR) (Alkan et al., 2004; 2006), colorimetric quantification (Becker and
Gerdemann, 1977), chitin and ergosterol analysis (Ekblad et al., 1998), specific fatty
acid analysis (Olsson et al., 1998), and gridline magnified intersect method
20
(Giovannetti and Mosse, 1980; McGonigle et al., 1990).
Using qPCR to detect AMF colonization is a relatively new method, considering its
first advent about only 30 years ago (Deepak et al., 2007). Therefore, it is limited by
some challenges. Firstly, it may take time to design suitable and reliable primer pairs
and protocols. Secondly, when colonization level is very low, qPCR might be
interfered by other substances in DNA extracts. Thirdly, root clearing before DNA
extraction may cause loss of AMF extraradical hyphae and underestimation of AMF
colonization level. Chitin and ergosterol combined measurement and specific fatty
acid analysis method can detect the total amount of living fungal biomass, but they
cannot distinguish the species (Ekblad et al., 1998; Olsson et al, 1998). Colorimetric
quantification is not suitable because phenolic compounds in dying roots could
influence the results. And dark non-mycorrhizal root which might contain yellow
pigment can affect the colorimetric signal (Becker and Gerdemann, 1977).
Gridline intersection method is commonly used to estimate the proportion of infected
root to the total root length without bias (Giovannetti and Mosse, 1980; Brundrett et
al., 1984; McGonigle et al., 1990). Within many visual stains, it has been reported
that Trypan Blue is one of the suitable stains for AMF visualization for its clarity and
accuracy (Abdel-Fattah, 2001; Vierheilig et al., 2005).
The gyrB gene, encoding the subunit B protein of DNA gyrase, can be used as a
phylogenetic marker to detect the presence of B. amyloliquefaciens in the soil
samples (Bavykin et al., 2004). B protein of DNA gyrase is universally distributed in
bacteria species because of its essential role in DNA replication process. The
concentration of B. amyloliquefaciens in the soil can be indirectly reflected by gyrB
gene content (Watt & Hickson, 1994; Huang, 1996). It has also been proven that,
compared with 16S rRNA gene, gyrB gene possesses higher resolution to distinguish
B. amyloliquefaciens from closely related B. subtilis group (Wang et al., 2007). To
detect gene content, qPCR is commonly used (reviewed by Heid et al., 1996).
21
3 Research objectives
The objectives of this research were to find out the survival and developmental
patterns of two artificially inoculated plant-growth-promoting micro-organisms, B.
amyloliquefaciens and G. intraradices, on a green roof in southern Finland. This
research could lay the foundation to further questions on whether the
micro-organisms can facilitate the survival and wellness of green roof plants, and to
what extend the added beneficial micro-organisms contribute to these processes. The
work might give a solution to maintain green roofs at a lower manpower, and enlarge
the selection of green roof plants.
22
4 Material and methods
4.1 Green roof installation and experimental design
The vegetation mats were installed on rooftop of Hong Kong Store in Kuninkaalantie
19, Vantaa. The location and layout of the green roof are presented in Fig. 4. The
total experimental area was 80 m2 including 12 experimental plots (1.5 m×1.5 m), to
which treatments were randomized. 4 plots were inoculated with G. intraradices (M,
abbreviation for MYC4000), 4 with B. amyloliquefaceins (R, abbreviation for
Rhizocell), and 4 were uninoculated as controls (C).
R1
C1
M1
C2
R2
65°
R3
M2
C3
M3
M4
C4
R4
60°
Fig.4. Green roof location and plot design on the green roof. M1-M4: MYC4000, R1-R4: Rhizocell,
C1-C4: Control.
The vegetation mats were purchased from Veg Tech and the seeds from Suomen
Niittysiemen. Soil type used in the mats is sandy loam mixed with gravels (2 mm to
40 mm). Plants on the green roof are stonecrops (Sedum acre/telphium), bluegrasses
(P. alpina), yellow rockets (Barbarea vulgaris), white clover (Trifolium repens),
mullein (Verbascum thapsus), and pennycress (Thlaspi arvense) (Detailed species list
23
is presented in Appendix I). The green roof is composed of 6 layers (Fig. 5).
Green roof vegetation
Green roof substrate (30 mm)
Substrate (30 mm)
Filter and moisture layer (10 mm)
Water retention and drainage (25 mm)
Root barrier (2 mm)
Fig.5. Six composition layers of vegetation mats on the green roof.
4.2 Plant material and inoculants
Various plants were growing on the green roof, and P. alpina was selected to detect
the colonization level of G. intraradices in plant root. Inoculant additives Rhizocell
C (B. amyloliquefaciens) and MYC4000 (G. intraradices) were purchased from
Lallemand Plant Care Company. Greenstim Flow from Oy Faintend Ltd was also
used in green roof soil to helps the plant to maintain proper liquid level in stressful
situations that normally leads to dehydration, drought, and salinity.
For every 1 m2 of green roof field, 1 ml of Greenstim Flow was diluted in 1.5 L
water and spread evenly on the green roof soil of each experimental plots. Similarly,
for every 1 m2 of green roof field, 10 g of Rhizocell and 2 g of MYC4000 were
separately diluted in 1.5 L water. Afterwards, the Rhizocell solution was spread
evenly in random selected R plots, and MYC4000 in M plots (Fig. 3). For control
plots, only the same amount of water was spread. Weather station (Vantage Pro 2,
Davis instruments) was installed to monitor the change of growth conditions on the
roof, including soil moisture, soil temperature, air moisture, air temperature, rainfall,
radiation and wind. Due to the battery failure, data were missing starting from July
3rd for both soil temperature and moisture measurements. Data from July 8th to July
24
30th were recorded by researchers of another project on the same green roof but
different experimental plots. Data from July 3rd to July 8th cannot be provided.
Soil samples were collected four times in each summer. One sample was collected at
5 different spots in one experimental plot, and mixed thoroughly. Roots of 5 P. alpina
plants were collected for observation of AMF colonization in the middle of August in
each plot (Table 2).
Table 2. Time schedule of field work on green roof
Items
Date
Tasks in brief
Green roof
21-22.5.2012
Sedum mat, filter and moisture layer, water
installation
tank layer and waterproof layer established
Irrigation
24-25.5.2012
Thorough irrigation
Inoculation
25.5.2012
Rhizocell (10 g/m2), MYC4000 (2 g/m2) and
Greenstim Flow (1 ml/m2) applied
Weather station
4.6.2012
Weather and soil condition data were recorded
Soil sampling
7.18, 8.9, 8.30, 9.19. 2012
Soil samples collected from six spots in one
5.28, 6.17, 7.8, 7.30. 2013
plot and mixed well before analyzing
18.8.2012
P. alpina plants roots collected from 5 plants
17.8.2013
per plot and stored in 70% ethanol.
Root sampling
4.3 Analysis of Glomus intraradices
4.3.1 Root treatment and staining
The root clearing and staining were carried out according to a modified protocol
(Philips & Hayman, 1970). From each experimental plot, root samples of P. alpina
were collected from five individual plants. After the roots were washed and brushed
so that they didn´t contain any soil, they were stored in 70% ethanol. The root could
be stored in the 70% ethanol for a long period. When needed, the root samples were
removed into 10% KOH solution for one day. After that they were placed in 1.5%
hydrogen peroxide containing 5 ml/l ammonia for two hours. Then the samples were
soaked in 1% hydrochloric acid for another two hours. Finally the samples were held
one hour in 0.02% Trypan Blue solution (solution was made of lactic acid containing
25
63 ml/l glycerol, 63 ml/l water, and 0.02% Trypan Blue) at +75 C˚. After staining the
samples were removed into clear glycerol for storage. Upon microscopic
examination, root samples were attached to the microscopic slide with polyvinyl
alcohol-lactic acid-glycerol (PVLG) solution, which is made of 10 ml/l water, 10 ml/l
lactic acid, 1 ml/l glyserol and 1.66 mg/l polyvinyl alcohol.
4.3.2 Microscopic examination of roots
Stained root samples were put on slides marked with a gridline at magnification
×200. Only the intersection of roots and the vertical crosshair were considered and
observed with caution. The method is shown in Fig. 6. When a vertical crosshair met
the root at point A, horizontal crosshair was rotated along the long axis of the root
until it was vertical to the root. The exit point B was taken as the point of intersection,
where observations were made. Records were made whether the vertical crosshair
cut any arbuscules, vesicles, or hyphae at point B. The categories were marked as
"negative”, “arbascules”, “vesicles”, and “hyphae only”. Afterwards, arbuscular
colonization (AC), vesicular colonization (VC) and hyphal colonization (HC) rate
can be calculated accordingly.
Root
Rotation
B
A
Horizontal
crosshair
Position to which
vertical crosshair
was moved
Vertical
crosshair
Fig.6. A diagram showing how to make observations according to the gridline magnified intersection
method.
26
4.3.3 Available phosphorus content measurement in the soil
Available phosphorous has been substantiated to negatively affect the survival and
growth of G. intraradices. Soil samples were collected in all 12 experimental plots
and sent to Finnish Forest Research Institution (METLA) for soil element content
analysis, using ammonium acetate extraction method (reviewed by Saarela, 2002).
4.4 Analysis of Bacillus amyloliquefaicens
4.4.1 Extraction of total DNA samples
0.25g of soil sample was added to the PowerBead tube containing 60 μl Solution C1
and vortexed vigorously for 15 min. The tube was centrifuged at 10000 ×g for 30 sec
and 450 μl supernatant was transferred to a new tube. Solution C2 (250 μl) was
added to the supernatant, incubated at 4 °C for 5 min, and centrifuged at 10000 ×g
for 1 min. Supernatant (600 μl) was transferred to a new tube and purified with 200
μl Solution C3 as the previous process. Thereafter, 700 μl of supernatant was mixed
with 1.2 ml Solution C4 and centrifuged at 10000 ×g for 1 min. The pellet was
washed with 500 μl Solution C5. The sedimental DNA was dissolved in 100 μl
nuclease free water (NFW). Before stored in -20 °C for further downstream
application, the DNA concentration and purity were measured with Nanodrop 2000
(Appendix II).
To formulate standard curve, total DNA of Rhizocell additive was extracted. 0.25g of
the additive was added into a centrifuge tube containing 400 μl 65 °C AP1 solution
and 4 μl RNase A, and vortexed vigorously. After incubating at 65 °C for 15 min,
130 μl AP2 lysis solution was added. The lysate was centrifuged at the highest speed
for 5 min, and supernatant was transferred to a QIAshredder Mini Spin column and
centrifuged to produce flowthrough. 375 μl of AP3/E solution was added into the
tube, transferred into the DNeasy mini spin column, and centrifuged. The pellet
remaining on the column membrane was washed twice with 500 μl Buffer AW. The
27
pellet was dissolved in 50 μl nuclease free water (NFW). The DNA concentration
was measured with Nanodrop 2000 before storage.
4.4.2 Detection of Bacillus amyloliquefaciens
To detect B. amyloliquefaciens content in the soil, total soil DNA was amplified with
gyrB gene primer pair using PCR, and the products were sent for sequencing to
compare with the gyrB gene sequence of B. amyloliquefaciens. The primer pair was
designed according to gyrB gene sequence from Rhizocell additive (Appendix III).
BaG3F (GTCGACCACTCTTGACGTTACGGTT) and BaG4R (CGATCACTTCAAGATCGGC
CACAG)
were chosen as the forward primer and reverse primer. The size of
amplification product was 94 bp. Phusion PCR reaction system was used to amplify
the DNA fragments. PCR protocol was carried out according to manufacturer’s
instruction (Appendix IV). Afterwards, the final PCR products were sent to
Haartman Institute for sequencing.
4.4.3 Quantification of Bacillus amyloliquefaciens
qPCR technique was used to quantify the amount of B. amyloliquefaciens in the soil
by detection gyrB gene expression. Before qPCR, the soil total DNAs were diluted to
10-14 ng/μl as required (Appendix II). The qPCR materials and protocol were
conducted according to Appendix V. The Rhizocell additive total DNA was diluted
to 1:10, 1:100, and 1: 1000, and then underwent the same qPCR protocol to
formulate the standard curve. For each soil/additive DNA sample, three replicates
were made. The melting curves of qPCR products were checked to make sure only
expected DNA amplicons were amplified. Ct values with melting curves other than
unimodal curves were omitted.
4.4.4 Calculation of Bacillus amyloliquefaciens content in the soil
A standard curve was drawn according to the Rhizocell additive total DNA qPCR
28
result. The equation of Ct value and log of DNA dilution are shown in Fig. 7. The
correlation coefficient equals 1, indicating that the data fits perfectly in the standard
line. Standard curve slope is –3.317, which is in the acceptable range according to
the manual and very close to the standard -3.321, suggesting very high qPCR
efficiency. The calculation process and examples are shown in Fig. 8.
Ct 30
25
20
y = -3,317x + 27,441
R² = 1
15
10
5
0
0
0,5
1
1,5
Log of DNA dilution
2
2,5
Fig.7. Real-Time PCR standard curve of B. amyloliquefaciens.
Data and values
Results
Steps
Calculate gyrB gene conc. in
Ct value
21.11
Soil DNA
diluted soil total DNA
81.03 ng/ul
according to the standard curve
equation
conc. used in
qPCR
10.80 ng/ul
Calculate the dilution rate
of soil total DNA
10.8÷42.7=25.29 %
Original soil
DNA conc.
42.70 ng/ul
Calculate gryB gene conc. in
the original soil total DNA
81.03÷0.2529=320.40 ng/ul
gyrB DNA concentration in the soil
Soil used in
total DNA
extraction
Calculate gyrB gene
conc. in the soil
320.4÷0.25=1281.60 ng/g
0.25 g
Fig.8. Calculation process examples of M2 collected on September 19th.
29
5 Results
5.1 Soil temperature/moisture on green roof during summers
Since the data from other group were obtained by different monitors, and the soil
temperature data from July 8th to July 11th were not authentic, the soil temperature
and moisture data from July 11th to July 30th were only shown in graphs without
further analysis.
5.1.1 Soil temperature
In 2012, soil temperature data were recorded for 71 days (Fig. 9, A). There were 18
days (25.4%) with temperature above 30 °C, 10 days (14.1%) above 35 °C, and 3
days (4.2%) above 40 °C. Daily highest temperature above 35 °C lasted continuously
for 5 days (July 25th-29th). The highest temperature was 41.1 °C on July 4th, 5th and
28th. Average daily highest temperatures of 7 days before each measurements were
26.2, 24.8, 24.1, and 16.8 °C.
In 2013, soil temperature data were recorded for 38 days (Fig. 9, B). There were 27
days (71.1%) with temperature over 30 °C, 22 days (57.9%) over 35 °C, and 11
(28.9%) days above 40 °C. Daily highest temperature above 35 °C lasted
continuously for 11 days (May 31th-June 10th). The highest temperature was 53.9 °C
on June 5th. Average daily highest temperatures of 7 days before two measurements
were 21.1, and 24.4 °C. The average soil temperature in 2013 was 22.5 °C, compared
with 16.8 °C in 2013.
27,05,13
28,05,13
30,05,13
31,05,13
01,06,13
02,06,13
03,06,13
04,06,13
06,06,13
07,06,13
08,06,13
09,06,13
10,06,13
12,06,13
13,06,13
14,06,13
15,06,13
16,06,13
18,06,13
19,06,13
20,06,13
21,06,13
22,06,13
23,06,13
25,06,13
26,06,13
27,06,13
28,06,13
29,06,13
01,07,13
02,07,13
03,07,13
45
40
35
30
25
20
15
10
5
0
-5
70
60
50
40
30
20
10
0
40,0
Date
11.7.2012
14.7.2012
16.7.2012
19.7.2012
21.7.2012
23.7.2012
25.7.2012
27.7.2012
29.7.2012
1.8.2012
3.8.2012
5.8.2012
7.8.2012
9.8.2012
11.8.2012
14.8.2012
16.8.2012
18.8.2012
20.8.2012
22.8.2012
24.8.2012
27.8.2012
29.8.2012
31.8.2012
2.9.2012
4.9.2012
6.9.2012
9.9.2012
11.9.2012
13.9.2012
15.9.2012
30
A
July 19th
Aug. 9th
May 28th
0,0
8.7.2013
13.7.2013
Aug. 30th
50,0
July 8th
18.7.2013
Date
July 30th). Soil sampling time is pointed out in this graph.
23.7.2013
Sep. 19th
Date
B
June 17th
Date
C
60,0
July 30th
30,0
20,0
10,0
28.7.2013
Fig.9. Soil temperature in (A) 2012; (B) 2013 (from May 27th to July 3rd) and (C) 2013 (July 8th to
31
5.1.2 Soil moisture
In 2012, soil moisture data were recorded for 75 days (Fig. 10, A). Centibar (cb) was
used as the unit to measure the water tension in the soil. The sensor can measure the
water tension from 0 to 200 cb. Lower value indicates a high water content in the soil,
and vice versa.
There were three periods when water tension reaching 50 to 70 cb, July 11th to 16th,
July 28th to 29th, and August 20th to 23rd. The graph A of Fig. 9 indicates a continuous
wet soil condition on the green roof. The highest value was 67.3 cb on July 14th.
Average daily soil moistures of 7 days before each measurements were 41.1, 8.5, 7.2,
and 7.2 cb.
In 2013, soil moisture data were recorded for 38 days (Fig. 10, B). Soil moisture was
assumed to be very high before the first measurement. But afterwards, the soil
tension increased steady from below 30 to 200 cb on June 3rd. The extremely
desiccated condition lasted for 11 days until June 14th when rain occurred. The soil
tension remained below 15 cb for 5 days until June 18th before it started to increase
again. The average daily soil moistures of 7 days before the two measurements were
24.3 and 102.1 cb. The average soil moisture in 2013 was 17.7 cb, compared with
94.1 cb in 2013.
32
A
cb
Soil water tension in 2012
60
50
40
July 18th
30
Aug. 9th
Aug. 30th
Sep. 19th
20
10
3.7.2012
5.7.2012
8.7.2012
11.7.2012
15.7.2012
17.7.2012
20.7.2012
23.7.2012
26.7.2012
28.7.2012
31.7.2012
3.8.2012
5.8.2012
8.8.2012
11.8.2012
13.8.2012
16.8.2012
19.8.2012
21.8.2012
24.8.2012
27.8.2012
29.8.2012
1.9.2012
4.9.2012
7.9.2012
9.9.2012
12.9.2012
15.9.2012
0
Date
B
Soil water tension in 2013 (5.27-7.3)
cb 250
200
May 28th
June 17th
150
100
50
27,05,13
28,05,13
30,05,13
31,05,13
01,06,13
03,06,13
04,06,13
05,06,13
07,06,13
08,06,13
09,06,13
11,06,13
12,06,13
13,06,13
15,06,13
16,06,13
17,06,13
18,06,13
20,06,13
21,06,13
22,06,13
24,06,13
25,06,13
26,06,13
28,06,13
29,06,13
30,06,13
02,07,13
03,07,13
0
Date
C
% 0,450
0,400
0,350
0,300
0,250
0,200
0,150
0,100
0,050
0,000
8.7.2013
Soil water content in 2013 (7.8-7.31)
July 30th
July 8th
13.7.2013
18.7.2013
Date
23.7.2013
D
28.7.2013
Fig.10. Soil moisture in (A) 2012; (B) 2013 (from May 27th to July 3rd) and (C) 2013 (July 8th to July
30th). Soil sampling time is pointed out in this graph. The moisture sensor for water tension can
measure in the range from 1 to 200cb. 200cb readings could mean the soil tension was even higher
than 200cb. Soil sampling time is pointed out in this graph.
33
5.2 Available phosphorus in the green roof soil
The available P contents in each experimental plot soil on the green roof were shown
in Table 3. The available P content ranged from 15.8 to 25.9 μg/g soil. The Data for
C3 is missing because the amount of soil sample sent to METLA was too small.
Table 3. Available P content in green roof soil samples.
Samples
P concentration (μg/g)
R1
21.7
R2
20.2
R3
22.6
R4
20.7
M1
23.2
M2
17.0
M3
21.7
M4
20.8
C1
15.8
C2
25.9
C4
22.9
Average
21.1±0.85
5.3 No mycorrhizal structures of Glomus intraradices were found
According to the gridline magnified intersect method, no typical G. intraradices
structures were detected in all groups. Fungus-colonized cells and unknown hyphae
were found in many root samples of both years, and they did not resemble AMF
structures (Fig. 11). It resembles microsclertia of the Phialophora, which is very
common among Poa species (Read & Haselwandter, 1981). Compared with standard
structures and organs of G. intraradices (Fig. 2), AMF was not established in the P.
alpina root system when the roots were collected.
34
Fig.11. Colonized cells found in P. alpina root system.
5.4 Content of Bacillus amyloliquefaciens in the soil
5.3.1 Detection of Bacillus amyloliquefaciens
After amplification, gel electrophoresis was conducted to check the amplification
efficiency and product size. The product size was a little smaller than 100 bp band,
and no other amplified fragments were detected. Rhizocell additive was used as a
positive control. In fact, the PCR products were also obtained from DNA samples of
C and M groups (Fig. 12).
Rhizocell
R
M
C
Ladder
100bp
Fig.12. Detection of B. amyloliquefaciens in soil samples and Rhizocell additive by PCR with gyrB
specific primer pair. The expected size of amplification product is 94 bp. Position of molecular marker
corresponding to 100 bp is indicated in the ladder of marker bands.
The PCR products were sent to Haartman Institute for sequencing using automated
35
chain termination method. The sequences were manually corrected according to
fluorescent graphs and all the four sequences were aligned using ClustalW2 program
(http://www.ebi.ac.uk/Tools/msa/clustalw2/). The results suggested that the PCR
products from all the DNA samples of soil and Rhizocell additive were gyrB gene
fragments (Fig. 13).
Fig.13. Alignment of soil sample DNA and Rhizocell additive DNA sequences.
5.3.2 Bacillus amyloliquefaciens content in soil samples
B. amyloliquefaciens content in the soil was calculated according to Fig. 7, and the
results are shown in Table 4 (detailed data in Appendix VI).
Table 4. B. amyloliquefaciens content (ng/g) in the soil samples.
18th July 2012
9th August2012
30th August2012
19th September 2012
C group
0.3606±0.1377a
0.5787±0.3340a
1.598±1.205a
4.672±2.118a
M group
0.3332±0.1046a
0.8721±0.2648a
2.301±0.9244a
2940±1324b
R group
1.575±0.7082a
0.2586±0.07455a
6.943±3.396a
952.4±251.1b
28th May 2013
17th June 2013
8th July 2013
30th July 2013
C group
5.453±1.450a
data missing
0.04900±0.01243a
0.02453±0.0043a
M group
1.416±0.2753b
0.06778±0.02649a
0.04819±0.008560a
0.02474±0.0068a
R group
0.8098±0.2755b
0.3170±0.1837a
0.1902±0.04762b
0.3839±0.1920b
C: Control group; M: G. intraradices group; R: B. amyloliquefaciens group
Data are presented as the mean±SE (n=4).
Values followed by the same lower-case letter did not differ significantly (p<0.05) by two sample t
test.
36
In 2012, B. amyloliquefaciens content in all soil samples showed increase at the
September 19th measurement to variable degrees, ranging from 10 to 10000 times
(Fig. 14). Non-inoculated control (C group) showed a much smaller increase than M
and R groups. Meanwhile no significant difference between M and R group was
detected (Table 4). B. amyloliquefaciens content in M and C groups exhibited steady
increase, while R group decreased at the second measurement (Fig. 14).
B
ng/g soil
ng/g soil
A
8
7
6
5
4
3
2
1
0
3,5
3
2,5
2
1,5
1
0,5
0
July 18th
July 18th
Aug. 9th Aug. 30th Sep. 19th
Aug. 9th
Aug. 30th
ng/g soil
C
12
10
8
6
4
2
0
July 18th
Aug. 9th
Aug. 30th
Fig.14. Development patterns of C (A), M (B) and R (C) groups in 2012. The last measurement of M
and R group were not shown in B and C graphs because of the extreme large values.
In 2013, B. amyloliquefaciens content in all soil samples besides C group showed a
decline after the winter according to the last measurement in 2012 and the first one in
2013. The decline was more than one thousand folds in M and R groups. The
following four measurements in 2013 showed that C and M groups decreased
continuously, while R group increased again from the third to the last measurement
37
(Fig. 15). B. amyloliquefaciens content in R group was significantly higher than M
and C groups at the P < 0.005 level in the last measurement, while M and C groups
were not significantly different (Fig. 16; Table 4).
ng/g soil
A
0,07
0,06
0,05
0,04
0,03
0,02
0,01
0
June 17th
July. 8th
July. 30th
C
ng/g soil
ng/g soil
B
0,1
0,08
1,2
1
0,8
0,06
0,6
0,04
0,4
0,02
0,2
0
0
June 17th
July 8th
May 28th June 17th July 8th July 30th
July 30th
Fig.15. Development patterns of C (A) and M (B) and R (C) in all measurements in 2013. Data of
June 17th was missing. The last measurements of C and M groups were not shown because of the
ng/g soil
extreme large values.
0,6
0,5
0,4
0,3
0,2
0,1
0
C1
C2
C3
C4
M1
M2
M3
M4
R1
R2
R3
R4
Samples
Fig.16. B. amyloliquefaciens content in all experimental plots at July 30th in 2013
38
5.3.3 Large increase of Bacillus amyloliquefaciens content in M group
In 2012 in M group where only AMF was inoculated, the B. amyloliquefaciens
content reached very high levels, ranging from 211 to 5912 ng/g soil, compared with
R group from 503 to 1672 ng/g soil. According to two sample t test analysis, there
was no significant difference between M and R group (Table 4), but both of them
exceeded control group tens to hundreds times.
39
6 Discussion
6.1 Hypothesis of absence of Glomus intraradices
No typical AMF structures (arbuscules, vesicles, and hyphae) were observed in all
three treatments. But some other micro-organisms resembling microsclertia were
found (Fig. 11). The possible explanations are high phosphorus content in the soil,
high soil temperature, low soil moisture, micro-organism competition, and unsuitable
sampling time.
6.1.1 High phosphorus content might inhibits AMF growth
According to the analysis results from METLA, the available P content in the green
roof soil ranged from 15.8 to 25.9μg/g during the summer in 2012, and the mean
value for 11 samples was 21.1±0.85μg/g (Table 3).
According to Sippola and Yli-Halla (2005), the mean available P content in
agricultural soils in Finland in the year 1998 was 4.9 μg/g. It increased 16% from the
year 1974 to 1987, and 22% from 1987 to 1998, due to heavy application of
fertilizers. It is understandable that in natural soil, the available P concentration can
be much lower. The data show that the P content in the soil on green roof is 3 to 5
times higher than P-rich, agricultural soil.
It has been accepted that high P concentration can negatively affect the colonization
level of mycorrhizal fungi (Abbott et al., 1984; Thomson et al., 1986; Miranda et al.,
1989; Miranda & Harris, 1994; Stevens et al., 2002). Miranda and Harris (1994)
found out the spore germination rate of G. etunicatum was significantly prohibited by
the available P content in the soil, decreasing from 60% to 30%, when the available P
content increased from 0 to 25 μg/g soil. Their experiment result supported the study
of Daniel and Trappe (1980), with a more concrete and meticulous design.
40
Furthermore, high available P content inhibited hyphal growth per spore by 30%
when the available P increased beyond 3 μg/g soil. As great as 83% decrease was
detected in high P concentration when available P increased beyond 25 μg/g soil
(Miranda et al., 1989). Additionally, the number of fungus entry points in root
diminished more than 75% for G. fasciculatum (Thomson et al., 1986).
Bolan’s (1984) studies found that increasing the soil P from severely deficient level
to a low level (1-4 μg/g) results in an increase of mycorrhzial infection before
reaching to a higher P content. It might be explained that, under different P level, the
quality and composition of root exudates are different (Ratnayake et al., 1978;
Schwab et al., 1983). When P content is insufficient, the quality and quantity of root
exudates do not elicit AMF colonization. When reaching desirable P content, the
quality and quantity of root exudates become attractive to AMF. But after reaching
higher P level where reverse effect takes place, the higher P content becomes a
limiting factor that restrains colonization of AMF. A hypothesis has been formulated
that low P content associated with decrease of phospholipid level in the membrane
results in higher root cell permeability. It has been further suggested that under
sufficient P level when low membrane permeability occurs, the root exudates are
reduced. Less root exudates attract less AMF, which leads to decrease of AMF
colonization (Ratnayake et al., 1978; Graham et al., 1981).
In some studies, it was found that higher P concentration increased not only the total
root length (TRL), but also the total AMF-infected root length (TARL) (Thomson et
al., 1986; Miranda et al., 1989). However, their increase was not proportional. TRL
increases more than TARL, which leads to the decrease of AMF colonization rate and
lower incidence to encounter AMF structures under microscope. Therefore, green
roof mat manufacturers might need to reduce P concentration, but still apply proper
amount of P fertilizers. Firstly, P loss through runoff of rainwater occurs naturally.
The depression of AMF by high P would finally be relieved. Secondly, when a green
roof is installed, green roof plants need nutrients to grow and establish. Proper
41
fertilization will help the plants to grow faster and better at their earlier development
stage. Thirdly, even a small amount of AMF colonization could perform plant
beneficial effects without much difference compared with high colonization level. So
reaching high colonization level is advisable but not very crucial (Caron et al., 1986;
Volkmar & Woodbury, 1989; Bødker et al., 1998).
6.1.2 Soil temperature could influences AMF growth
Soil physical conditions not only directly affect growth and development of
micro-organisms, but also indirectly affect them by influencing the well-being of
host plants through plant-microbe interaction. Moisture and temperature are two
main factors that we looked into. And they are not acting solely to affect AMF but
interactively as cofactors (Lekberg & Koide, 2008).
Even though the air temperature scarcely surpassed 25 °C in 2012 and 28 °C in 2013,
the soil temperature constantly reached beyond 35 °C. Especially in the year 2013,
soil temperature over 40 °C lasted for 11 day before the second measurement, and
the highest temperature reached 53.9 °C (Fig. 9). Such a high temperature could be
detrimental to both plants and AMF.
One study conducted with G. intraradices showed that AMF colonization was almost
completely inhibited at 10 °C (10.0%), and remarkably lower at 15 °C (17.4%)
compared with 23 °C (59.2%). Sporulation was reduced significantly at both 10 and
15 °C, along with spore metabolic activity (Liu et al., 2004). Similar results were
obtained in Gavito’s (2005) research.
Negative effect of AMF colonization on plant growth under suboptimal temperature
was reported in scientific papers (Hayman, 1974; Liu et al., 2004). Under suboptimal
temperature condition, the well-being of plants decreases, including decreased root
exudates and carbohydrates. Also the AMF itself could not perform benefits towards
42
host plants under such low temperature (Bowen et al., 1975; Hayman, 1977).
Meanwhile, AMF still utilize limited nutrients from host plants without contribution,
which further enhances the negative effect (Hetrick & Bloom, 1984). It supports the
idea that any adverse environmental conditions affecting the hosts can affect the
AMF.
When temperature exceeds 30°C, root colonization by AMF often decreases (Bowen,
1987). Transformed carrot root organ cultures inoculated by G. intraradices DC1
exhibited steady rise in both colonization rate and extraradical hyphae growth, when
temperature increased from 6 to 30 °C (Gavito et al., 2005). But the effect of higher
temperature on AMF colonization (> 35 °C) was not studied in much detail. Martin
and Stutz (2004) found out G. intraradices colonization reduced dramatically from
42.3% at moderate temperature (minimum 20.7 °C in the dark period and maximum
25.4 °C in the light period) to 20.1% at high temperature (minimum 32.1 °C in the
dark period and maximum 38.0 °C in the light period). Studies showed that G.
intraradices spores can withstand heat treatment of 54-60 °C for up to 1 hour, but
high temperature can seriously reduce root colonization over 40 °C (Schenck et al.,
1975; Bendavid-Val et al., 1997).
6.1.3 Soil moisture could influences AMF growth
Because of the thin layer of soil on the green roof and no shading, it was very
common to get very dry after a long period of sunny days, or get saturated when
there was enough precipitation. In 2013, the drought seemed to be more severe and
lasted for a longer time, when extremely dry condition (> 200 cb) lasted for 11 days
at the beginning of June.
The spore germination is affected significantly by moisture, decreasing from 60%
(moisture level of 50%) to nearly 0% (moisture level of 10%) (Daniels &Trappe,
1980). Shukla et al. (2013) found out that under low P concentration (2.5 μg/g soil),
43
G. intraradices performed higher colonization under full-field capacity (FC = 16%)
than half (FC = 8%) or double (FC =32) field capacity. This result is in accordance
with findings of other research groups who reported too wet or too dry condition
could both reduce AMF colonization (Escudero & Mendoza, 2005).
In a wet land environment, inundation has been found to reduce AMF colonization
level (Stevens et at., 2002; Lumini et al., 2011; Matsumura et al., 2014). Excessive
water in the rhizosphere increases mobility and availability of P, whose ability to
suppress AMF has been profoundly studied. But controversial results have also been
reported that mycorrhization occurs commonly in wetland, and AMF are tolerant to a
wide range of soil moisture (Bohrer et al., 2004). This inconsistence might be
explained in the way that indigenous plant species and/or AMF species that were
adapted to wetland environment.
Under water stress, constrained colonization by AMF was observed (Shukla et al.,
2002; Wu & Xia, 2006; Lekberg & Koide, 2008). It has been suggested that the
decreased entry point for AMF caused by water deficiency could be the underlying
reason (Wu & Xia, 2006). But no studies about the effect of extreme drought
condition on mycorrhization were found. Suggested irrigation based on soil water
suction for different plants were presented, indicating that most plants need to be
irrigated when the soil suction reaches 40 to 60 cb. Otherwise, drought beyond that
range would damage the plants (Taylor, 1965; Hanson et al., 2000). When plants are
not maintaining good physiological condition, AMF growth would be affected by
decreasing nutrients supplied by plants. However, plant species with different water
requirements was suggested as another factor of colonization efficiency in Shukla’s
(2013) work.
6.1.4 Micro-organism competition might leads to AMF absence
The absence of G. intraradices might be attributed to micro-organism competition.
44
However, it needs further examination to find out whether competition with other
micro-organisms contributed to the absence of G. intraradices, or the absence of G.
intraradices led to colonization of other micro-organisms.
6.1.5 AMF was not observed due to the unsuitable sampling time
The roots were collected in the middle of August in both years, when the plants were
almost wilted. It is said that appearance of arbuscules indicates the dead-end growth
of AMF (Bonfante & Perotto, 1995). They senesce and disappear after 4 to 10 days
of symbiosis. When plants wilt, they could not provide enough nutrients to support
the AMF (Sanders et al., 1977). So the absence of G. intraradices might be caused by
unsuitable sampling time.
6.2 Development pattern of Bacillus amyloliquefaciens content
6.2.1 Existence of Bacillus amyloliquefaicens in non-inoculated soil
In C and M groups, B. amyloliquefaicens was detected throughout the experiment,
which suggests either pre-existence of B. amyloliquefaicens in the soil mats or spread
from the R group. Bacillus species are widely distributed in natural soils (Idriss et al.,
2002; Rückert et al., 2011). According to Vardharajula et al. (2011), Bacillus spp.
was detected in non-inoculated soils, but only 1/10000-1/100000 compared with
inoculated ones. Bacterial movement is mostly associated with soil moisture. When
water is adequate in the soil, water film around soil particles is thicker, which
facilitates the movement of soil bacteria, and vice versa (Ongena & Jacques, 2008).
On the green roof, the soil layer is only 3cm, which means that it is very easy to get
saturated after rain. There is a possibility that B. amyloliquefaciens from R plots
moved and drained to other plots. To avoid that situation, when designing experiment,
plots with different inoculants should be kept away from each other, which however
might make comparison of results more difficult if conditions in the whole
experimental areas cannot be controlled and maintained uniform.
45
6.2.2 Soil temperature affects Bacillus amyloliquefaciens reproduction
In 2012, average daily soil highest temperatures of 7 days before each measurements
were 26.2, 24.8, 24.1, and 16.8 °C. It showed a steady decrease at first three
measurements and a sudden drop from the third to the last one. This finding
coincides with the change of B. amyloliquefaciens content in the soil. The content
increased slowly and continuously during the first three measurements. However,
from the third to the fourth measurement, Bacillus content increased 2.9, 1278.3, and
137.2 fold respectively in C, M, and R groups.
In 2013, the average soil temperature was 22.5 °C compared with 16.8 °C in 2012.
The green roof soil temperature reached over 35 °C for 15 days between the first and
second measurement. During the period, B. amyloliquefaciens content dropped from
1.416 to 0.068 ng/g in M group and from 0.810 to 0.317 ng/g in R group (data for the
second measurement in C group was missing). It can be concluded that soil
temperature might be a key factor influencing B. amyloliquefaciens content in the
soil.
It has been established that soil temperature has significant influence on sporulation
and development of soil micro-organisms, including Bacillus species. Aslim et al.
(2002) studied the growth conditions for six Bacillus species isolated from grassland
soil. Optimum growth temperature for them ranged from 28 to 37 °C. The growth
temperature for B. sublitis, a closely related species to B. amyloliquefaciens, was
37 °C. Ahmed et al. (2007) found the optimal growth temperature for B. boroniphilus
sp. nov. was 30 °C, while growth was not observed when temperature exceeded
45 °C. Besides, lower temperature (16 °C) could also inhibit growth.
A research group in South Korea conducted experiments on factors influencing
carboxymethylcellulase production by B. amyloliquefaciens DL-3. They found in
three different cultivation temperatures (32, 37, and 42 °C) that DL-3 grew at 32 °C
46
showed the highest bacterial cell concentration, about twice as much as at 42 °C (Jo
et al., 2008).
Soil temperature in 2012 seldom exceeded 40 °C, but in 2013, the temperature
continuously surpassed 40 °C in the daytime, especially between the first and second
measurements (for 10 days). It might be one of the reasons that B. amyloliquefaciens
content dropped significantly between the first two measurement in 2013 (Table 4).
Microbial biomass in soil experiences major shift between summer and winter,
mainly because of temperature change (reviewed by Lipson et al., 2004). It might be
the reason for the large decrease detected between September 19th 2012 and May 28th
2013.
6.2.3 Soil moisture affects Bacillus amyloliquefaciens reproduction
In 2012, when the soil was sampled, the soil was high in water content. Before each
of the first three measurements, the soil water tension reached only slightly above 50
cb (Fig. 10). It might be under this high moisture level that Bacillus content grew
steadily in the first three measurements. In the last measurement, the soil kept wet
(<15 cb) for 28 days. This long period of moist soil condition might give rise to the
huge increase of Bacillus. The average soil moisture of 7 days before each
measurements were 32.9, 8.2, 6.7, and 6.6 cb.
In 2013, before the first measurement, the soil water tension remained low (<30 cb).
Then, it reached 200 cb for over 11 days, indicating a long period of extremely dry
soil condition. This might explained Bacillus content drop between first and second
measurements.
Along with soil temperature, soil moisture plays an important role in growth and
development of soil micro-organisms (Kilian, et al., 2000; Babalola, 2010; Song &
Lee, 2010). Under natural conditions, the survival and germination of spores of
47
Bacillus species do not show any decrease at low soil moisture, due to their extreme
resistance towards desiccation (Petras & Casida, 1985; Koike et al., 1992). But the
growth and development of Bacillus is depended upon water availability.
Some Bacillus species could tolerate minimal water potential at -0.73 Mpa (-730 cb).
Vardharajula et al. (2011) found a tenfold decrease in population of a drought
resistant B. amyloliquefaciens species, HYD-B17, under drought-stress condition at
46.6% water holding capacity (WHC) as compared with non-drought stress at 75%
WHC (soil type unknown). Even inoculated with drought resistant Bacillus species,
they could not survive beyond nine days. In solid state fermentation industry,
maintaining high moisture level is crucial for B. amyloliquefaciens production
(Prakasham et al., 2006).
6.2.4 Resistant Bacillus amyloliqufaciens from Rhizocell additive survived through
winter
In 2012, the B. amyloliquefaicens content in both R and M group increased
dramatically at the last measurement, even though no B. amyloliquefaicens was
inoculated in M group. But in the second year, only R group inoculated with
Rhizocell additive increased again, and the absolute values were significantly higher
than C and M groups (Fig. 16).
The hypothesis is formulated that B. amyloliqufaciens from commercial Rhizocell
additive might be more resistant to environmental stress than the existing ones in the
soil. During the winter, B. amloliquefaciens population in all groups declined
because of the cold temperature and water unavailability. But the environmental
pressure selected more robust species, strains or individuals. So in the following
spring and summer, the indigenous B. amyloliquefaciens in C and M group could not
survive through winter and then decreased continuously. Meanwhile, some of the
resistant ones from Rhizocell additive in R group survived through the winter,
48
fluctuated in its content and increased again in the last two measurements in 2013.
6.2.5 Synergetic effect between the two micro-organisms
At the last measurement in 2012, both M and R groups showed an enormous increase
in B. amyloliquefaciens content, even though no Rhizocell was added in M group.
Synergistic effect of micro-organisms might hold the key to the answer.
It has been reported repeatedly that AMF and nitrogen fixing bacteria have either
beneficial or negative effect towards each other to various extent when they are
inoculated and established within the same host plant (Mansfeld-Giese et al., 2002;
Medina et al., 2003; Artursson et al., 2006).
On one hand, laboratory experiments have revealed that Bacillus species can
promote fungal growth, spore germination, and root colonization of AMF (Artursson
et al., 2006). For instance, B. cereus was observed attaching closely to hyphae of G.
dussii at a significantly higher level than non-mycorrhizal zone. But the underlying
mechanism was unspecific (Artursson & Jansson, 2003). On the other hand, AMF
has been claimed to promote nitrogen fixing bacteria such as Bacillus species.
Medina et al. (2003) found that when co-inoculated with three Glomus species,
Sinorhizobium meliloti developed more nodules. Within the three Glomus species, G.
intraradices showed the highest performance. The efficiency of synergistic effect
largely depends on the combination of AMF and bacteria, and even the
fungi-bacteria-plant three way interactions.
49
7 Conclusions and future perspectives
The absence of AMF in the plants of P. alpina was probably attributed to various
factors including high P level, extreme soil temperature and moisture,
micro-organism competition, and unsuitable sampling time. Even though no AMF
structures were found in the experiment, it does not necessarily mean that there was
no AMF colonization at all. Further studies should be carried out with some
modifications in order to assess AMF development in the third year. Firstly, with the
passage of time, the P is slowly flowing away through runoff water. This process
might relive AMF from growth inhibited situation imposed by high P concentration.
Secondly, soil temperature and soil moisture situation could be significantly different
in each year, as the weather data in 2012 and 2013 shows. Thirdly, multiple
samplings at earlier time may increase the chance of AMF detection.
The survival and preliminary establishment of B. amyloliquefaciens in green roof soil
was confirmed, even though large fluctuation between the year 2012 and 2013 was
recorded. As high soil temperature and low moisture were crucial factors affecting
the abundance of B. amyloliquefaciens, irrigation on the green roof is suggested to
relieve heat and drought condition. Thicker soil layer could also be applied, if
possible, to mitigate sudden saturation and desiccation. Moreover, if the synergistic
effect between B. amyloliquefaciens and G. intraradices is verified in the future
studies, co-inoculation of both micro-organisms might result in higher colonization
level of both micro-organisms.
In the detection of B. amyloliquefaciens content in soil, the qPCR standard curve was
formulated with Rhizocell additive DNA instead of pure bacterial culture DNA. The
DNA mixture extracted from the additive contained other DNAs including yeast. In
that case, the outcome of the B. amyloliquefaciens content was overestimated. It
might be the reason why some of the target DNA concentrations were higher than the
50
original input DNA concentration (Fig. 8). However, it still gave a relative value for
comparison, conclusion, and formulation of development patterns.
In experimental design, micro-organism inventory should have been conducted in
beforehand to see whether B. amyloliquefaciens and G. intraradices were existing in
the green roof soil. It helps to explain background noise. The distance between plots
should be larger to avoid micro-organisms moving to control groups. And, in order to
achieve AMF survival under extreme roof conditions, strains or species resistant
towards drought, wet, high temperature/solar-radiation, and freeze should be
considered before inoculation.
After confirming the existence of inoculated beneficial micro-organisms, the next
work is to study whether the micro-organisms contribute to the well-being of green
roof plants, and if they do, to what extent.
51
8 Acknowledgements
First of all, I would like to extend my gratitude to my master thesis supervisor, Prof.
Jari Valkonen, for introducing me to the Fifth Dimension Group so that I can work in
this very interesting and thought-provoking project. As the Fifth Dimension Group is
a multidisciplinary research team, I could work with experts from different fields
including agro-engineering, ecology, environmental science, social science and
economics. I would like to thank Jari, Susanna Lehvävirta and Marja Mesimäki for
their instructions and suggestions on my experiment. They helped me with
establishing the outline and direction of the thesis, which was the first step towards
success. I am so thankful for the efforts and concern Jari put in my thesis even
though he was quite occupied himself.
Secondly, I would like to say thank you to my master program professor Paula
Elomaa and her strawberry research team. She instructed me in building up
fundamental knowledge that was basis for me to conduct this master thesis. In her
strawberry research group, led by Timo Hytönen, I gained lab work experience
during summer training in 2012. My thanks go to Takeshi, and Elli for their teaching
and help in the summer training with patience, even when I made mistakes and
increased the workload. They let me realize that lab work should not be done without
caution and devotion.
Thirdly, I am very thankful to KPAT group members, Eeva, Delfia, Layla, Mikko,
Tian, Shahid, Bahram, and Iman, who offered their help and knowledge to me to
solve my numerous problems during the experiments. Also many thanks goes to Dr.
Sari Timonen who offered plenty of help in identification of mycorrhizal fungi from
my samples. Moreover, she provided me with PhD recruiting information and
encouraged me to march forward on the academic road. Even though I failed her and
did not get the position, I am still appreciated all she did for me, and I will never give
52
up on my way to academic research.
Finally, words fail to express my love to my beloved parents, relatives, classmates
and my love Zhao Chen for having always been keeping me company regardless of
ups and downs. They are my strong support.
Thanks to all of you who appeared in my life. Hope you have a great and productive
life.
53
References
Abbott, L., Robson, A. & De Boer, G. 1984. The Effect of Phosphorus on the
Formation of Hyphae in Soil by the Vesicular-Arbuscular Mycorrhizal Fungus,
Glomus fasciculatum. New Phytologist 97: 437-446.
Abdel-Fattah, G. M. 2001. Measurement of the viability of arbuscular-mycorrhizal
fungi using three different stains; relation to growth and metabolic activities of
soybean plants. Microbiological Research 156: 359-367.
Ahmed, I., Yokota, A. & Fujiwara, T. 2007. A novel highly boron tolerant
bacterium, Bacillus boroniphilus sp. nov., isolated from soil, that requires boron
for its growth. Extremophiles 11: 217-224.
Akimoto, H. 2003. Global air quality and pollution. Science 302: 1716-1719.
Alkan, N., Gadkar, V., Coburn, J., Yarden, O. & Kapulnik, Y. 2004. Quantification
of the arbuscular mycorrhizal fungus Glomus intraradices in host tissue using
real-time polymerase chain reaction. New Phytologist 161: 877-885.
Alkan, N., Gadkar, V., Yarden, O. & Kapulnik, Y. 2006. Analysis of quantitative
interactions between two species of arbuscular mycorrhizal fungi, Glomus
mosseae and G. intraradices, by real-time PCR. Applied and Environmental
Microbiology 72: 4192-4199.
Allen, M. F., Moore Jr, T. S. & Christensen, M. 1980. Phytohormone changes in
Bouteloua gracilis infected by vesicular-arbuscular mycorrhizae: I. Cytokinin
increases in the host plant. Canadian Journal of Botany 58: 371-374.
Aroca, R., Porcel, R. & Ruiz-Lozano, J. M. 2007. How does arbuscular mycorrhizal
symbiosis regulate root hydraulic properties and plasma membrane aquaporins in
Phaseolus vulgaris under drought, cold or salinity stresses? New Phytologist
173: 808-816.
Artursson, V., Finlay, R. D. & Jansson, J. K. 2006. Interactions between arbuscular
mycorrhizal fungi and bacteria and their potential for stimulating plant growth.
Environmental Microbiology 8: 1-10.
Artursson, V. & Jansson, J. K. 2003. Use of bromodeoxyuridine immunocapture to
identify active bacteria associated with arbuscular mycorrhizal hyphae. Applied
and Environmental Microbiology 69: 6208-6215.
54
Aslim, B., Yuksekdag, Z. & Beyatli, Y. 2002. Determination of some properties of
Bacillus isolated from soil. Turkish Journal of Biotechnology 26: 41-48.
Babalola, O. O. 2010. Beneficial bacteria of agricultural importance. Biotechnology
Letters 32: 1559-1570.
Bago, B., Cano, C., Azcón-Aguilar, C., Samson, J., Coughlan, A. P. & Piché, Y.
2004. Differential morphogenesis of the extraradical mycelium of an arbuscular
mycorrhizal fungus grown monoxenically on spatially heterogeneous culture
media. Mycologia 96: 452-462.
Bago, B., Azcón-Aguilar, C. & Piché, Y. 1998. Architecture and developmental
dynamics of the external mycelium of the arbuscular mycorrhizal fungus Glomus
intraradices grown under monoxenic conditions. Mycologia 90: 52-62.
Balestrini, R., Romera, C., Puigdomenech, P. & Bonfante, P. 1994. Location of a
cell-wall hydroxyproline-rich glycoprotein, cellulose and β-1, 3-glucans in apical
and differentiated regions of maize mycorrhizal roots. Planta 195: 201-209.
Bavykin, S. G., Lysov, Y. P., Zakhariev, V., Kelly, J. J., Jackman, J., Stahl, D. A. &
Cherni, A. 2004. Use of 16S rRNA, 23S rRNA, and gyrB gene sequence analysis
to determine phylogenetic relationships of Bacillus cereus group
micro-organisms. Journal of Clinical Microbiology 42: 3711-3730.
Becker, W. & Gerdemann, J. 1977. Colorimetric quantification of vesicular
-arbuscular mycorrhizal infection in onion. New Phytologist 78: 289-295.
Bendavid-Val, R., Rabinowitch, H., Katan, J. & Kapulnik, Y. 1997. Viability of
VA-mycorrhizal fungi following soil solarization and fumigation. Plant and Soil
195: 185-193.
Benitez, L. B., Velho, R. V., de Souza, M. A., Segalin, J. & Brandelli, A. 2012.
Antimicrobial factor from Bacillus amyloliquefaciens inhibits Paenibacillus
larvae, the causative agent of American foulbrood. Archives of Microbiology
194: 177-185.
Bernstein, J. A., Alexis, N., Barnes, C., Bernstein, I. L., Nel, A., Peden, D.,
Diaz-Sanchez, D., Tarlo, S. M. & Williams, P. B. 2004. Health effects of air
pollution. Journal of Allergy and Clinical Immunology 114: 1116-1123.
Biermann, B. & Linderman, R. 1983. Use of vesicular-arbuscular mycorrhizal roots,
intraradical vesicles and extraradical vesicles as inoculum. New Phytologist 95:
97-105.
55
Blancaflor, E., Zhao, L. & Harrison, M. 2001. Microtubule organization in root cells
of Medicago truncatula during development of an arbuscular mycorrhizal
symbiosis with Glomus versiforme. Protoplasma 4: 217.
Bledsoe, C., Klein, P., & Bliss, L. C. 1990. A survey of mycorrhizal plants on
Truelove Lowland, Devon Island, NWT, Canada. Canadian Journal of Botany 68:
1848-1856.
Bødker, L., Kjøller, R. & Rosendahl, S. 1998. Effect of phosphate and the arbuscular
mycorrhizal fungus Glomus intraradices on disease severity of root rot of peas
(Pisum sativum) caused by Aphanomyces euteiches. Mycorrhiza 8: 169-174.
Bohrer, K. E., Friese, C. F. & Amon, J. P. 2004. Seasonal dynamics of arbuscular
mycorrhizal fungi in differing wetland habitats. Mycorrhiza 14: 329-337.
Bolan, N., Robson, A. & Barrow, N. 1984. Increasing phosphorus supply can
increase the infection of plant roots by vesicular-arbuscular mycorrhizal fungi.
Soil Biology and Biochemistry 16: 419-420.
Bolund, P. & Hunhammar, S. 1999. Ecosystem services in urban areas. Ecological
Economics 29: 293-301.
Bonfante, P. & Perotto, S. 1995. Tansley Review No. 82 Strategies of arbuscular
mycorrhizal fungi when infecting host plants. New Phytologist 130: 3-21.
Bothe, H., Klingner, A., Kaldorf, M., Schmitz, O., Esch, H., Hundeshagen, B. &
Kernebeck, H. 1994. Biochemical approaches to the study of plant-fungal
interactions in arbuscular mycorrhiza. Experientia 50: 919-925.
Bowen, G. D., Bevege, D. I., Mosse B. 1975. Phosphate physiology of vesicular
arbuscular mycorrhizas. In: Sanders, F. E., Mosse, B., Thinker, P. B. (eds.).
Endomycorrhizas. Academic, New York, USA. 241-260.
Bowen, G. D. 1987. The biology and physiology of infection and its development. In
Safir G. R. (ed.). Ecophysiology of VA mycorrhizal plants. CRC, Boca Raton,
Florida, USA. 27-70.
Brundrett, M., Piche, Y. & Peterson, R. 1984. A new method for observing the
morphology of vesicular-arbuscular mycorrhizae. Canadian Journal of Botany
62: 2128-2134.
Bücking, H. & Shachar-Hill, Y. 2005. Phosphate uptake, transport and transfer by
the arbuscular mycorrhizal fungus Glomus intraradices is stimulated by
increased carbohydrate availability. New Phytologist 165: 899-912.
56
Burkett-Cadena, M., Kokalis-Burelle, N., Lawrence, K. S., van Santen, E. &
Kloepper, J. W. 2008. Suppressiveness of root-knot nematodes mediated by
rhizobacteria. Biological Control 47: 55-59.
Caron, M., Fortin, J. & Richard, C. 1986. Effect of phosphorus concentration and
Glomus intraradices on Fusarium crown and root rot of tomatoes.
Phytopathology 76: 942-946.
Chen, Y., Cao, S., Chai, Y., Clardy, J., Kolter, R., Guo, J. & Losick, R. 2012. A
Bacillus subtilis sensor kinase involved in triggering biofilm formation on the
roots of tomato plants. Molecular Microbiology 85: 418-430.
Cole, M. A. & Neumayer, E. 2004. Examining the impact of demographic factors on
air pollution. Population and Environment 26: 5-21.
Compant, S., Christophe C., and Angela S. 2010. Plant growth-promoting bacteria in
the rhizo-and endosphere of plants: their role, colonization, mechanisms
involved and prospects for utilization. Soil Biology and Biochemistry 42:
669-678.
Cripps, C. L. & Eddington, L. H. 2005. Distribution of mycorrhizal types among
alpine vascular plant families on the Beartooth Plateau, Rocky Mountains, USA,
in reference to large-scale patterns in arctic-alpine habitats. Arctic, Antarctic, and
Alpine Research 37: 177-188.
Dalal, R. 1977. Soil organic phosphorus. Advances in Agronomy 29: 83-117.
Daniel, B. A., & Trappe, J. M. 1980. Factors affecting spore germination of the
vesicular-arbuscular mycorrhizal fungus. Glomus Epigaeus Mycologia 72:
457-471.
Deepak, S., Kottapalli, K., Rakwal, R., Oros, G., Rangappa, K., Iwahashi, H., Masuo,
Y. & Agrawal, G. 2007. Real-time PCR: revolutionizing detection and
expression analysis of genes. Current Genomics 8: 234-251.
DeMars, B. G. & Boerner R. E. J. 1995. Mycorrhizal status of Deschampsia
antarctica in the Palmer Station area, Antarctica. Mycologia 87: 451-453.
DeNardo, J., Jarrett, A., Manbeck, H., Beattie, D. & Berghage, R. 2005. Stormwater
mitigation and surface temperature reduction by green roofs. Transactions of the
ASAE 48: 1491-1496.
Desoignies, N., Schramme, F., Ongena, M., Legrève, A. & Merz, U. 2011. Control of
Polymyxa betae by Bacillus amyloliquefaciens lipopeptides. In: Merz, U. (ed.).
57
(Proceedings of the 8th Symposium of the International Working Group on Plant
Viruses with Fungal Vectors, 6-8 July, 2011, Louvain-La-Neuve, Belgium).
Zurich, Germany: Plant Pathology, ESci Journals Publishing Ltd. s. 59-62.
Ekblad, A., Wallander, H. & Näsholm, T. 1998. Chitin and ergosterol combined to
measure total and living fungal biomass in ectomycorrhizas. New Phytologist
138: 143-149.
Errington, J. 2003. Regulation of endospore formation in Bacillus subtilis. Nature
Reviews Microbiology 1: 117-126.
Escudero, V. & Mendoza, R. 2005. Seasonal variation of arbuscular mycorrhizal
fungi in temperate grasslands along a wide hydrologic gradient. Mycorrhiza 15:
291-299.
Feng, G., Song, Y., Li, X. & Christie, P. 2003. Contribution of arbuscular
mycorrhizal fungi to utilization of organic sources of phosphorus by red clover
in a calcareous soil. Applied Soil Ecology 22: 139-148.
Flora of North America Editorial Committee. 2007. Flora of North America: North
of Mexico Volume 24: Magnoliophyta: Commelinidae (in part): Poaceae, part 1.
Oxford, UK: Oxford University Press. 944 s.
Fox, K. & Eder, B. D. 1969. Comparison of survivor curves of Bacillus subtilis
spores subjected to wet and dry heat. Journal of Food Science 34: 518-521.
Fukumoto, J. 1943. Studies on the production of bacterial amylase. I. isolation of
bacteria secreting potent amylase and their distribution. Journal of the
Agricultural Chemical Society of Japan 19: 643-650.
Gaur, A. C. & Ostwal, K. P. 1972. Influence of phosphate dissolving Bacilli on yield
and phosphate uptake of wheat crop. Indian Journal of Experimental Biology 10:
393-394.
Gavito, M. E., Olsson, P. A., Rouhier, H., Medina-Peñafiel, A., Jakobsen, I., Bago,
A. & Azcón-Aguilar, C. 2005. Temperature constraints on the growth and
functioning of root organ cultures with arbuscular mycorrhizal fungi. New
Phytologist 168: 179-188.
Genre, A. & Bonfante, P. 1998. Actin versus tubulin configuration in
arbuscule-containing cells from mycorrhizal tobacco roots. New Phytologist 140:
745-752.
58
Genre, A. & Bonfante, P. 1997. A mycorrhizal fungus changes microtubule
orientation in tobacco root cells. Protoplasma 199: 30-38.
Giovannetti, M., & Mosse, B. 1980. An evaluation of techniques for measuring
vesicular arbuscular mycorrhizal infection in roots. New phytologist 84:
489-500.
González-Guerrero, M., Melville, L. H., Ferrol, N., Lott, J. N., Azcon-Aguilar, C. &
Peterson, R. L. 2008. Ultrastructural localization of heavy metals in the
extraradical mycelium and spores of the arbuscular mycorrhizal fungus Glomus
intraradices. Canadian Journal of Microbiology 54: 103-110.
Gordon, R., Haynes, W., & Pang, C. 1973. The genus bacillus. US Department of
Agriculture handbook 427: 109-26.
Graham, J. H., Leonard, R. T. & Menge, J. A. 1981. Membrane-mediated decrease in
root exudation responsible for phorphorus inhibition of vesicular-arbuscular
mycorrhiza formation. Plant Physiology 68: 548-552.
Grimm, N. B., Faeth, S. H., Golubiewski, N. E., Redman, C. L., Wu, J., Bai, X. &
Briggs, J. M. 2008. Global change and the ecology of cities. Science 319:
756-760.
Hanson, B., Orloff, S. & Peters, D. 2000. Monitoring soil moisture helps refine
irrigation management. California Agriculture 54: 38-42.
Hayman, D. 1974. Plant growth responses to vesiculararbuscular mycorrhiza. New
Phytologist 73: 71-80.
Hayman, D. 1977. Mycorrhizal effects on white clover in relation to hill land
improvement. ARC Research Review 3: 82-85.
Heid, C. A., Stevens, J., Livak, K.J. & Williams, P.M. 1996. Real time quantitative
PCR. Genome Research 6: 986-994.
Henry, A. & Frascaria-Lacoste, N. 2012. The green roof dilemma–Discussion of
Francis and Lorimer. Journal of Environmental Management 104: 91-92.
Hetrick, B. D. & Bloom, J. 1984. The influence of temperature on colonization of
winter wheat by vesicular-arbuscular mycorrhizal fungi. Mycologia 76: 953-956.
Hildebrandt, U., Regvar, M. & Bothe, H. 2007. Arbuscular mycorrhiza and heavy
metal tolerance. Phytochemistry 68: 139-146.
59
Hu, H., Li, X. & He, H. 2010. Characterization of an antimicrobial material from a
newly isolated Bacillus amyloliquefaciens from mangrove for biocontrol of
Capsicum bacterial wilt. Biological Control 54: 359-365.
Huang, W. M. 1996. Bacterial diversity based on type II DNA topoisomerase genes.
Annual Review of Genetics 30: 79-107.
Huang, X., Zhao, N. & Zhang, K. 2004. Extracellular enzymes serving as virulence
factors in nematophagous fungi involved in infection of the host. Research in
Microbiology 155: 811-816.
Idris, E. E., Iglesias, D. J., Talon, M. & Borriss, R. 2007. Tryptophan-dependent
production of indole-3-acetic acid (IAA) affects level of plant growth promotion
by Bacillus amyloliquefaciens FZB42. Molecular Plant-Microbe Interactions 20:
619-626.
Idriss, E. E., Makarewicz, O., Farouk, A., Rosner. K., Greiner, R., Bochow, H.,
Richter, T. & Borriss, R. 2002. Extracellular phytase activity of Bacillus
amyloliquefaciens FZB45 contributes to its plant-growth-promoting effect.
Microbiology 148: 2097-2109.
Jo, K., Lee, Y., Kim, B., Lee, B., Chung, C., Nam, S., Kim, S. & Lee, J. 2008.
Pilot-scale production of carboxymethylcellulase from rice hull by Bacillus
amyloliquefaciens DL-3. Biotechnology and Bioprocess Engineering 13:
182-188.
Kasim, W. A., Osman, M. E., Omar, M. N., El-Daim, I. A. A., Bejai, S. & Meijer, J.
2013. Control of drought stress in wheat using plant-growth-promoting bacteria.
Journal of Plant Growth Regulation 32: 122-130.
Katsouyanni, K. & Pershagen, G. 1997. Ambient air pollution exposure and cancer.
Cancer Causes & Control 8: 284-291.
Khalil, S., Loynachan, T. E. & Tabatabai, M. A. 1994. Mycorrhizal dependency and
nutrient uptake by improved and unimproved corn and soybean cultivars.
Agronomy Journal 86: 949-958.
Kilian, M., Steiner, U., Krebs, B., Junge, H., Schmiedeknecht, G. & Hain, R. 2000.
FZB24® Bacillus subtilis-mode of action of a microbial agent enhancing plant
vitality. Pflanzenschutz-Nachrichten Bayer 53: 72-93.
Kim, J. J., Smorodinsky, S., Lipsett, M., Singer, B. C., Hodgson, A. T. & Ostro, B.
2004. Traffic-related air pollution near busy roads: the East Bay Children's
60
Respiratory Health Study. American Journal of Respiratory and Critical Care
Medicine 170: 520-526.
Koide, R. & Kabir, Z. 2000. Extraradical hyphae of the mycorrhizal fungus Glomus
intraradices can hydrolyse organic phosphate. New Phytologist 148: 511-517.
Koike, J., Oshima, T., Koike, K., Taguchi, H., Tanaka, R., Nishimura, K. & Miyaji,
M. 1992. Survival rates of some terrestrial micro-organisms under simulated
space conditions. Advances in Space Research 12: 271-274.
Kloepper, J. W., Ryu, C. M., & Zhang, S. 2004. Induced systemic resistance and
promotion of plant growth by Bacillus spp. Phytopathology 94: 1259-1266.
Krasilnikov, M. 1962. On the role of soil bacteria in plant nutrition. Journal of
General and Applied Microbiology 7: 128-144.
Lekberg, Y. & Koide, R. 2008. Effect of soil moisture and temperature during fallow
on survival of contrasting isolates of arbuscular mycorrhizal fungi. Botany 86:
1117-1124.
Li, B., Xu, L., Lou, M., Li, F., Zhang, Y. & Xie, G. 2008. Isolation and
characterization of antagonistic bacteria against bacterial leaf spot of Euphorbia
pulcherrima. Letters in Applied Microbiology 46: 450-455.
Lian, L. H., Tian, B. Y., Xiong, R., Zhu, M. Z., Xu, J., & Zhang, K. Q. 2007.
Proteases from Bacillus: a new insight into the mechanism of action for
rhizobacterial suppression of nematode populations. Letters in applied
microbiology 45: 262-269.
, D. A. & Schmidt, S. K. 2004. Seasonal changes
in an alpine soil bacterial community in the Colorado Rocky Mountains. Applied
and Environmental Microbiology 70: 2867-2879.
Liu, A., Wang, B. & Hamel, C. 2004. Arbuscular mycorrhiza colonization and
development at suboptimal root zone temperature. Mycorrhiza 14: 93-101.
Liu, Z., Budiharjo, A., Wang, P., Shi, H., Fang, J., Borriss, R., Zhang, K. & Huang,
X. 2013. The highly modified microcin peptide plantazolicin is associated with
nematicidal activity of Bacillus amyloliquefaciens FZB42. Applied
Microbiology and Biotechnology 97: 10081-10090.
Logan, N. & Berkeley, R. 1984. Identification of Bacillus strains using the API
system. Journal of General Microbiology 130: 1871-1882.
61
López, D., Fischbach, M. A., Chu, F., Losick, R., & Kolter, R. 2009. Structurally
diverse natural products that cause potassium leakage trigger multicellularity in
Bacillus subtilis. Proceedings of the National Academy of Sciences 106:
280-285.
Lumini, E., Vallino, M., Alguacil, M. M., Romani, M. & Bianciotto, V. 2011.
Different farming and water regimes in Italian rice fields affect arbuscular
mycorrhizal fungal soil communities. Ecological Applications 21: 1696-1707.
Mage, D., Ozolins, G., Peterson, P., Webster, A., Orthofer, R., Vandeweerd, V. &
Gwynne, M. 1996. Urban air pollution in megacities of the world. Atmospheric
Environment 30: 681-686.
Maget-Dana, R., Thimon, L., Peypoux, F. & Ptak, M. 1992. Surfactin/iturin A
interactions may explain the synergistic effect of surfactin on the biological
properties of iturin A. Biochimie 74: 1047-1051.
Maheshwari, D. K. 2011. Bacteria in agrobiology. New York, USA: Springer Ltd.
370 s.
Makarewicz, O., Dubrac, S., Msadek, T., & Borriss, R. 2006. Dual role of the PhoP∼
P response regulator: Bacillus amyloliquefaciens FZB45 phytase gene
transcription is directed by positive and negative interactions with the phyC
promoter. Journal of bacteriology188: 6953-6965.
Mansfeld-Giese, K., Larsen, J. & Bødker, L. 2002. Bacterial populations associated
with mycelium of the arbuscular mycorrhizal fungus Glomus intraradices.
FEMS Microbiology Ecology 41: 133-140.
Margosch, D., Ehrmann, M. A., Buckow, R., Heinz, V., Vogel, R. F. & Gänzle, M.
G. 2006. High-pressure-mediated survival of Clostridium botulinum and Bacillus
amyloliquefaciens endospores at high temperature. Applied and Environmental
Microbiology 72: 3476-3481.
Margosch, D., Gänzle, M. G., Ehrmann, M. A. & Vogel, R. F. 2004. Pressure
inactivation of Bacillus endospores. Applied and Environmental Microbiology
70: 7321-7328.
Martin, C. A. & Stutz, J. C. 2004. Interactive effects of temperature and arbuscular
mycorrhizal fungi on growth, P uptake and root respiration of Capsicum annuum
L. Mycorrhiza 14: 241-244.
Matsumura, A., Hattori, R., Yamawaki, K., Tarui, A. & Daimon, H. 2014. Excess
soil water impact on colonization and extraradical mycelium biomass production
62
of arbuscular mycorrhizal fungi in soybean field. International Journal of
Agricultural Policy and Research 2: 41-48.
McGonigle, T., Miller, M., Evans, D., Fairchild, G. & Swan, J. 1990. A new method
which gives an objective measure of colonization of roots by
vesicular-arbuscular mycorrhizal fungi. New Phytologist 115: 495-501.
Medina, A., Probanza, A., Mañero, F. G. & Azcón, R. 2003. Interactions of
arbuscular-mycorrhizal fungi and Bacillus strains and their effects on plant
growth, microbial rhizosphere activity (thymidine and leucine incorporation) and
fungal biomass (ergosterol and chitin). Applied Soil Ecology 22: 15-28.
Mendoza, A. R., Kiewnick, S. & Sikora, R. A. 2008. In vitro activity of Bacillus
firmus against the burrowing nematode Radopholus similis, the root-knot
nematode Meloidogyne incognita and the stem nematode Ditylenchus dipsaci.
Biocontrol Science and Technology 18: 377-389.
Miranda, J. D. & Harris, P. 1994. Effects of soil phosphorus on spore germination
and hyphal growth of arbuscular mycorrhizal fungi. New Phytologist 128:
103-108.
Miranda, J. D., Harris, P. & Wild, A. 1989. Effects of soil and plant phosphorus
concentrations on vesicular-arbuscular mycorrhiza in sorghum plants. New
Phytologist 112: 405-410.
Morancho, A. B. 2003. A hedonic valuation of urban green areas. Landscape and
Urban Planning 66: 35-41.
Murphy, J. F., Zehnder, G. W., Schuster, D. J., Sikora, E. J., Polston, J. E. &
Kloepper, J. W. 2000. Plant growth-promoting rhizobacterial mediated
protection in tomato against Tomato mottle virus. Plant Disease 84: 779-784.
Myllys, L. Stenroos, S. & Huhtinen, S. 2013. Sienten luokittelu ja evoluutio. In:
Sienten biologia, Eds. Timonen S & Valkonen J. Gaudeamus, Helsinki: 11-47.
Nautiyal, C. S., Srivastava, S., Chauhan, P. S., Seem, K., Mishra, A. & Sopory, S. K.
2013. Plant growth-promoting bacteria Bacillus amyloliquefaciens NBRISN13
modulates gene expression profile of leaf and rhizosphere community in rice
during salt stress. Plant Physiology and Biochemistry 66: 1-9.
Nicholson, W. L., Munakata, N., Horneck, G., Melosh, H. J. & Setlow, P. 2000.
Resistance of Bacillus endospores to extreme terrestrial and extraterrestrial
environments. Microbiology and Molecular Biology Reviews 64: 548-572.
63
Nowak, D. J., Crane, D. E. & Stevens, J. C. 2006. Air pollution removal by urban
trees and shrubs in the United States. Urban Forestry & Urban Greening 4:
115-123.
Nurmi, V., Votsis, A., Perrels, A., & Lehvävirta, S. 2013. Cost-benefit analysis of
green roofs in urban areas: case study in Helsinki. Finnish Meteorological
Institute Reports 2. Helsinki, Finland: 58 s.
Oberndorfer, E., Lundholm, J., Bass, B., Coffman, R. R., Doshi, H., Dunnett, N.,
Gaffin, S., Köhler, M., Liu, K. & Rowe, B. 2007. Green roofs as urban
ecosystems: ecological structures, functions, and services. Bioscience 57:
823-833.
O'donnell, A., Norris, J., Berkeley, R., Claus, D., Kaneko, T., Logan, N. & Nozaki,
R. 1980. Characterization of Bacillus subtilis, Bacillus pumilus, Bacillus
licheniformis, and Bacillus amyloliquefaciens by pyrolysis gas-liquid
chromatography, deoxyribonucleic acid-deoxyribonucleic acid hybridization,
biochemical tests, and API systems. International Journal of Systematic
Bacteriology 30: 448-459.
Olsson, P., Francis, R., Read, D. & Söderström, B. 1998. Growth of arbuscular
mycorrhizal mycelium in calcareous dune sand and its interaction with other soil
micro-organisms as estimated by measurement of specific fatty acids. Plant and
Soil 201: 9-16.
Ongena, M. & Jacques, P. 2008. Bacillus lipopeptides: versatile weapons for plant
disease biocontrol. Trends in Microbiology 16: 115-125.
Park, K., Ahn, I. & Kim, C. 2001. Systemic resistance and expression of the
pathogenesis-related genes mediated by the plant growth-promoting
rhizobacterium Bacillus amyloliquefaciens EXTN-1 against anthracnose disease
in cucumber. Mycobiology 29: 48-53.
Peck, S. W., Callaghan, C., Kuhn, M. E. & Bass, B. 1999. Greenbacks from green
roofs: forging a new industry in Canada. Ottawa, Canada: CMHC/SCHL. 78 s.
Petras, S. F. & Casida, L. E. 1985. Survival of Bacillus thuringiensis spores in soil.
Applied and Environmental Microbiology 50: 1496-1501.
Philips, J. M. & Hayman, D. S. 1970. Improved procedures for clearing roots and
staining parasitic and vesicular-arbuscular mycorrhizal fungi for rapid
assessment of infection. Transactions of the British Mycological Society 55:
158-161.
64
Prakasham, R., Rao, C. S. & Sarma, P. 2006. Green gram husk-an inexpensive
substrate for alkaline protease production by Bacillus sp. in solid-state
fermentation. Bioresource Technology 97: 1449-1454.
Prathuangwong, S. & Buensanteai, N. 2007. Bacillus amyloliquefaciens induced
systemic resistance against bacterial pustule pathogen with increased phenols,
phenylalanine ammonia lyase, peroxidases and 1, 3-β-glucanases in soybean
plants. Acta Phytopathologica Et Entomologica Hungarica 42: 321-330.
Priest, F. G. 1977. Extracellular enzyme synthesis in the genus Bacillus.
Bacteriological Reviews 41: 711.
Priest, F., Goodfellow, M., Shute, L. & Berkeley, R. 1987. Bacillus
amyloliquefaciens sp. nov., nom. rev. International Journal of Systematic
Bacteriology 37: 69-71.
Ratnayake, M., Leonard, R. & Menge, J. 1978. Root exudation in relation to supply
of phosphorus and its possible relevance to mycorrhizal formation. New
Phytologist 81: 543-552.
Read, D. & Haselwandter, K. 1981. Observations on the mycorrhizal status of some
alpine plant communities. New Phytologist 88: 341-352.
Regvar, M., Gogala, N. & Zalar, P. 1996. Effects of jasmonic acid on mycorrhizal
Allium sativum. New Phytologist 134: 703-707.
Reva, O. N., Dixelius, C., Meijer, J. & Priest, F. G. 2004. Taxonomic
characterization and plant colonizing abilities of some bacteria related to
Bacillus amyloliquefaciens and Bacillus subtilis. FEMS Microbiology Ecology
48: 249-259.
Rodríguez, H., & Fraga, R. 1999. Phosphate solubilizing bacteria and their role in
plant growth promotion. Biotechnology advances 17: 319-339.
Rückert, C., Blom, J., Chen, X., Reva, O. & Borriss, R. 2011. Genome sequence of
B. amyloliquefaciens type strain DSM7 reveals differences to plant-associated B.
amyloliquefaciens FZB42. Journal of Biotechnology 155: 78-85.
Ruiz-Lozano, J., Azcon, R. & Gomez, M. 1996. Alleviation of salt stress by
arbuscular-mycorrhizal Glomus species in Lactuca sativa plants. Physiologia
Plantarum 98: 767-772.
65
Ruiz-Lozano, J., Azcón, R. & Gomez, M. 1995. Effects of arbuscular-mycorrhizal
Glomus species on drought tolerance: physiological and nutritional plant
responses. Applied and Environmental Microbiology 61: 456-460.
Ryu, C., Farag, M. A., Hu, C., Reddy, M. S., Kloepper, J. W. & Paré, P. W. 2004.
Bacterial volatiles induce systemic resistance in Arabidopsis. Plant Physiology
134: 1017-1026.
Saarela, I. 2002. Phosphorus in Finnish soils in the 1900s with particular reference to
the acid ammonium acetate soil test. Agricultural and food science in Finland 11:
257-271.
Sanders, F., Tinker, P., Black, R. & Palmerley, S. 1977. The development of
endomycorrhizal root systems: I. Spread of infection and growth-promoting
effects with four species of vesicular-arbuscular endophyte. New Phytologist 78:
257-268.
Schachtman, D. P., Reid, R. J. & Ayling, S. M. 1998. Phosphorus Uptake by Plants:
From Soil to Cell. Plant Physiology 116: 447-453.
Schenck, N. C., Graham, S. O., & Green, N. E. 1975. Temperature and light effect on
contamination and spore germination of vesicular-arbuscular mycorrhizal fungi.
Mycologia 67: 1189-1192.
Schöttelndreier, M. & Falkengren-Grerup, U. 1999. Plant induced alteration in the
rhizosphere and the utilisation of soil heterogeneity. Plant and Soil 209: 297-309.
Schwab, S., Menge, J. & Leonard, R. 1983. Comparison of stages of
vesicular-arbuscular mycorrhiza formation in sudangrass grown at two levels of
phosphorus nutrition. American Journal of Botany 70: 1225-1232.
Seki, T., Oshima, T. & Oshima, Y. 1975. Taxonomic study of Bacillus by
deoxyribonucleic acid-deoxyribonucleic acid hybridization and interspecific
transformation. International Journal of Systematic Bacteriology 25: 258-270.
Setlow, P. 1988. Resistance of Bacterial Spores to Ultraviolet Light. Comments on
Molecular and Cellular Biophysics 5: 253-264.
Shen, L., Wang, F., Qian, Y. & Yang, J. 2010. Tobacco growth promotion effect and
TMV resistance of Bacillus amyloliquefaciens Ba33. Journal of Jilin Agricultural
University 32: 383.
66
Shukla, A., Kumar, A., Jha, A., Salunkhe, O., & Vyas, D. 2013. Soil moisture levels
affect mycorrhization during early stages of development of agroforestry plants.
Biology and fertility of soils 49: 545-554.
Shute, L., Gutteridge, C., Norris, J. & Berkeley, R. 1984. Curie-point pyrolysis mass
spectrometry applied to characterization and identification of selected Bacillus
species. Journal of General Microbiology 130: 343-355.
Sippola, J. & Yli-Halla, M. 2005. Status of Soil Mapping in Finland. Research
Report 9. Jokioinen, Finland: European Soil Bureau. 417 s.
Song, U. & Lee, E. J. 2010. Environmental and economical assessment of sewage
sludge compost application on soil and plants in a landfill. Resources,
Conservation and Recycling 54: 1109-1116.
St-Arnaud, M., Hamel, C. & Fortin, J. 1994. Inhibition of Pythium ultimum in roots
and growth substrate of mycorrhizal Tagetes patula colonized with Glomus
intraradices. Canadian Journal of Plant Pathology 16: 187-194.
St-Arnaud, M., Hamel, C., Vimard, B., Caron, M. & Fortin, J. 1996. Enhanced
hyphal growth and spore production of the arbuscular mycorrhizal fungus
Glomus intraradices in an in vitro system in the absence of host roots.
Mycological Research 100: 328-332.
Stevens, K., Spender, S. & Peterson, R. 2002. Phosphorus, arbuscular mycorrhizal
fungi and performance of the wetland plant Lythrum salicaria L. under
inundated conditions. Mycorrhiza 12: 277-283.
Strack, D., Fester, T., Hause, B., Schliemann, W. & Walter, M. H. 2003. Review
paper: arbuscular mycorrhiza: biological, chemical, and molecular aspects.
Journal of Chemical Ecology 29: 1955-1979.
Takebayashi, H. & Moriyama, M. 2007. Surface heat budget on green roof and high
reflection roof for mitigation of urban heat island. Building and Environment 42:
2971-2979.
Taylor S. A. 1965. Managing irrigation water on the farm. Transactions of the
American Society of Agricultural Engineers 8: 6.
Thomson, B., Robson, A. & Abbott, L. 1986. Effects of phosphorus on the formation
of mycorrhizas by Gigaspora calospora and Glomus fasciculatum in relation to
root carbohydrates. New Phytologist 103: 751-765.
67
Timonen, S. & Peterson, R. L. 2002. Cytoskeleton in mycorrhizal symbiosis. Plant
and Soil 244: 199-210.
Tsuru, D. 1962. Inhibitory effect of glycine on the prodution of amylase and
proteinase by Bacillus subyilis. I. Effect of glycine and glycine derivatives on the
enzyme production by washed cells. Agricultural and Biological Chemistry 30:
225-420.
United Nation 2004. World uribanization prospect: the 2003 revision.
http://www.un.org/esa/population/publications/wup2003/WUP2003Report.pdf.
New York, USA: Department of Economic and Social Affairs of the United
Nation. Visited 10.3.2014.
United Nation 2012. World uribanization prospect: the 2011 revision.
http://esa.un.org/unup/CD-ROM/Urban-Rural-Population.htm. New York, USA:
Department of Economic and Social Affairs of the United Nation. Visited
10.3.2014.
Van Aarle, I. M., Olsson, P. A. & Söderström, B. 2002. Arbuscular mycorrhizal
fungi respond to the substrate pH of their extraradical mycelium by altered
growth and root colonization. New Phytologist 155: 173-182.
Van Loon, L. C. & Glick, B. R. 2004. Increased plant fitness by rhizobacteria.
Ecological Studies 170: 177-206.
Vardharajula, S., Zulfikar Ali, S., Grover, M., Reddy, G. & Bandi, V. 2011.
Drought-tolerant plant growth promoting Bacillus spp.: effect on growth,
osmolytes, and antioxidant status of maize under drought stress. Journal of Plant
Interactions 6: 1-14.
Vazquez, P., Holguin, G., Puente, M., Lopez-Cortes, A. & Bashan, Y. 2000.
Phosphate-solubilizing micro-organisms associated with the rhizosphere of
mangroves in a semiarid coastal lagoon. Biology and Fertility of Soils 30:
460-468.
Vierheilig, H., Schweiger, P. & Brundrett, M. 2005. An overview of methods for the
detection and observation of arbuscular mycorrhizal fungi in roots. Physiologia
Plantarum 125: 393-404.
Vitullo, D., Di Pietro, A., Romano, A., Lanzotti, V. & Lima, G. 2012. Role of new
bacterial surfactins in the antifungal interaction between Bacillus
amyloliquefaciens and Fusarium oxysporum. Plant Pathology 61: 689-699.
68
Volkmar, K. & Woodbury, W. 1989. Effects of soil temperature and depth on
colonization and root and shoot growth of barley inoculated with
vesicular-arbuscular mycorrhizae indigenous to Canadian prairie soil. Canadian
Journal of Botany 67: 1702-1707.
Wang, L., Lee, F., Tai, C. & Kasai, H. 2007. Comparison of gyrB gene sequences,
16S rRNA gene sequences and DNA–DNA hybridization in the Bacillus subtilis
group. International Journal of Systematic and Evolutionary Microbiology 57:
1846-1850.
Watt, P. M. & Hickson, I. D. 1994. Structure and function of type II DNA
topoisomerases. Biochemical Journal 303: 681.
Welker, N. & Campbell, L. L. 1967. Unrelatedness of Bacillus amyloliquefaciens
and Bacillus subtilis. Journal of Bacteriology 94: 1124-1130.
Wu, Q. & Xia, R. 2006. Arbuscular mycorrhizal fungi influence growth, osmotic
adjustment and photosynthesis of citrus under well-watered and water stress
conditions. Journal of Plant Physiology 163: 417-425.
Yang S., Yang G., Ren G., Wang L., Wu Y. & Chen P. 2010. A comparative study
on drought resistance and frost resistance of five kinds of introduced grasses.
Acta Agriculturae Boreali-Occidentalis Sinica 19: 91-95.
Yang, J., Yu, Q. & Gong, P. 2008. Quantifying air pollution removal by green roofs
in Chicago. Atmospheric Environment 42: 7266-7273.
Yu, G., Sinclair, J., Hartman, G. & Bertagnolli, B. 2002. Production of iturin A by
Bacillus amyloliquefaciens suppressing Rhizoctonia solani. Soil Biology and
Biochemistry 34: 955-963.
Zehnder, G. W., Yao, C., Murphy, J. F., Sikora, E. R. & Kloepper, J. W. 2000.
Induction of resistance in tomato against Cucumber mosaic cucumovirus by
plant growth-promoting rhizobacteria. Biocontrol 45: 127-137.
69
Appendices
Appendix I: Plant species found on the green roof sedum mats
Alopecurus geniculatus
Plantago major
Arenaria serpyllifolia
Polygonum aviculare
Artemisia vulgaris
Ranunculus repens
Barbarea vulgaris
Rumex acetosella
Cerastium fontanum
Spergula arvensis
Chaenorhinum minu
Thlaspi arvense
Geranium sanguineum
Trifolium repens
Medicago lupulina
Verbascum thapsus
Origanum vulgare
Veronica serpyllifolia
Plantago lanceolata
Veronica verna
70
Appendix II: DNA concentration of soil samples and used in qPCR
Sample ID
Nucleic Acid Conc. (ng/μl)
260/280
Conc. used in qPCR (ng/μl)
C1 18.7.12
33.6
1.71
11.6
C2 18.7.12
33.7
1.77
11.4
C3 18.7.12
35.7
1.76
12.6
C4 18.7.12
42.3
1.79
11.9
M1 18.7.12
39.7
1.83
11.6
M2 18.7.12
45.6
1.78
11.7
M3 18.7.12
43.6
1.80
13.0
M4 18.7.12
48.2
1.86
13.6
R1 18.7.12
33.8
1.78
10.0
R2 18.7.12
35.2
1.76
11.7
R3 18.7.12
29.4
1.75
10.5
R4 18.7.12
33.2
1.82
14.1
C1 9.8.12
27.0
1.8
12.3
C2 9.8.12
34.8
1.94
12.1
C3 9.8.12
46.7
1.85
13.6
C4 9.8.12
44.5
1.81
12.5
M1 9.8.12
44.8
1.89
12.5
M2 9.8.12
42.2
1.81
12.8
M3 9.8.12
47.1
1.74
12.3
M4 9.8.12
47.3
1.77
13.0
R1 9.8.12
46.0
1.84
13.6
R2 9.8.12
40.4
1.80
13.7
R3 9.8.12
40.7
1.76
12.5
R4 9.8.12
42.8
1.76
12.8
C1 30.8.12
47.1
1.82
12.2
C2 30.8.12
45.3
1.77
14.2
C3 30.8.12
44.9
1.80
12.6
C4 30.8.12
42.5
1.86
12.4
M1 30.8.12
52.1
1.78
14.0
M2 30.8.12
50.8
1.79
11.0
M3 30.8.12
48.1
1.84
10.7
M4 30.8.12
45.9
1.84
13.1
R1 30.8.12
46.8
1.85
12.7
R2 30.8.12
51.8
1.81
12.6
R3 30.8.12
47.0
1.80
12.5
R4 30.8.12
44.9
1.76
12.1
71
Sample ID
Nucleic Acid Conc.(ng/μl)
260/280
Conc. used in qPCR (ng/μl)
C1 19.9.12
39.8
1.80
12.3
C2 19.9.12
36.7
1.80
13.4
C3 19.9.12
40.2
1.78
12.3
C4 19.9.12
38.6
1.81
13.0
M1 19.9.12
42.8
1.78
13.6
M2 19.9.12
42.7
1.82
10.8
M3 19.9.12
41.7
1.84
13.8
M4 19.9.12
45.7
1.82
11.7
R1 19.9.12
42.7
1.84
13.6
R2 19.912
33.8
1.79
12.7
R3 19.9.12
39.6
1.83
13.1
R4 19.9.12
41.3
1.84
13.1
C1 28.5.13
32.2
1.93
12.3
C2 28.5.13
45.1
1.94
11.5
C3 28.5.13
36.1
1.96
12.1
C4 28.5.13
35.1
1.97
12.1
M1 28.5.13
26.7
1.99
12.5
M2 28.5.13
35.7
1.98
11.3
M3 28.5.13
36.2
1.95
12.6
M4 28.5.13
34.2
1.94
14.1
R1 28.5.13
35.4
1.96
10.5
R2 28.5.13
35.7
2.00
13.6
R3 28.5.13
28.5
1.97
10.7
R4 28.5.13
28.1
1.97
11.7
C1 17.6.13
29.0
1.96
12.2
C2 17.6.13
missing
missing
missing
C3 17.6.13
31.8
1.93
11.5
C4 17.6.13
79.0
1.89
12.5
M1 17.6.13
54.5
1.89
12.5
M2 17.6.13
44.0
1.96
12.2
M3 17.6.13
24.5
1.90
12.7
M4 17.6.13
28.2
1.85
13.4
R1 17.6.13
31.8
1.89
12.3
R2 17.6.13
25.4
1.92
12.0
R3 17.6.13
37.5
1.94
11.7
R4 17.6.13
33.4
1.95
12.5
72
Sample ID
Nucleic Acid Conc (ng/μl)
260/280
Conc. used in qPCR (ng/μl)
C1 8.7.13
34.2
1.92
12.4
C2 8.7.13
52.0
1.95
12.5
C3 8.7.13
38.9
1.96
11.7
C4 8.7.13
36.3
1.91
11.6
M1 8.7.13
43.5
1.94
12.2
M2 8.7.13
38.0
1.92
10.0
M3 8.7.13
35.6
1.92
12.5
M4 8.7.13
37.9
1.91
12.5
R1 8.7.13
48.8
1.92
11.6
R2 8.713
38.0
1.90
13.3
R3 8.7.13
33.1
1.92
12.2
R4 8.7.13
38.4
1.92
13.6
C1 30.7.13
41.0
1.93
11.8
C2 30.7.13
55.1
1.93
12.2
C3 30.7.13
46.3
1.93
12.5
C4 30.7.13
43.2
1.97
12.4
M1 30.7.13
47.8
1.93
12.2
M2 30.7.13
40.1
1.94
11.9
M3 30.7.13
37.4
1.94
12.0
M4 30.7.13
37.1
1.98
12.2
R1 30.7.13
53.4
1.85
11.6
R2 30.7.13
38.1
1.91
12.0
R3 30.7.13
46.5
1.93
13.2
R4 30.7.13
52.4
1.94
11.7
73
Appendix III: gyrB gene sequence from Rhizocell additive
TCGTAAACGCCTTGTCGACCACTCTTGACGTTACGGTTCATCGTGACGGAAAAATCCATTA
TCAGGCGTACGAGCGCGGGTACCTGTGGCCGATCTTGAAGTGATCGGCGAAACTGATAAG
ACCGGAACGATTACGCACTTCGTTCCGGACCCGGAAATTTTCAAAGAAACAACTGTATATG
ACTATGATCTGCTTTCAAACCGTGTCCGGGAATTGGCCTTCCTGACAAAAGGCGTAAACAT
CACGATTGAAGACAAACGTGAAGGACAAGAACGGAAAAACGAGTACCACTACGAAGGC
GGAATCAAAAGCTATGTTGAGTACTTAAACCGTTCCAAAGAAGTCGTTCATGAAGAGCCG
ATTTATATCGAAGGCGAGAAAGACGGCATAACGGTTGAAGTTGCATTGCAATACAACGACA
GCTATACAAGCAATATTTATTCTTTCACAAATAATATCAACACATACGAAGGCGGCACGCAC
GAGGCCGGATTTAAAACCGGTCTGACCCGTGTCATAAACGACTATGCAAGAAGAAAAGGG
ATTTTCAAAGAAAATGATCCGAATTTAAGCGGGGATGATGTGAGAGAAGGGCTGACTGCC
ATTATTTCAATTAAGCACCCTGATCCGCAATTCGAAGGGCAGACGAAAACCAAGCTCGGC
AACTCCGAAGCGAGAACGATCACTGATACGCTGTTTTCTTCTGCGCTGGAAACATTCCTTC
TTGAAAATCCGGACTCAGCCCGCAAAATCGTTGAAAAAGGTTTAATGGCCGCAAGAGCGC
GGATGGCGGCGAAAAAAGCCCGGGAA
74
Appendix IV: PCR protocol for identifying gyrB gene
Table 1. PCR materials
Reagents
×1
qPCR master mix
4 μl
10mM dNTPs
0.4 μl
10mM BaG3F
0.5 μl
10mM BaG4F
0.5 μl
Phusion
0.2 μl
Template DNA/water(control)
3 μl
Water
11.4 μl
Total Volume
20 μl
Table 2. PCR temperature program
Steps
Temperature
Time
Cycle
Initial denaturation
98 °C
30 s
1
Denaturation
98 °C
10 s
Annealing
68 °C
20 s
Extension
72 °C
5s
Final extension
72 °C
7 min
35
1
75
Appendix V: qPCR protocol for quantification of B. amyloliquefaciens
Table 1. qPCR materials
Reagents
×1
SYBR Green Mater Mix
5 μl
5mM BaG3F
0.8 μl
5mM BaG4F
0.8 μl
Template DNA/water(control)
3 μl
Water
1.4 μl
Total Volume
11 μl
Table 2. qPCR temperature program
Steps
Temperature
Time
Cycle
Pre-incubation
95 °C
5 min
1
amplification
95 °C
10 s
62 °C
10 s
72 °C
10 s
95 °C
5s
65 °C
1 min
97 °C
Continuous
40 °C
30 s
Melting curve
Cooling
45
1
1
76
Appendix VI: B. amyloliquefaciens content in soil samples
Table 1. B. amyloliquefaciens content in soil samples in 2012 (ng/g soil).
Date
July 18th
Aug. 9th
Aug. 30th
Sep. 19th
C1
0.1889
0.0329
0.5061
2.6118
C2
0.2826
1.5520
0.2546
10.9990
C3
0.2019
0.3950
0.4224
2.0714
C4
0.7690
0.3349
5.2092
3.0066
M1
0.3746
0.7302
4.4514
210.9910
M2
0.1805
0.4199
0.4715
1281.4938
M3
0.1651
1.6385
1.0843
4356.5773
M4
0.6126
0.6999
3.19710
5911.8335
R1
0.4678
0.3204
14.2967
1671.6971
R2
0.8274
0.3578
1.6460
796.4444
R3
3.6234
1.1294
11.1149
838.1156
R4
1.3805
1.2108
0.7126
503.3066
Table 2. B. amyloliquefaciens content in soil samples in 2013 (ng/g soil).
Date
May 28th
June 17th
July 8th
July 30th
C1
6.7903
2.5860
0.0649
0.0184
C2
8.2027
missing
0.0229
0.0189
C3
5.3509
0.1231
0.0750
0.0239
C4
1.4672
0.0214
0.0332
0.0369
M1
1.7622
0.1178
0.0468
0.0102
M2
1.7856
0.0251
0.0720
0.0416
M3
1.5049
0.1092
0.0317
0.0180
M4
0.8474
0.0190
0.0422
0.0291
R1
1.5201
0.8352
0.3161
0.5471
R2
0.2577
0.0765
0.1029
0.4867
R3
0.5198
0.3198
0.1323
0.3000
R4
0.9416
0.0366
0.2095
0.2634