ISOLATION AND CHARACTERIZATION OF ACT1NOMYCETES FROM
ARTEMISIA TRIDENTATA (SAGEBRUSH) RHIZOSPHERE
A POTENTIAL SOURCE FOR ANTIFUNGAL ANTIBIOTICS
Aida Elisabet Jimenez Esquilin
B.S. University of Great Falls, 2001
A thesis submitted to the
University of Colorado at Denver
in partial fulfillment
of the requirements for the degree of
Masters of Arts
This thesis for the Masters of Arts
Aida E Jimenez Esquilin
has been approved
Jimenez Esquilin Aida E (M.A. Biology)
Isolation and Characterization of Actinomycetes from Artemisia tridentata
(Sagebrush) Rhizosphere A Potential Source for Antifungal Antibiotics.
Thesis directed by Assistant Professor Timberley M. Roane
Actinomycetes are known to be major antibiotic producers. As of today 42% of known and
used antibiotics have been isolated from actinomycetes. Actinomycetes are commonly present in soil,
but are present in high numbers in rhizosphere associated soil where they interact in vivo with the
plants to provide protection against fungal pathogens. By isolating these actinomycetes we can
screen for the production of antifungal agents both in vivo and in vitro. In this study we isolated
actinomycetes from Artemisia tridentata (Sagebrush) rhizosphere associated soil. Five actinomycete
isolates, 6.5W5, SBR2, SBP3, SBG4, SBA showed in vitro antagonism against the fungi Armillaria
mellea, Aspergillus niger, and Altemaria altemata. Isolates 6.5W5, SBR2, and SBP3 are members
of the Streptomyces genus. Also we isolated total rhizosphere soil DNA to begin to characterize the
actinomycete community using molecular based methods. We hypothesize that this ecological niche
is full of new actinomycete strains yet to be described from which new antibiotics can be isolated
that could be used to prevent biodegradation of materials by fungi.
This abstract accurately represents the content of the candidates thesis. I recommend its publication.
I dedicate this thesis to my parents Esther Esquilin and Luis Jimenez for their love and support and
for believing that I can do anything. Thanks.
Thanks to my advisor Dr. Timberley Roane for all of her help and support and for teaching me to
question everything. My advisor Martin Gonzalez for his help and encouragement and for his sense
of humor. Thanks to my advisor Dr. Ellen Levy for her expertise and for inspiring me to be the
educator she is. Special thanks to Duried Kassab for being like a brother to me and to Katie Atchison
for her ffienship. Both of you thanks for welcoming me into the lab and making me feel at home.
Thanks to Dr. Don Crawford for his help and to the US Department of Defense for providing funding
for this research. Thanks to Dr. Toranzos for introducing me to the awe-inspiring field of
Environmental Microbiology and to Aaron Mondeau for his love and support. Finally, thanks to
other people in my life for letting me go so that I could follow my dreams.
Rationale and Significance...............................................2
2. LITERATURE REVIEW...........................................................5
The Problem: Degradation of Materials.......................................5
Fungal Degradation of Materials..........................................6
Fungal Degradation of Materials of Military Importance..................10
Mechanisms of Biodegradation............................................10
Current Treatments to Prevent Degradation...............................14
A Feasible Solution.....................................................15
The Actinomyecetes: A General Review.......................................16
Developmental Biology and Growth Kinetics of Actinomycetes.................22
Symbiotic and Parasitic Relationships in the Rhizosphere................28
Actinomycetes in the Rhizosphere........................................29
Rhizosphere in Desert Ecosystems........................................34
Antifungal Secondary Metabolites...........................................38
Types of Antifungal Compounds...........................................40
Mechanisms of Resistance................................................41
3. SAMPLING SITE AND SOIL CHARCTERIZATION.....................................43
Collection of Soil Samples..............................................45
Analysis of Soil Samples................................................45
Results and Discussion..................................................47
4. ISOLATION OF ACTINOMYCETES INHABITING THE RHIZOSPHRE OF
Culture Based Methods for Isolation........................................49
Culture Media Preparation...............................................49
Dilution and Spread Plate Technique.....................................49
Results and Discussion..................................................50
5. SCREENING FOR ANTAGONISTIC ACTINOMYCETE ISOLATES...........................58
In vitro Antifiingal Assays................................................58
Results and Discussion..................................................62
6. IDENTIFICATION OF ANTAGONISTIC ISOLATES....................................77
Identification of Antagonistic Actinomycetes...............................77
16S rDNA PCR and Sequencing.............................................77
Fatty Acid Analysis.....................................................78
Results and Discussion..................................................79
7. COMMUNITY CHARACTERIZATION STUDIES.........................................89
Molecular Methods for Community Characterization...........................89
Total Soil DNA Extraction...............................................89
Amplification by Polymerase Chain Reaction..............................90
Results and Discussion..................................................91
8. CONCLUDING REMARKS........................................................ 98
Future Studies........................................................... 100
2.1. Hypothetical model for the degradation of an organic polymer by a microbial community..7
2.2. Humic acid molecule................................................................... 8
2.3. Mechanisms of biodegradation and biodeterioration of materials by microbial communities.13
2.4. Colonies of Streptomyces lividans pIJ702 growing on R2A media.........................19
2.5. Potato scab disease caused by Streptomyces scabies....................................21
2.6. Growth curve for Streptomyces spp. and its relationship to antibiotic production......23
2.7. The rhizosphere.......................................................................27
2.8. Cellulose degrading fungi Armillaria mellea...........................................30
2.9. Streptomyces lydicus WYE108 colonizing the surface of the roots of the pea plant (Pisum
2.10. Artemisia tridentata (big sagebrush) inhabiting the western slope of the continental divide
2.11. Distribution of sagebrush throughout the western United Stated as of 2000 ..............36
3.1. Sampling sites........................................................................44
4.1. Microbial rhizosphere counts..........................................................51
4.2. Culturable actinomycetes numbers on YCED and WYE media................................53
4.3. Actinomycete fall and summer counts in the rhizosphere (Wolcott samples)..............54
5.1. Set up for in vitro antifungal plate assays............................................60
5.2. Antagonism against A. mellea of various actinomycete isolates from sagebrush
5.3. Antagonism against A. niger of various actinomycete isolates from sagebrush
5.4. Antagonism against A. alternata of various actinomycete isolates from sagebrush
5.5. Inhibiton of A. niger growth by actinomycete isolate SBG4.................................67
5.6. Inhibiton of A. niger growth by actinomycete isolate SBA..................................68
5.7. Reduction of A. niger spores after simultaneous incubation of fungus and isolate SBR2
for a total of 96 hours........................................................................69
5.8. Growth curve of the fungus A. niger co-cultured with isolate SBR2 for 96 hours............71
5.9. Reduction of A. niger spores after incubation for 48 hours with isolate SBG4..............73
5.10. Reduction of A. niger spores after incubation for 48 hours with isolate SBA.............74
5.11. Confluent streak assay for isolate SBG4..................................................75
6.1. Fatty acid methyl ester profile for isolate 6.5W5.........................................80
6.2. Fatty acid methyl ester profile for isolate SBR2..........................................81
6.3. Fatty acid methyl ester profile for isolate SBP3..........................................82
7.1. DNA isolated from sagebrush rhizosphere soil and bulk soil................................92
7.2 DNA isolated from the rhizosphere of sagebrush (plant A) using various extraction
2.1. Some characteristics of actinomycete groups............................................18
2.2. Effects of microbial colonization of the rhizosphere...................................31
2.3. Some antibiotics isolated from actinomycete species currently used clinically
and in research..........................................................................39
3.1. Characteristics of the rhizosphere associated soil and bulk soil ofboth sampling sites
4.1. Actinomycete and bacterial counts from sagebrush rhizosphere soil and bulk soil during fall
4.2 Actinomycete and bacterial counts form sagebrush rhizosphere soil and bulk soil during
summer of 2002...........................................................................56
6.1. Characteristics of the antagonistic actinomycetes isolated from sagebrush
6.2. Metabolic profile of the antagonistic actinomycete isolates............................86
7.1. Summarized data for DNA extraction from sagebrush rhizosphere and bulk soil........... 94
7.2. Summarized data for various DNA extraction protocols used in this project..............95
The microbial ecology of the desert plant rhizosphere is poorly understood and has
not been completely characterized as of today. However, rhizospheres are thought to
harbor microorganisms with unique metabolic pathways. We hypothesize that this
ecological niche is full of new actinomycete strains yet to be described, and that these
novel strains of bacteria are also capable of producing secondary metabolites that are
useful against fungal pathogens and fungal biodegraders. Current antifungals include
products like Imazail (Fungazil A), Azithromycin, Amphotericin B, and
Cycloheximide. Some of these antibiotics are used against biodegradation (Fungazil
A) and some are of medical importance (Azithromycin and Amphotericin B).
However, most of these antifungals are no longer effective due to antimicrobial
resistance. Fungi, including, Aureobasidium pullulans, Altemaria alternata, and
Aspergillus niger, can colonize and degrade plastics, such as plasticized polyvinyl
chloride, rendering the material non-useful. Other treatments for materials include the
use of organic compounds, such as polyanhydrides to protect textiles. The need for
new antifungal substances to prevent biodeterioration of materials, such as fabrics and
plastics, is increasing because the currently used ones (mentioned above) can be severe
on the materials themselves and because of antimicrobial resistance. Some of these
biodegrading fungi were also plant pathogens.
To address this need, we propose to find actinomycete strains that will produce
antifungal compounds in the unexplored environment of the rhizosphere. The
objectives of this research are (1) to characterize and identify the actinomycete
community that inhabit the rhizosphere of desert plants, and (2) to characterize
actinomycetes capable of producing antifungal compounds.
In order to achieve these objectives, we employed traditional physiological
methods in conjunction with molecular techniques to assess the actinomycete diversity
associated with sagebrush rhizosphere in desert shrublands in Colorado. We isolated
actinomycetes by the serial dilution-spread plate method and then identified them by
analysis of their 16S rDNA gene, as well as using fatty acid analysis. We also
performed an in vitro antifungal assays to screen for antifungal metabolite secretion by
each isolate. We also attempted in this project to extract total community DNA from
the rhizosphere to try to characterize the actinomycete community. In the search of
new secondary metabolites, we have started elucidating the ecology of sagebrush
Rationale and Significance
We hypothesize that in these uncharacterized arid environments there are
actinomycete strains that are capable of producing new antimicrobials (in particular
antifungals) while interacting in vivo with the plants. Some of these antifungals can be
used against fungi that cause biodeterioration of materials, such as plastics and fabrics,
and also against fungal plant pathogens. Usually fungal plant pathogens can also
degrade cellulose and other compounds used in the manufacturing of textiles and
plastics, and therefore finding antifungal compounds to target these microorganisms is
important industrially and agriculturally. By isolating these actinomycetes, we can
screen for the production of antifungal agents in vitro. Also, because the microbial
ecology of desert plant rhizospheres is poorly understood and has not been
characterized, we hypothesize that this ecological niche is full of new actinomycete
strains yet to be described.
Fungi, such as Aureobasidium pullulans, Alternaria altemata, and Aspergillus
niger, can colonize and degrade plastics such as plasticized polyvinyl chloride (pPVC).
Alternaria altemata is also a plant pathogen. Webb et al. (2002) found that growth of
A. pullulans is critical in the establishment of a microbial community on pPVC
because it produces an extracellular enzyme that degrades the plastic into more simple
organics that the rest of the microbial community can use. This presents a problem
because these materials can be degraded and made non functional. To prevent
deterioration by biodegraders like A. pullulans,ox plant disease by A. altemata,
materials and plants must be treated with antifungal compounds, such as Imazail and
sodium phenylacetate (produced by Streptomyces species). Because of the problem
with antimicrobial resistance, however, novel antimicrobial substances must be
discovered and/or developed and we speculate that actinomycetes will be the most
The specific objectives of this research are:
(1) To characterize the actinomycete communities which inhabit the rhizosphere of
sagebrush, a desert plant.
(2) To isolate actinomycetes with the capability of producing antifungal
metabolites and characterize and isolate these antifungal metabolites.
These objectives complement each other since as we isolate new actinomycete strains
to screen for antifungal compounds, we can also characterize the actinomycete
communities of the desert plant rhizosphere, especially regarding actinomycetes.
Eventually, a culture collection of desert plant rhizosphere actinomycetes will result
from this study.
This research will examine the actinomycetes isolated from desert plant
rhizosphere. Because there is little knowledge about the characteristics of the
microbial communities of the desert plant rhizosphere, this type of research will help
to better understand and elucidate the composition of such communities and at the
same time we will be able to describe these communities in terms of their
The Problem: Biodegradation of Materials
As our society becomes more aware of how susceptible our ecosystems are to
the pollution we introduce, new technologies to prevent catastrophic damages to our
environment are being explored. Among these technologies is the creation and use of
biodegradable materials including as plastics, textiles, and other synthetic polymers
that are readily degraded into non-distinguishable and useful organic matter, once they
are no longer needed. Biodegradation is defined as the use of biological systems to
break larger substances into smaller subunits (Prescott et al., 2002). In this review,
biodegradation and biodeterioration will be used as equal terms. The way these
biodegradable materials are made is by using components that are easily susceptible to
microbial degradation (Flemming, 1998). For example, using cellulose instead of
nylon in the manufacturing of textiles (cellulose is readily degraded by
microorganisms, especially fungi, into individual glucose molecules, which can then
be used as a carbon source, whereas nylon is not). This technology has relieved some
of the stress our environment experiences when non-degradable materials are disposed
of by humans as they persist in the environment for decades if not centuries polluting
and damaging, sometimes irreversibly, various ecosystems. However, this technology
also ensures that any materials that are made with microbially susceptible components
will not last as long in the environment. In turn, biodegradable materials have to be
replaced more often than those that are more recalcitrant, which is less cost effective.
The conflict between duration of materials and environmental protection is therefore
Besides microbial degradation of materials that we have modified to be more
susceptible to microbial attack, there is also intrinsic microbial degradation due to the
growth and metabolic flexibilities of microorganisms. The difference is that those
materials that are biodegradable are degraded at a faster rate that those that are not.
Almost any organic compound can be used by microorganisms as a carbon source,
which implies that in a microbial community many organisms can work together to
degrade any particular carbon based polymer as long as other necessary grow
requirements (eg., N and P) are available.
Fugal Degradation of Materials
Although bacteria, algae, protozoa, and fungi can degrade organics, fungi are the
principal organic degraders in the environment. In most cases the degradation of a
particular organic polymer will be initiated by the fungi and followed bacteria. Figure
2.1 shows a hypothetical model for the degradation of an organic polymer by a
In nature, fungi can degrade very complex molecules such as humics (see Figure
2.2), and because of their degrading capabilities, fungi have been implicated in the
deterioration process of a variety of materials, rendering them no longer useful.
Examples of fungal degradation of various polymers are: degradation of plasticized
Figure 2.1: Hypothetical model for the degradation of an organic polymer by a microbial community.
The larger polymer composed of many different monomers is degraded by many different species
producing smaller subunits that can further be degraded by other microorganisms in the community
Figure 2.2: Humic acid molecule. These type of molecules are prevalent in soil and fungi are famous for being
able to initiate their degradation. Carbon and Nitrogen released from these molecules is used in metabolism.
(From Stevenson, 1982).
polyvinalchloride (pPVC) by Aerobasidium pollulans (Webb et al., 2000; Robert and
Davison, 1986), degradation of cellulose, polyester and polycarbonates (both use the
manufacturing of textiles) by Aspergillus sp. (Vries and Visser, 2001, Pranamuda et
al., 1999), and degradation of paints by Penicillum, Altemaria and Aspergillus species
(Ciferri, 1999). Most of the fungal species that degrade textiles are also known plant
pathogens. The ability to degrade cellulose makes a fungi as a potential plant pathogen
and cellulose is a component of many textiles (Vries and Visser, 2001). Therefore, this
problem of biodegradation has an agricultural as well as industrial component. The
review will focus on fungal degradation of materials in particular those of importance
to the military.
Fungal Degradation of Materials of Military Importance
Millions of dollars are lost every year to the degradation of materials by fungi and
other microorganisms (Crawford, 2001). The United States armed forces spend many
millions of dollars every year replacing equipment, such as soldier tents, computer
circuit boards, and other electrical equipment, that are susceptible to microbial
degradation. For example, in the middle east, were the United States is currently at
war with Iraq, the main type of ecosystem is desert and desert-like, and even though
this implies low humidity and high dissecation, these conditions can be optimal for the
growth of many fungal species. Since fungi have spores, they are highly resistant to
dissecation and so can persist on materials for long periods of time (Prescott et al.,
During World War II, the military began to use equipment made out of plastics and
textiles. These materials were less heavy than their metal counterparts and therefore
easier to transport. They were also supposed to be durable. However, after some time,
troops stationed in the Pacific started to report many types of damages due to fungal
growth; damage ranging from moldy raincoats, allergies to the fungal spores, to failure
of electrical equipment (Flemming, 1998). These types of reports were and are
common in the military forces, and because of the extensive damage to the equipment
and health hazards, research in the prevention of biodeterioration of materials is
currently being underway.
Mechanisms of Biodegradation
Microbial communities, in particular biofilms, are the culprits in the degradation of
materials. Biofilms are organized microbial communities, consisting of layers of
microbial cells (fungi, bacteria, viruses, and other microbes) surrounded by
exopolymeric substances (EPS) (Prescott et al., 2002; Flemming, 1998). Biofilms are
always associated with surfaces and have chemical and physical gradients (oxygen,
pH, pressure, and nutrients), which give the biofilm complex structural and functional
flexibility. Because of the viscosity associated with the EPS layer, these biofilms form
easily on inanimate surfaces, such as bathroom tiles, glass, and plastics.
When biofilms form in the surface of some material, for example, a plastic, there is
the potential for the community to survive by using the plastic as a substrate. All is
needed is one member of the community to have the enzymes necessary to initiate the
degradation process and the rest of the community follows with the degradation of the
smaller subunits (as seen in Figure 2.1). There are usually four w'ays materials can be
degraded by biofilms: (1) biofouling, (2) degradation of leaching components, (3)
corrosion, and (4) hydration and penetration (Flemming, 1998).
Biofouling involves unwanted deposition and growth of microorganisms on
surfaces. This is the initial stage of actual biofilm formation. During fouling not much
degradation occurs, but because of the biofilm EPS layers, the characteristics of the
surfaces change (eg. hydrophobicity of a surface decreases due to biofilm formation).
Degradation of leaching components occurs when additives and monomers from the
surface of the material leach to the biofilm-surface interface and the microorganisms
can utilize them as substrates. This can cause embrittlement as well as loss of stability
of the material. Another way materials can be degraded is by corrosion, which occurs
when radicals resulting from metabolism of the additives and monomers leach into the
material causing degradation. Corrosion also causes loss of stability of the material.
Hydration and penetration involves increasing surface conductivity of the material as
well as penetration of the surfaces by the fungal hyphae. Because biofilms are up to
95% water, a biofilm growing on a surface can increase the conductivity of the surface
and can cause short circuits. This is how failure of electrical equipment occurs.
Another problem with biofilms is penetration of surfaces. The hyphae of the fungi
inhabiting the biofilms can penetrate any surface once it has been weakened. This may
cause swelling of the surface and loss of stability of the material. Once fungi penetrate
a surface, other microorganisms follow. Turgor pressure by fungal hyphae in surfaces
can be as great as 8 MPa (this is 78 times greater than atmospheric pressure). Figure
2.3 shows the various mechanisms of biodegradation of materials by biofilm formation
Besides structural damage done to the materials colonized by these biofilms, there
are health hazards related to fugal spore release from the surfaces of these materials
(Flemming, 1998).One extreme example of the extent of the damage of fungal
degradation occurred in 1984, when a German freight vessel hit a gale in the middle of
the South Atlantic Ocean and sank. Before the boat sank all the crew members were
able to escape in the life boats but they did not know that all of the rescue radio
equipment on the life boats had fungal colonization on the plastic isolators. Short
circuits resulting from conductivity caused the radios to stop functioning properly. The
crew could not ask for help on time and they died (Frank, 1984).
Deterioration of materials due to fungal colonization is not a trivial problem as the
damages can be as simple a deterioration of raincoats and as severe as being life
threatening. However extreme the damage is, millions of dollars are spend in the
replacement of materials lost to biodeterioration.
Biofouling Degradation of Corrosion
Effects -Change in surface -Embrittlement -Deterioration -Changes in conductivity
hydrophobicity -Loss of stability -Loss of stability -Swelling
Figure 2.3: Mechanisms of biodegradation and biodeterioration of materials by microbial communities.
Current Treatments to Prevent Degradation
Current treatments of materials to prevent antifungal deterioration are limited to the
use of antimicrobial additives, including Imazil A and Fosetyl-AI. These antifimgals
are classified as azoles (refer to page 40 for a complete explanation of mechanisms of
action of azoles). As antifungal resistance spread, however, the need for new
antifungals increases. Later this review will look at the mechanisms of antifungal
Other treatments involve chemical treatment of the materials, such as the use of
H3PO4. This acid is used as a disinfectant and is applied over the surfaces of materials.
No antifungal resistance is seen in when H3PO4 is used, however this acid is caustic,
and sometimes deterioration of the material is due to the acid itself rotting the surfaces
making it susceptible to microbial degradation and weathering. Textiles can be
protected by the use of metals that are toxic to microorganisms. Some fabrics can be
preserved by treating them with a 3.5mM solution copper hydroxide. The metal in the
solution replaces some of the fabric, fibers enough to have an antimicrobial effect but
; not to change the characteristics of the textiles (Chen et al., 1998). This is also done
with other metals, such as mercury. The problem with the mineralization of textiles to
preserve them is that the metal can also be toxic to humans even at low concentrations.
Currently available methods of preservation of materials work to some extent, but
because of resistance and because some of these chemicals can have toxic effects to
humans, new technologies are necessary.
A Feasible Solution
Of all of the available treatments to prevent biodegradation, the treatment that is
most beneficial for the environment and is cost effective is the use of antibiotics with
specific antifungal activity, such as Imidazole. By treating these materials with
antibiotics we can prevent the growth of microorganisms and, in turn, the deterioration
of these materials. Because a certain dosage of the antimicrobial is used in the
treatment process, once this dosage is no longer active, then the materials become
susceptible to microbial deterioration and eventual degradation. So while the materials
are in use, they can be periodically treated with the antibiotic of choice and before
they are to be disposed of, the antimicrobial will be either mechanically removed by
washing or no longer applied to the material. However, there is the problem of
resistance and new antifungals are needed. Since antibiotics are produced by a group
of bacteria called actinomycetes, we propose to search for actinomycetes in
unexplored desert ecosystems, such as the sagebrush rhizosphere. Here we expect to
find actinomycete isolates that may be able to produce antifungal antibiotics to be used
for the prevention of biodeterioration of materials.
The Actinomvcetes: A General Review
Actinomycetes belong to the high G+C content group of gram-positive bacteria.
Yet, these bacteria have filamentous hyphae that produce asexual spores, resembling
fungi. For the most part actinomycetes are aerobic (Prescott, et al. 2002; Ensign,
2000). Table 2.1 summarizes some of the characteristics of the actinomycete family.
Gross morphology of the actinomycete colony and color of aerial spores are some of
the tools used to identify actinomycetes isolates. As shown in Figure 2.4, aerial spores
in an actinomycete colony have distinct characteristic colors. For example
Streptomyces lividans has blue aerial spores and Streptomyces griseus has grey aerial
The ecological niche of most actinomycete species is the aerobic zone of the soil
(O and A horizons) (Kutzner, 2000). Actinomycetes of various genera have also been
found in fresh and seawater sediments and associated with sea animals, such as the sea
sponge Orodabiles spp. (Webster, et al 2001). In soil, actinomycetes live as symbiots
on or near the roots of desert plants, where presumably they provide the plant with
protection from pathogens and nutrients and receive, in turn, organic and inorganic
substances from the exudates (Elkan, 2001). The plant also removes water from soil,
reduces O2 tension and releases CO2, increasing soil acidity, which is particularly
beneficial for actinomycete colonization as they prefer acidic environments.
Actinomycetes are also free living in bulk soil but in bulk soil they are present in low
numbers when compared to the rhizosphere (Sylvia et al. 1998). Most actinomycete
isolates, regardless of their ecological niche, secrete antibiotics into the environment
(Ensign, 2000; Prescott et al. 2002; Kutzner, 2000), and even though the ecological
importance of antibiotic production is still a matter of debate, the ecological
importance of actinomycetes themselves is relevant to any soil ecosystem,
(this will be explored later in this review). Actinomycetes can produce a myriad of
compounds (besides antibiotics) that can enhance their ability to inhabit terrestrial
environments. Some of these compounds are: chitinases and other extracellular
enzymes, such as cellulases, that can degrade complex humics present in soil (Ensign,
2000; Bormann et al. 1999). Because of their degrading capabilities, actinomycetes are
being studied for their bioremediation potential (Crawford, 1988; Piret and Demain,
The process by which actinomycetes produce antibiotics is via secondary
metabolism. Secondary metabolism results when the microorganism produces
compounds that are not involved in the growth process of the organism itself (no
energy or biomass gain is directly related to the production of secondary metabolites)(
Martin and Demain, 1980; Cavalier-Smith, 1992). However, secondary metabolites
may assist with survival of the organism. Examples of such metabolites include
antibiotics, antitumor agents, and plant and animal growth promotants (Bushell, 1988).
Forty two percent of the microbial metabolites isolated as of 1999 come from
actinomycetes strains isolated from soil (Ensign, 2000). Even though the majority of
the antibiotics isolated from actinomycete strains have been used in the medical and
research fields, recently attention has been paid to the use of antimicrobial agents to
prevent biodeterioration of materials.
Besides their unique ability to produce antibiotics, actinomycetes have very
complex morphologies and very complex life cycles. Quorum sensing and pheromone-
like protein utilization are important factors for actinomycete growth and
Table 2.1: Some Characteristics of Actinomycete Groups
Group Wall Type* Sugar Pattern** Sporangia Spore Mol % G+C Arrangement Genus
Streptomycetes I (-) Chains of 5 to 50 spores 69-78 Streptomyces Sporichtya
Maduromycetes III B,C (+) or (-) Varies Varies Actinomadura
Nocardioforms I, iv, VI A (-) Varies 59-79 Nocardia, Rhodococcus
Thermonospora III C or B (-) Varies 64-73 Nocardiopsis
Actinomplanetes II D usually (+) Varies 71-73 Micronospora
*Wall types ^Characteristic Sugar in cell wall
I- L,L diaminopimelic acid, no characteristic sugar, glycine in interpeptide bridge A- Arabinose, galactose
II- Meso diaminopimelic acid, no characteristic sugar, glycine in interpeptide bridge B- Madurose
III- Meso diaminopimelic acid, no characteristic sugar C- None
IV- Meso diaminopimelic acid, arabinose, galactose D- Arabinose, xylose
Modified from Prescott et al. 2002
Figure 2.4: Colonies of Streptomyces lividans pIJ702
growing on R2A media. Each colony morphology represents
the same strain producing a different antibiotic. (From: ASIRC
(http://www. cbs. umn. edu/asirc/lib/pict/sl702. html)
development. Other worth-mentioning characteristics of some actinomycetes are
nitrogen fixation by Frankia species in non-leguminous plants and production of
flagellated motile spores by members of the Actinoplanaceae family (Prescott et al,
2002). Most actinomycete species are non-pathogenic soil bacteria, however, some
members of the Nocardia, Actinomyces and Mycobacterium genera are important
human pathogens. For example, Nocardia brazilensis, causes nocardiosis and
Mycobacterium tuberculosis causes tuberculosis (Ensign, 2000).
Other actinomycete species, such as Streptomyces scabies and Streptomyces
ipomoea, are known plant pathogens responsible of the potato scab disease as shown
in Figure 2.5.
Figure 2.5: Potato Scab disease caused by
Developmental Biology and Growth Kinetics of Actinomvcetes
The morphological diversity of actinomycete colonies is rather complex. One of
the reasons for such complexity is that these bacteria share many bacterial and fungal
characteristics (e.g., they are filamentous like fungi and but like bacteria they are gram
positive). Nonetheless, actinomycete developmental biology has been extensively
studied because the changes occurring during their life cycle are directly related to
secondary metabolism and therefore to antibiotic production. Most of the research
done on actinomycete cell development has been done with Streptomyces species.
Actinomycetes, albeit prokaryotic, do not follow the typical bacterial growth curve
seen in the laboratory. A typical bacterial curve has a lag, an exponential, a stationary,
and a death phase. Actinomycete growth, however, can be separated into three events:
(1) substrate mycelia formation, (2) aerial mycelia formation, and (3) sporogenesis
(Bushell, 1988; Prosser and Tough, 1991). It is difficult to determine a true
exponential phase in a pure culture of actinomycetes because these bacteria are
filamentous. In most cases, actinomycetes grow by polar elongation, where the cell
wall synthesis takes place at both ends of the growing hyphal fragment (Prosser and
Growth stages are directly related to antibiotic production. Figure 2.6 shows the
growth stages of a typical actinomycete colony compared to antibiotic production.
Note that antibiotic production is not maximal until aerial mycelia is produced, and it
continues until sporogenesis occurs.
On solid media, the different stages of growth can be identified using microscopy
techniques. As the growing hyphal fragments become a colony, storage of compounds
Figure 2.6: Growth curve for Streptomyces spp. and its relationship to antibiotic
production. *Modified from Brana et al. 1986.
promote formation of the substrate mycelia. The compounds stored in the substrate
mycelia migrate towards the upper layers of the colony to form aerial mycelia. The
next step in differentiation of an actinomycete colony is the formation of spores, which
is supported by polysaccharides stored by the aerial mycelium (Mendez, 1985;
Once a colony forms, the further spread of colonies over the agar occurs in a
similar fashion to that of filamentous fungi (Prosser and Though, 1991). The rough
surface characteristic of actinomycete colonies is a result of packed aerial spores
(Figure 2.4). Fungal and actinomycete colonies have four different functional and
structurally different zones. The first zone is the peripheral growth zone in which the
biomass increases exponentially. The second zone is the productive zone, in which the
antibiotic production occurs. The third zone is the fruiting zone where sporogenesis
occurs. Last, there is an aged zone in the colony which consists of dead hyphae and
substrate mycelia (Bushell, 1988). All four zones exist simultaneously in one colony.
In liquid culture, growth kinetics are very different. Due to the hydrophobicity of
actinomycete spores, the spores tend to aggregate forming a pellet. Pelleted growth
inhibits antibiotic production (Bushell, 1988; Martin and Demain, 1980; Higgs et al.,
2001). In order to achieve maximal antibiotic production, a well-dispersed mycelial
culture is necessary. For this, pharmaceutical industries have devised bioreactors with
the components necessary to allow growth of the actinomycete of interest to the stage
in which antibiotic is produced. In liquid culture, however, the changes in morphology
are not easily seen as separate events (Higgs, et al. 2001).
In the context of antibiotic production, actinomycetes usually undergo two
different phases. The first phase, called the trophophase, is characterized by rapid
growth of the microorganism resulting in nutrient exhaustion. Interestingly, some
sugars have been found to interfere with antibiotic production, a type of carbon
catabolite repression. For example, if culturing in a glucose rich medium, antibiotics
will not be produced or will be produced in very small amounts (Martin and Demain,
1980). The second phase is called the idiophase. The idiophase is characterized by
very slow to no growth, and it is here where antibiotics and other secondary
metabolites are produced. During the idiophase, the aerial mycelium appears and
sporulation occurs. According to studies done by Mendez et al. (1985), compounds
such as polysaccharides stored in the substrate mycelia will serve as energy sources
during the formation of aerial mycelia and spore formation. Therefore, rapid
accumulation of storage compounds, such as glycogen, marks the beginning of
sporulation events (Brana et al., 1982, 1986).
Antibiotic production does not occur until some metabolites that repress antibiotic
production are exhausted or until full vegetative growth is achieved. However, the
mechanisms of antibiotic production are not well understood. Many scientists over the
years have studied the metabolism of actinomycetes and of antibiotic production with
the hope to elucidate the mechanisms of secondary metabolism so that they can be
exploited in biotechnology.
As mentioned above, actinomycetes are ubiquitous to the rhizosphere, where
they have an important ecological role. About 80 percent of all terrestrial plants have
root systems that depend on other organisms for proper function and survival (Levine,
1994). Microorganisms, such as bacteria and microscopic fungi, are the major
colonizers of the rhizosphere (rhizo means root). The term rhizosphere has been
used to define the portion of soil that is under immediate influence of the plant root
(Hiltner, 1904). The term was modified eventually to include the plant, the soil
surrounding the roots, and the organisms inhabiting that environment (Lynch, 1987).
In this study, we will refer to the rhizosphere as the area of soil extending from 1 to 10
mm around the plant root as seen in Figure 2.7, where exudates released from the root
result in increased numbers and diversity of microorganisms, according to the
definition provided by Lynch et al .(1987).
As by-products of metabolism, the plant releases a myriad of nutrients and
chemical compounds that make the rhizosphere an optimal place to live. Plant roots
release compounds that include simple sugars, amino acids, organic acids, hormones,
vitamins, polymeric carbohydrates, and enzyme lysates that include whole cells, cell
walls, and volatile compounds, such as ethanol, C02, and acetoin (Dandurand and
Knudsen, 1997). Martin et al. (1977) determined that 17.3 % of the total carbon
assimilated by wheat roots is lost back to the soil in the form of exudates, the majority
of which is from amino acids. The quality and quantity of exudates depend on the age
of the roots, the abiotic characteristics of the soil, such as water content and pH, plant
genetics, and whether or not the soil has been amended with fertilizers (Bruehl, 1987).
Root with attached hyphae
Rhizosphere Associated Soil ~10 mm
Figure 2.7: The Rhizosphere. Defined as the few mm of soil under direct influence of the plant root.
Because of the variety of exudates released by the roots, the composition of the soil
adjacent to roots is different than that of non-rhizosphere soil and that, in turn, affects
the microbial community structure of the rhizosphere (Crawford et al., 1993).
Consequently, rhizosphere bacterial counts are usually higher than bulk soil counts,
with R (Rhizosphere): S (bulk soil) ratios ranging from 10:1 to 20:1 (Bruehl, 1987).
Symbiotic and Parasitic Relationships in the Rhizosphere
The organisms that colonize the rhizosphere can have a direct or indirect effect on
plant growth and endurance. A direct effect on plant growth implies that the plant is
provided with a compound or substance to enhance the uptake of nutrients, which in
turns, enhances growth. Plant growth promotant rhizobacteria (PGPR) are a specific
group of root colonizing bacteria that enhance seed germination and plant growth
(Tokala et al. 2002). Among these are Azotobacter and Azospirillum species. These
species in addition to fixing nitrogen for the plant, produce hormones such as
indoleacetic acid, that enhance plant growth. Another example of growth promotion is
microbial siderophore synthesis, which facilitates plant uptake of iron compounds
from soil (Gottschalck, 1988). An indirect effect on plant growth implies that the plant
is provided with a secondary metabolite, such as an antibiotic, that exerts antagonistic
effects on pathogens that might pose a risk to the plant. The plant, in turn, offers the
microorganism a habitat with plenty of nutrients, reduced oxygen tension, increased
soil acidity due to C02 release, and removal of water that prevents anaerobic
conditions (Dandurand and Knudsen, 1997).
Symbiotic relationships between plants and microbial root colonizers exist.
Mycorrhizae are root colonizing fungi which are obligate symbiots of plants including
oaks, beaches, and conifers, and offer the plant increased surface area for nutrient
absorption. In return the plant provides the fungus with exudates. In the absence of this
fungal relationship, the plant has a decreased chance of survival in soils with poor
nutrient availability (Varma and Hock, 1995). The nitrogen fixing bacteria, rhizobia,
are another example of beneficial rhizosphere microorganisms. In this relationship, the
plant is provided with NH4+ for growth while the rhizobia are provided with nutrients
and protection (Prescott et al. 2002; Kennedy, 1998; Papavizas and Davey, 1961).
The plant-microbe relationship exists on many levels. The mutualistic relationships
have been discussed above in detail, but there are also parasitic and antagonistic
relationships between plants and some microorganisms in which plant disease is often
the outcome. As seen in Figure 2.8, pathogenic fungi, such as Armillaria mellea,
degrade cellulose and lignin causing plant disease (Varma and Hock, 1995). Another
example is the actinomycete pathogen Streptomyces scabies, that causes potato scab
disease (Ensign, 2000). In other instances, there is more of a competitive relationship
between plants and microorganisms. For example, nitrifying bacteria and plants
compete agamst each other for the ammonia present in soil. The bacteria, because of
their high rates of metabolism, outcompete the plant oxidizing the ammonia present in
soil and in fertilizers into nitrate and nitrite (Prescott et al. 2002). This process, of
course, can be a nuisance to agriculture. Table 2.2 summarizes the benefits and
problems of rhizosphere colonization by microorganisms.
Actinomvcetes in the Rhizosphere
Among the microorganisms inhabiting the rhizosphere, actinomycetes are present
in relatively high number when compared to bulk soil. Actinomycetes can readily
Figure 2.8: Cellulose degrading fungus Armillaria mellea. A piece
of bark peeled back showing the fungal hyphae. (courtesy of Robert L.
Anderson, USDA Forest Service).
Table2.2: Effects of Microbial Colonization of the Rhizosphere.
BENEFICIAL NEUTRAL HARMFUL
Nitrogen fixation Biocontrol Growth Promotion Soil Stabilization Nutrient Uptake Symbiosis Antibiosis Free Enzyme Release Nutrient flux Disease Phytotoxicity Competition Alleopathy
Modified from Pepper et al., 1999.
Figure 2.9: Streptomyces lydicus WYE 108 colonizing the surface of the
roots of the pea plant (Pisum savitum). (A) shows the fungal hyphae in
contrast with the actinomycete hyphae. (B) shows bacteroid formation in the
root. (From Tokala et al., 2002)
colonize plant roots. Figure 2.9 shows actinomycetes colonizing the roots of a pea
plant (Tokala et al. 2002). Actinomycetes account for 10 to 30 percent of the total
microflora in the rhizosphere (Sylvia et al. 1998). The function of actinomycetes in the
rhizosphere is currently being investigated, but data indicates that they are important
for biocontrol. Preliminary greenhouse experiments indicate that some actinomycetes
can protect germinating lettuce seeds against damping-off, a disease caused by the
fungal pathogen Pythium ultimum (Yuan and Crawford, 1995; Hultberg et al., 2000)
The mechanisms of actinomycete protection against fungal pathogens are not well
understood, but in 1993, Crawford et al., demonstrated that secretion of secondary
metabolites vnth antifungal activity by actinomycetes living in the rhizosphere may be
responsible for protection. Since actinomycetes are known for their remarkable
capability of producing an unlimited number of antibiotics it is possible that their role
in the rhizosphere involves protection of the plant host against fungi and other
Recently, Tokala and coworkers (2002) have demonstrated that actinomycetes can
function as plant growth promoting rhizobacteria by developing favorable
relationships with nitrogen fixing bacteroids nodules in the roots of leguminous plants.
They have shown that Streptomyces lydicus WYE 108 enhances nodule growth and
bacteroid differentiation and aids in the assimilation of iron and other organic nutrients
from the soil. The result of this relationship is enhanced growth of the plant. They have
proposed that the most likely mechanisms for facilitation of iron uptake and enhanced
nodule formation is through the production of iron and molybdenum chelators that
assimilate and then transfer metals to the bacteroids (Tokala et al. 2002). According to
Tokala et al., (2002) this relationship although it has not been reported before, might
be very common in the environment making actinomycetes species more important
than previously thought.
Rhizospheres in Desert Ecosystems
The bacterial community structure of the rhizosphere is highly regulated and
influenced by the abiotic characteristics of the soil in which the plant lives (Bruehl,
1987). Desert and desert-like ecosystems (such as shrublands) occur where
precipitation is less than 25 cm per year and is greatly exceeded by evaporation. In
shrublands, the dry season is marked by little or no rain, the winters are long and dry,
and the dominant plants are shrubs that range from less than 1 meter to 5 meters in
height. The soil in these ecosystems is composed of a layer of topsoil that contains
little or no humus, and contains subsurface layers of sand, clay, silt, and minerals. It is
in the subsoil where desert plant root systems are concentrated (Levine, 1996; Jones
and Smith, 1992).
One of the most common plants in the desert and desert-like ecosystems west of
the continental divide is Sagebrush seen in Figure 2.10. As it is shown in Figure 2.11,
sixty million hectares in the Western United States are occupied by Big Sagebrush
(Artemisia tridentata) (Wambolt et al., 2000). This plant is arguably one of the most
important plant species in desert ecosystems since it has been demonstrated that
reduction in sagebrush communities by fire affects many native wild life species,
including the obligate symbiot sage grouse (Welch and McArthur, 1979; Wamboldt,
2000; Welch, 1999).
Figure 2.10: Artemisia tridentata (Big Sagebrush) inhabiting the Western slope of the Continental Divide
Potential Natural Vegetation Groups
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The characteristics of the rhizosphere soil of plants in desert and desert-like
ecosystems will therefore act as selective agents for the growth of bacteria and fungi
that favor such conditions. Actinomycetes, although common in all kinds of soil, favor
sandy soils, with neutral to slightly acidic pH (7.5-6.0) and low organic content
(Crawford et al., 1993). Consequently, actinomycetes are thought to be present in
higher numbers in the desert soils versus other kinds of soils (Asirc, 2002). Deserts
and desert-like environments, which cover about 85 % of the Western United States,
are an unexplored source of actinomycetes species and therefore a potential source of
Antifungal Secondary Metabolites
Below is a brief review to summarize the currently available antifungal compounds
used nowadays and the mechanisms of resistance used by the targeted fungi.
Actinomycetes are known for their ability to produce antimicrobials. They can also
produce anti-tumor agents, insecticides, and anti-nematode agents. Table 2.3 shows
some of the most commonly used antibiotics produced by actinomycetes and their
mode of action (Actinomycetes and Streptomyces Internet Resource Center, 2002).
Some examples are Streptomycin, produced by Streptomyces griseus, which targets
protein synthesis in bacteria, and Azythromycin, an antifungal produced by
Streptomyces nodosus, which targets fungal membrane sterols. Most of these
antibiotics are used clinically and as expected, resistance has already developed
against them. This brings up an important point for this projects rationale: there is an
increasing need for new antibiotics because of the microbial resistance to the currently
Of particular importance are antifungal compounds. Most of the antibiotics
currently identified and used are antibacterial, whereas the search and study of
antibiotics that specifically target fungi has lagged behind until very recently. This new
interest in antifungal compounds is mostly due to an increase in fungal infections after
therapies that repress the immune system, specially with AIDS and Cancer patients
(Ghannoum and Rice, 1999). However, even the antifungal compounds currently used
for therapy are in most cases rendered useless due to antifungal resistance mechanisms
developed by the targeted fungi.
Table 2.3: Some Antibiotics Isolated from Actinomycetes Currently Used
Clinically and in Research
Antibiotic Mode of Action Isolated from:
Protein synthesis (30S)
Protein synthesis (30S)
Protein synthesis (50S)
Protein synthesis (50S)
Artfi fungal antibiotics
The use of antifungal antibiotics is not, however, limited to clinical use. Fungi besides
being pathogenic can also be biodegraders (as stated earlier in this review),
which means that they have the ability to degrade materials, such as plastics and
textiles, rendering these materials useless. For example, Phaenerochaete
crysosporium, Aspergillus niger and Areobasidium pollutions are problematic fungal
degraders (Webb et al., 2000). These fungi release extracellular compounds that
breakdown complex organic molecules into individual subunits which the fungi can
use as growth substrates. Antifungals are needed to treat these materials to protect
them against biodegrading fungi.
Types of Antifungal Compounds
Currently used antifungals are categorized in three groups: azoles, polyenes, and
allylamines. All of these compounds act on membrane sterols, in particular, ergosterol
(Nisbet, 1992). Ergosterol acts as regulator of membrane fluidity similar to cholesterol
in other eukaryotic cell membranes. In order to synthezise ergosterol from its
precursor, 14-methylfecosterol, the enzyme 14- a-demethylase is needed. Antifungal
compounds that target membrane components, such as Imidazole, inhibit this enzyme
and therefore prevents the production of ergosterol which leads to cell death
(Ghannoum and Rice, 1999).
Actinomycetes are currently being explored for alternative antifungals. Chitinases
(enzymes that degrade chitin, the principal component of fungal cell walls) have been
found in the supemantants of Streptomycete tendae cultures, which suggests that
actinomycetes also produce these enzymes that serve as antifungal compounds in the
environment (Bormann et al. 1999). In 2002, Woo et al., isolated from Streptomycetes
sp. a protein called SAP that shows extraordinary antifungal activity against the fungus
Pythium porphyrae. The SAP protein is not a chitinase and is not active towards
human cells. Hwang et al. (2001) found that Streptomyces humidus produces
phenylacetic acid and sodium phenylacetate (two very simple organic compounds) that
have antifungal activity that compares to that of the azole fosetyl-AI and of H3PO4.
Mechanisms of Resistance
Mechanisms of resistance to these antibiotics include but are not limited to:
elimination of ergosterol from membrane composition, efflux pumps to remove the
drug from the cytoplasm, degradation of die antifungal, overexpression of the
antifungal target, and chemical inactivation of the antifungal (Ghannoum and Rice,
1999). Note that these mechanisms are not different than the mechanisms used by
bacterial cells to resist antibacterial antibiotics.
However, resistance is not the only problem with current antifungals. Some
antifimgals are also toxic to other biological systems, including humans. With the
exception of azoles (e.g. Fluconazole, Imidazole), most antifungal antibiotics present
some level of toxicity to humans, as well as affecting fungi, due to the similarities in
the cell membrane composition (Morrisey and Osbourn, 1999). New antifungals will
need to be specific only to fungal targets, such as cell wall components including
chitin, mannan, and a and P glucans (Ghannoum and Rice, 1999). Work is also being
done on using base analogs to inhibit fungal DNA synthesis. (Ghannoum and Rice,
Even though we have discovered (and understand their mechanisms of action)
more antifungal antibiotics in die last 20 years than ever before, novel more efficient
antifungals are needed to supply the need in both the clinical and industrial settings.
Also, if we can identify more actinomycete isolates that secrete efficient antifungal
compounds there is also potential to use them for biocontrol of plant fungal pathogens.
The applications are endless and the best source of novel antifungals are actinomycetes
from terrestrial environments.
Based on the current knowledge, actinomycetes isolated from soil are by far the
best source of antimicrobial agents available. By isolating actinomycetes and
screening them in vitro we can discover new antimicrobials, in particular new
antifungal compounds to which resistance has not yet developed, to prevent fungal
biodeterioration of materials. The soil in the shrublands of western Colorado are a
good source of these actinomycetes.
SAMPLING SITE AND SOIL CHARACTERIZATION
Artemisia tridentata (Sagebrush) is widely distributed on the western slope of
the continental divide in Colorado. Sagebrush communities occur in sub-alpine desert
like ecosystems, most of the time in association with other brushes and grasses. The
first sampling site chosen for this research was a sagebrush community located
approximately 1.5 miles west from US-131 to the junction of Horse Mountain Rd and
Milk Creek Rd. This community is located near the town of Wolcott, CO, on the west
side of the Continental Divide at an elevation of 7,500 ft with a dry and desert like
climate. Two major species of plants are present at the collection site: Sagebrush
(Artemisia tridentata) and Winterfat (Krascheninnikovia lanata). Only sagebrush will
be examined due to its prevalence at the site. The second site chosen was a sagebrush
community located in the town of Silverthome, CO, near the Green Mountain
Reservoir, located approximately 10 miles north from the intersection of 1-70 and US-
9 and an elevation of 7,800 feet. The climate and the weather at this site was similar to
the sampling site in Wolcott. The major plant species was Artemisia tridentata. Figure
3.1 shows the location of both sampling sites.
Figure 3.1. Sampling sites. Locations are marked on the map (Red) Wolcott, CO and (yellow)
Collection of Soil Samples
Three sagebrush specimens were collected as aseptically as possible and with care
to not disturb the soil associated with the root system. Sampling at the Wolcott
site occurred on September 15,2002 (Fall sample) and June 6, 2002 (Summer sample)
and sampling at the Silverthome site occurred on January 25, 2003 (Winter sample).
All specimens were immediately brought back to the laboratory for processing.
Rhizosphere soil from sagebrush specimens were collected by aseptically scrapping
the soil from the roots and placing the soil in sterile whirlpak bags. Bulk soil was
also collected at each site for each plant specimen to serve as non-rhizosphere control.
All samples were kept at 4 C until analysis.
Analysis of Soil Samples
To determine some of the general characteristics of the soil samples from each site,
the following analyses were performed: particle size analysis, water content and pH.
Particle Size Analysis. Particle size analysis was done according to the methods of
McGeeham and Naybor (1989). First, a 50 g sample of soil was oven dried for 24
hours at 110 C. The sample of soil was mixed with 10 ml of 5% sodium
hexametaphosphate in a 250 ml beaker and then 166 ml of distilled water was added.
The mixture was then stirred for four minutes and transferred to a 1L graduated
cylinder. A hydrometer was lowered into the cylinder and distilled water was added to
bring the volume (including the hydrometer) to 1130 mL. The hydrometer was
removed and the solution stirred to disperse all soil particles. The hydrometer was
lowered carefully into the cylinder at 40 seconds and the density reading recorded. At
this point the temperature of the solution was also recorded. The cylinder was left
undisturbed for 120 minutes at which point the hydrometer was lowered again and the
hydrometer density reading and temperature recorded. The rationale behind this
analysis is that as soil particles are dispersed in the water column, the density of water
changes. As time passes, the heavier particles (sand and silt) settle to the bottom of the
column and the lighter ones (clay, smaller silts) remain suspended. The hydrometer
measures the changes in water density as the heavy particles settle with time. Using the
formula offered by McGeeham and Naybor, (1989) and the standardized textural
triangle, the percent clay, silt and sand was calculated for each soil sample.
Soil Water Content Analysis. To determine the soil water content or moisture of
the soil, a sample of soil was pre-weighed and placed in pre-weighed foil tins in an
oven for 24 hours at 110 C to evaporate the water. Following heating, the tin were
weighed again. This experiment was repeated in duplicate so an average soil water
content was obtained using the following mathematical formula:
(3.1) Percent SWC= (wt of soil + tin) (wt dry soil + tin) x 100
(wt dry soil + tin)- (wt tin)
Determination of Soil pH. To determine the pH of the soil, 10 g of soil were placed
in a 250mL beaker and suspended on 10 mL of distilled water. The suspension was
mixed thoroughly with a stirring rod and left standing for 10 minutes at room
temperature. The pH sensor of a pre-calibrated pH meter was lowered into the solution
and the reading recorded.
Results and Discussion
Table 3.1 summarizes the characteristics of the rhizosphere associated soil as
well as of the bulk soil at both sampling sites. The soil (rhizosphere and bulk soil) used
in this study were sandy with sand percentages raging from 95-97%. The soils pH of
were neutral to slightly basic (7.2-8.3) with a soil water content ranging from 7.9 to
12%. These characteristics are similar to those of desert and sub-alpine ecosystems.
According to the literature (Asirc, 2002), actinomycetes favor sandy soils and desert
ecosystems with slightly acidic to neutral pH, all characteristics of both sampling sites
and therefore these soils were determined as adequate for actinomycete isolation.
Table 3.1: Characteristics of the rhizosphere associated soil and bulk
soil of both sampling sites in Colorado.
Sampling Site Percent Rhizosphere Bulk Soil Percent
Sand Silt Clay pH pH SWC
Wolcott, CO 97.56 0.21 2.24 7.20 8.02 7.985 1.3
Silverthome, CO 95.26 0.61 4.13 7.82 8.10 12.33 2.5
ISOLATION OF ACTINOMYCETES INHABITING
THE RHIZOSPHERE OF SAGEBRUSH
Culture Based Methods for Isolation
Culture Media Preparation
Selective media were used to isolate actinomycetes from sagebrush rhizosphere
associated soil and bulk soils. The media used was Yeast Casaminoacids Extract and
Dextrose Agar (YCED) and Water Yeast Extract Agar (WYE). According to the
literature, these media provide enough nutrients to favor the growth of actinomycetes
and retard the growth of fungi and other bacteria. YCED contained per liter: 0.3 g of
glucose, 0.3 g of casaminoacids, 0.3 g yeast extract and 2.0 g of dibasic potassium
phosphate and 18.0 g of agar. WYE contained per liter: 0.3 g of yeast extract, 0.5 g
dibasic potassium phosphate, and 18.0 g of agar. The pH of the media was then
adjusted to 6.5 and 7.4 using 2 M HC1 and autoclaved. Cyclohexamide (lOOpg/ml)
was added to the media following autoclaving (cool down to 55 C) to reduce fungal
Dilution and Spread PlateTechnique
Dilution and plating was used to determine culturable microbial counts from
sagebrush rhizosphere associated and bulk soils. Soil (5.3 g dry wt) was mixed with
50 ml of 0.1% glycerophosphate, and homogenized in a rotator for 30 minutes at 40
rpm. This was done to ensure that soil aggregates were dissolved and bacteria
associated with individual particles released. Follow this treatment, serial dilutions
were made (1 O'2 to 10'5) of the soil slurry in 0.9 ml dilution blanks (0.1%
glycerophosphate) and plated onto YCED and WYE agar plates at pH 6.5 and 7.4.
Each dilution was plated in duplicate, and the plates were incubated at 25 C for 14
days or until visible actinomycete colonies were observed. Counts for bacteria, fungi,
and actinomycetes that were able to grow on these media were obtained.
Once the actinomycete colonies were visible, colonies were picked and plated onto
YCED and WYE agar plates (pH 6.5 and 7.4) to obtain pure cultures. Actinomycete
isolates were distinguished based on general appereance, colony morphology, and
color of the aerial spores. Isolates were incubated at 25 C for 14 days or until
sporulation was visible. After sporulation, actinomycetes were kept at 4 C until
analyzed. Long term stocks were made of the isolates by suspending actinomycete
colonies in 10% glycerol and freezing at -20 C until needed.
Results and Discussion
Culture based methods were used to determine the culturable numbers of
bacteria, actinomycetes, and fungi present in the sagebrush rhizosphere. Although the
media used YCED and WYE were intended to be selective for actinomycetes, counts
of other bacteria and fungi were obtained. Microbial numbers were higher in the soil
associated with sagebrush rhizosphere when compared to non-rhizosphere soil. Figure
4.1 shows that bacterial numbers were in the range 107-108cells/g soil followed by
Figure 4.1: Microbial rhizosphere counts. Bacterial numbers were 107 to 108
cells per g soil (+/- 2.23 x 105), actinomycete numbers were 105to 106 cells
per g of soil (+/-2.0 xl 05), and fungal numbers were 103 cells per g soil
(+/- 2.3 x 105) in each plant specimen.
fungi and actinomycetes at 105-106 cells /g soil. These numbers drop significantly in
the bulk soil with bacterial numbers in the range of 105cells/g soil, actinomycete
numbers at 103cells/g soil and fungi at 103cells/g soil, as expected since it has been
well demonstrated that actinomycetes are present in higher numbers in the rhizosphere
when compared to bulk soil (Sylvia et al., 1998). These numbers represent averages of
the counts obtained for both types of media at both pHs. In further studies,
was added to prevent fungal contamination and therefore fungal numbers are not
available for the summer and winter microbial counts.
The media utilized in these studies was designed to select or favor the growth
of actinomycetes. Originally, the media was designed with a neutral pH of 7.4, but it
was noted that a more acidic pH of 6.5 seemed to favor even more the growth of
actinomycetes. As seen in Figure 4.2, WYE medium at pH of 7.4 gave the highest
numbers of actinomycetes (1.20 xl06cells/g soil) and at pH of 6.5, the YCED medium
allowed for more actinomycete growth (1.40 x 107 cells/g soil).
Our data also indicated that during the fall months actinomycete numbers are high
(8.91 x 105 cells/ g soil) when compared to the summer months 4.60 x 103 cells/g soil.
Figure 4.3 shows that the numbers of actinomycetes present in sagebrush rhizosphere
were lower (but not significantly lower) during the summer than during the fall
months. Tables 4.1 and 4.2 summarize the data for Figure 4.3. Since the numbers of
actinomycetes during the fall months was higher, it is not surprising that diversity was
also higher in the fall season. A total of 122 actinomycete isolates were recovered and
screened for the production of secondary metabolites.
Number of Actinomycetes in Soil (cells/g soil) Number of Actinomycetes in Soil (cells/ g soil)
Figure 4.2: (A) Culturable actinomycete numbers on YCED and
WYE media. Growth on WYE media at pH 7 4 was optimal
(averagelO6 cells per g (+/- 2.2 xlO5)), whereas growth on YCED
(B) was better at pH 6 5 (averagelO5 cells per g soil (+/- 1.35 x 104)).
Number of Actinomycetes in Soil (cells/ g soil) Number of Actinomycetes in soil (cells/ g soil)
Figure 4.3: Actinomycete Fall and Summer counts in the rhizosphere
(Wolcott samples). (A) shows actinomycete numbers in media
at pH 7.4. (B) shows numbers in media at pH 6.5. Note that
overall actinomycete numbers are higher in the fall than in the summer
(+/- 2.5 x 104 in summer and +/- 3 xlO3 in the fall)
Table 4.1: Actinomycete and bacterial counts from sagebrush rhizosphere soil and bulk soil during Fall of 2001.
Note how ratios of bacteria to actinomycete increase in the bulk soil when compared to the ratios in rhizosphere
associated soil in media at pH 7.4.
Rhizosphere Bulk Soil
Media at pH 7.4 Media at pH 6.5 Media at pH 7.4 Media at pH 6.5
Bacteria 4.58 x 107 1.54 x 106 -------------- 3.21 x 103 1.17 x 105 ------------------
Actinomycetes 8.92 x 105 3.65 x 10s 1.66 x 105 3.69 x 104 3.54 x 103 1.31 x 103 2.12 x 103 1.01 x 103
Bacteria: Actinomycetes 51:1 -------------- 126:1 ------------------
R/S ratio: 186:1
Data not available
Table 4.2: Actinomycete and bacterial counts from sagebrush rhizosphere soil and bulk soil during
summer of 2002. Note how ratios of bacteria to actinomycete increase in the bulk soil when compared to
the ratios in rhizosphere associated soil.
Media at pH 7.4 Rhizosphere Media at pH 6.5 Media at pH 7.4 Bulk Soil Media at pH 6.5
Bacteria 1.35 x 105 5.17 x 104 9.24 x 104 2.54 x 104 2.21 xl05 1.75 xlO4 3.56 x 105 1.17 x 104
Actinomycetes 4.66 x 103 9.57 x 102 6.83 xl03 3.27 xlO2 5.63 xl02 1.75 xlO2 5.32 x 102 2.23 x 102
Ratio 13:1 29:1 39:1 66:1
R/S ratio (10:1)
The differences observed in microbial numbers in fall and summer months may be
related to the fluctuations in temperature between the fall and the summer months at
the Wolcott site. In the year 2002, Colorado experienced a dry summer season.
Temperature affects precipitation, and this, in turns, affect plant biomass, which
directly affects exudate concentration and composition (Martin et al., 1977).
Our results for culture based methods support the current hypothesis that the
rhizospheres of plants are occupied by a myriad of microorganisms, in particular
actinomycetes. Tables 4.1 and 4.2 show that actinomycetes were present in the
rhizosphere soil in higher numbers than in the bulk soil. The average rhizosphere/bulk
soil ratio (R/S) of our samples was 186:1 (Fall) and 10:1 (Summer). The numbers of
bacteria and fungi were also higher in rhizosphere associated soil. These results also
support our hypothesis that desert soils will be occupied by many actinomycetes and
that there is potential for the isolation of novel secondary metabolites.
SCREENING FOR ANTAGONISTIC ACTINOMYCETE ISOLATES
In vitro Antifungal Assays
A total of 122 isolates collected on each medium at each pH were screened in vitro
for the production of antifungal secondary metabolites. Antagonism against fungi was
assessed by two different methods: plate assays and liquid antifungal assays. To
determine antifungal production, actinomycete isolates were challenged with three
different fungal isolates. Aspergillus niger was chosen because it is a well
characterized biodegrader of plastics. Altemaria altemata and Armariella mellea are
both cellulose degraders and plant pathogens. Degrading fungal isolates were used as
the goal of this study was to isolate antifungal producing actinomycetes for the long
term goal of protecting materials from fungal degradation.
Actinomycete Spore Suspensions. In order to challenge fungi with each
actinomycete isolate in the liquid antifungal assays, spore suspensions were prepared
using a modified version of Braiia et al., (1986). Twenty to thirty sterile 10 mm glass
beads were added to the plate cultures of sporulated actinomycete isolates. The beads
were rolled gently over the colonies for about 10 seconds. Following this, the
spores covered beads were washed using a 0.9% NaCl: 0.85%Tween 80 sterile
solution. The resulting spore solution was filtered using No. 2 Whatman paper to
eliminate hyphae. The spore suspension was then centrifuged at 14,000 x g and the
pellet resuspended in 3 mL of 0.9% sterile saline. These suspensions were kept at -20
C until use.
Fungal Cultures. The fungal cultures used in this study (Aspergillus niger,
Altemaria altemata and Armillariella mellea) were kept on Com Meal Agar, at pH of
6.0, and kept at 4 C until needed.
Bacterial Cultures. Cultures of Bacillus megaterium and Micrococcus luteus were
grown on nutrient broth for 24 hours at 37C.
Plate Assays. Antifungal plate assays were done according to the methods of
Crawford et al., (1993). Each actinomycete isolate was streaked onto one half of a com
meal agar plate and incubated at 25 C for 7 days or until sporulation had occurred. A
0.5 cm2 agar plug with actively growing fungal mycelia was then placed next to the
actinomycete growth on the plate, which was then incubated for an additional 48
hours. Figure 5.1 shows the experimental set up for the plate assays. Antagonism was
determined as the distance between actinomycete growth and fungal growth. Growth
of fungi at 7.5 to 20 mm apart from actinomycete colonies was considered strong
antagonism. According to the literature, values for antagonism of well characterized
antibiotic producing isolates is 20 mm (Quiroga et al., 2001; Hwang et al., 2001;
Oudouch et al., 2001).
A modified version of this method was also used. Briefly, each actinomycete was
streaked in a confluent line in the center of a YCED or WYE plate and incubated at 25
Figure 5.1: Set up for in vitro antifungal plate assays
C for 3-4 days or until sporulation occurred. After sporulation, A. niger, and of
bacteria Bacillus megaterium and Micrococcus luteus were streaked perpendicular to
the actinomycete growth and incubated for another 48 hours (see Figure 5.10 for set
up). Antagonism was determined as above. Growth of fungi and bacteria 20 mm apart
was considered antagonism.
Liquid Antifungal Assays. Isolates that showed antagonism in the plate assays
were further examined by using the methods of Yuan and Crawford (1995). Triplicate
plates with 15 ml of liquid YCED media (pH 6.0) enriched with 20mM glucose were
seeded with a 0.5 mm agar plug of actively growing fungal mycelia. The
fungi was grown at 25 C for 48 hours at which point 1 ml of each actinomycete spore
suspension (105 spores/ mL) were added to the actively growing fungi. The fungi and
actinomycetes were incubated further for 48 to 120 hours and spores were collected
every 24 hours. This was done by filtration using a vacuum pump and pre weighed
No.2 Whatman filter paper. The filtrates were oven dried at 60 C overnight and then
re-weighed to determine the amount of the fungal spores present in each sample. These
experiments were run together with control plates to which no actinomycete spores
were added. By using the following formulas,
(5.1) S = (Dry Spores+ Paper wt) (Paper wt)
(5.2) AS S coDtrolS experimental
we determined the amount of fungal spores present in each plate and the differences
between the control and the experimental cultures. AS values were obtained and
plotted against time (48, 72, 96 and 120 hours).
Using analysis of variance and student t tests, the p- values associated with
the treatment groups at each time interval were obtained in order to statistically
support the data collected in these experiments.
Results and Discussion
Plate assays and liquid antifungal assays were performed to screen for in vitro
antifungal antagonism. Figures 5.2, 5.3, and 5.4 show the preliminary results for the
plate assays performed fof each actinomycete isolate challenged by either Armillaria
mellea,, Aspergillus niger, or Altemaria altemata, respectively. Antagonism in this
experiment was defined by measuring the distance between the fungal growth and the
actinomycete colonies, assuming that the farther apart they grow the more antagonistic
the actinomycete isolates are to the fungi. Strong antagonism was defined by a
distance of 7.5 mm or more in the plate assays. Although 122 isolates were tested for
in vitro antagonism, only 21 isolates showed some level of antagonism to the fungi.
Four isolates showed antagonism against .4. mellea, eight to A. niger and nine isolates
to A. altemata.
A. mellea is an extremely slow grower; this makes it difficult to assess full
antagonism because it is difficult to distinguish between no growth due to a slow
growth rate and no growth because of antagonism. However, isolates SBP3, SBR2,
and 6.5W5 showed strong antagonism against .4. mellea since the fungus grew no
closer than 10 mm from the actinomycete isolate colonies following 120 hours of
incubation, as seen in Figure 5.2.
Figure 5.2: Antagonism against A. mellea of various actinomycete isolates
from sagebrush rhizosphere. Isolate SBP2 shows poor antagonism,
whereas isolates 6.5W5, SBR2 and and SBP3 show strong antanonism.
Distance of Fungal Growth from Actinomycete Colonies (cm)
6.5W3 6.5W5 6.5W8 6.5R2 SBR2 S8W14 S8G4 SBA
Figure 5.3: Antagonism against A. niger of various actinomycete isolates
from sagebrush rhizosphere. Isolates 6.5W5, 6.5W8, and 6.5R2,
show poor antagonism, whereas isolates SBW14 and SBG4 show moderate
antagonism and isolates 6.5W3, SBR2 and SBA show strong antanonism.
Figure 5.4: Antagonism against A. altemata of various actinomycete isolates
from sagebrush rhizosphere. Isolates 6.5W3,6.5W5,6.5W8,6.5R2,
6.5R4,6.5C3, and SBP3 show poor antagonism, whereas isolates SBP2
and SBR2 show mild and strong antanonism, respectively.
There were 8 isolates that showed some kind of antagonism against the fungus
A.niger. A. niger is a fast growing fungus (grows and sporulates in 48 hours on
Com meal agar) and antagonism assessment was relatively easy to assess. Isolates
SBR2, SBG4, and SBA were strongly antagonistic against A. niger because the fungus
grew 30, 10 and 20 mm away from each isolate actinomycete isolate respectively.
Figure 5.3 shows the results for the plate assays for isolate antagonistic against A.
niger. The rest of the isolates were not studied further because compared to SBR2,
SBG4 and SBA their antagonism was weak. For A. altemata, also a fast grower, only
isolate SBR2 was strongly antagonistic as seen in Figure 5.4.
Isolates SBP3, SBR2, 6.5W5, SBG4, and SBA were further studied in the liquid
antifungal assays and identified because they showed the strongest antagonism in the
plate assays against the fungi. Figure 5.5 and 5.6 show examples of the antagonistic
isolates SBG4 and SBA, respectively, against the fungus A. niger.
However, since the plate assays were observational studies, strong inferences
cannot be made based on the plate antifungal assays. To quantitatively analyze the
antagonism of these isolates, liquid antifungal assays were performed. A. niger only
was used because it is a fast growing fungus and because most isolates yielded positive
antagonism results against A. niger, except SBP3 and 6.5W5. The assumption behind
this assay was that if some kind of antagonistic antifungal compound was being
produced by the actinomycete, this would stall or negatively affect the growth of
fungus in the liquid medium. The assumption that the number of spores was directly
proportional to fungal growth was made.
Our results showed that isolate SBR2 could significantly slow or prevent the
growth of A. niger after incubating the fungus with the isolate for 48 to 72 hours.
Figure 5.5 (A): Inhibition of A niger growth by actinomycete isolate SBG4.
The fungus A. niger grew approximately 15 mm away from SBG4 colonies.
Figure 5.5 (B): Inhibition of A.niger growth by actinomycete isolate SB A.
48 hours 72 hours 96 hours 120 hours
Time in Hours
Figure 5.7: Reduction of Aspergillus rtiger spores after simultaneous
incubation of the fungus and isolate SBR2 for a total of 96 hours. At 48
and 72 hours reduction is significant (p values 0.004 and 0.009 respectively).
After 48 hours reduction is not significant (p values 0.8765). Dark blue bars
represent A. niger alone and light blue bars represent the fungus co-cultured
with isolate SBR2.
Figure 5.7 shows that after 48 hours of inoculation with SBR2 the average amount
of spores was approximately 9 mg and without the treatment (no actinomycete) the
average amount of fungal spores was approximately 18.3 mg. This resulted in a 50.8 %
inhibition of fungal growth with a p value of 0.004 (less than 0.05 was considered
significant). After 72 hours of fungal growth in the presence of isolate SBR2, the
average amount of spores was approximately 27.6 mg whereas in the control the
average amount of spores was approximately 39 mg. This resulted in 29.2 % inhibition
of fungal growth with a p value of 0.009. What these results implied was that the
fungus by itself could grow normally in broth culture but when it was grown in
combination with isolate SBR2, the growth of this fungi was negatively affected, seen
by the reduced amount of spores produced by the fungus. Figure 5.8 shows a growth
curve of the fungus A. niger compared to a growth curve of the actinomycete isolate,
and both cultures growing together. In this Figure we can observe that the fungal
growth is stopped or significantly reduced when co-cultured with isolate SBR2. After
96 and 120 hours of incubation, there was a reduction in the amount of the fungal
spores (50.6% and 36.9%, respectively), although no longer statistically significant (p
value = 0.8765). However, this did not mean that these observations were not
biologically significant because the ineffectiveness of the antifungal at this point might
be due to inactivation of the drug by an environmental condition (e.g., incubation
temperature). The results at 96 and 120 hours can also be explained by a low
concentration of the antibiotic in the culture or maybe a low concentration of
actinomycetes in the culture (we know that antibiotic level and activity increases for a
period of time and then levels off). Also, if nutrients in the medium were exhausted by
fungal growth, then the spores could not germinate and grow to produce antibiotics.
Time in hours
Figure 5.8: Growth curve of the fungus A. niger co-cultured with isolate SBR2
for 96 hours.Note the reduction on the amount of spores is significant for
72 hours (p value = 0.009) and it is non-significant at 96 and 120 hours
(p value = 0.8765).
Nonetheless, reduction of the amounts of spores was still occurring even after 120
These results, however preliminary, suggested that isolate SBR2 might be secreting
a secondary metabolite that affects the growth of A. niger. The liquid antifungal assays
for isolates SBG4 and SBA were performed for only 48 hours and 5 replicates were
used instead of 3. The results shown in Figure 5.9 show that isolate SBG4 was
successful in reducing the fungal spores of A. niger by 51.9%, (p value = 0.0186), with
an average amount of spores in the experimental set of 10 mg whereas in the control
set there were 20.8 mg of fungal spores. However the reduction of spores by isolate
SBA was not statistically significant (p value = 0.8765) ( Figure 5.10).
Several mathematical formulas were used to determine the effectivity of the
presumed antifungal produced by actinomycete SBR2. Percent inhibition was
determined by the following formula:
(Average wt of spores! ____,i
1 Average wt of spores) control
where the experimental shows the treatment effect (inoculation with actinomycete
isolate) and the control shows the non treatment effect (no actinomycete challenge).
Also, paired Student t tests were used to determine the p values for the liquid
antifungal assays. This statistical analysis was appropriate because of the clustering
effect of the samples (experimental and control) and because equal variances were
assumed for both groups. However, some log transformations were done and t test
analysis repeated to confirm that the p values were accurate.
Figure 5.9: Reduction of A. niger spores after incubation for 48 hours
with isolate SBG4. In every replicate there is reduction in the amount of
fungal spores and the reduction is significant (p value = 0.0186).
Figure 5.10: Reduction of A. niger spores after incubation for 48 hours
with isolate SB A. In every replicate there is reduction in the amount of
fungal spores howeverreduction is not statistically significant (p value =
Figure 5.10: Confluent streak assay for isolate SBG4. The fungus A. niger
grew more than 20 mm away from the SBG4 colonies, B. megaterium and
M. luteus seemed unaffected by the actinomycete.
Based on this data and the statistical analyses, there was enough evidence to
suggest that the actinomycete isolates were secreting possible antifungal secondary
metabolites. An additional plate assay was performed with isolate SBG4 to determine
if the possible antifungal also had some antibacterial effect. Figure 5.11 shows a
confluent streak of isolate SBG4 and perpendicular to it there are streaks of A. rtiger,
Bacillus megaterium and Micrococcus luteus. The last two bacteria do not seem to be
affected by this metabolite, however, the fungus does grow away from the
actinomycete colonies, confirming that the compound is antifungal.
Based on their antifungal activity in vitro, these isolates (SBP3, SBR2, 6.5W5,
SBG4, and SB A) were identified using FAME analysis and 16S rDNA sequencing to
determine whether or not they were novel isolates or already characterized isolates
known to produce antibiotics. These isolates were also characterized based on the
carbon source utilization using BIOLOG technology.
IDENTIFICATION OF ANTAGONISTIC ISOLATES
Identification of Antagonistic Actinomvcetes
Each isolate that showed antagonism in the in vitro antifungal assays was
identified by using 16S rDNA sequencing with primers specific for actinomycetes.
Isolates were also identified using FAME (fatty acid methylester) analysis.
16S rDNA PCR and Sequencing
DNA Extraction of Isolates. Each antagonistic isolate was grown on YCED or
WYE broth medium at each respective pH (based on media used in isolation) with
shaking at 30 C for 3-4 days. A 100 pi sample of each culture was heat lysed at 98
C for 10 minutes. After heat lysis, the cultures were centrifuged at 14,000 x g for
45 seconds to remove cell debris.
Amplification by Polymerase Chain Reaction. The 16S rDNA of individual
isolates was amplified by using actinomycete specific primers. The primer set
included R513GC 5 CGG CCG CGG CTG CTG GCA CGT A 3, and F243 5
GGA TGA GCC CGC GGC CTA 3 (Huer et al., 1997). The reaction mixture of
one 50pl reaction tube included 5pl of 10X PCR buffer, 3pl of 50 mM MgCl2,
lOpl of 2.5 nM (ea) dNTP mix, 5pl Dimethyl Sulfoxide, 2pl BSA (4mg/ml), lpl
Taq Polymerase, 19.2 jal dH20 and 1.4 pi of each primer diluted IX (lOOpg/ml). 2 pi of
template were added to each tube and the mixture was amplified using a Perkin Elmer
2400 Thermocycler for 30 cycles as follows: 2 minutes denaturation at 94 C, 1
minute primer annealing at 63 C and 2 minutes of primer extension at 72 C followed
by one final step at 72 C for 10 minutes and cooling at 4 C (Huer et al., 1997).
Sequencing. Following amplification, the samples were run on a 1.5% agarose gel
along with a 123 bp ladder to verify the 16S rDNA amplification product, which was
approximately 250-300 bp long. After confirmation of the PCR product by
electrophoresis, each sample was purified using the Wizard PCR Preps Purification
Kit according to the manufacturers instructions and sent for sequencing to at the
UCHSC Cancer Center DNA sequencing Core. Sequences were blasted on the
National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov)
database for identification of the isolate.
Fatty Acid Analysis
Isolates were also identified using FAME (fatty acid methyl ester) analysis. Those
isolates showing strong antagonism against the fungi included in this experiment were
isolated in pure culture, streaked onto nutrient agar media and incubated at 28 C until
sporulation occurred. The isolates were then sent to Microbial ID Systems Inc. (MIDI)
(Newark, DE) for identification.
Spore suspensions were made of each antagonistic isolate as explained above and
used to inoculate BIOLOG plates according to the manufacturers instructions. The
plates were incubated at 25 C for 3 days and the optical density was measured at 590
nm. An absorbance greater than 0.05 was recorded as positive (visible purple color).
Results and Discussion
Isolates that were found to be antagonistic to A. mellea, A.altemata, and A.
niger were identified using the following methods: FAMEs (Fatty acid methyl ester)
and 16S rDNA sequencing. FAME analysis allowed for the identification of
microorganisms based on the fatty acid composition of their cell membrane. A profile
of the fatty acid composition of the bacterial membrane was generated by gas
chromatography and the resulting profile was compared to a standardized database.
This method is reliable as an identification tool as the composition and relative
amounts of fatty acids is different for each microorganism. The sensitivity of this assay
allowed for identification at the species level. When comparing the unknown species
to the database, a similarity index was assigned to the isolate that ranged from 0 to
0.8. A similarity index of 0.6 was considered a good match (98% confidence).
The results of the FAME analysis for isolates 6.5W5, SBR2, and SBP3 is
shown in Figures 6.1, 6.2, and 6.3, respectively. As seen in these Figures, each fatty
acid profile was distinct for each isolate, although these all appear to be members of
the Streptomyces genus. According to the MIDI database, isolate 6.5W5 was
Streptomyces anulatus with a similarity index of 0.589, isolate SBR2 was
Streptomyces amakusaensis with a similarity index of 0.641, and isolate SBP3 was
Streptomyces scabies with a similarity index of 0.636. At the time of this assay,
Figure 6.1: Fatty acid methyl ester profile for isolate 6.5W5. Each piece of the pie represents a different
fatty acid in the membrane. According to this profile, isolate 6.5W5 is Streptomyces anulatus (SIM =
Figure 6.2: Fatty acid methyl ester profile for isolate SBR2. Each peak represents a different fatty acid in the
membrane. According to this profile, isolate SBR2 is Streptomyces amakusaensis (SIM = 0.641)
Figure 6.3: Fatty acid methyl ester profile for isolate SBP3. Each peak represents a different fatty acid in the
membrane. According to this profile, isolate SBP3 is Streptomyces scabies (SIM = 0.636).
isolates SBG4 and SBA were not available as they were found in the summer
Even though FAME analysis is a reliable tool for identification of microorganisms,
molecular based methods, such as gene sequencing, are more reliable because these do
not depend on the conditions for growth as the FAMEs analysis does. Isolates 6.5W5,
SBR2, and SBP3 were identified using 16S rDNA sequencing using actinomycete
specific primers. The 16S rDNA gene is widely used for the identification of
prokaryotic microorganisms because this gene is highly conserved within species.
According to the sequences obtained, isolate 6.5W5 was Streptomyces laceyi,
isolate SBR2 was Streptomyces lateritus and isolate SBP3 was also Streptomyces
lateritus. All isolates match the respective sequences with 99% similarity. Currently,
isolates SBG4 and SBA are being identified by their 16S rDNA sequences.
Note that the FAMEs analysis and the 16S rDNA analysis differ in the
identification of the isolates at the species level although they both agree at the genus
level classifying the isolates in the Streptomyces genus. One reason for this
discrepancy might be the size of the PCR product. The 16S rDNA of actinomycetes is
approximately 1100 bp long. Only 300 bp were amplified using primers F243 and
R153GC. Such a short sequence might not provide enough information to identify the
isolates at the species level. This was also seen in the fact that isolates SBR2 and SBP3
were identified as the same isolate, when they display distinct differences in their gross
morphology as well as their antagonism to different fungi.
Regardless of the discrepancies of both identification methods, all of the isolates
belong to the Streptomyces genus. This result was not unexpected as 90% of soil
actinomycetes belong to this genus (Kutzer, 2000) and almost all of the antibiotic
producing species of actinomycetes are Streptomyces species (Ensign, 2000). Table
6.1 summarizes some of the characteristics of all of the antifungal producing isolates
To characterize these antagonistic isolates a metabolic fingerprint of each isolate
was produced using BIOLOG plates. A BIOLOG plate is a 96 well plate that
contains 95 different organic substrates and one negative well, along with other
nutritional requirements (eg. N and P) and a dye called tetrazolium chloride. If the
isolate is capable of utilizing a particular carbon source present on the plate, electrons
released during metabolism will reduce the tetrazolium dye from clear to a purple
color. The optical density of each well was read at 560 nm and an absorbance greater
than 0.05 was recorded as positive (visible purple color). The intensity of the purple
color is related to how well the substrate is utilized. A metabolic profile was done for
each isolate and the results are shown in Table 6.2. Note the different patterns of
substrate utilization of isolates 6.5 W5, SBR2, SBP3 and SBG4. Also, note that some
of the compounds (Arabinose, Tween 40 and 80, Mannitol, Pyruvic acid methyl ester
and others) are favored by all of the isolates, which was not surprising as they all
belong to the same genus. All of the isolates also seemed to disfavor similar
compounds, such as glucose and mannose. The differences in carbon utilization
suggested that these were different isolates and that the discrepancies seen on the 16S
rDNA assays were probably related to the length of the obtained product as explained
Table 6.1: Characteristics of the antagonistic actinomycetes isolated from sagebrush rhizosphere.
Isolate Spore Color .Antagonism against Identification according to FAMEs 16S rDNA Sequencing Similarity index/ Percent similarity
6.5W5 White/Grey A. mellea S. anulatus S. laceyi 0.589/99%
SBR2 Pink A. mellea, A. alternata, and S. amakusaensis S. laleritus 0.641 /99%
SBP3 SBG4 Purple Dark Grey A. mellea A. niger S. scabies S lateritus 0.636/99%
SBA White A. niger
Table 6.2: Metabolic profile of the antagonistic actinomycete isolates.
Organic Acids Amino Acids Nucleotides Others
D-Psicose Pyruvic acid L-Asparagine Thymidine Tween 40
D-Fructose Acetic acid L-Glutamic acid Tween 80
L- Erythritol Cis-Aconitic acid Histidine
D-Melibiose Citric Acid L-Phenylalanine
(3-Methyl-D-Glucoside Formic Acid Gluconic acid Hydroxyphenylacetic acid L-Proline
D-Psicose Cis-Aconitic acid L-Aspartic acid L-pyroglutamic
L-Arabinose Citric Acid L-GIutamic acid acid
D-Galactonic Acid L-leucine Tween 40
Pyruvic acid Succinic acid Hydroxybutyric acid Lactic acid Bromosuccinic acid Itaconic acid L-Phenylalanine L-Proline Tween 80
TabJe 6.2. Continued.
Isolate Sugars Organic Acids Amino Acids Nucleotides Others
SBP3 L-Arabinose Pyruvic acid L-Histidine L-Pyroglutamic
D-Fructose Cis-Aconitic acid L-Hydroxyproline acid
D-Galactose Citric acid L-Proline Camithine
D-Glucose m-Inositol D- Mannitol D-Psicose D-Sorbitol Sucrose D-Trehalose D-Galactonuc acid D-Galacturonic acid D-Gluconic acid D-Glucuronic acid Hydroxybutyric acid Itaconic acid Lactic acid Malonic acid Propionic acid D-Saccharic acid Quinic acid Succinic acid L-Serine Glycerol Putrescine Inosine Tween 40 Tween 80
SBG4 D-Psicose Pyruvic acid L-Alanine Plienyletlrylamiriv
L-Arabinose Succinic acid Acetic acid Cis Aconitic acid Citric acid D-Galactonic acid Itaconic acid Lactic acid L-Aspartic acid L-Glutamic acid L-Leucine L-Phenylalanine L-Proline Tweert 40 Tween 80
Table 6.2 Continued.
Organic Acids Amino Acids Nucleotides Others
SBA D-Glucose D-Galactonuc acid L-Serine ........ Glycerol
oo , m-Inositol D-Galacturonic acid Putrescfne
D- Mannitol D-Gluconic acid Inosine
D-Psicose D-Glucuronic acid Tween 40
Pyruvic acid Tween SO
COMMUNITY CHARACTERIZATION STUDIES
Molecular Methods for Community Characterization
In an attempt to assess the composition of the microbial community inhabiting
sagebrush rhizosphere, in particular the actinomycete community, molecular
techniques such as 16S rDNA sequencing from total soil DNA using actinomycete
specific primers were employed.
Total Soil DNA Extraction
Total soil DNA was extracted by a modified version of Zhou et al, 1996. A 5 g soil
sample was place in a 15 mL Falcon tube and suspended in 13.5 mL of DNA
extraction buffer (100 mM sodium EDTA [pH=8.0], lOOmM sodium phosphate
[pH=8.0], 1.5 M NaCl, 1% CTAB). One hundred microliters of Proteinase K
(lOmg/mL) were added and the suspension was incubated at 37 C with shaking (225
rpm). After the shaking treatment, the samples were treated three different ways to
determine what method yielded the most DNA. Treatment (1) did not include any
SDS or Lyzozyme, treatment (2) included the addition of lOOpl (10 mg/ml) of
Lyzozyme and 10 ml of 20% SDS, treatment (3) included addition of SDS alone, and
treatment (4) included the addition of Lyzozyme alone. The samples were incubated
for 2 hours at 65 C with gentle inversions every 20 minutes and then centrifuged at
6,000 x g for 10 minutes at room temperature to collect the supernatant. The
supernatant was then transferred to a clean tube. The soil pellet was extracted two
more times by adding 4.5 mL of extraction buffer and repeating the procedure above
but the 65 C incubation time was 10 minutes instead. Supernatants of all 3 cycles
were pooled and mixed with an equal volume of chloroform: isoamyl alcohol (24:1
vol/vol) and centrifuged at 6,000 x g for 5 minutes. The aqueous phase was recovered
and precipitated with 0.6 volumes of isopropanol at room temperature for 1 hour. The
DNA pellet was obtained by centrifuging at 16,000 x g for 20 minutes and wasched
with 70% ice-cold ethanol. The pellet was resuspended in 500 pi deonized water.
DNA was extracted from rhizosphere associated soil as well as bulk soil following this
protocol and the samples were stored at -20C until analyzed using a UV-Vis
Spectrophotometer (Absorbance at 280 (protein) and 260 (DNA) nm) and on a 1%
agarose gel ran at 120 V with TBE buffer (100 mM Tris, 30 mM Boric acid and 0.2
mM EDTA, pH =8).
DNA was also extracted from soil using the SoilMaster DNA Extraction Kit
(Epicentre (Madison, Wisconsin)), according to the manufacturers instructions and
stored and analyzed as above.
The agarose gels were analyzed using the NIH image computer program
(http://rsb.info.nih.gov/nih-image). Briefly, the NIH image program allows us to
normalize a scanned image of a gel based the intensity of the bands in a gel and on the
intensities of standards of know concentrations. This was done to determine how much
DNA was extracted from rhizosphere associated soil and bulk soil.
Amplification by Polymerase Chain Reaction
Amplification of the 16S rDNA operon from total soil DNA was amplified by
using actinomycete specific primers. The primer set included R513GC 5 CGG CCG