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ISOLATION AND CHARACTERIZATION OF MEMBERS OF
THE PHYLUM ACIDOBACTERIA FROM SOILS

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STEPHANIE A. EICHORST

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ISOLATION AND CHARACTERIZATION OF MEMBERS OF
THE PHYLUM ACIDOBAC T ERIA FROM SOILS

by

Stephanie A. Eichorst

A DISSERTATION

Submitted to
Michigan State University
In partial fiJlfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY
Department of Microbiology & Molecular Genetics

2007

ABSTRACT
Isolation and characterization of members of the phylum Acidobacteria from soils
by

Stephanie A. Eichorst

The soil environment is an abundant source of microbial life; recent studies have
estimated that in one gram of soil, there are over one million species. These microbes are
essential to the environment as major contributors to biogeochemical cyles (e.g. carbon
and nitrogen). Unfortunately only 0.1 to 1% of the total microbial community has been
cultivated, leaving a wealth of unexplored diversity; one such phylum is the
Acidobacteria. The phylum Acidobacteria is an ubiquitous group of microorganisms
found in various soil and sediment environments. Historically, it contains eight
monophyletic subdivisions and only three cultivated representatives. This dissertation
explored members of this unknown phylum by using molecular and growth-based
approaches to increase our understanding of acidobacteria ecology in the soil
environment.

A phylogenetic survey of soil taken from the KBS LTER revealed the presence of
subdivisions 1, 3, 4, 5, and 6, with members of subdivision 4 being more dominant in the
conventional agriculture treatment and subdivisions 1 and 6 being more dominant in the
successional community treatment. Additionally, the acidobacteria community
composition changed in relation to edaphic properties such as soil moisture (subdivision
3), carbon concentration (subdivision 4), soil pH (subdivision 1), methane fluxes
(subdivision 1), and nitrous oxide fluxes (subdivision 4). These trends were used to

isolate strains of acidobacteria and helped to provide insight into their physiology.

Successful attempts were made to cultivate members of this phylum with soil
collected from the KBS LTER using cultivation strategies designed to mimic the native
soil environment. After ca. 30 days of growth, total recoveries ranged from ca. 3 to 6%
of the total microbial community; soils containing high moisture content had significantly
increased total recovery. In order to screen for the presence of acidobacteria on
cultivation plates, a facile high-throughput method called Plate Wash PCR (PWPCR) was
developed to rapidly screen enrichment plates using phylum-specific 16S rRNA gene
primers. Additionally, PWPCR revealed that acidobacteria were more frequently
detected with elevated levels of carbon dioxide (significantly), the presence of the
catalase enzyme, low nutrients, and low oxygen concentrations.

The use of these cultivation strategies along with PWPCR was instrumental in
isolating more than a dozen members of the phylum Acidobacteria from subdivisions 1
and 3. Colonies of these strains were approximately 1 mm in diameter and either white,
pale yellow or pink in color, the latter due to a carotenoid that was synthesized
preferentially under 20% as compared to 2% oxygen. Strains were Gram negative,
aerobic, chemoorganotrophic, nonmotile rods that produced an extracellular matrix
causing cells to clump in liquid culture. All strains contained either 1 or 2 copies of the
16S ribosomal RNA encoding gene, which along with a relatively slow doubling time
suggests an oligotrophic lifestyle. Genotypic, physiological, and morphological data
revealed the presence of a novel genus in subdivision 1, T erriglobus, which contained
pigmented strains of acidobacteria. The physiological and nutritional characteristics of

these acidobacteria are consistent with their potential widespread distribution in soil.

DEDICATION

This work is dedicated to my dearly beloved grandparents and great aunts: Edward J.
Schultz, Constance B. Schultz, Anna Grien, Leonardo Ciesla, John H. Eichorst, and Rose
M. Eichorst.

iv

ACKNOWLEDGEMENTS

Throughout this journey, I have grown as a scientist, with the guidance and
support of my co-advisors, John Breznak and Tom Schmidt. I am forever grateful to
them for their support, encouragement, intellectual challenges, opportunities, and
intoxicating enthusiasm for teaching and science. I am truly thankful. I would like to
thank members of my graduate thesis committee, Terry Marsh, Mike King, Frank Dazzo,
and Phil Robertson, for their advice and insight into my project during our rewarding
meetings. I would like to thank members of the Microbiology & Molecular Genetics
department who have all made this experience memorable, and the staff at the Kellogg
Biological Station, especially Drew Corbin who was extremely helpful in the soil
sampling process. Lastly, I would like to thank the College of Natural Science and the
Lyman Bridges School for giving me the opportunities to teach. My most rewarding
teaching experience was Honors Biology with John Urbance. He has been a true
teaching mentor to me and has led by example.

My time here at Michigan State University has been quite a journey. From the
first day of my rotation in the Schmeznak lab, Joseph R. Graber, Joel A. Klappenbach,
Les A. Defihlefson, Joel Hashimoto, Kwi S. Kim, Daniel H. Buckley, Kristin M.
Huizinga, and Bradley S. Stevenson, welcomed and introduced me to the exciting
research world of microbial ecology. I would like to thank them for showing me the
ropes and making research both a fun and rewarding experience. Additionally, the past
and present members of the Schmeznak lab, Dion Antonopoulous, Zarraz Lee, Brian
Carnpbell, Uri Levine, Brendan Keough, John Wertz, and Jorge Rodriguez have provided

support, assistance, great discussions, and camaraderie in the lab. Specifically, I would

like to thank Kristin for all of her help with soil sampling at the KBS LTER, and Dion,
Jorge, and Brian for our coffee trips and great talks about life and science. You all have
been great mentors to me.

While at Michigan State University, I have made some ever-lasting friendship
with Jennifer Gray, Kristin Huizinga, Julie Hotopp Dunning, Zarraz Lee, Kristi
Whitehead, and Daniel Whitehead. Our friendship over the years has meant so much to
me, and I would not have made my way through the labyrinth of graduate school without
your support, both scientifically and socially. I am looking forward to the next chapter of
our life.

None of this would have been possible without the love and support of my family.
First and foremost, I would like to thank my parents, John and Janet; words cannot
express what your love means to me. The challenges and obstacles throughout this
journey allowed me to change, grow, and discover who I am, and I would not have
survived if it wasn’t for your love, support, and guidance. My brother and sister, John
and Stacy, kept me grounded and were always a phone call away when I was homesick.
To the rest of the Eichorst and Schultz clans, I love you all very much and will be home
soon. My Polish/German heritage has been a huge part of my life, with roots stemming
from the back-of-the-yards of Halsted St. in Chicago to the polkas on Euclid Ave. in
Berwyn which I brought to work with me everyday. I will never forget the wisdom, love,
and integrity that my dearly, beloved grandparents and great aunts -- Edward J. Schultz,
Constance B. Schultz, Anna Grien, Leonarda Ciesla, John H. Eichorst, and Rose M.
Eichorst -- bestowed upon me. I love and dearly miss you all very much. All that I am

is because of you, and I hope you are all proud of me up there.

vi

TABLE OF CONTENTS

LIST OF TABLES ........................................................................................................... ix

LIST OF FIGURES .......................................................................................................... x

CHAPTER 1: MICROBIAL DIVERSITY IN SOIL AND THE PHYLUM

ACIDOBACTERIA
INTRODUCTION .......................................................................................................... 1
The soil environment ...................................................................................................... 2
Microbial diversity and function in soil .......................................................................... S
Cultivating the uncultivables .......................................................................................... 8
The phylum Acidobacteria ........................................................................................... 1 1
Thesis Outline ............................................................................................................... 18
Experimental Approach ................................................................................................ l 8
REFERENCES ............................................................................................................. 21

CHAPTER 2: ACIDOBACTERIA COMPOSITION FROM MIGHIGAN SOILS
IN CONJUNCTION WITH NOVEL CULTIVATION AND SCREENING
METHODS FOR THEIR SUCCESSFUL ISOLATION

INTRODUCTION ........................................................................................................ 3 1
MATERIALS and METHODS ..................................................................................... 34
RESULTS and DISCUSSION ...................................................................................... 43
SUMMARY .................................................................................................................. 63
ACKNOWLEDGEMENTS .......................................................................................... 66
REFERENCES ............................................................................................................. 67

CHAPTER 3: ISOLATION AND CHARACTERIZATION OF SOIL BACTERIA
THAT DEFINE TERRIGLOBUS GEN. NOV., IN THE PHYLUM
ACIDOBACTERIA

INTRODUCTION ........................................................................................................ 75
MATERIALS and METHODS ..................................................................................... 78
RESULTS and DISCUSSION ...................................................................................... 86
ACKNOWLEDGEMENTS ........................................................................................ 107
REFERENCES ........................................................................................................... 109

CHAPTER 4: DESCRIPTION OF NOVEL MEMBERS OF SUBDIVISION 1 AND
3 IN THE PHYLUM ACIDOBACTERIA FROM AGRICULTURAL SOIL

INTRODUCTION ...................................................................................................... l 17
MATERIALS and METHODS ................................................................................... 118
RESULTS and DISCUSSION .................................................................................... 125
ACKNOWLEDGEMENTS ........................................................................................ 146
REFERENCES ........................................................................................................... 147

vii

 

 

CHAPTER 5: THE EXPLORATION OF METHANOTROPHY AND
METHYLOTROPHY AMONG MEMBERS OF THE PHYLUM

ACIDOBACTERIA
INTRODUCTION ...................................................................................................... 154
MATERIALS and METHODS ................................................................................... 158
RESULTS ................................................................................................................... 162
DISCUSSION and FUTURE DIRECTION ............................................................... 167
ACKNOWLEDGEMENTS ........................................................................................ 174
REFERENCES ........................................................................................................... 175

CHAPTER 6: SUMMARY and SIGNIFICANCE ................................................... 179

APPENDICES

A P P E N D I X A: MEDIA COMPOSITIONS ......................................................... 185

A P P E N D I X B: RAW DATA FROM SOIL PLATING EXPERIMENTS ........... 189

A P P E N D I X C: LIST OF FUNCTIONAL & 16S rRNA GENE PRIMERS ........ 199

A P P E N D I X D: BACTERIAL GROWTH EFFICIENCY: UNDERSTANDING
ECOLOGICAL STRATEGIES OF COPIOTROPHS & OLIGOTROPHS ................... 203

viii

LIST OF TABLES

Table 1.1. Location, description, and sample type of various environments in which

members of the phylum Acidobacteria reside ............................................... 12-14
Table 2.1. Description of soil samples from the KBS LTER ................................. 35
Table 2.2. Soil pH, percent soil moisture, carbon concentration, and nitrogen
concentration ......................................................................................... 37
Table 2.3. Medium additives used for cultivation experiments .............................. 41
Table 3.1. Phenotypic characteristics of T erriglobus, Acidobacteriaceae, and
Acidobacterium strains .............................................................................. 97

Table 3.2. Whole-cell fatty acid composition (%) of T erriglobus, Acidobacteriaceae, and
Acidobacterium strains grown in GYE medium. The most abundant fatty acids that
distinguish the T erriglobus strains from A. capsulatum are presented in bold font.102-103

Table 4.1. Summary of growth and end products of KBS 83 & KBS 96 under anoxic
conditions with nitrate ............................................................................. 131

Table 4.2. Table 4.2. Phenotypic characteristics of Terriglobus, Acidobacteriaceae, and
Acidobacterium strains ........................................................................................... 134-136

Table D.1. Efficiency experimental strains and their growth characteristics ............. 210

Table D2. Intrinsic metabolic features and environmental factors that influence bacterial
growth efficiency .................................................................................. 224

ix

LIST OF FIGURES
Images in this thesis/dissertation are presented in color.

Figure 1.1. Representation of the soil environment. Gradients of pH, light, oxygen,
nutrients, temperature, and carbon dioxide are indicated on the left-hand side, and the soil
profile on the right-hand side. An outline of the carbon and nitrogen cycle are
represented. Microbial and viral communities are depicted pictorially ....................... 4

Figure 1.2. Common phyla of bacteria found in the soil environment. Each gray bar
represents the number of clones in each phylum; the black, shaded region represents the
percentage of the phylum that has been cultivated. Data was obtained from the
Ribosomal Database Proj ect-II in January 2007 .................................................. 9

Figure 1.3. Maximum-likelihood tree of the Acidobacteria subdivisions 1, 2, 3, 4, 5, 6,
and 7 (indicated to the right of the group) based on the 168 rRNA gene using sequences
obtained from cultivated representatives and environmental clones. Geothrixfermentans
and Holophagafoetida of subdivision 8 were used as an outgroup (not shown). The scale
bar indicated 0.10 changes per nucleotide ................................................... 16-17

Figure 2.1. Plate Wash PCR method used to detect the presence of the target
organism(s) on the three treatments (A, B, & C). Treatment C indicates a positive
detection for the target organism (66) ................................................................................ 42

Figure 2.2. Distribution of Acidobacteria subdivisions among the different agricultural
regimes: (A) T1 R1 0-7 cm; (B) T1 R2 0-7cm; (C) T7 R1 O-7cm; (D) T7 R4 0-7cm; (E)
T8 R1 0-7cm; and (F) T8 R2 0-7cm .............................................................. 45

Figure 2.3. Neighbor joining phylogenetic tree of the phylum Acidobacteria with 16S
rRNA gene derived from DNA extracted from KBS LTER soil. Geothrixfermentans and
H010phaga foetida of subdivision 8 were used as an outgroup (not shown). All boxes in
black and gray represent clones (partial sequence ca. 500 bp) fiom upper layers of T8 and
T1, respectively. The subdivision is indicated to the right of the phylogenetic tree. Bar
graphs are illustrated with the number of clones fiom each subdivision (SD#) with
respect to the two treatments. Scale bar indicates 0.10 changes per nucleotide ........ 46—47

Figure 2.4. Relationship between relative percentage of acidobacteria subdivision 3 and
% soil moisture .......................................................................................... 49

Figure 2.5. Relationship between relative percentage of acidobacteria subdivision 1 and
soil pH ................................................................................................... 50

Figure 2.6. Relationship between the relative percentage of subdivision 1 and methane
fluxes. Methane data was averaged for each treatment from historical data obtained fiom
the KBS LTER website for methane flux data collected between 1992 and 2001; a
detailed description of the sampling can be found in a previous study by Robertson and
collegues (55). 51

Figure 2.7. Relationship between relative percentage of acidobacteria subdivision 4 and
carbon concentration. Carbon data was averaged for each treatment from historical data
obtained from the KBS LTER website for carbon data collected between 1989 and 2001;
a detailed description of the sampling can be found at the KBS LTER website
(http://lter.kbs.msu.edu/) ................................................................................................... 53

Figure 2.8. Relationship between the relative percentage of subdivision 4 and nitrous
oxide fluxes. Nitrous oxide data was averaged for each treatment from historical data
obtained from the KBS LTER website for nitrous oxide flux data collected between 1992
and 2001; a detailed description of the sampling can be found in a previous study by
Robertson and collegues (55)54

Figure 2.9. Summary of cultivation experiments for total recovery of colonies per gram
of soil (dry weight) (Panel A) and acidobacteria detection (% of 11) (Panel B). Panel A
represents the average CFU recovery from soil +/- the standard error; the number of times
a particular treatment was employed (n) is listed at the base of each bar. Panel B
represents the percentage of treatments that yielded a positive test for acidobacteria using
PWPCR from the plates of panel A. The asterisks “*”indicates treatments that yielded
significant difference (panel A, AN OVA or = 0.05; panel B, chi square goodness of fit, or
= 0.10) ........................................................................................................................... 55-56

Figure 2.10. Relationship between total recovery of aerobic, heterotrophic soil bacteria
from T8 soil and % soil moisture. Panel A depicts a significant increase in total recovery
up to 21% soil moisture, whereas panel B illustrates the limit of soil moisture influence
on total recovery. Panel C represents the effect of recovery on aerobic heterotrophs from
KBS LTER treatment 8 soil (black boxes) and a garden plot (gray boxes) (68) ....... 58-59

Figure 2.11. Maximum likelihood tree of the Acidobacteria subdivisions 1, 3, 4, 5, and 6
(indicated to the right of the group) based on the 16S rRNA gene using sequences
obtained from cultivated representatives and environmental clones. Geothrix fermentans
and Holophaga foetida of subdivision 8 were used as an outgroup (not shown). Strains
from this study are in bold text. Each gray trapezoid represents a different subdivision.
The scale bar indicates 0.10 changes per nucleotide ............................................ 62

Figure 2.12. Summary of acidobacteria community composition and the edaphic
properties that influence the composition of subdivisions (SD) 1, 4, and 6... ... ......64

Figure 3.1. Summary of acidobacterial 16S rRNA gene clone libraries survey. Panel A
depicts the percent abundance of acidobacteria subdivision 1 out of the total
acidobacteria] community at various soil pHs. Data from the KBS LTER are depicted as
gray boxes ( I ), whereas data from published Soil surveys are depicted as black boxes (
I ). If a range in pH was given in these studies, the average of that range was used for
this analysis. The equation of the line is y = -13.1x + 103 with an R2 value = 0.38 and p-
value < 0.004. Panel B depicts the average percent abundance of acidobacteria
subdivisions 1, 3, 4, 5, & 6 (subdivisions are indicated next to each pie wedge) from

xi

replicate plots of the never-tilled, successional community with herbaceous plants at the
KBS LTER, depth of 0-7cm ....................................................................... 88

Figure 3.2. Summary of cultivation experiments for total recovery of colonies per gram
of soil (dry weight) (Panel A) and acidobacteria detection (% of 11) (Panel B). Panel A
represents the average CFU recovery from soil +/- the standard error; the number of times
a particular treatment was employed (n) is listed at the base of each bar. Panel B
represents the percentage of treatments that yielded a positive test for acidobacteria using
PWPCR from the plates of panel A. The asterisks “*”indicates treatments that yielded
significant difference (panel A, ANOVA or = 0.05; panel B, chi square goodness of fit, or
= 0.10) ............................................................................................. 90-91

Figure 3.3. Electron micrographs of thin sections of strains TAA 43 (a), TAA 48 (b),
TAA 166 (c), and KBS 63 ((1) grown in R2A. Negatively stained cell of strain KBS 63
(e) and a scanning electron micrograph of strain KBS 89 (f). Bars: (a-e) 500 nm, and (f)
2 pm ................................................................................................... 94

Figure 3.4. Strain KBS 63 grown on R2A under 20% and 2% oxygen (vol/vol) of
nitrogen & 5% C02 mix) (panel A). Absorption spectra of strain KBS 63 cells extracted
in acetonezmethanol (7:2) after growth in 20% ( I ) and 2% (I) oxygen on R2B (panel
B) ...................................................................................................... 95

Figure 3.5. Southern hybridization of digested genomic DNA from strains KBS 62, KBS
63, KBS 68, KBS89, KBS 112, TAA 43, TAA 48, & TAA 166. In panel A, isolates were
cut with EcoRV(lanes 2, 4, & 6), Sma 1 (lane 3, 5, 7, & 9), and Rsa I (lane 8). In panel B,
isolates were cut with Sma I (lanes 2, 4, 6, 8, & 10) and Apa I (lanes 3, 5, 7, 9, & 11).
The lambda DNA Hind III marker (lane 1 & 10 (panel A) and lanes 1 & 12 (panel 12))
provided size estimates ........................................................................... 100

Figure 3.6. Euclidian distance dendrograrn for T erriglobus, Acidobacteriaceae, and
Acidobacterium strains generated from the fatty acid profiles .............................. 104

Figure 3.7. Maximum-likelihood tree of the Acidobacteria subdivisions 1, 3, 4, 5, and 6
(indicated to the right of the group) based on 16S rRNA gene using sequences obtained
from cultivated representatives and environmental clones. Geothrix fermentans and
Holophagafoetida of subdivision 8 were used as an outgroup (not shown). Strains from
this study are in bold font. Each gray trapezoid represents a different subdivision with
approximately 8 representative sequences. Internal nodes supported by a bootstrap value
of >95% are indicated with a filled circle ( o ) and of >75 with an open circle ( o ). The
scale bar indicates 0.10 changes per nucleotide ................................................ 105

Figure 4.1. Summary of cultivation experiments. Panels A & B depict total recovery 1:
standard error after 30 days of growth from plates incubated under oxic conditions with
the media at a pH of 5.5 and 6.5, respectively. Panels C & D depict total recovery after
30 days for growth from plates incubated under hypoxic conditions with media at a pH of
5.5 and 6.5, respectively ............................................................................. 127

xii

Figure 4.2. Phase contrast and transmission electron micrograph of KBS 83 (panels A &
B) and KBS 96 (panels C & D). Arrow in panel A indicates the capsular material layer
produced by KBS 83. Scale bar indicates 1 pm (panels B & D) and 200 nm (panels A &
C) ..................................................................................................... 129

Figure 4.3. Summary of growth rates for KBS 83 under different concentrations of
oxygen. Note growth rate for an oxygen concentration of 0% was obtained from the
studies assessing the capacity to use nitrate .................................................... 138

Figure 4.4. Southern hybridization of digested genomic DNA from strains KBS 83 &
KBS 96. Strains were cut with EcoRI (lanes 2 & 5), EcoRVI (lane 3 & 6), and HindIII
(lane 4 & 7). The lambda DNA Hind III marker (lane 1 & 8) provided size
estimates ............................................................................................. 140

Figure 4.5. Neighbor-joining phylogenetic tree depicting the number of 16S rRNA gene
clone sequences generated from treatments 1 and 8 (black and gray boxes, respectively)
at the KBS LTER in the clade harboring strain KBS 83 ..................................... 143

Figure 4.6. Maximum-likelihood tree of the Acidobacteria subdivisions 1, 3, 4, 5, and 6
(indicated to the right of the group) based on 16S rRNA gene using sequences obtained
from cultivated representatives and environmental clones. Geothrix fermentans and
Holophagafoetida of subdivision 8 were used as an outgroup (not shown). Strains fiom
this study are in bold font. Each gray trapezoid represents a different subdivision with
approximately 5 representative sequences. Internal nodes supported by a bootstrap value
of >95% are indicated with a filled circle ( 0 ) and of >70 with an open circle ( o ). The
scale bar indicates 0.10 changes per nucleotide ................................................ 144

Figure 5.1. Simplified overview of the pathway for the oxidation and assimilation of one-
carbon compounds into cellular material using the RuMP, RuBP, or serine
pathways ............................................................................................ 156

Figure 5.2. Acidobacteria clones obtained after incubation of acidic oak forest soil
samples with 350 mM methanol (26)157

Figure 5.3. Maximum likelihood tree of the phylum Acidobacteria, subdivision 1.
Sequences were obtained from Genbank and accession number are listed parenthetically.
Sequences obtained from Radajewski and colleagues (26) are in bolded text. The scale
bar indicates a 0.10 change per nucleotide ..................................................... 164

Figure 5.4. Methanol consumption by KBS LTER replicate soil microcosms, replicate l
(gray boxes) and replicate 2 (black boxes). Each point represents the mean :1: standard
deviation fiom triplicate measurements of methanol in the headspace as measured by GC-

xiii

Figure 5.5. Neighbor-joining tree of the Acidobacteria subdivisions 1, 3, 4, 5, and 6
based on 16S rRNA gene using sequences obtained from cultivated representatives and
environmental clones. Geothrixfermentans and Holophagafoetida of subdivision 8 were
used as the outgroup (not shown). Strains from this study are in bolded text. JAB 971 in
subdivision 1 is a partial 16S rRNA gene sequence from cultivation experiments using
soil from the methanol microcosms as a source of inoculurn. The scale bar indicates 0.10
changes per nucleotide ............................................................................ 168

Figure 5.6. Neighbor-joining tree of partial sequences of 16S rRNA gene clones in the
phylum Acidobacteria obtained before and after incubation of KBS LTER treatment 8
soil with 350 mM methanol. Acidobacteria] 'clones before and after the addition of
methanol are represented in light gray and dark gray boxes, respectively. White boxes
with black outlines represent Acidobacteria 16S rRNA gene clones obtained fiom oak

forest soil incubated with l3C-methanol, previously (26) ............................... 169-170
Figure 5.7. G + C content of 12C- and 13C- DNA in relation to its buoyant density on a
cesium chloride gradient .......................................................................... 172
Figure D. 1. Classic organization of the rrn operon ........................................... 206

Figure 0.2. Methodology for trapping ”C-labeled carbon dioxide in the bacterial growth
efficiency experiments. Approximately 5mL 'of a 10% (vol/vol) tricholoracetic acid
solution was added to the 30 mL of culture to acidify, thereby allowing the dissolved
carbon dioxide to convert into a gaseous form. The gaseous carbon dioxide was trapped
with a carbon dioxide trap (phenylethylamine and methanol, 1:1 (vol/vol)) upon venting
the headspace with nitrogen gas ................................................................. 212

Figure D.3. Neighbor-joining phylogenetic tree of the 16S rRNA gene of strains used to
determine bacterial growth efficiency. Strains from this study are in bolded text; bolded
in black represent copiotrophs (n=4) whereas bolded in gray represents oligotrophs (n=4).
The rrs copy number and major phylogenetic affiliations are illustrated to the right of the
phylogenetic tree. The scale bar indicates 0.10 changes per nucleotide. ...................... 215

Figure D.4. Relationship between bacterial growth rate and rrs copy number. Each point
represents the average growth rate from two replicates grown in the VSB-6,7 with
2.5mM glucose and Sug/mL yeast extract at room temperature ............................ 217

Figure D.5. Panel A depicts the relationship between bacterial growth efficiency and cell
yield. Panel B depicts the relationship between bacterial growth efficiency and rrs copy
number .............................................................................................. 218

Figure D.6. Percent of ATP-dependent proteins (out of the total proteins) versus rrs copy

number for non-symbiotic heterotrophs. Data obtained from 133 sequenced genomes
from the TransportDB (http://www.membranetransport.org) (31) .......................... 220

xiv ‘

Figure D.7. Percent of response regulators in two-component systems not associated
with chemotaxis, versus rrs copy number for bacteria and archaea. Data obtained 187
sequenced genomes obtained from TIGR
(http://www.ncbi.nlm.nih.gov/Complete Genomes/RRcensus.html) ............................. 223

Figure D.8. Proposed model for future bacterial growth efficiency experiments. Panel A
depicts the proposed relationship between bacterial growth efficiency (BGE) versus
dilution rate. Panel B depicts the proposed linear relationship between dilution rate, a
proxy for resource concentration indicating rate at which the organism reaches its
maximum growth efficiency (Panel A) measured in a chemostat, versus rrs copy
number .............................................................................................. 227

Figure D.9. Proposed model for future bacterial growth efficiency experiments. Panel A
depicts the proposed relationship for maintenance energy for oligotrophs and copiotrophs
as illustrated by Pirt’s double reciprocal plots of the inverse of yield (1/Y) versus the
inverse of growth rate (1/ u) at steady state. Panel B depicts the proposed linear
relationship between maintenance energy of oligotrophs and copiotrophs grown at steady
state dilution rates versus rrs copy number .................................................... 228

XV

CHAPTER 1

MICROBIAL DIVERSITY IN SOIL AND THE PHYLUM ACIDOBAC T ERLA

INTRODUCTION

Soil is a dynamic, three-dimensional environment containing an abundant source
of life. This environment encompasses an area of ca. 1.23 x 1014 m2 on Earth (93), which
is home to a multitude of our planet’s biodiversity. It is also the foundation of our society
supporting our way of life primarily as a growth meditun for food such as corn, soybean,
and wheat to sustain both animal and human life. It is ironic that we know and
understand very little about this diverse environment, since we are highly dependent on
the community living within the soil environment.

In this work I explored a major, but poorly understood group of soil bacteria
called the Acidobacteria. Acidobacteria are in high abundance in the soil environment
based on rRNA studies suggesting that they are essential to the soil environment. My
underlying goal was to understand the ecological role(s) of the phylum Acidobaceria in
soil. I addressed this goal by combining molecular and growth—based approaches. This
research is necessary to build a strong foundation of knowledge and understanding about

this bacterial group. It justifies and demands the need for further research addressing the
impact the Acidobacteria have on nutrient cycling, biogeochemistry, and agricultural

practices.

The soil environment

Soil is a complex habitat of living and non-living components. Air and water
make up the pore space, and mineral and organic matter make up the solid space (7). The
organic matter includes nutrients, living microbiota, and organic matter in various stages,
whereas the mineral fraction is classified as being either larger sized particles of sand,
silt, or smaller sized particles of clay (86). Depending on the soil, the relative proportions
of these components vary which in turn can affect the composition of the microbial
communities (29, 90).

Solid soil particles are in association with eachother due to interactions with
factors such as humic substances, bacterial polysaccharides, plant roots, and fungal
hyphae (67) which together form soil aggregates. The formation of these aggregates and
the ability of these particles to withstand change defines the overall soil structure (32).
Soil aggregates are divided up into two main categories based on composition and size:
microaggregates and macroaggregates. Microaggregates, defined as having a diameter of
< 250 um, are composed of humic molecules, clay minerals, microbes, and amorphous Al
and Fe oxides, whereas macroaggregates typically have a diameter of > 250 um and are
composed of the smaller sized microaggregates often adhering to plant roots (32, 67).
The organic matter in the soil allows these different sized aggregates to withstand
external forces, and often there is a linear relationship observed between aggregates
stability and organic matter content of the soil (32). Within soil aggregates exists a
gradient of oxygen (7, 67, 91), creating anoxic microsites, which are ideal for metabolic
processes such as denitn'fication and fermentation as well as providing transition zones

between oxic and anoxic conditions (37, 79). Methods have been developed to study the

communities of these aggregates by separating them based on class size (81) as well as
analyzing the outer- and inner- microaggregate communities (64). In general, microbial
diversity increases with soil aggregate size which is in turn influenced by soil
management (57, 62); in this way aggregate size affects the overall microbial community
composition (63).

Aggregates create a three-dimensional structure amidst gradients of carbon,
nitrogen, phosphorous, sulfur, oxygen, carbon dioxide, pH, moisture, and temperature
(Figure 1.1). The foremost gradient in this environment is the soil profile made up of
different horizons with the O horizon at the soil-air interface followed by the A, B, and C
horizons atop the underlying bedrock. The soil content changes with depth from the
surface (0 horizon), which primarily consists of organic matter, to clays, oxides, and
carbonates (A, B, and C horizons), and eventually transforms into salts and minerals
(bedrock) (7).

Soil is one of the major reservoirs of organic carbon on the planet (93) with most
of the carbon resulting from the degradation leaves, roots, fauna, and microbes; however
it is typically defined as carbon-limiting since the carbon is not present in a readily usable
form. Microbes are responsible for the breakdown of this recalcitrant carbon, which is an
astronomical feat taking into account that this community comprises only ca. 5% of the
overall available space in soil (39). Organic carbon in soil is typically composed of high
molecular weight organic molecules known as hurrric substances; for example, in mineral

soils humic substances account for ca. 70 to 80% of the total organic matter (32). The

 

 

 

 

 

Figure 1.1. Representation of the soil environment. Gradients of pH, light, oxygen,
nutrients, temperature, and carbon dioxide are indicated on the left-hand side, and the soil
profile on the right-hand side. An outline of the carbon and nitrogen cycle are
represented. Microbial and viral communities are depicted pictorially.

amount of soil organic matter in a particular soil is influenced by soil management. Soil
organic matter is of the utmost importance to this environment as it contributes to soil
fertility by acting as a reservoir for nutrients such as nitrogen, sulfur, and phosphorus;
source of water; maintenance of proper cation exchange; and ultimately is essential in the

formation and stability of the soil structure (32).

Microbial diversity and function in soil

The soil has an abundant wealth of life including members of the Domains
Archaea, Bacteria, and Eukatjya. Eukaryotic life found in soil includes, but is not limited
to, algae, nematodes, protozoans, lichens, earthworms, insects, arthropods, plants, and
fungi. Fungi, an often overlooked group, are responsible for maintaining the cycle of
nutrients through the decomposition of plant material such as root exudates, lignin, and
cellulose; soil stabilization; and balancing of other microbial populations through
predation on certain species (18). There is a great deal of untapped fungal diversity, as
seen by a recent survey of basidiomycetes in Michigan agricultural soils which revealed
multiple, unidentified species (58).

Another poorly studied group in the soil environment is viruses. Viruses,
typically lytic phages, are a source of mortality for soil microbes (13), causing a release
of nutrients as well as genetic material into the surrounding environment. Determination
of virus diversity is typically based on morphotype as viewed by electron microscopy
with such groups as tailless, Podoviridae, filamentous, Siphoviridae, and Myoviridae.

Viruses are influenced by land use, moisture, water content, and soil organic matter and

 

have been observed with a high abundance, ca. 1.9 x 109 virus-like particles/gram soil
(dry weight), in forested and agricultural soils (94).

The bacteria and archaea encompass a great wealth of diversity and importance.
One study estimated that the worldwide soil environemnt contains ca. 2.6 x 1029 bacteria
and archaea (93). The importance of these communities becomes clear when estimating
their enormous carbon contributions of ca. 5 to 7% to the planet (93). Additionally their
contributions to the total amount of nitrogen and phosphorous are similar to that of plants
(93).

Bacteria and archaea play a major role in the carbon and nitrogen cycles. The
majority of the carbon in the soil environment results from the bacterial and archaeal
decomposition of plant polymers and sugars. Plant polymers are hydrolysed and
degraded into simple sugars by a variety of microorganisms, oftentimes by a
collaborative effort of different microbial groups. This carbon is converted into
microbial biomass, and ultimately degraded into carbon dioxide. Carbon dioxide can be
used to produce biomass via autotrophic pathways, methane by the methanogenic
archaeal communities which is further oxidized by nearly methanotrophic communities,
or released into the atmosphere.

Nitrogen, another important, limiting nutrient in the soil environment, is
introduced to the soil environment in one of three ways: (i) nitrogen fixation exclusively
performed by bacterial and archaeal communities, (ii) addition of fertilizer or deposition
of nitrogen, and (iii) decomposition of plant and microbial biomass. Microbial
decomposition of plant and microbial biomass releases various forms of nitrogen, such as

ammonium, into the surrounding community. Bacteria oxidize ammonium to nitrite and

nitrate through a process known as nitrification. Nitrification is predominantly performed
by the y-Proteobacteria and B-Proteobacteria in soil, however it was recently discovered
that members of the Kingdom Crenarchaeota have this capacity (46). Nitrate and nitrite,
under anoxic conditions, can be used as a terminal electron acceptor whereby these forms
of nitrogen are sequentially reduced to nitric oxide, nitrous oxide, and dinitrogen gas
which is referred to as denitrification. This is a heterotrophic, facultative trait common to
organisms including the proteobacteria, gram positive bacteria, archaea, and fungi (99).

The community of soil bacteria is very diverse based on 16S rDNA studies; some
have even described this unique environment as being a “microbial rain forest” (9),
containing exotic species of bacterial life. Members of the phyla Proteobacteria,
Acidobacteria, Verrucomicrobia, Cytophagales, Actinobacteria, F irmicutes, and
Planctomycetes are dominant members in the soil environment with their abundance
varying depending on the environmental condition. In addition to these bacterial phyla,
nontherrnophilic members of the Kingdom Crenarchaeota are diverse and abundant
members of the soil environment accounting for as much as 1.42 :t 0.42% of the total
community (10).

Over the years, numerous methods have been developed to study the soil
microbial diversity. Common techniques include rDNA (typically the 16S/18S small
subunit rRN A) based analyses such as terminal restriction fragment length
polymorphism, sequencing of the 16S or 18S rRNA gene, amplified ribosomal DNA
restriction analysis (ARDRA) or restriction fragement length polymorphism (RFLP),
while other techniques used are determination of the G + C content, BIOLOGTM profiles,

stable isotope probing, fatty acid analysis, nucleic acid reassociation curves, and DNA

microarrays (44, 87). Depending on the application and question, a basic picture of the
structure and/or function of specific microbial community can be obtained with these

methods; however these techniques yield no new cultivated representatives.

Cultivating the uncultivables

Soil is a plentiful source of life. Based on total microscopic counts, the soil
environment contains approximately 109 to 1010 microorganisms per gram of soil dry
weight. The soil community is estimated to contain over a million species per gram of
soil (27); as a result soil harbors more genera and species of microorganisms than any
other habitat on the planet (91).

However, only a small percentage 0.1 to 1% of the total soil microbial community
has been cultivated (88, 89) based on direct counts. This phenomenon is referred to as
the “great plate count anomaly” (83) owing to the observation that there are ofientimes
orders of magnitude differences between counts from cultivation plates and direct counts
from the native environment. The lack of cultivated representatives of common soil
bacterial phyla becomes evident when one compares the number of cultivated sequences
to the total number of sequences (Figure 1.2). It has been suggested that these cells have
not been cultivated not because they are uncultivable; rather microbiologists are not
providing the proper grth conditions for these bacteria. Cells are viable but as a result
of various environmental stresses, they enter a dormant state where they cannot be
cultivated under standard conditions (6); oftentimes referred to as the viable but non-
culturable state (V BNC). This argument has been used to explained the uncultivability of

such organisms as Enterococcusfaecalis (33) and Vibrio vulnificus (66).

   
 
   
 
   
   
   
 

Alpha
Beta Proteobactcria
Gamma Proteobacteria
Deha
Epsilon

Plantomyces

Bacteroidetes
Cyanobacteria
0 10000 20000 30000 40000 50000 60000 70000
Number of sequences
Figure 1.2. Common phyla of bacteria found in the soil environment. Each gray bar
represents the number of clones in each phylum; the black, shaded region represents the

percentage of the phylum that has been cultivated. Data was obtained from the
Ribosomal Database Proj ect-II in January 2007.

Recalcitrant microorganisms such as members in the phyla Proteobacteria,
Acidobacteria, Verrucomicrobia, Plantomyces, Actinobacteria, Firmicutes, and
Bacteriodetes continue to evade our cultivation experiments, and as a result our
understanding of these microorganisms’ roles is remaining obscure. The need to retrieve
this “not-yet-cultured majority” (8, 69) has become evident with the footprint of these
microbial communities becoming more and more defined with molecular techniques.
Our poor understanding of microorganisms’ biochemistry and a lack of patience to grow
these presumed, slow growing microbes (52) is to blame for our low cultivation recovery,
since typical cultivation experiments include short incubation times on nutrient rich
medium, under oxic atmospheres. Recently, microbiologists have attempted to address
the “great plate count anomaly” by developing new, novel cultivation methods such as
diffusion chambers to isolate aerobic organoheterotrophs from intertidal marine
sediments (43), extinction culturing with low-nutrient media from oligotrophic marine
environments (14), encapsulation of cells in gel microdroplets (96), and novel cultivation
strategies combined with facile, high throughput methods for screening such as plate
wash PCR (PWPCR) (85).

In addition to these growth-based techniques, the innovative field of
metagenomics (31) has developed in recent years as a culture-independent means to gain
insight into the physiology of novel groups of microorganisms by discovering novel
genes and revealing their physiology from such enviromnents as soil (23, 34, 35, 70),
seawater (4, 5, 15, 84), and marine sponges (77). Data obtained from metagenomic
projects in conjunction with the patience and desire to isolate microorganisms in pure

culture will be the synergism needed to push the scientific community forward in the 21St

10

century. The combination of patience with the satisfying reward of isolating novel
groups of microorganisms was summarized best by Peter H. Janssen:
“Successful approaches to culturing these microorganisms require patience, but
the outcomes are immensely satisfizing to microbiologists who enjoy the challenge
and savor the reward of observing colony on the plate, seeing the cells of the pure
culture under the microscope, elucidating the bacterium ’s physiology, or

releasing its genome sequence into public databases ” (40).

The phylum Acidobacteria

A group of bacteria that are prevalent in the soil environment, but have few
cultivated representatives are members of the phylum Acidobacteria. This phylum is
defined by a large collection of 16S rRNA gene sequences (>3,000 in the Ribosome
Database Project) (12) retrieved from diverse environments including: soils and
sediments (3, 21), soil crusts of sand dunes (82), wastewater (16, 50), water distribution
systems (60), peat bogs (l9), acid mine drainage (45), hot springs (36), shallow
submarine hydrothermal vents (80), and on the surface of Paleolithic cave paintings and
catacombs (74-76, 97, 98) (Table 1.1). In situ hybridization with acidobacteria-specific
probes has also confirmed the presence of intact acidobacteria in many environments, and
revealed multiple cellular morphotypes, including cocci, short rods, and thin filaments

(56).

Historically, the phylum Acidobacteria has been comprised of three organisms,
Acidobacterium capsulatum, Holophaga foetida, and Geothrz'x fermentans. The name

A cidobacterium was first introduced in the early 1990’s after a group of Japanesse

ll

Table 1.1. Location, description, and sample type of various environments in which
members of the phylum Acidobacteria reside.

12

 

Table 1. Distribution of members of the phylum Acidobacteria from environmental samples

 

Location Description Sample Type Reference
Soils
Sunset Crater. Ariz. Volcanic Cinders (3, 21, 48)
Sunset Crater, Ariz. Pinyon rhizosphere Cinders (3, 21, 48)
Cosnino, Ariz. Pinyon woodland Sandy loam (3, 21, 48)
Cosnino, Ariz. Pinyon rhimsphere Sandy loam (3, 21, 48)
Las Cruces, N. Mex. Chile Field Sandy loam (3, 21, 48)
Raleigh, NC. Meadow Clay loam (3, 21, 48)
Elk River, Minn. Garden black soil Organic (3, 21, 48)
Paineville, Ohio Agricultural Sandy loam (3, 21, 48)
Elk River, Minn. Aged cow manure Organic (3, 21, 48)
Oslo, Norway Boreal forest Organic (3, 21, 48)
Antelope Valley, Calif. Citrus grove Sandy loam (3, 21, 48)
Dugway, Utah Semiarid desert Sandy clay loam (3, 21, 48)
Elk River, Minn. Gravel Pit "white peat" Silt organic (3, 21, 48)
Niceville, Fla. Pine Forest Sandy (3, 21, 48)
Poncha Pass, Colo. Roadside Hill Sandy (3, 21, 48)
Berkeley Heights, NJ. Hardwood forest Organic (3, 21, 48)
Ohio Cranben'y bog Muck; rifle peat (3, 21, 48)
Cologne, Germany Oak-hombeam forest no data (71)
Drentse A, Netherlands Dutch Grasslands loamy fine sand (24-26)
Ellinbank, Victoria, Australia Dairy Research Institute basaltic clay loam (17, 4042’ .72, 73)
Kellogg Biological Station Long Term Ecological Research Site Kalamazoo sandy loam (22, 85)
Colorado Rocky Mountains Niwot Ridge Long Term Research Site Skeletal-loamy pergaelic (55)
Cape Cfiggéfifligse ashore, Sand dunes Crusted sands (82)
Yellowstone National Park Ragged Hills, Norris Geyser Basin No data (65)
Madison, Wisconsin West Madison Agricultural Station No data (54)
Canyonlandlsjtljlational Park. Arid grasslands Calcareous loamy sand (49)
Western, Morroco Agronmsiiii‘ifiziigggnigl;Egghfihcenate Vertisol (I)
Yorkshire, England Gisbum Forest Experiment Acidic oak forest soil (68)
Ultuna, Sweden Ultuna long-term field experiment No data (78)
Eschikon, Switzerland Monoculture soil Clay loam (59)
Border Region, Scotland Sourhope Research Station Rhizosphere soil (61)
Sediments and mats
Los Alamos, N. Mex. Marsh (anaerobic) Organic (3, 21, 48)
Abiquiu, N. Mex. River (anaerobic) Organic (3, 21, 48)

 

l3

 

Table 1.] continued

Location Description Sample Type Reference
Sediments and mats continued
Los Alamos, N. Mex. Shallow Lake Organic (3, 21, 48)
Santa Rosa 1., Fla. Shallow Marine Sandy (3, 21, 48)
Pine Barrens, NJ. Cedar Swamp Organic (3, 21, 48)
Pine Barrens, NJ. Hardwood forest Streambed (3, 21, 48)
Yellowstone National Park 71°C hot spring Sediment (3, 21, 4s)
Yellowstone National Park 670C hot spring Mat (3, 21, 48)
Yellowstone National Park 640C hot spring Mat (3, 21, 48)
Aiken, S.Carolina Rainbow Bay no data (95)
Vercelli, Italy Rice paddles (anoxic) no data (92)
Other
Santillana del Mar, Spain Altamira Cave no data (74)
Ribadesella, Spain Tito Bustillo Cave no data (75)
Rome, Italy Saint Callixtus Catacombs Epilithic biofilms (97)
Aegean Sea, Milos, Greece Hydrothermal vent no data (80)
Wastewater , Indiana Bioreactors no data (51)
Queensland, Australia Wastewater batch reactors no data (16)
West Siberia, Russia Peat samples Bakchar, Plotrrikovo Sphagnum peat bog ( 19)
North Wale}? United Myny d d Parys & Cae Coch mines Acidic, metal-rich mine (30)
lngdom waters

 

14

scientists isolated eight novel strains (termed, Biogroup 5) from an acidic mineral
environment. This was the first report of an isolation of an acidophilic heterotrophic
bacteria inhabiting acidic mineral environments other then Acidiphilium (45, 47). The

phylum was named Acidobacteria after the only validated species at the time (56).

Shortly after the isolation and characterization of A. capsulatum, H. foetida was
isolated and described as a new homoacetogenic bacterium capable of degrading
methoxylated aromatic compounds (53). Given the high sequence similarity of its 16S
rRNA gene to A. capsulatum (ca. 81.6%), it was placed in the phylum Acidobacteria.
Additionally, G. fermentans, a bacterium capable of Fe (III) reduction, was added to this
new phylum based on the high sequence similarity of its 16S rRNA gene (ca. 94%) to H.

foetida (11).

The phylum Acidobacteria is now officially recognized in the Bergey’s Manual of
Systematic Bacteriology (28) and includes three genera with cultured representatives:
Acidobacterium (45), Geothrix (11), and Holophaga (53). Currently, there are three

proposed genera in this phylum; Solibacter (mutajgi.doe.gov), Terriglobus (22), and

 

Edaphobacter (WWVM’flCbLnithV). This phylum contains eight recognized

 

monophyletic subdivisions (38) (Figure 1.3) that encompass the molecular diversity
originally recognized as Acidobacteria (48), along with additionally unnamed and mostly
uncharacterized cultivars in subdivisions 1, 2, 3, and 4 (17, 19, 20, 30, 40-42, 71-73, 85).
Recently, a survey of 16S rRNA genes in microbial communities associated with
uranium contaminated subsurface sediment uncovered additional novel acidobacteria,

expanding the number of subdivisions to as many as twenty-six (2).

15

Figure 1.3. Maximum-likelihood tree of the Acidobacteria subdivisions 1, 2, 3, 4, 5, 6,
and 7 (indicated to the right of the group) based on the 16S rRNA gene using sequences
obtained from cultivated representatives and environmental clones. Geothrixfermentans
and Holophagafoetida of subdivision 8 were used as an outgroup (not shown). The scale
bar indicates 0.10 changes per nucleotide.

16

nv. 3220 (295713) 7
env. iii1e8 (Z9572 9)

env. Bai138 (A10014Z4)

nv. mb512l28 )(295733)
env. R1327 (Z95

 
   
  
   
   
    

env AzC028 (A 013527) 6
env. RBZ4(Z 957
cm RB4en0) (Z957

v.pB1 1214 (U86489)
env kb2426 (2:115732)
env.AzCI 12(A0F013534)
env. DA023(Y 758

env. Asz43 (U68625)
env.clone (AF431450)

ll 7) 5
UBA519393 ((AJS 19393)
e.nv clone (AY214798)
env. clone (AY150893)
_ env. clone en(AY150 99)
v1.124 (295708
eennv. vAzSI II (AF013560)
E11in6099 (AY234751)
env.AmzP12 (U68646)
env.1125(295 09)
env.AzC105 (AF013530) 4
cnv.3211 (29571RI)
Vean 20)
A5027: (AZFOI7 3554)
env.AzSO213V(AF0135)50)
env. RB41 (Z957

 

 

     
   
         
  
   

 

 

env.Asz26 (U68612) ] 2
env. DA052 (Y07646)

env. AmzPIS (U68648)

27 (X 466)
env. WDZS4(WA1295282)
env 05 (M29257l)
EIIin37I (AF498753)

b)

 

env.SJA I49 (AJ009495)

 

(AF050548)

 

Thesis Outline

The biodiversity of the soil environment is poorly understood, although we rely
on it quite extensively as a society. The exploration of this untapped microbial diversity
allows us to understand the role microorganisms play in the environment, which will in
turn enhance our understanding of nutrient eyeling, biogeochemistry, and the effect of
agricultural practices. This research was a union of both molecular and growth-based
approaches, which generated information pertaining to a ubiquitous and abundant group

of soil bacteria called the Acidobacteria.

My underlying, long-range objective was to understand the ecological role(s)
members of the phylum Acidobacteria have in, the soil. In order to address this long-

range objective, I have divided my thesis into three main goals:

1. Determine the composition of the phylum Acidobacteria in the soil environment,
particularly assessing the influence of edaphic properties on the distribution of the

different monophyletic subdivisions.

2. Develop novel cultivation strategies and screening methods to cultivate members

of the phylum Acidobacteria.
3. Characterize novel members of the phylum Acidobacteria from a genotypic,
physiological, and morphological perspective as a first step towards

understanding this interesting and diverse group of bacteria to ultimately

understand their ecological role(s) in the soil environment.

Experimental Approach

18

These goals were addressed with soil collected from Michigan State University’s
WK. Kellogg Biological Station Long Term Ecological Research Site (KBS LTER).
The KBS LTER is a 48-hectare research site established in 1989 to study ecological
processes in agroecosystems. The dominant soil series are Kalamazoo (fine-loamy,
mixed, mesic Typic Hapludalfs) and Oshtemo (coarse-loamy, mixed, mesic Typic
Hapludalfs). A detailed description of this site can be accessed at

http://lter.kbs.msu.edu/.

 

Chapter 2 describes a phylogenetic survey of the phylum Acidobacteria from
various treatments at the KBS LTER as well as novel cultivation strategies and screening
methods used to isolate members of this phylum. The phylogenetic survey revealed
information linking the different subdivisions with various edaphic properties. In
addition, trends linking members of subdivision 1 to soil pH were consistent with data
obtained during the characterization experiments in Chapters 3 and 4. PWPCR, a rapid,
high-throughput method (85), as well as novel cultivation strategies employed with
particular attention to the conditions found in the native soil environment (22), were
developed and instrumental in the successful isolation of members of subdivisions 1 and
3.

In Chapters 3 and 4, the newly isolated members of subdivisions 1 and 3 were
characterized with particular attention to genotypic, physiological, and morphological
properties which helped to provide insight into their presumed ecology in the soil
environment. During these experiments, data revealed that a group of newly isolated

members of subdivision 1 warranted the creation of a new genus, T erriglobus.

l9

Finally, in Chapter 5, I explored the capacity of acidobacteria to oxidize one-
carbon compounds. Previous work with the use of stable isotope probing with 13C-
methanol indicated that members of the phylum Acidobacteria from oak-forest soil
samples were potentially capable of oxidizing methanol, since a portion of the 13 C DNA
isolated was acidobacterial (68). Molecular and growth—based approaches were used to

explore this interesting physiology in acidobacteria.

20

10.

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30

CHAPTER 2

ACIDOBACTERIA COMPOSITION FROM MIGHIGAN SOILS IN
CONJUNCTION WITH NOVEL CULTIVATION AND SCREENING METHODS

FOR THEIR SUCCESSFUL ISOLATION

Some of these results have been published in the article: Stevenson, B. S., S. A. Eichorst,
J. T. Wertz, T. M. Schmidt, and J. A. Breznak. 2004. New Strategies for Cultivation and
Detection of Previously Uncultured Microbes. Appl. Environ. Microbiol. Vol. 70, No. 8,

pgs. 4748-4755.

The raw data for the cultivation experiments from the KBS LTER can be found in

Appendix B.

INTRODUCTION

Soil is a heterogeneous environment containing over one million different species
per gram (26), harboring more species of microorganisms than any other habitat on the
planet (72). Additionally, this community of soil microbes is very diverse based on 16S
rDNA studies; some have even described this unique environment as being a “microbial
rain forest” (10), containing exotic species of microbial life. Members of the phyla
Proteobacteria, Acidobacteria, Verrucomicrobia, Cytophagales, Actinobacteria,
F irmicutes, and Planctomycetes are dominant members in the soil environment with their

abundance varying depending on the environmental condition.

31

Unfortunately, only a small percent, 0.1 to 1%, of the total microbial community
has been cultivated from soil (39, 49), revealing a glimpse of the diversity in culture.
Excitingly, the soil microbial diversity continues to grow with every 16S rRNA gene
analysis uncovering additional novel, uncultivated phyla such as OPll (5, 42), TM7 (5),
and W86 (7 5), subsequently reducing our cultivation percentage more and more. These
recalcitrant microorganisms continue to evade our cultivation experiments primarily
because of our poor understanding of their biochemistry and a lack of patience to grow
these presumed, slow-growing microorganisms (46). Recently, rrricrobiologists have
attempted to address our inadequate cultivation techniques by developing new, novel
methods such as a diffusion chamber to isolate aerobic organoheterotrophs from
intertidial marine sediments (39), extinction culturing with low-nutrient media from
oligotrophic marine environments (16), extending incubation times to accommodate
slow-growing bacteria (37, 57), encapsulation of cells in gel microdroplets (74), and
novel cultivation strategies (25) combined with facile, high throughput methods for
screening such as plate wash PCR (PWPCR) (66).

Not only is there a lack of cultivated representatives in such phyla as
Proteobacteria, Acidobacteria, Verrucomicrobia, Cytophagales, Actinobacteria,
F irmicutes, and Planctomycetes, there is little to no information linking their abundance
and distribution to edaphic properties. Ofientimes scientists are faced with the
conundrum of trying to cultivate an organism having little to no information about their
physiology. Information generated from soil survey(s) linking edaphic properties to the
abundance and/or composition of these phyla provides a roadmap to bring about their

successful isolation as well as insight into their physiology. This knowledge can be used

32

to develop strategies, such as media compositions and incubation conditions, to
successfully isolate these poorly cultivated phyla. When members of these phyla have
been successfully isolated, we can determine if the trends observed in the soil surveys are
consistent with their physiology.

Members of the phylum Acidobacteria are a prime example of a dominant group
of soil bacteria with few cultivated representatives and no information linking their
abundance and/or composition to edaphic properties. The Acidobacteria is a phylum of
soil bacteria that are known to be present in soil in high abundance in various types of
environments including soils and sediments (3, 24), soil crusts of sand dunes (64),
wastewater (18, 45), water distribution systems (48), peat bogs (23), acid mine drainage
(41), hot springs (33), shallow submarine hydrothermal vents (63), and on the surface of
Paleolithic cave paintings and catacombs (58-60, 76, 77). Acidobacteria is a large,
monophyletic group of Bacteria historically harboring eight different subdivisions with
few cultured representatives (21, 23, 29, 36-3 8, 56, 57, 66) with recent evidence of
eighteen additional subdivisions, some unique to uranium contaminated subsurface
sediments (2). Their common occurrence and high abundance suggests they play an
ecologically significant role in the soil environment which remains unknown due to the
lack of cultivated representatives. In addition to the lack of cultivated representatives,
there is little to no information linking these eight monophyletic subdivisions of the
Acidobacteria to various edaphic properties such as moisture, pH, carbon content,
nitrogen content, or soil type, which could provide insight into new methods for their

successful isolation as well as features unique to a particular subdivision.

33

The purpose of this study was three-fold: (i) determine the acidobacteria
composition in the different treatments at the Kellogg Biological Station Long Term
Ecological Research (KBS LTER) site; (ii) assess relationships between the acidobacteria
composition and edaphic properties; and (iii) increase the recoverability of soil
microorganisms, specifically acidobacteria, using recognized supplements for increased
recovery such as low nutrients in the form _ of organic carbon (16, 50), extended
incubation times (3 7, 57), inclusion of quorum sensing compounds (N-acyl homoserine
lactones) (9, 12) as well as novel methods such as the catalase enzyme and anthraquinone

disulfonate and incubations under atmospheric gas concentrations native to soil.

MATERIALS and METHODS
Molecular Survey
Soil collection Soil samples were collected August 2005 from the Michigan State
University WK. Kellogg Biological Station Long Term Ecological Research Site (KBS
LTER, Hickory Comets, MI, USA). The KBS LTER is a 48-hectare plot designed to
study ecological processes in agroecosystems, which prior to its establishment in 1989
was uniformly farmed for more than 50 years. A description of this site can be accessed

at httL/Mwwnkbsmsuedu. The soil type is defined as a Typic Hapludalf, consisting of

 

sandy to silty clay loam with moderate fertility. Two series of soil cores (2.5 cm
diameter) were collected from the upper 7 centimeters of five sampling stations in two
replicate l-ha plots from early succssional, mid succesional, and conventional
agricultural soils (Table 2.1), as well as additional un-replicated samples from the upper 7

centimeters of the deciduous forest site and loWer 13 to 20 centimeters of treatments 7

34

 

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35

and 8. These pooled soil cores from each plot were homogenized with a 2-mm sieve, and
portions were used to determine soil pH and soil moisture (54) (Table 2.2); the remaining

soil was frozen in liquid nitrogen and stored at -80°C.

DNA Extraction and PCR DNA was extracted using the UltraClean Fecal DNA
MoBio DNA Extraction Kit (MoBio Lab. Inc., Carlsbad, CA) as per the protocol, further
purified using the Promega Wizard DNA Clean-Up System (Promega, Madison, WI), and
quantified with a Perkin Elmer UV/V is Spectrometer Lambda 14. The 163 rDNAs were
amplified using the polymerase chain reaction (PCR) with the Acidobacteria specific
forward primer ACD31F (3) and general bacterial broadly inclusive reverse primer
1492R (43, 44). Each 25 ul PCR reaction contained 1x PCR Buffer, 1 mM magnesium
chloride, 0.03 mM of each dNTP, 0.2 uM each primer, and 5U taq polymerase
(Invitrogen, Carlsbad, CA). The PCR program consisted of the following steps: (1) 95°C
for 3 minutes, (2) 95°C for 30 seconds, 56°C for 30 seconds, 72°C for 45 seconds
(repeated for 30 cycles), (3) 72°C for 10 minutes, and (4) a hold at 4°C. Purified genomic
DNA from A. capsulatum (ATCC 51196) was used as a positive control. PCR reactions
were electrophoresed on a 1% agarose gel and stained with GelStar Nucleic Acid Stain
(BioWhittaker Molecular Applications, Rockland, MA). Gel images were captured using
a Kodak Electrophoresis Docrunentation and Analysis System (EDAS) 290 fitted with an
ethidiurn bromide (590 nm bandpass) filter. PCR products were cloned into the
Invitrogen TOPO TA Cloning Kit for Sequencing (Invitrogen, Carlsbad, CA) as per the

protocol. The PCR product was then re-amplified

36

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.couebaoocoo cowobm: new .cozebcooeoo .8930 65568 :8 Bop—om in zom .~.N 03¢...

37

using the modified-M13F and modified-M13R (40) primers with the PCR conditions as
described above. PCR products were purified with ExoSAP-IT (USB, Cleveland, OH)
upon a 1:8 dilution of the enzyme, incubated at 37°C for 30 minutes and 80°C for 15
minutes, and submitted for sequencing with the 531R primer (5' TAC CGC GGC TGC
TGG CAC 3’). Sequencing reactions were sent to the Michigan State University
Research Technology Support Facility (RTSF) (http://genomics.msu.edu/index.html).
Sequences were aligned using ARB Software (47). Other sequences were downloaded
from Genbank and the Ribosomal Database Project (RDP) (15) for the generation of

phylogenetic trees.

Data analysis These acidobacterial 16S rRNA gene clone libraries (50 clones per
library) were analyzed using the ARB Software (47). Known representatives of the eight
monophyletic subdivisions were used to determine the relative percentage of soil clones
in each subdivision. The relationship between the relative percentages of the subdivision

and various edaphic properties was assessed using regression analysis.

Cultivation Experiments

Soil Sampling for Cultivation Soil samples were collected at various times
between August 2001 and October 2002 from the KBS LTER, primarily treatment 8
along with treatments 1 and 7, and the KBS deciduous forest site. Soil cores (2.5 cm
diameter x 10 cm depth) were collected from replicate plots and stored at 4°C until
extraction typically no more than 48 hours after sampling. Five soil cores were

homogenized under microoxic (vol/vol: 2% oxygen, 5% carbon dioxide, 93% nitrogen)

38

conditions and approximately 30 grams of soil was added to 100 mL of a phosphate-
buffered salts solution (NaCl, 137 mM; KCI, 2.7 mM; NazHPO4, 10 mM; KHzPO4, 2
mM), buffered to a pH of 7.0, and amended with 2.24 mM sodium pyrophosphate
(Na4PzO7*10HzO) added as a dispersal agent and 1 mM dithiothreitol (Sigma-Aldrich, St.
Louis, MO) added as a reducing agent. The microoxic atmosphere was maintained
within a flexible vinyl chamber fitted with an oxygen sensor/controller (Coy Laboratory
Products, Grass Lake, MI). The soil suspension was stirred with a magnetic stir bar for
30 minutes, large soil particles were allowed to settle for 30 minutes, and an aliquot was
used for lO-fold decimal dilutions in the same buffer used for the initial soil suspension.
Aliquots (50 ul) of selected dilutions were then spread onto media of various
compositions (below) with at least four replicate plates per dilution.

Total direct counts to determine the number of cells/gram of soil (dry wt.) were
made by using 5-(4,6-dichlorotriazine-2-yl) aminofluorescein (DTAF) (4) with an
epifluorescence microscope (Axioskop, Carl Zeiss Inc., Oberkochen, Germany) at 1000X
magnification under blue illumination (BP 450-490 exciter filter, FT 510 nm beam
splitter, and LP 520 nm barrier filter) in a dark room. Soil moisture was determined as
described above and was used with total direct counts to estimate the number of

cells/gram (dry wt.) of soil.

Media Composition and Incubation Conditions The isolation medium consisted of
vitamins (excluding thiamine), inorganic salts, and trace elements as described previously
(66); hereafter, this medium will be referred to as VSB-# (VSB- “vitamins and salts

base”; # indicates the respective pH of the basal solution). VSB was buffered to pH 7.0

39

with 10 mM HEPES (VSB-7) and amended with one or more of the following additives,
prepared in distilled water unless otherwise noted: soil extract, 10 % (vol/vol); a mixture
of one-carbon compounds (methanol, 5 mM; methylamine, 3 mM; dimethylamine, 1
mM); washed agar; the humic acid analog anthraquinone-2,6-disulfonic acid disodium
salt (AQDS; final concentration 25mM) as a potential electron acceptor; a rrrixture of N-
acylhomoserine lactones (N -(butyryl, heptanoyl, hexanoyl, B-ketocaproyl, octanoyl, and
tetradecanoyl)-DL-homoserine lactones) (Sigma-Aldrich Co., St. Louis, MO) prepared in
ethyl-acetate acidified with 0.1% (vol/vol) acetic acid at a final concentration in the
media of 1 pM each as possible growth-promoting signals; catalase, added directly to
plates at a concentration of 65 U/ml or to cooled, molten agar just prior to pouring plates
at 130 U/ml to protect cells from peroxides; and a mixture of yeast extract, Bacto
protease peptone #3, casamino acids, and dextrose at 0.05 g/l each (Table 2.3). Plates
were incubated for 20 to 30 days at 12°C or 23°C under one of the following
atmospheres: COz-enriched hypoxia (vol/vol: 2% 02, 5% C02, and bal N2); COz-enriched
air (vol/vol: 95% air, 5% C02), or air. Plates were screened for the presence of
acidobacteria using the Plate Wash PCR (PWPCR) procedure described below. A chi-
square test was used to assess the significance of treatments that increased the frequency
with which acidobacteria colonies were detected on the plates. ANOVA was used to

assess the significance of treatments on the total number of colonies formed.

Plate Wash PCR Replicate agar plates were selected for screening with group-

specific I6S rRNA gene primers using the Plate Wash PCR (PWPCR) (66) (Figure 2.1).

Plates were flooded with 2 mL of Bead Solution from the UltraClean Dry Soil DNA kit

40

Table 2.3. Medium additives used for cultivation experiments.
__ . e -. . . 1... .4 .,_______ r 4 ”--.... -...-- m ...- -..... - . __,__ -. I!

'T'I'I." ...—...}..—

‘ Medium Additive Rationale g,

 

I
* gr'h .

N-acyl homoserine lactones (aHSL) Quorum sensing compounds

 

Anthraquinone disulfonate (AQDS) Humic acid analogue

 

 

 

 

 

 

 

 

 

 

 

Low temperature (12°C) F acultative psychrophiles
Catalase Protection fiom peroxides
. Mimic soil environment,
Low light . .
prevent perox1de formation
Elevated carbon dioxide concentrations Requirement for carbon dioxide for
(5% CO2/95% air) growth; CO2 fixation
Microoxic Oxygen Concentrations (2% . . . .

oxygen, 5% C 02’ bal. N2) Microaerophrlrc lifestyle

Extended incubation times Slow gr owmg nncroorgamsms
(low rrn copy number)

Low nutrients (organic carbon) Mum}: 8011 environment, mmnc
oligotrophic envrronment
Cl compounds Methylotrophy
. Provide nutrients, cofactors, additives
Sorl extract
necessary for growth
Remove excess nutrients, mimic
Washed agar . . .
oligotrophic envrronment

 

 

 

41

lnoculum

Medium &
incubation
conditions

.’ m ‘.
B Transfer
.- ._; isolated

colonies to

 

 

 

 

 

 
 
    

 

  

 

 

 

 

Id t' f bl '
en 1fy avora e Screen and identify

I I I
Extract DNA, - -
screen with culture conditions targeted organisms
group-specific wrth group-specrfic
P$R afimevrs PCR primers
A B C

 

 

Plate Wash PCR method used to detect the presence of the target

Figure 2.1.
organism(s) on the three treatments (A, B, & C). Treatment C indicates a positive

detection for the target organism (66).

42

for fecal samples (MoBio Laboratories, Carlsbad, CA) and a sterile spreader was used to
scrape off colony material. The colony slurry was added to a dry bead tube from the soil
DNA kit along with solutions S1 and IRS. To further aid in cell lysis, a 50 ul aliquot of
Iysozyme (50 mg/ml, Sigma-Aldrich, St. Louis, MO) was added to the resuspended
colony material, and the tube was incubated for 45 minutes in a 56°C water bath. After
this incubation step, cell lysis and DNA extraction were carried out per the
manufacturer’s protocol. The concentration and purity of each extracted DNA sample
was estimated spectrophotometrically by determining the absorbance at wavelengths

between 220 to 320 nm.

Isolation of novel acidobacteria from soil The plates were screened for the
presence of acidobacteria using PWPCR (Figure 2.1) (66). Once media and atmospheres
which supported the growth of acidobacteria were identified, all isolated colonies and
patches of colonies were transferred to plates containing fresh medium and incubated
under the same conditions. These isolated eclonies or patches of cell growth from
individual sectors were then grouped and screened for the presence of target organisms.

Isolation plating ensued for patches of cell growth that were positive for target organisms.
RESULTS and DISCUSSION
Acidobacteria composition The specific phylogenetic affiliations of partial

sequences (ca. 500 nucleotides) of 100 cloned l6S rRNA genes (2 replicate libraries,

consisting of 50 clones each) from 0 to 7 cm of treatments 1, 7, and 8 revealed the

43

presence of subdivision 1, 3, 4, 5, and 6 (Figure 2.2). Members of subdivisions 2, 7, and
8 were not detected because the 16S rRNA gene sequences of these subdivisions are not
amplificable with the specific forward primer, ACD 31F (3).

When the relative percentage of each subdivision was represented as a pie chart,
there was a clear indication that the subdivisions were responding to soil management
(Figure 2.2) revealing that particular treatments selected for different subdivisions. The
mid-successional treatment with no history of tillage (T8) contained members of
subdivision 1 and 6 with their average percent composition of ca. 27% and 38%,
respectively. The acidobacteria community from the mid-successional treatment with a
history of tillage (T7) revealed that subdivision I became less dominant, while
subdivision 4 and 6 became more dominant, ca. 35% and 43%, respectively. Finally, the
conventional agricultural treatment (T1) was dominated by subdivision 4 with ca. 44% of
the total acidobacterial community.

In summary, the mid-successional soil (T8) was dominated by members of
subdivision(s) l and 6 whereas the conventional agriculture soil (T1) was dominated by
members of subdivision 4 (Figure 2.3). The shift in the relative percentages of
subdivision 4 from the mid-successional (T8) to the conventional agriculture (T1) soil
were statistically significant (p < 0.03) with a Student’s t-test; however the shifts in the
composition of subdivision(s) l and 6 were not. These data taken together demonstrate
that the management of agriculture resulted in a dramatic shift in the acidobacteria

composition, with members of subdivision 4 becoming more dominant.

.44

C .
1 ,g\ 46% E) o /10%
\

32%
16%

   

B) 0%
sat/L Law» 47\

X 28%

     

_ subdivision 1
- subdivision 3
subdivision 4
I: subdivision 5
i: subdivision 6

Figure 2. 2. Distribution of Acidobacteria subdivisions among the different agricultural
regimes: (A) T1 RI 0- 7 cm; (B) Tl R2 0- 7cm, (C) T7 R1 0- 7cm; (D) T7 R4 0- 7cm; (E)
T8 R1 0- 7cm; and (F) T8 R2 0- 7cm.

45

Figure 2.3. Neighbor joining phylogenetic tree of the phylum Acidobacteria with 16S
rRNA gene derived from DNA extracted from KBS LTER soil. Geothrixfermentans and
Holophagafoetida of subdivision 8 were used as an outgroup (not shown). All boxes in
black and gray represent clones (partial sequence ca. 500 bp) from upper layers of T8 and
T1, respectively. The subdivision is indicated to the right of the phylogenetic tree. Bar
graphs are illustrated with the number of clones from each subdivision (SD#) with
respect to the two treatments. Scale bar indicates 0.10 changes per nucleotide.

46

 
 

 

1
T
w. W m. w.
W. N... w.
.m n n. to.
8
m T
<0. w m m w 0 <0. m m m m 0 w w m 0 w m m w m w 0
am 8:20.? a am we muse—owe n momma 8:20 we a momma Eco—owe & ammo muse—owe &
6 4. 5 3 l

 

  

:____,____.____._______:_ ______._,___

  

A, , L
: L r , L , —
i. L 7 U
L , . , _ i ,

 

 

0.10

47

Influence of soil properties The relative percentage of clones of each
subdivision varied with depth and treatment, suggesting edaphic properties may influence
the distribution of the various subdivisions. Members of subdivision 3 correlated
positively to soil moisture (p < 0.09, a = 0.10) (Figure 2.4) suggesting that wetter soils
are preferred; 32% of the increase in relative percentage of subdivision 3 can be
explained by soil moisture at the time of sampling. This suggests that members of
subdivision 3 might be capable of growing under low or no oxygen conditions, given
their prevalence in wetter soils that are typically oxygen depleted. Additionally, in situ
higher soil moisture can influence the activity of the microbial community by enhancing
the diffusion of nutrients (13) thus influencing the community structure (6, 7, 11) and
activity (20, 65) of soil microbes.

The relative abundance of subdivision 1 clones in each treatment appeared to be
related to soil pH, with the greatest abundance occurring in mildly acidic soils (Figure
2.5). This is consistent with recent publications regarding the physiology of members of
subdivision 1. Members of subdivision 1 formed more colonies in media with pH
between 5.5 and 7.0 (56) and recent characterization of subdivision 1 strains in the genus
Terriglobus (25), indicated that the optimal pH range for this subdivision is between 5.0
and 6.0 (Chapter 3) (25). Taken together, these results suggest that members of
subdivision 1 of the Acidobacteria have a preference for mildly acidic pH conditions.

Previous studies have implicated members of subdivision 1 with the capacity to
oxidize one-carbon compounds such as methanol (51). There was a negative correlation
between the net influx of methane into the soil and the relative percentage of subdivision

1 (p < 0.07, a = 0.10) (Figure 2.6). A larger proportion of the acidobacterial community

48

OJ
C
1

     

M

t:

.9.

.59. 25‘

.2

'U

..D

a

g, 20-

O

G.)

DD

(6

’a’ 15-

8

s.

‘3 10- . . y=0.80x-0.27
g J . R2=0.32
g 5_ . . p<0o9

 

 

IfiIJI'I"I'I'I'I

10 ' 12 14 16 18 20 22 24 26 '
% soil moisture ((g HzO/g dry soil)* 100)

Figure 2.4. Relationship between relative percentage of acidobacteria subdivision 3 and
% soil moisture.

49

50"

y2= -22.7x + 165.1
R = 0.49
p < 0.02

  
   

20‘

10‘

Relative percentage of subdivision 1

 

 

0 1 I I I I I I
5.8 6.0 6.2 6.4 6.6 6.8 7.0 7.2

Soil pH

Figure 2.5. Relationship between relative percentage of acidobacteria subdivision 1 and
soil pH.

I

50

 

Relative percentage of subdivision 1

 

 

0 ' I ' r I ' I . '
-12 -10 -8 -6 -4 -2 0
CH4-C (g ha'l d")

if

Figure 2.6. Relationship between the relative percentage of subdivision 1 and methane
fluxes. Methane data was averaged for each treatment from historical data obtained from
the KBS LTER website for methane flux data collected between 1992 and 2001; a
detailed description of the sampling can be found in a previous study by Robertson and
collegues (55).

51

contained members of subdivision 1 when there was approximately a 6-fold increase in
the total methane flux into the soil. The affinities for known methane-oxidizing
microorganisms and environmental concentrations of methane do not correlate
suggesting that there could be an unexplored population of methanotrophs which is
responsible for this discrepancy (17); perhaps members of subdivision 1 are a bacterial
group responsible for this discrepancy or they could respond favorably to nearby
methanotrophic communities.

A statistically significant negative correlation was observed between the relative
percentage of acidobacteria subdivision 4 and the carbon concentration of each site (p <
0.0008) (Figure 2.7). Ninety-one percent of the increase in the relative abundance of
subdivision 4 can be explained by the lower carbon concentrations suggesting that
members of subdivision 4 might be oligotrophic, preferring lower carbon conditions for
growth. Additionally, these data suggest that members of subdivision 4 are better
competitors for carbon since they were found more prevalently in low carbon
environments which coincided with environments with the higher nitrous oxide fluxes.
Their percentages correlated positively to these fluxes (p < 0.01) (Figure 2.8) suggesting
preliminarily that they either responded favorably to the nearby denitrifying community
or are capable of denitrification since typically denitrifiers are good competitors for

carbon, which is a major factor responsible for their distribution (71).
Cultivation experiments The average recovery of soil microorganisms was 3.5 to 5 x

107 CPU/gram of soil (dry wt), which based on total direct counts, is 2.5 to 6% of the

total microbial community (Figure 2.9, panel A). These recoveries were similar to

52

50- I

40-

      

y = -36.4x + 69.8
R2 = 0.91
p < 0.0008

30-

Relative percentage of subdivision 4

 

 

1

0.6 0.8 1.0 1.2 1.4 1.6 1.8 2.0
% carbon concentration» ((g C/g dry soil)*100)

Figure 2.7. Relationship between relative percentage of acidobacteria subdivision 4 and
carbon concentration. Carbon data was averaged for each treatment from historical data
obtained from the KBS LTER website for carbon data collected between 1989 and 2001;
a detailed description of the sampling can be found at the KBS LTER website
(http://lter.kbs.msu.edu/). ‘

53

507

40‘

30'

 

20 ‘
y = 43.4x - 42.9

R2 = 0.72
p<001

   

10‘

Relative percentage of subdivision 4

 

 

O r ' r ' I ' I ' T ' I
1.0 1.2 1.4 ' 1.6 1.8 2.0

NZO-N (g ha-l d-l)

Figure 2.8. Relationship between the relative percentage of subdivision 4 and nitrous
oxide fluxes. Nitrous oxide data was averaged for each treatment from historical data
obtained from the KBS LTER website for nitrous oxide flux data collected between 1992
and 2001; a detailed description of the sampling can be found in a previous study by
Robertson and collegues (55).

54

Figure 2.9. Summary of cultivation experiments for total recovery of colonies per gram
of soil (dry weight) (Panel A) and acidobacteria detection (% of 11) (Panel B). Panel A
represents the average CFU recovery from soil +/- the standard error; the number of times
a particular treatment was employed (n) is listed at the base of each bar. Panel B
represents the percentage of treatments that yielded a positive test for acidobacteria using
PWPCR from the plates of panel A. The asterisks “*”indicates treatments that yielded
significant difference (panel A, AN OVA or = 0.05; panel B, chi square goodness of fit, or
= 0.10).

55

CPU (107) /gram of soil (dry wt)

Acidobacteria detection (% of n)

50

C02

+ -
aHSL

+ -

catalase

+ -

catalase

*>I<

 

2% 21% + - 23C 12C + -
02 nutrients temperature AQDS

Treatments

2% 21%
02
Treatments

56

 

+ - 23C 12C + -
nutrients temperature AQDS

cultivation experiments with low nutrients and extended incubation times (16, 37). All
treatments were identical in terms of recovery with the exception of the inclusion of
AQDS or were incubated at 12°C. This suggests that the extended incubation time was
responsible for the increased recovery. Although the use of our novel cultivation strategy
of incubation under atmospheric gas concentrations native to soil did not significantly
increase total recovery, it was instrumental in isolating members of the phylum
Acidobacteria, specifically the elevated levels of carbon dioxide (below).

While increased recovery resulting from 'low nutrient concentrations and extended
incubation times were more of a confirmatory nature, it was interesting to note that soil
moisture had a significant effect on increasing total recovery, suggested only once
previously in the mid-19305 (69). Soil moisture had a positive effect on the total
recovery of aerobic, heterotrophic soil microorganisms after 20 to 25 days of growth
from treatment 8 soil (Figure 2.10, panel A), similar to a previous study with garden soil
(Figure 2.10, panel C, gray boxes). As soil moisture increased from 7% to 21%, the
number of colony forming units significantly increased ca.10-fold (p < 0.009), but
recovery reached an asymptote above a soil moisture of 21% (Figure 2.10, panel B). The
asymptotic nature of the curve above 21% soil moisture suggests that there is a limit of
soil moisture’s effectiveness on total recovery.

Previously soil moisture has been shown to influence the community structure (6,
7, 11) and activity (20, 65) of soil microbes. In situ, higher soil moisture can influence
the activity of the microbial community by enhancing the diffusion of nutrients (13) and

decreasing oxygen tension (53, 62). Conversely, the low recoveries resulting from low

57

Figure 2.10. Relationship between total recovery of aerobic, heterotrophic soil bacteria
fiom T8 soil and % soil moisture. Panel A depicts a significant increase in total recovery
up to 21% soil moisture, whereas panel B illustrates the limit of soil moisture influence
on total recovery. Panel C represents the effect of recovery on aerobic heterotrophs from
KBS LTER treatment 8 soil (black boxes) and a garden plot (gray boxes) (69).

58

.>

CFU (107) /gram of soil (dry wt.)

99

CFU (107) /gram of soil (dry wt.)

.0

CFU (107) /gram ofsoil (dry wt.)

 

 

 

 

 

1003
10.:
1
1's
j y=0.08x-0.7l
j R2=0.98
- p<0.009
0.1..r,....fi.vt
5 10 15 20 25 30 35
% soil moisture
1003
103
*1: 'I'
'1.
0.1 , . , . . , . 1 . r .
5 10 15 20 25 30 35
% soil moisture

100.:

JL All.

 

 

y=0.04x-0.18
R2=0.62
p<0.0005
0.1 . , 4 , . , . .

 

(It-
—I
O
—l
Lit

20 25
% soil moisture

59

I V l

30 35

moisture soil inoculum suggests that osmotic stress may have been detrimental to these
cells. Drier soils typically have a low water potential, meaning it is harder for an
organism to extract water from the environment, therefore water can become limiting. In
order for organisms to maintain their cell integrity, they either produce compatible
organic solutes or take up ions from surrounding environment (19) to create a high
intercellular solute concentration to promote an inward diffusion of water. When taken
fiom this low water potential environment and introduced to a high water potential
environment, such as the creation of a soil suspension solution for plating, these cells
could be more susceptible to cell lysis resulting from the rapid influx of water to their
high intercellular solute concentration. This scenario could drastically reduce their
culturability as compared to cells originating from a high water potential environment.
Osmotic stress has been shown previously to be responsible for decreased culturability of
pure cultures, especially in gram-negative bacteria (30). Our results suggest that soil
moistures above 21% produce a sufficiently low water potential environment conducive
with the creation of our soil suspension.

Although the treatments tested did not reveal significant increases in total
recovery of aerobic heterotrophs from soil, the inclusion of N-acyl homoserine lactones
(Figure 2.9, panel A, aHSL) and incubation under microoxic conditions (Figure 2.9,
panel A, 2% 02) did increase the total recovery. The inclusion of N-acyl homoserine
lactones increases the recoverability of soil microorganisms ca. 1.2-fold, as seen
previously (9, 12), perhaps due to gene activation for such activities as siderophore
production (28), biofilm formation (22), starvation survival (70), or utilization (35, 73).

The concentrations of oxygen were manipulated during incubation to better simulate

60

conditions in soil aggregates, and it increased the total recovery ca. 1.4-fold. Although
upland soils like those at the KBS LTER are generally well-aerated, they include pockets
of anoxia within soil aggregates, and hence also include transitions zones between oxic

and anoxic conditions (34, 61).

Detection and isolation of acidobacteria with PWPCR The PWPCR method, in
addition to isolating novel members of the acidobacteria (Figure 2.11), revealed insight
into this phylum through the detection of acidobacteria over the multitude of treatments
(Figure 2.9, panel B). Across treatments used for isolation, acidobacteria were detected
more frequently under atmosphere(s) containing elevated levels of carbon dioxide
(0.05<p<0.10), suggesting that they might be capable of using elevated levels of carbon
dioxide perhaps through autotrophic pathways.

Additionally, acidobacteria were more. frequently detected under low carbon
concentrations and along with extended incubation times could have provided the ideal
conditions to isolate potential oligotrophic acidobacteria (25). The use of low carbon
concentrations has been shown previously to improve the recovery of bacteria from soils
(31, 32, 67, 68), marine environments (16), heterotrophic planktonic bacteria from lake
water (12), and more diverse communities of pseudomonas from soil (1). The inclusion
of the catalase enzyme as well as incubation under microoxic conditions increased the
frequency of acidobacteria detection by ca. 3.3- and 1.3-fold, respectively. Taken
together, this result suggests that some acidobacteria are microaerophiles, preferring low
levels of oxygen. Furthermore, the catalase enzyme decreased overall recoveries of soil

microorganisms (Figure 2.9, panel A), but increased acidobacteria detection

61

Acbt.capsl (D26171)
F env.WD277 (AJ292577)

 

 

 

 

Ellin5225 (AY234576)

Ellin5017 (AY234434)

KBS89 (AY587227)

env.TM2 (X97098)

Ellin351 (AF498733)

TAA166 (AY587230)

Ellin337 (AF498719)

r— TAA43 (AY587228)

TAA48 (AY587229)
KBSllZ (DQ660895)
KBS63 (DQ660892)
KBS62 (DQ660893)
KBS68 (DQ660894) _

    
  
 
  

 

 

 

 

 

 

_L/ 1

 

 

 

 

 

7

 

/

 

 

 

 

 

1_____/

 

 

Figure 2.11. Maximum likelihood tree of the Acidobacteria subdivisions 1, 3, 4, 5, and 6
(indicated to the right of the group) based on the 16S rRNA gene using sequences
obtained from cultivated representatives and environmental clones. Geothrix fermentans
and Holophaga foetida of subdivision 8 were used as an outgroup (not shown). Strains
from this study are in bold text. Each gray trapezoid represents a different subdivision.

The scale bar indicates 0.10 changes per nucleotide.

62

(Figure 2.9, panel B), thereby potentially selecting for microaerophilic acidobacteria.
Previous studies suggest that catalase-negative microorganisms are abundant in the soil
environment (14, 27), and the inclusion of the catalase enzyme would provide protection
for cells from dangerous peroxides resulting from the oxidation of the low concentrations
of added carbon to the plates. Micro-zones of elevated carbon dioxide and/or lower
levels of oxygen located in the soil matrix provide a great range of habitats and niches

which are ideal for soil microorganisms.

SUMMARY

Soil is a heterogeneous environment Containing over one million species of
bacteria per gram (26), however, these microorganisms are poorly understood, since they
are not well represented in culture, such as members of the phylum Acidobacteria.
Before the start of these experiments, there was little to information available on this
diverse phylum, except for the information on the historical strains, A. capsulatum, H.
foetida, G. fermentas and generic soil surveys illustrating the phylum’s ubiquity. The
acidobacteria 16S rRN A gene survey in this study generated useful information
pertaining to the composition of the acidobacteria community among the KBS LTER
treatments along with an assessment of the impact certain edaphic properties have on the
eight, monophyletic subdivisions.

Figure 2.12 represents a summary for the changes in the acidobacteria community
composition in relation to edaphic properties observed at the KBS LTER. Subdivisions
1, 3, 4, 5, and 6 were present in the soils at the KBS LTER, with members of subdivision

4 being the dominant subdivision in the conventional agriculture treatment and

63

ACIDOBACTERIA COMMUNITY COMPOSITION SUMMARY

CH 4 CH4
CH 4

 

CH4

       

CH4

C?
. ca. 12% SD4 ' 35% “44% sod
. ‘g ‘ I_ I g ‘ . ‘ . / g . - .3; f.-‘ _. V131,»): jKlf-‘w

,1" i v

ca. 15% SD]
I ' fr

. ca27%SD1 . 7% $151 ‘ Q ‘ ,
O‘Zw‘v—v a .' 1’ i "1. 1 - 1 w ’. " 1 /x
1_ ca.38%sn6‘ ' ‘ 'f ' ’ 43% ' j ’ * 23%SD6“
D'eatment 7
no tillage history of tillage conventional agriculture
herbaceous plants herbaceous plants com/soybean/wheat

Figure 2.12. Summary of acidobacteria community composition and the edaphic
properties that influence the composition of subdivisions (SD) 1, 4, and 6.

subdivisions 1 and 6 being dominant in the mid-sucessional community with no history
of tillage. Additionally, the subdivisions responded to soil moisture (subdivision 3),
carbon concentration (subdivision 4), soil pH (subdivision 1), methane fluxes
(subdivision 1), and nitrous oxide fluxes (subdivision 4). These data, along with general
knowledge about the soil environment, were instrumental in the development of
cultivation methods used to isolate these strains as well as additional strains (Chapter 4).
Moreover, these trends may help to bear insight into the physiology of members of this
phylum. When members of this phylum have been successfully isolated, we can
determine if the trends observed in the soil surveys are consistent with their physiology.

Media composition and incubation conditions were designed to mimic the soil
environment, specifically through incubations under native soil atmosphere gas
concentration under low light, low nutrients, and extended incubation periods to select for
potential, slow-growing microorganisms. Extended incubation time, low concentrations
of nutrients, and soil inoculum with high moisture (significantly) improved total recovery
of aerobic heterotrophs from soil. A facile, high- throughput method called Plate Wash
PCR (PWPCR) was developed to rapidly screen a multitude of plates for the target
organism(s) using group-specific primers. These methods lead to the successful isolation
of Acidobacteria as well as providing trends linking acidobacteria detection to our
various amendments and treatments which helped to bear insight into their physiology.

In conclusion, the microbial community in the soil is an area we know very little
about. The need to retrieve this not yet cultured majority (8, 52) is becoming more and
more evident with the footprint of the soil community becoming more defined with

molecular surveys. The retrieval of these recalcitrant microbes requires patience and an

65

understanding of environmental factors that affects their composition. These studies
suggest that a combination of molecular and growth-based approaches such as those

realized in this study are the strategies needed to isolate these “uncultivables.”

ACKNOWLEDGEMENTS
This work was funded by the USDA (#2001-35107-09939), NSF (#MCB-
0135880), and the NSF Long-Term Ecological Research Program at the Kellogg
Biological Station and the Michigan Agricultural Experiment Station.
I would like to thank Leslie A. Dethlesfson for his input on the statistical analysis

and Kristin M. Huizinga for her help with soil sampling.

66

10.

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74

CHAPTER 3

ISOLATION AND CHARACTERIZATION OF SOIL BACTERIA THAT

DEFINE T ERRIGLOB US GEN. NOV., IN THE PHYLUMACIDOBACTERLA

These results have been published in the article: Eichorst, S.A., J .A. Breznak, and TM
Schmidt. 2007. Isolation and characterization of soil bacteria that define, T erriglobus

gen. nov., in the phylum Acidobacteria. Appl. Environ. Microbiol. Vol. 73, No. 8,

pgs. 2708-2717.

INTRODUCTION
Soils typically contain 109-1010 microorganisms per gram (dry weight) that may
represent more than a million bacterial species (21). However, characterization of the
small fraction of microbes that has been cultivated provides only a glimpse of their
physiological capacity and potential impact on soil ecosystems. The absence of pure
cultures or genome sequences makes it difficult to ascertain the roles of specific microbes
in soil environments: this is particularly true for bacteria in the phylum Acidobacteria,

which are broadly distributed in soils but poorly represented in culture.

The phylum Acidobacteria is defined by a large collection of 16S rRNA gene
sequences (>1,500 in the Ribosome Database Project) (10) retrieved fi'om diverse
environments including: soils and sediments (3, 17), soil crusts of sand dunes (69),
wastewater (13, 41), water distribution systems (49), peat bogs (15), acid mine drainage

(33), hot springs (26), shallow submarine hydrothermal vents (67), and on the surface of

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Paleolithic cave paintings and catacombs (62-64, 75, 76). In situ hybridization with
acidobacteria-specific probes has also confirmed the presence of intact acidobacteria in
many environments, and revealed multiple cellular morphotypes, including cocci, short

rods, and thin filaments (46).

Acidobacterium capsulatum was the first described member of the phylum
Acidobacteria. It was among eight strains isolated from acidic mine drainage and acidic
muds in the early 1990’s (33, 34). This was the first report of the isolation of acidophilic,
heterotrophic bacteria from an acidic mineral environment other than Acidiphilium (33).
By the mid-1990’s, the growing collection of ribosomal RNA gene sequences from
molecular surveys of diverse environments resulted in the recognition of A. capsulatum
as a member of a large, deeply branching, monophyletic lineage within the Bacteria (25).
The phylum Acidobacteria was named after the only described species at the time (46).

Shortly after the characterization of A. capsulatum, Holophaga foetida was
isolated and described as a novel, homoacetogenic bacterium capable of degrading
methoxylated aromatic compounds (43). Based in part on the 81.6% divergence in the
16S rRNA gene sequences of these two cultivars, the phylum was sometimes referred to
as Holophaga/Acidobacterium. Geothrix fermentans, a strain capable of Fe(III)
reduction, was later placed in the Holophaga/Acidobacterium phylum based on the
similarity of its 16S rRNA gene sequence to that of H. foetida (ca. 94%) (9).

The phylum Acidobacteria is now officially recognized in the Bergey’s Manual of
Systematic Bacteriology (22) and includes three genera with cultured representatives:
Acidobacterium (33), Geothrix (9), and Holophaga (43). The genus Solibacter was

recently proposed as the fourth genus in this phylum (wwwj gi.doe. gov). There are

 

76

currently eight recognized monophyletic subdivisions within this phylum (28) that
encompasses the molecular diversity first recognized as Acidobacteria (38), and
additional unnamed and mostly uncharacterized cultivars in subdivisions 1, 2, 3, and 4
(14, 29-31, 60, 61, 70). A recent survey of 23S rRNA genes in microbial communities
associated with Paleolithic paintings uncovered additional novel acidobacteria, expanding

the number of subdivisions to as many as eleven (76).

The breadth of divergence of 16S rRNA gene sequences in the phylum
Acidobacteria (ca. 77% based on >1000 nearly full length sequences, this study) is
similar to that within the metabolically diverse phylum Proteobacteria (28), suggesting
the capacity for extensive metabolic diversity. Acidobacteria are oftentimes the most
abundant bacteria represented in molecular surveys of soil environments: as many as half
of all clones from Arizona soil samples clustered in the phylum Acidobacteria (16), as
did more than 40% of PCR-amplified and cloned rRNA-encoding genes in soils of alpine
ecosystems (44). In a comprehensive review of acidobacterial abundance in soil
communities (29), acidobacteria averaged ca. 20% of the total bacterial community.
Although the metabolic potential of acidobacteria is poorly described, their abundance
suggests a major impact on nutrient cycling in soil environments.

To learn more about the metabolic properties and potential ecological roles of
members of this poorly explored phylum, we sought to cultivate and characterize new
strains from terrestrial habitats. Using incubation conditions and media designed to
mimic their natural environment, eight strains of the phylum Acidobacteria were isolated
from soil as well as from the hindguts of soil-dwelling termites. These strains were

characterized, with emphasis on properties that may bear on their ecological roles in soil.

77

Results from the physiological and phylogenetic characterization warrant creation of a

new genus, Terriglobus.

MATERIALS and METHODS
Phylogenetic survey and isolation of strains Soil samples were collected between
August 2001 and August 2005 from the Michigan State University WK. Kellogg
Biological Station Long Term Ecological Research (KBS LTER) site. The KBS LTER is
a 48-hectare research site established in 1989 to study ecological processes in
agroecosystems. The dominant soil types are of the Kalamazoo (fine-loamy, mixed,
mesic Typic Hapludalfs) and Oshtemo (coarse-loamy, mixed, mesic Typic Hapludalfs)

series. A detailed description of this site can be accessed at http://wwwkbsmsuedu.

 

For the survey of acidobacterial diversity, soil cores (2.5cm diameter x 20 cm
depth, divided into 0-7cm and 13-20cm) were collected fiom four different treatments at
the KBS LTER: treatment 1 (T1) is managed as a conventional agriculture site with a
rotation of corn/soybean/wheat; treatment 7 (T7) is a successional plant community on
historically tilled soil —- tillage was abandoned after spring plowing in 1989; treatment 8
(T8) is a successional plant community on never-tilled soil; and the fourth treatment is
native deciduous forest. Five soil cores per plot were pooled and homogenized using a 2-
mm sieve, portions of the sieved soil were used to determine soil pH and soil moisture
(58), and the remaining soil was frozen immediately in liquid nitrogen, and stored at -80 °
C until use. DNA was extracted from soil samples using the UltraClean Fecal DNA
MoBio DNA Extraction Kit (MoBio, Carlsbad, CA). The 16S rRNA genes were

amplified from the DNA by using the PCR with an acidobacterial-specific forward

78

primer (ACD31F: 5’ GAT CCT GGC TCA GAA TC 3’) (3) and a broadly inclusive
bacterial reverse primer (1492R: 5’ GGT TAC CTT GTT ACG ACT T 3’) (39, 40).
Each 25 pl PCR reaction contained 1x PCR buffer, lmM MgCl2, 0.03mM of each dNTP,
0.2uM each primer, and 5U T aq DNA polymerase (Invitrogen, Carlsbad, CA). Thermal
cycling consisted of the following steps: (1) 95°C for 3 minutes; (2) 95°C for 30 seconds,
56°C for 30 seconds, 72°C for 45 seconds (repeated 30X); (3) 72°C for 10 minutes.
Genomic DNA purified from A. capsulatum (ATCC Number 51196) was used as a
positive control. PCR products were electrophoresed through a 1% agarose gel in 0.5x
tris-borate-EDTA (TBE) and visualized with GelStar Nucleic Acid Stain (BioWhittaker
Molecular Applications, Rockland, MA).

PCR products were cloned with the Invitrogen TOPO TA Cloning Kit for
Sequencing (Invitrogen Carlsbad, California) and the inserts were re-amplified using the
modified-M13F (F2) (5’ CAG TCA CGA CGT TGT AAA ACG ACG GC 3’) and
modifed-M13R (F4) (5’ CAG GAA ACA GCT ATG ACC ATG 3’) (32) with the PCR
conditions described above. The PCR products for sequencing were treated with
ExoSAP-IT (U SB, Cleveland, OH) using a modification of the manufacturer’s protocol

(ExoSAP enzyme was diluted 1:8 and the reaction was incubated at 37°C for 30 minutes,
followed by an 80°C incubation for 15 minutes) and submitted for sequencing with the
531R (5' TAC CGC GGC TGC TGG CAC 3') primer. Sequencing was performed at the
Michigan State University Research Technology Support Facility

(genomicsmsuedu/index.html). Sequences were aligned with other sequences

 

downloaded from GenBank using an integrated aligner (ARB Software) (47) as well as

manual corrections based on secondary structure models of the 16S rRNA gene.

79

For isolation attempts, five cores (2.5 cm diameter x 10 cm depth) from the never
tilled, successional treatment (T8) were collected and homogenized with a sterile spatula
in a 500mL beaker under a hypoxic atmosphere (vol/vol: 2% O2, 5% CO2, 93% N2)
within a flexible vinyl chamber fitted with an oxygen sensor/controller (Coy Laboratory
Products, Grass Lake, MI). Portions of the homogenized soil were used to determine soil
moisture (58). Approximately 30 grams of soil were added to 100 mL of a phosphate-
buffered salts solution (137 mM NaCl, 2.7 mM KCl, 10mM Na2HPO4 and 2 mM
KH2PO4); pH was adjusted to 7.0, and supplemented with 2.24 mM Na4P2O7-10H2O as a
dispersal agent, and dithiothreitol (lmM) as a reducing agent (73). The soil suspension
was stirred vigorously with a magnetic stir bar for 30 minutes. Denser soil aggregates
were then allowed to settle for 30 minutes, after which an aliquot of the supernatant
fraction was used for total direct cell counts after staining with 5-(4,6-dichlorotriazine-2-
yl) aminofluorescein (DTAF) (4). Another aliquot was used to prepare 10-fold decimal
dilutions in the same buffer for subsequent inoculation of the isolation medium consisting
of vitamins (excluding thiamine), inorganic salts, and trace elements as described
previously (70); hereafter, this composition will be referred to as VSB-# (VSB- “vitamins
and salts base”; # indicates the respective pH of the basal solution). VSB was buffered to
a pH 7.0 with 10mM HEPES (VSB-7) and amended with one or more of the following
additives, prepared in distilled water unless otherwise noted: the humic acid analog
anthraquinone-2,6-disulfonic acid disodium salt (AQDS; final concentration 25mM) as a
potential electron acceptor; a mixture of N-acylhomoserine lactones prepared in ethyl-
acetate acidified with 0.1% (vol/vol) acetic acid at a final concentration in the media of 1

uM each N-(butyryl, heptanoyl, hexanoyl, B-ketocaproyl, octanoyl, and tetradecanoyl)-

80

DL-homoserine lactones (Sigma-Aldrich Co.) as possible growth-promoting signals;
catalase, added directly to plates at a concentration of 65 U/ml or to cooled, molten agar
just prior to pouring plates at 130 U/ml to protect cells from peroxides; and a mixture of
yeast extract, Bacto protease peptone #3, casamino acids, and dextrose at 0.05 g/l each.
Cultivation conditions for samples from the hindguts of soil-dwelling termites,
Reticulitermesflavipes, have been described previously (70).

Plates were incubated for 20-30 days at 12°C or 23°C under one of the following
atmospheres: CO2-enriched hypoxia (vol/vol: 2% O2, 5% CO2, and bal N2); CO2-enriched
air (vol/vol: 95% air, 5% CO2), or air. Plates were screened for the presence of
acidobacteria using Plate Wash PCR (PWPCR) (70). A chi-square test was used to
assess the significance of treatments that increased the frequency with which
acidobacteria colonies were detected on the plates. ANOVA was used to assess the

significance of any treatments on the total number of colonies formed.

Optimization of Growth Rate VSB was amended with a mixture of organic
compounds (yeast extract, protease peptone, casamino acids, and glucose - 0.15 g/l each)
to assess capacity for growth under a range of conditions. This complex medium
(designated VSM-#, where # = pH) was used to determine the effect of pH on growth
rate. Strains were grown at room temperature (23°C) between pH 2.0 and 7.5 at intervals
of 0.5 pH units: MES was used to buffer at pH ranges between 5.0-6.0, 4-
morpholinepropanesulfonic acid (MOPS) was used at pH ranges between 6.5-7.5, and
citric acid was used as a buffer below pH 5.0, all at a final concentration of 10mM.

Culture tubes (BellcoTM cat. no. 2048-18150) containing 5-10 mL of medium, were

81

stoppered under an air atmosphere and held on a reciprocating shaker operating at ca. 190
strokes/min. to disrupt cellular flocs. Optical density at 600nm was monitored
periodically with a ThermoTM Spectronic model 20D+ spectrophotometer. Inocula
consisted of a 1:100 dilution of cultures in mid/late log phase growth (ca. 1x108 cells/mL)
in R2B - a complex medium containing yeast extract, amino acids, peptone,
carbohydrates, pyruvate, and inorganic salts (56). After identifying the optimal pH for
growth, the range of growth-permissive temperatures was determined by assessing
growth rate at 4°C, 12°C, 23°C, or 37°C in VSM-6 for all strains except KBS 89 which
was grown in VSM-S.

To determine if growth rate was influenced by the elevated concentration of
carbon dioxide used during isolation, strains were grown on VSM-6 in triplicate culture
tubes and were incubated with either 0.05% CO2 (vol/vol, balance air) or 5% CO2
(vol/vol, balance air). Sodium bicarbonate (7 mM) was used as an additional buffer
under elevated levels of carbon dioxide. To control for the possible influence of increased
concentration of sodium when sodium bicarbonate was added, growth was also
monitored in a medium containing 7 mM sodium chloride without a CO2-enriched
atmosphere. Analysis of variance (ANOVA) was used to assess if culture conditions
resulted in significant differences in growth rate (SAS System version 8e software; SAS

Institute Inc., Cary, NC).
Colony and cell morphology Bacterial colonies on plates of R2A medium were

examined under a Nikon SMZ-2T dissecting microscope at 10-15X for size,

pigmentation, form, elevation, and margin (68). Gram stain reactions were performed as

82

described previously (65). Gross cell morphology and motility were assessed by using
phase contrast microscopy with a Zeiss Axioskop Microscope (Carl Zeiss Inc.,
Thomwood, NY). Cell shape, length, and width were measured with the CMEIAS image
analysis software (45). Transmission and scanning and electron microscopy (using the
polylysine procedure), including whole cell negative staining (using 2% uranyl acetate in
water), were completed at the Michigan State University Center for Advanced

Microscopy (wwwceomsuedu).

 

Carbon source utilization The ability of strains to use various organic compounds for
aerobic growth was tested in replicate 25mL Erlenmeyer flasks containing 5 mL of VSB-
6 for all strains except KBS 89 (VSB-5) amended with the following carbon sources at a
final concentration of 10mM (unless indicated otherwise): D-glucose, D-fructose, D-
galactose, D-mannose, D-ribose, D-xylose, L-arabinose, D-mannitol, D-sorbitol, sucrose,
D-maltose, D-raffinose, D-cellobiose, methyl cellulose (0.1% v/v, final conc.),
carboxymethyl cellulose (0.1% v/v, final conc.), sodium acetate, sodium pyruvate,
sodium formate, sodium succinate, a mixture of organic acids (sodium citrate, sodium
pyruvate, sodium fumarate, sodium D-L-lactate, and malic acid; 0.02% each, final conc.),
a mixture of purines and pyrimidines (adenine, guanine, thymine, cytosine, and uracil;
10ug/mL each, final conc.), D-glucuronate, D-galacturonate, D-gluconic acid,
trimethoxybenzoate, syringate, ferulate, vanillate, sodium benzoate, resorcinol, and tannic
acid (0.1% v/v, final conc.). Flasks were incubated at room temperature (ca. 23°C) and
shaken on an orbital shaker (ca. 190 rotations/min). A positive result was defined as

visible turbidity after incubation for ca. 14 days.

83

Strains were also tested for their ability to grow anaerobically by fermentation of
glucose, lysine, or omithine (BBLTM Enterotube II, Sparks, MD), or by anaerobic
respiration with nitrate (10mM), iron citrate (10mM), or AQDS (8mM) as the electron
acceptor with glucose (10mM) as the sole carbon and energy source in VSB-6 for all
strains except KBS 89 (VSB-5) with the addition of 7 mM bicarbonate. The headspace
gas for these tests was 85% nitrogen, 10% hydrogen, and 5% carbon dioxide, except for

samples with nitrate, which had a headspace of helium.

Carotenoid characterization Replicate 50 mL cultures of the pigmented strains in
250 mL side-arm flasks were grown in R2B to late-log phase under elevated carbon
dioxide (5% carbon dioxide) and either 20% oxygen or 2% oxygen in the dark. Cultures
were normalized to the same optical density at 600 nm and then cells were harvested by
centrifugation at 5,000 x g for 25 minutes. The. cell pellet was resuspended in a mixture
of acetonezmethanol (7:2 vol/vol), incubated overnight at 4°C in the dark (18),
centrifuged to remove cell debris, and the absorption spectra of the extracts were
determined between 220 and 800 nm with a Perkin Elmer model Lambda 14 scanning
UV/V is Spectrometer. Carotenoids were redissolved in chloroform for comparison of

their spectra with previously characterized carotenoids (7).

16S rRNA gene phylogeny Nearly full length sequence of the 16S rRNA gene (ca.
1500 nt) was generated. Briefly, the 16S rRNA gene was amplified using the
acidobacteria-specific forward primer, ACD31F and the broadly inclusive bacterial

reverse primer 1492R using the conditions described above. PCR products were cloned

84

and prepared for sequencing as described above. Ten sequencing primers were used to
obtain at an average coverage ranging from 4 to 8x of the 16S rRNA gene are as follows:
ACD31F, 1492R, 338F (5' ACT CCT ACG GGA GGC AGC 3'), 338R (5' GCT GCC
TCC CGT AGG AGT 3'), 531R, 810R (5' GGC GTG GAC TIC CAG GGT ATC T 3'),
776F (5' AGC AAA CAG GAT TAG ATA CCC TGG 3'), 1087F (5' GGT TAA GTC
CCG CAA CGA 3'), modified-M13F, and the modified-M13R. Each primer was used in
duplicate sequencing reactions. Sequences were assembled using the DNA Star
LaserGene Software (Madison, WI) and aligned using the ARB Software (47), and
compared to acidobacterial 16S rRNA gene sequences downloaded from GenBank. The
phylogeny algorithms in ARB were used for the generation of the phylogenetic trees;
PAUP* Version 4.0b10 was used for bootstrapping analysis (72). The 16S rRNA gene
sequences of the Acidobacteria strains described in this study have been deposited in
GenBank with accession numbers AY587227 through AY587230 and DQ660892

through DQ660895.

Characterization of genomic DNA The mole percent G + C content of genomic DNA
from strains KBS 63, KBS 89, TAA 43, and TAA 166 was determined as described
previously (51). Briefly, genomic DNA was extracted using a Qiagen Genomic DNA
Extraction Kit (Qiagen, Valencia, CA), and approximately 5 ug of DNA was digested
with P1 nuclease and alkaline phosphatase. The nucleosides were separated and
quantified using a ShimadzuTM High Pressure Liquid Chromatograph fitted with a UV

detector and a VP Series Alltirna C18 column (250 x 4.6mm; particle size 5pm) (Alltech

85

Associates, Inc., Deerfield, IL). Genomic DNA purified from A. capsulatum (ATCC
Number 51196) was used as a positive control.

The number of 16S rRNA-encoding genes was determined by non-radioactive
Southern hybridization after restriction endonuclease digestion of genomic DNA, as

described previously (35, 36) (http://rrndb.cme.msu.edu/rmdb/servlet/controller). A

 

DIG-labeled, acidobacteria-specific 16S rRNA gene probe was made targeting regions
between 31 and 531 (E. coli numbering). Genomic DNA purified from A. capsulatum

was used as a positive control.

Other physiological tests Catalase and oxidase tests were performed using standard
methods (65, 68). Escherichia coli strain REL 607, a derivative of E. coli B/r (42), was
used as a positive control for the catalase test and a negative control for the oxidase test.
Pseudomonas aeruginosa ATCC 10145 was used as a positive control for the oxidase
test. Fatty acid profiles were generated and analyzed by Microbial ID (Newark, DE;

www.microbialidcom) after all strains, including A. capsulatum, were grown on glucose

 

yeast extract (GYE) medium adjusted to within their optimal pH range.

RESULTS and DISCUSSION
Occurrence and isolation of acidobacteria Acidobacteria are abundant members
of the microbial community in soils at the KBS LTER; their rRNAs account for ca. 1 to
6% of the total bacterial rRNA (8). The specific phylogenetic affiliations of these
acidobacteria were determined from samples collected during August 2005 from different

management treatments and soil depths at the KBS LTER. Partial sequences (ca. 500

86

nucleotides) of 50 cloned 16S rRNA genes from each depth and treatment (total number
of clones was 550) revealed the presence of acidobacteria from subdivisions 1, 3, 4, 5,
and 6. The relative abundance of clones of each subdivision varied with depth and
treatment, suggesting that physical and chemical properties of the soil may influence the
distribution of the various subdivisions. In particular, the percentage of acidobacteria in
subdivision 1 was correlated with soil pH, generally increasing in mildly acidic soils
(Figure 3.1, panel A, gray boxes). To determine if the relationship between soil pH and
relative abundance of subdivision 1 acidobacteria was significant for other terrestrial
environments as well, previously published sequences from a variety of soil
environments (15, 23, 38, 44, 50, 53, 54, 59, 69) were analyzed. The combined data sets
reveal a significant correlation (R2 = 0.38, p < 0.004; Figure 3.1, panel A) between pH
and the percentage of acidobacteria associated with subdivision 1. This result is
consistent with a previously determined relationship indicating that as soil pH decreased,
there was an increased abundance of subdivision 1 acidobacteria in relation to the total
bacterial community (60).

The strains characterized in this study were isolated fi'om soils under a never-
tilled, successional community with herbaceous plants (T8) with an average soil pH of
6.1 and a high percentage of subdivision 1 acidobacteria (27%; Figure 3.1, panel B).
Media and incubation conditions were designed .to mimic the microenvironments thought
to be experienced by microbes in these soils. In particular, relatively low concentrations
of organic carbon in conjunction with extended incubation periods were used to

accommodate oligotrophic, slow-growing acidobacteria (35). The concentrations of

87

,'>

 

Percentage of acidobacteria in subdivsion 1

 

 

12%

160 i)

 

1

Figure 3.1. Summary of acidobacterial 16S rRNA gene clone libraries survey. Panel A
depicts the percent abundance of acidobacteria subdivision 1 out of the total
acidobacterial community at various soil pHs. Data from the KBS LTER are depicted as
gray boxes ( I ), whereas data from published soil surveys are depicted as black boxes (
I ). If a range in pH was given in these studies, the average of that range was used for
this analysis. The equation of the line is y = -13.1x + 103 with an R2 value = 0.38 and p-
value < 0.004. Panel B depicts the average percent abundance of acidobacteria
subdivisions 1, 3, 4, 5, & 6 (subdivisions are indicated next to each pie wedge) from
replicate plots of the never-tilled, successional community with herbaceous plants at the
KBS LTER, depth of 0-7cm.

88

oxygen and carbon dioxide during incubation were also manipulated to better simulate
conditions in soil aggregates. Although upland soils like those at the KBS LTER are
generally well-aerated, they include pockets of anoxia within soil aggregates, and hence
also include transitions zones between oxic and anoxic conditions (27, 66). Moreover,
metabolism of heterotrophic microbes in soil leads to increased concentrations of carbon
dioxide (20), which has the potential to influence the pH of microenvironments.
Accordingly, hypoxic and CO2- enriched incubation atmospheres were included, as was
the incorporation of catalase in the media to minimize the damaging effects of hydrogen
peroxide produced during aerobic respiration.

With the exception of decreased recoveries at lower temperature and in the
presence of AQDS, the recovery of soil microorganisms under the various conditions
tested was between 3.5 to 5 x 107 CF U/gram of soil (dry wt), which was 2.5 to 6% of the
direct cell count (Figure 3.2, panel A). Previous experiments done on three different types
of solidifying agents (Bacto Agar and washed Bacto agar in VSB-7 and agarose in
distilled water) indicated a decrease in total CFU recovery as the purity of the solidifying
agent increased: agar, ca. 5.2 x 107 CF U/gram of soil dry weight washed agar, ca. 4.4 x
107 CFU/gram of soil dry weight; and agarose, ca. 0.3 x 107 CFU/gram of soil dry weight
when grown under oxic conditions. Agar was chosen as a solidifying agent to increase
our recovery of soil microorganisms, thereby presumably increasing our chances of
isolating member of the phylum Acidobacteria. Across treatments used for isolation,
acidobacteria were detected more frequently under atmosphere(s) containing elevated

levels of carbon dioxide (0.05<p<0.10) (Figure 3.2, panel B). This was apparently due to

89

Figure 3.2. Summary of cultivation experiments for total recovery of colonies per gram
of soil (dry weight) (Panel A) and acidobacteria detection (% of n) (Panel B). Panel A
represents the average CFU recovery from soil +/- the standard error; the number of times
a particular treatment was employed (n) is listed at the base of each bar. Panel B
represents the percentage of treatments that yielded a positive test for acidobacteria using
PWPCR from the plates of panel A. The asterisks “*”indicates treatments that yielded
significant difference (panel A, ANOVA a = 0.05; panel B, chi square goodness of fit, or
= 0.10).

90

CFU (107) /gram of soil (dry wt)

Acidobacteria detection (% of n)

 

C02

+ -
aHSL

+ -

catalase

+ -

catalase

**

2% 21% + - 23C 12C + -
02 nutrients temperature AQDS
Treatments

 

2% 21% + - 23C 12C + -

02 . nutrients temperature AQDS
Treatments

91

a mild acidification of the medium under 5% carbon dioxide (plates incubated under 5%
CO2 had a pH ca. 6.5), which favored growth of these acidobacteria, rather than to a
requirement by cells for exogenous carbon dioxide per se (see below).

Although overall recoveries of bacteria here were similar to other studies that
used low nutrient concentrations and extended incubation times (12, 14), the specific
conditions used in this study resulted in the isolation of several novel, pink-pigmented
strains of acidobacteria (strains, KBS 62, KBS ‘63, KBS 68, and KBS 112). All were
isolated from plates that contained a mixture of organic carbon substrates and that were
incubated for 20-30 days under CO2-enriched air.

Previous isolation efforts with homogenized hindguts of Reticulitermes flavipes
soil-dwelling termites under similar cultivation conditions resulted in the isolation of
three acidobacteria strains (strains TAA 43, TAA 48, and TAA 166), which appeared to
be allocthonous contaminants from nearby soil (J .T. Wertz, B.S. Stevenson, and J.A.
Breznak, Abstr. 103rd Gen. Meet. Am. Soc. Microbiol. 2003, abstr. N-223, 2003) and
strain KBS 89 from soils at the KBS LTER. These strains are also characterized further

in the current study.

Colonial and cell morphology and pigmentation Colonies of all acidobacteria were
small, approximately 1mm in diameter (alter 7 day incubation, except for KBS 89 which
took about 14 days), and had a circular form with a convex elevation and entire margin
when grown on VSM-6, -5(KBS89) or R2A. With the exception of KBS 89, all isolates
took approximately 3-5 days before visible, smooth, butyrous colonies were seen on the

surface of the plate of the VSM-6, -5(KBS 89) while KBS 89 took approximately 14-16

92

days to form visible colonies. All strains achieved maximal growth rates in complex
medium, such as RZB, at room temperature (ca. 23°C) with doubling time between 10
and 15 hours.

Cells of all strains are short, plump, Gram negative, nonmotile rods measuring 1
um x 0.6 pm when grown on R2A (Figure 3.3). Cells possessed exaggerated convoluted
outer membranes when viewed by TEM as is typical of a Gram-negative type cell wall.
All strains produced an extracellular matrix of as yet unknown chemical composition, but
which apparently caused cells to stick together tightly (Figure 3.3, panel 1) in colonies
and form visible flocs in liquid culture, qualitatively to the same extent under microoxic
and oxic conditions. The formation of such extracellular material in soils may serve as a
form of protection from predation, or a web to trap water or nutrients (57), and may be
involved in the formation of soil aggregates.

Four strains (KBS 62, 63, 68, and 112) were pink, owing to the presence of a
pigment(s), whose solvent extractability and absorption spectrum were typical of
carotenoids. Interestingly, the pigmentation was more pronounced in cells grown under
20% 02 than under 2% 02 (Figure 3.4, panel A). The ratio of the absorbance maxima of
the two dominant peaks in acetone:methanol (A493 and A523) were nearly identical, 1.2 :t
0.0 and 1.3 d: 0.1, respectively, suggesting the same carotenoid(s) was produced under
each oxygen regime (Figure 3.4, panel B). The visible absorption spectrum of strain
KBS 63 pigment(s) in chloroform had maxima at 473, 505, and 539 nm, which are most

similar to those of spirilloxanthin (475, 505, and 543 nm) (7).

93

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94

 

Absorbance

 

 

 

330 400 4'50 '_ s00 ' 5'50 ' 600
Wavelength (nm)

Figure 3.4. Strain KBS 63 grown on R2A under 20% and 2% oxygen (vol/vol) of
nitrogen & 5% CO2 mix) (panel A). Absorption spectra of strain KBS 63 cells extracted
in acetonezmethanol (7:2) after growth in 20% ( I ) and 2% (I) oxygen on R28 (panel B).

95

Carotenoids are naturally occurring accessory pigments that typically absorb
visible light maximally between 450 and 550 nm (48), and are known to play roles in
light-harvesting for photosynthesis and protection from photooxidative damage (6), as
well as quenching singlet oxygen (11). Carotenoids are found not only in phototrophic
bacteria, but in numerous, phylogenetically diverse genera of heterotrophic soil bacteria,
including Streptomyces, Myxococcus, Flavobacterium, and Erwim'a (2). Carotenoid
synthesis has long been known to be regulated by two key environmental factors, oxygen
and light (2). While light had no measurable influence on the carotenoid(s) content of the
T erriglobus strains, the concentration of carotenoid(s) in T. roseus increased in response
to oxygen. We suggest that increased production of carotenoid(s) under 20% oxygen as
compared to 2% oxygen concentration represents a response to oxidative stress. The
potential antioxidant property of carotenoids is well known: B-carotene is effective in
quenching singlet oxygen (11) and the addition of carotenoid genes to E. coli inhibits
peroxidation (1). Specifically in heterotrophs, carotenoids can influence growth rate and
survival (52) presumably through the inactivation of reactive oxygen species generated
during respiration. Differential expression of carotenoids might also be advantageous in
the soil environment, where oxygen gradients change in response to soil moisture and
depths, with higher concentrations of oxygen occurring in upper layers of soil that are
also subject to more intense solar radiation where carotenoids would also provide
protection against the damaging effects of ultraviolet radiation.

The frequency of acidobacteria detection was notably higher in the presence of
catalase (Figure 3.2, panel B), suggesting that acidobacteria are sensitive to damage by

reactive oxygen species. All strains, except for KBS 89, tested positive for the presence

96

Table 3.1.

Acidobacterium strains.

Phenotypic characteristics

of Terriglobus,

Acidobacteriaceae, and

 

 

 

 

Terriglobus Strains Acidobacteriaceae Strains Acidobacterium
Characteristics KBS 63* TAA 43* KBS 89 TAA 166 A.capsu1atum'
Origin soil my; soil termite hindgut afig‘g‘g"
Cell Shapeb Rod Rod Rod Rod Rod
Length (um) 1.1i0.2 1.24:0.2 ca. 1 1.1 :1:0.3 1.1-2.3
Width (um) 0.6 :t 0.1 0.6 i 0.1 ca. 0.5 0.6 :1: 0.0 0.3 - 0.8
Pigment pink white white white orange
rrs copy number 2 1 1 1 1
81:19, fitment 59.8 e 0.5 58.1 a: 0.04 59.7 e 1.8 54.7 a: 0.7 “£8682; (1:91
Catalase + + - + +
Oxidase - - - - -
Motility - - - - +
pH range
(optimum) 5 — 7 (6) 5 — 6.5 (5) ND 5 — 7(6) 3 — 6
Growth at 4°C - - - - ND
Growth at 12°C + + + + ND
Growth at 25°C + + + + ND
Growth at 37°C - - - - +
Differences
Carbon
Utilization:
D-Ribose - - + - ND
D-Sorbitol - - + - ND
Purrne & _ + _ _ ND

Derivative Mix°

 

 

 

 

*Data are identical for strains KBS 62, KBS 63, KBS 68, and KBS 112.
* Data are identical for strains TAA 43 and TAA 48.
u The mole percent G+C content was only done for strains KBS 63, KBS 89, TAA 43,

and TAA 166.

a Reference: characteristics described from a previous study (3 3). ND “not determined”
b Cell length and width was determined from a culture in mid-log phase on RZB medium.
° The mix of adenine, guanine, thymine, cytosine, and uracil had a final concentration of

10 ug/ml (total concentration of sugar derivative was 50ug/ml).

 

of catalase (Table 3.1), and some strains produced a carotenoid(s) which was
preferentially expressed under higher oxygen concentrations (Figure 3.4). It would be
beneficial for a soil microorganism to have mechanisms to deal with the damaging effects
of reactive oxygen species, since there are oxygen gradients in the three-dimensional soil
structure that change with properties such as soil moisture and nearby microbial

communities.

Physiological properties of strains Physiological properties of all strains are
tabulated in Table 3.1. The growth rates of all strains grown in VSM- 5 (KBS 89) or
VSM-6 (all others) were highest (avg. u z 0.15) at room temperature under air. Growth
was observed over a pH range of 4.5 to 7.0. It is not surprising that the pH optima of the
strains were similar to that of their native environment. Most soils are slightly acidic due
to the presence of humic substances, which make up 70-80% of the soil organic matter
(24). Sait et a1. recently suggested that there is an effect with pH on the isolation and
distribution of subdivision 1 acidobacteria. Specifically, representatives of subdivision 1
formed more colonies in media having a pH 5.5 as compared to a pH of 7.0 after a 4
month incubation (60). Furthermore, the results of our molecular-based survey reveals
that subdivision 1 acidobacteria are found in a higher abundance in mildly-acidic to
acidic soil environments (Figure 3.1, panel A). Taken together, these results suggest that
members of subdivision 1 of the Acidobacteria have a preference for mildly acidic pH
conditions, which is consistent with their ubiquitous distribution in soil.

We examined the effect of CO2 on the growth of isolated strains to explore the

greater frequency at which acidobacteria formed colonies on plates incubated under 5%

98

CO2 (Figure 3.2, panel B). There was no significant difference in growth rate of strains
KBS 63, TAA 43, and TAA 166 incubated under an atmosphere of air (ca. 0.05% CO2
vol/vol) or 5% CO2 (balance air) (p-value = 0.83, 0.98 and 1.0, respectively). Therefore,
the increased frequency of appearance of acidobacteria on plates incubated under 5%
CO2 appears to be a result of mild acidification of the isolation medium. The degree of
acidification that can occur on plates of media without added bicarbonate and incubated
under elevated levels of carbon dioxide (pH ca. 6.5) is consistent with the optimal pH of
the strains (Table 3.1).

The relatively slow growth rate of the acidobacteria strains is consistent with the
presence of one (strains TAA 43, 48, 166, and KBS 89) or two (strains KBS 62, 63, 68,
and 112) genes coding for the 16S rRNA gene (Figure 3.5). Previous studies have
indicated that the number of rRNA operons correlates with the rate at which bacteria
respond to resource availability and how efficiently those resources are utilized (35, 71).

The metabolism and growth of bacteria in soil environment are frequently limited
by carbon availability, but subject to periodic fluctuations in resource resulting from
events including rain or spring snow melt (74). Populations of Acidobacteria are more
abundant in soil environments where resource availability is low (19), and can be
described as oligotrophic bacteria because of this distribution in soil, their relatively slow
growth rate, and possession of either one or two copies of the 16S rRNA encoding gene —
a genomic marker of oligotrophic bacteria (35). Although the acidobacteria characterized
in this study do not fit the definition of an obligate oligotroph because they are capable of
growth on rich media, they do fit into the broader notion of an oligotroph as being an

organism adapted to low-nutrient environments (3 7).

99

b
KBS63 KBS62 'KBS68 KBSIIZ E

A

— Marker

2345678910

23,130 bp —

9,4l6bp —
6,557 hp -

4,361 hp —

2,322 bp -
2,027 hp _

 

' h
TAA48 TAA43 K3589 TAA166 A.“ '15
I I I I I I I l I l 2

B

'- Marker

2 3 4 S 6 7 8 9 10 11 12
23,130bp—

9,416 bp —
6,557 bp -

4,361 bp -

2,322 bp —
2,027 hp _

 

Figure 3.5. Southern hybridization of digested genomic DNA fi'om strains KBS 62, KBS
63, KBS 68, KBS89, KBS 112, TAA 43, TAA 48, & TAA 166. In panel A, isolates were
cut with EcoRV(lanes 2, 4, & 6), Sma I (lane 3, 5, 7, & 9), and Rsa I (lane 8). In panel B,
isolates were cut with Sma I (lanes 2, 4, 6, 8, & 10) and Apa I (lanes 3, 5, 7, 9, & 11).
The lambda DNA Hind III marker (lane 1 & 10 (panel A) and lanes 1 & 12 (panel 12))
provided size estimates.

100 ‘

All strains were able to grow in a 2% oxygen atmosphere as well as atmospheric
concentrations of oxygen, with or without elevated levels of carbon dioxide, but they
could not grow anaerobically with nitrate, iron (Fe(III)), or AQDS as external electron
acceptor, nor could they grow by anaerobic fermentation of glucose, lysine, or omithine
(Table 3.1). Growth under low levels of oxygen would be advantageous in soil, since
oxygen is depleted in the soil environment, particularly in soil aggregates as soil moisture
increases (5, 55, 74).

There were nominal differences in the carbon utilization profile; all strains were
able to oxidize D-glucose, D-fructose, D-galactose, D-mannose, D-xylose, D-arabinose,
sucrose, D-maltose, D-raffinose, D-cellobiose, sodium succinate, D-glucuronate, and D-
gluconic acid; however they were not able to oxidize D-mannitol, methyl cellulose,
carboxymethyl cellulose, sodium acetate, sodium pyruvate, sodium formate, maleic acid,
the organic acid mix, D-galacturonate, monomeric constituents of lignins, or humic acids.
Only two differences in the carbon utilization pattern were noticed, strain KBS 89 was
able to oxidize ribose and sorbitol while strains TAA 43 and TAA 48 were the only
strains capable of oxidizing the mixture of purines and pyrimidines (Table 3.1).

The whole-cell fatty acid compositions of the newly isolated strains and A.
capsulatum are listed in Table 3.2 and Figure 3.6. The dominant fatty acids for the new
strains are 15:0 iso (36.33-47.09%) and 16:1 (070/15 iso 2-OH (summed feature 3)
(26.18-30.24%). These results were compared to the Microbial ID database, but did not
correspond to any known species. A. capsulatum was sent as a source of comparison, and

its dominant fatty acids were 15:0 iso (40.28%) and 18:1 (0% (21.21%).

101

Table 3.2. Whole-cell fatty acid composition (%) of Terriglobus, Acidobacteriaceae, and
Acidobacterium strains grown in GYE medium. The most abundant fatty acids that
distinguish the T erriglobus strains from A. capsulatum are presented in bold font.

102

 

 

 

 

 

 

 

 

 

 

 

Terriglobus Acidobacteriaceae Ac Ida ba cter tum
Fatty acid Strains Strains
KBS 63 TAA 43 KBS 89 TAA 166 A. capsulatum
Saturated
14:0 3.42 4.25 1.16 3.68 1.58
15:0 0.64 0.74 0.00 0.60 0.57
16:0 7.77 9.14 7.33 8.71 4.29
16:0 N alcohol 0.52 3.54 0.50 5.01 4.61
17:0 0.70 0.00 0.93 0.00 1.85
18:0 0.38 0.68 0.75 0.59 2.34
20:0 0.76 0.00 4.07 0.00 0.00
Unsaturated
14:1 (05¢ 1.07 0.67 1.00 0.57 0.00
15:1 (06c 0.83 0.92 0.43 0.71 0.00
16:1 (05c 0.43 0.00 0.00 0.56 0.00
17:1 018C 0.00 0.00 0.00 0.00 3.97
18:1 (09¢ 0.00 0.00 0.00 0.00 21.21
18:1 (07c 0.00 0.84 0.00 1.27 0.45
18:1 (05c 1.05 0.00 0.00 0.00 2.24
19:0 cyclo (08c 0.00 0.00 0.00 1.10 0.00
20:2 006,9c 0.67 0.00 0.00 0.00 0.00
Methyl-branched
13:0 iso 4.02 0.00 4.12 0.00 0.00
15:0 iso 43.88 36.33 47.09 37.06 40.28
15:0 anteiso 0.48 0.70 0.00 1.03 0.00
16:0 iso 0.00 0.00 0.00 0.58 0.00
iso 17:1 009C 0.00 0.00 1.25 0.00 0.90
iso 17:1 (05c 4.24 7.27 2.33 0.00 7.40
17:0 iso 0.53 0.79 1.60 0.78 2.15
17:0 anteiso 0.27 0.60 0.00 0.74 0.49
19:0 anteiso 0.00 1.82 0.00 0.00 0.00
Hydroxy
15:0 iso 3-OH 0.00 0.00 0.58 0.00 0.00
17:0 iso 3-OH 0.00 0.00 0.00 0.00 0.77
Summed Feature
15:1 iso 1-1/13:0 3-OH 0.59 0.00 0.36 0.00 0.32
16:1 007e/15 iso 2-OH 27.08 30.24 26.18 26.99 4.30
17:1 iso i/anteiso b 0.00 0.00 0.00 8.07 0.00
18:2 (06,9c/18:0 anteiso 0.64 1.47 0.31 1.96 0.00
Unknown
Unknown 11.799 0.00 0.00 0.00 0.00 0.28

 

103

 

 

TAA 166

 

TAA 43

 

 

 

 

 

A. capsulatum

1 1 1
0 5 10

Euclidian Distance

Figure 3.6. Euclidian distance dendrogram for T erriglobus, Acidobacteriaceae, and
Acidobacterium strains generated from the fatty acid profiles.

104

Acbt.capsl (D2617l)
env.WD277 (A1292577)
Ellin5225 (AY234576)
Ellin5017 (AY234434)
KBSS9 (AY587227)
env.TM2 (X97098)
Ellin351 (AF498733)
TAA166 (AY587230) 1
Ellin337 (AF498719)
TAA43 (AY587228)
TAA48 (AY587229)
KBSllz (DQ660895)
KBS63 (DQ660892)
KBS62 (DQ660893)
__ KBS68 (DQ660894) .

   
   
 
  
 
 
   

 

 

 

 

 

 

 

 

 

 

 

 

 

1 . / ]3
___+ / ]4
—C7 ]5

0.10

 

Figure 3.7. Maximum-likelihood tree of the Acidobacteria subdivisions 1, 3, 4, 5, and 6
(indicated to the right of the group) based on 16S rRNA gene using sequences obtained
from cultivated representatives and environmental clones. Geothrix fermentans and
Holophaga foetida of subdivision 8 were used as an outgroup (not shown). Strains fiom
this study are in bold font. Each gray trapezoid represents a different subdivision with
approximately 8 representative sequences. Internal nodes supported by a bootstrap value
of >95% are indicated with a filled circle ( 0 ) and of >75 with an open circle ( o ). The
scale bar indicates 0.10 changes per nucleotide.

105

Six of the strains (KBS 62, KBS 63, KBS 68, KBS 112, TAA 43, and TAA 48)
are sufficiently similar to one another, but distinct from previously isolated acidobacteria
(Figure 3.7), to warrant creation of a new genus. The similarity of the 16S rRNA gene
sequences within these six strains is 96% and, on average, only 92% similar to
Acapsulatum. Moreover, unlike A. capsulatum these new strains do not require
extremely low pH conditions (pH 3.0-6.0), they cannot grow at 37°C, and they contain
16:1 (07c/15:0 iso 2-OH as a dominant fatty acid and not 18:1 (1)96, which is dominant in
A capsulatum (Table 3.2, Figure 3.6). Hence, we propose that these six strains be
regarded as members of a new genus, T erriglobus, with KBS 63 as the type strain of the
new species, roseus, which also includes the other pink-pigmented strains (KBS 62, KBS
68, and KBS 112). The nomenclature and description of the genus and species are given
below. We are postponing application of specific epithets to strains TAA 43 and TAA 48
until further work is done to determine their relatedness to strains comprising the species,
T. roseus. Although the characteristics of strains KBS 89 and TAA 166 are consistent
with the description of T erriglobus, there is insufficient metabolic information currently
available about the previously isolated and closely related strains Ellin 351 and Ellin 337

(61) to include all four strains in the genus T erriglobus at this time.

Description of T erriglobus gen. nov. T erriglobus gen. nov. (Ter.ri.glo' bus. L.
fem n. term, earth; L. masc. n. globus ball, clump; N.L. masc. n. T erriglobus clump of
earth) literally translated as “clump of earth” in the phylum Acidobacteria. Gram
negative, short, plump, rods (ca. 1 um x 0.6 pm) when grown in R2B medium. Moderate

acidophiles (pH range for growth is 5.0 - 7.0, optimal for KBS 63 is 6.0 and TAA 43 is

106

5.0). Produce an extracellular matrix which apparently facilitates cell clumps in liquid
media incubated under an atmosphere containing either 2% or 20% molecular oxygen.
G+C content ca. 59 mol% (Table 3.1); 15:0 iso and 16:1 (07c/ 15 :0 iso 2-OH as dominant
fatty acids (Table 3.2). Members of the phylum Acidobacteria, subdivision 1. The type

species is Terriglobus roseus.

Description of T erriglobus roseus sp. nov. T erriglobus roseus sp. nov. (ro'
se.us. L. masc. adj. roseus rose-colored, pink), “a pink T erriglobus” (type strain KBS 63).
Pink-pigmented colonies due to the presence of carotenoids; catalase positive; oxidase
negative; and two copies of the rrs. Isolated from soils in a never-tilled, successional
plant community at the KBS LTER. The type strain KBS 63 was deposited into the
USDA Agricultural Research Service Culture Collection (NRRL B-41598T) and

Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSM 18391).

ACKNOWLEDGEMENTS

This work was funded by the USDA (#2001-35107-09939), NSF (#MCB-
0135880), and the NSF Long-Tenn Ecological Research Program at the Kellogg
Biological Station and the Michigan Agricultural Experiment Station.

The authors would like to thank Kwi S. Kim for her assistance with determining
the rrs copy number, Les Dethlefsen for his help with the statistical analysis, Frank B.
Dazzo for his assistance with the CMEIAS software, Kristin M. Huizinga and Jorge L.M.
Rodrigues for their help with the soil collection, Joseph R. Graber with his help on G+C

mole percent determination, Zachary Blount for his helpful suggestions on the

107

manuscript, and John P. Eichorst and Hans G. Triiper for their help with the

nomenclature and Latinized specific epithets.

108

10.

11.

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uncultured microbes. Appl. Environ. Microbiol. 70:4748-4755.

Stevenson, B. S., and T. M. Schmidt. 2004. Life history implications of rRNA
gene copy number in Escherichia coli. Appl. Environ. Microbiol. 70:6670-6677.

Swofford, D. L. 2003. PAUP“. Phylogenetic analysis using parsimony (* and
other methods). Version 4.0b10. Sinauer Associates, Sunderland, Massachusetts.

van Elsas, J. D., and K. Smalla. 1997. Methods for sampling soil microbes, p.
383-390. In C. J. Hurst, G. R. Knudsen, M. J. McInemey, L. D. Stetzenbach, and
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van Elsas, J. D., J. T. Trevors, and E. M. H. Wellington. 1997. Modern soil
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115

75.

76.

Zimmermann, J., J. M. Gonzalez, and C. Saiz-Jimenez. 2006. Epilithic
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Zimmermann, J., J. M. Gonzalez, C. Saiz-Jimenez, and W. Ludwig. 2005.
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Geomicrobiol. J. 22:379-388.

116

CHAPTER 4

DESCRIPTION OF NOVEL MEMBERS OF SUBDIVISION 1 AND 3 IN THE

PHYLUM ACIDOBAC T ERL4 FROM AGRICULTURAL SOIL

INTRODUCTION

Members of the phylum Acidobacteria are abundant and diverse members of the
soil environment. They have been retrieved from various environments including: soils
and sediments (5, 15), soil crusts of sand dunes (51), wastewater (10, 29), water
distribution systems (32), peat bogs (13), acid mine drainage (24), hot springs (18),
shallow submarine hydrothermal vents (49), and on the surface of Paleolithic cave
paintings and catacombs (45-47, 62, 63). It is a diverse phylum, historically harboring
eight, monophyletic subdivisions with recent evidence of eighteen additional
subdivisions, some unique to uranium contaminated subsurface sediments (4). Their
common occurrence and high abundance suggests they play an ecologically significant
role(s) in the soil environment.

Recently, scientists have made successful attempts to cultivate members of
subdivision 1, 2, 3, and 4 (11, 13, 14, 16, 20-22, 42-44, 52) from soil and sediment
environments. To date, cultivation efforts have been most successful with members of
subdivision 1, with more than forty isolates obtained fiom this subdivision; however,
there are only a few cultivated representatives from subdivisions 2, 3, and 4. In order to
cultivate new strains in these poorly represented subdivisions, the information generated

from the acidobacteria soil survey (Chapter 2) was used to develop additional cultivation

117

strategies such as the influence of soil pH (subdivision 1), soil moisture (subdivision 3),
and low carbon concentrations (subdivision 4). Soil from the conventional agriculture
site at the KBS LTER (treatment 1) was used as the source of inoculum in an attempt to
isolate members of subdivision 4, given their high prevalence in this soil (Chapter 2).
These experiments were successful at isolating additional members of subdivision
1 as well as a novel strain in subdivision 3 using medium with plant polymers as the
carbon source at low pHs. As such, the characterization of newly isolated strains had
particular emphasis on determining the ability of these strains to use xylan and more
recalcitrant carbon sources such as benzoate, syringate, cellulose, and resorcinol as well
as their ability to use nitrate since the inoculum was fiom a conventional agricultural soil,
which has high inputs of nitrates. Microorganisms that harbor active enzymes to degrade
recalcitrant carbon sources are essential for maintaining an active carbon flow in the soil,
since typically these compounds are slow to decompose. Nitrate as well as nitrite, under
anoxic conditions, can be used as an electron acceptor for anaerobic respiration and be
reduced to gaseous forms of nitrogen such as nitric oxide, nitrous oxide, and dinitrogen
gas, a process commonly referred to as denitrification. This process results in a loss of
fixed nitrogen, ca. 20 to 30% of the added nitrogenous fertilizers to agricultural fields
(56), as well as producing the dangerous greenhouse gas, nitrous oxide. Results from the
physiological and phenotypic characterization warrant the creation of a new genus in

subdivision 1 and in subdivision 3.

MATERIALS and METHODS

118

Cultivation experiments Soil samples were collected from the conventional
agriculture plot (treatment 1) and never tilled suCcessional plot (treatment 8) in July 2006
from the Michigan State University WK. Kellogg Biological Station Long Term
Ecological Research (KBS LTER) site. The KBS LTER is a 48-hectare research site
established in 1989 to study ecological processes in agroecosystems. The dominant soil
types are of the Kalamazoo (fine-loamy, mixed, mesic Typic Hapludalfs) and Oshtemo
(coarse-loamy, mixed, mesic Typic Hapludalfs) series. A detailed description of this site

can be accessed at http://wvtwkbs.msu.edu.

 

For isolation attempts, five cores (2.5 cm diameter x 10 cm depth) from the
conventional agricultural plot (T1) were collected and homogenized with a 2 mm sieve
on the bench top. Portions of the homogenized soil were used to determine soil moisture
(40). Approximately 30 grams of soil were added to 100 mL of a phosphate-buffered
salts solution (137 mM NaCl, 2.7 mM KCl, 10mM Na2HPO4 and 2 mM KH2PO4); pH
was adjusted to 7.0, and supplemented with 2.24 mM Na4P2O7-10H2O as a dispersal
agent, and dithiothreitol (lmM) as a reducing agent (57). The soil suspension was stirred
vigorously with a magnetic stir bar for 30 minutes on the bench top. Denser soil
aggregates were then allowed to settle for 30 minutes, after which an aliquot was used to
prepare lO-fold decimal dilutions in the same buffer for subsequent inoculation of the
isolation medium consisting of VL55 medium (22) with either R2B carbon (V L55-R2B),
a mixture of plant polymers (V L55-PP), or xylan from birchwood (V L55-xylan);
hereafter, this composition will be referred to as VL55-(R2B, PP, or xylan) - # “VL55
medium with either carbon as R2B, plant polymer mix, or xylan”; # indicates the

respective pH of the basal solution. The plant polymer mix contained 0.05 grams/liter

119

of the following components: xylan (derived from oat spelts), xanthan, pectin, xylan
(derived from birchwood), methyl cellulose, and xylan (derived from larch) (Sigma-
Aldrich Co. St. Louis, MO). The R2B carbon mixture contained 0.05 grams/liter of the
following components: yeast extract, protease peptone #3, casamino acids, dextrose, and
soluble starch. The xylan mixture contained 0.05 gram/liter of xylan from Birchwood
(Sigma-Aldrich Co.). The final pH of the plates was buffered to either 5.5 or 6.5 with 10
mM of MES, and they were incubated for 30 days at 23°C under one of the following
atmospheres: CO2-enriched hypoxia (vol/vol: 2% O2, 5% CO2, and bal N2); CO2-enriched
anoxia (vol/vol: 85% nitrogen, 10% hydrogen, 5% CO2), or air. Plates were screened for

the presence of acidobacterial colonies after ca. 30 days of growth using PWPCR (52).

Isolation and standard growth medium During the isolation of these strains, an
alternative medium was sought that would provide better growth of the strains, KBS 83
and KBS 96: this provided to be modified hyphomicrobium medium “337” (36) (MHM).
MHM contained (per liter): NaCl 1.0 g; MgCl2-6H2O 0.4 g; CaC12o2H2O, 0.1 g; NH4C1,
0.25 g; KH2PO4, 0.2 g; KCl, 0.5 g; Na2SO4, 0.28 g; KNO3, 5 g; KPO4, 0.15 mM, 20 mM
respective buffer; 10 mM designated carbon source (for KBS 83) or a mixture of organic
compounds (below) (for KBS 96); along with 1 mL SL-10 trace element solution; 1 mL
vitamin B12 (50mg/mL); and 1 mL vitamin mix as described previously (16). All other
strains were successfully isolated and routinely grown on R23 - a complex medium
containing yeast extract, amino acids, peptone, carbohydrates, pyruvate, and inorganic

salts (3 8).

120

Optimization of growth rate MHM was amended with 10 mM glucose (KBS
83) or a mixture of organic compounds (yeast extract, protease peptone, casamino acids,
and glucose - 0.15g/l each) (KBS 96) to assess capacity for growth under a range of
conditions. This complex medium (designated MHM-#, where # indicates pH) was used
to determine the effect of pH on growth rate. Strains were grown at room temperature
(23°C) between pH 3.0 and 7.0 at intervals of 0.5 pH units: MES was used to buffer at
pH ranges between 5.0 to 6.0, 4-morpholinepropanesulfonic acid (MOPS) was used at pH
ranges between 6.5 to 7.0, and citric acid was used as a buffer below pH 5.0, all at a final
concentration of 20 mM. Culture tubes (BellcoTM cat. no. 2048-18150) containing 5 mL
of medium, were stoppered under an air atmosphere and held on a reciprocating shaker,
horizontally, operating at ca. 50 strokes/min. to disrupt cellular flocs. Optical density at
600nm was monitored periodically with a ThermoTM Spectronic model 20D+
spectrophotometer. Inocula consisted of a 1:100 dilution of cultures in mid/late log phase
growth (ca. 1x108 cells/mL) in MHM-5 with 10 mM glucose (KBS 83) or MHM-5 with
0.3x R2B (KBS 96).

Strain KBS 83 was tested for its ability to grow under hypoxic conditions with the
following concentrations of oxygen in the headspace (vol/bal. He): 2%, 4%, 8%, 16%,
and 21% on MHM-5.5. A final concentration of 2.5 mM glucose was used to ensure an
excess of oxygen in the headspace of the 750 mL sealed, side-arm flasks. Similar
experiments were performed for strain KBS 96, however this strain was grown on MHM-
5 with a mixture of organic compounds (yeast extract, protease peptone, casamino acids,
and glucose - 0.05 g/l each) due to some unknown auxotrophic requirement for better

growth. The headspace was purged every third day of growth and oxygen re-added at the

121

respective concentration. Strains were grown at room temperature, on an orbital shaker
at 150 RPM, and growth was monitored as described above. Inocula consisted of a 1:100
dilution of cultures in late log/early stationary phase growth (ca. 1x108 cells/mL) in

MHM-5 with respective carbon source, presumed to be depleted in oxygen.

Carbon utilization Strains KBS 83 & KBS 96 were tested for their ability to oxidize
various carbon sources. Strains were grown under oxic conditions on the following
carbon sources for ca. 30 days, with a 10 mM final concentration on MHM-5.5: D-
glucose, D-fructose, D-galactose, D-mannose, D-ribose, D-xylose, L-arabinose, D-
mannitol, D-sorbitol, sucrose, D-maltose, D-raffrnose, succinate, acetate, formate,
pyruvate, benzoate, ferulate, resorcinol, syringate, and trimethoxybenzoate. Methyl
cellulose, pectin, cellulose, and a xylan mix, containing xylan fiom oat spelts, birchwood,
and larch were used with MHM-5.5 at a final concentration of 0.1% (wt/vol). Culture
vessels and incubation were identical to the pH conditions (above). Due to an unknown,
auxotrophic requirement of strain KBS 96 for faster growth, a small amount of yeast
extract (5 ug/mL) was added to the medium for growth; grth for each respective
carbon source was determined after the amount of growth on 5 ug/mL yeast extract alone

was subtracted.

Capacity to utilize nitrate Strains KBS 83 & KBS 96 were tested for their ability to
grow anaerobically and reduce nitrate by determining the concentration of ammonium,
nitrite, and nitrous oxide while simultaneously monitoring the quantity of nitrate at the

beginning and end of growth. Strains were grown in MHM-5.5 with 10 mM glucose in a

122

headspace of helium. Culture vessels and incubation were identical to the pH conditions
(above). Nitrate, nitrite, and ammonium were measured at the Michigan State University
Soil Testing Laboratory using the Lachat QuickChem automated flow injection ion
analyzer. Nitrous oxide, if present, and carbon dioxide were measured using a
ShimadzuTM thermal conductivity gas chromatograph series GC-2014 with the following
settings: flow rate, 60 mL/min; column temperature, 55°C; injector temperature, 100°C;
detector temperature, 100°C; and a thermal conductivity of 175 mA. Glucose
concentrations were determined using the Glucose Assay Kit (Sigma Aldrich Co., St.

Louis, MO).

16S rRNA gene phylogeny Nearly full length sequence of the 16S rRNA gene (ca.
1,500 nt) was generated by amplifying it with the broadly inclusive bacterial forward
primer 8F and bacterial reverse primer 1492R (28). Each 25111 PCR reaction contained
1x PCR buffer, 1mM MgCl2, 0.03mM of each dNTP, 0.23M each primer, and 5U Taq
DNA polymerase (Invitrogen, Carlsbad, CA). Thermal cycling consisted of the
following steps: (1) 95°C for 3 minutes; (2) 95°C for 30 seconds, 56°C for 30 seconds,
72°C for 45 seconds (repeated 30X); (3) 72°C for 10 minutes. Genomic DNA purified
from A. capsulatum (ATCC Number 51196) was used as a positive control. PCR
products were electrophoresed through a 1% agarose gel in 0.5x tris-borate-EDTA (TBE)
and visualized with GelStar Nucleic Acid Stain (BioWhittaker Molecular Applications,
Rockland, MA). PCR products were cloned with the Invitrogen TOPO TA Cloning Kit
for Sequencing (Invitrogen Carlsbad, California) and the inserts were re-amplified using
the modified-M13F (F2) (5’ CAG TCA CGA CGT TGT AAA ACG ACG GC 3’) and

modifed-M13R (F4) (5’ CAG GAA ACA GCT ATG ACC ATG 3’) (23) with the PCR

123

conditions described above. The PCR products for sequencing were treated with
ExoSAP-IT (USB, Cleveland, OH) using a modification of the manufacturer’s protocol
(ExoSAP enzyme was diluted 1:8 and the reaction was incubated at 37°C for 30 minutes,

followed by an 80°C incubation for 15 minutes) and submitted for sequencing with
ACD31F, 1492R, 338F (5' ACT CCT ACG GGA GGC AGC 3'), 338R (5' GCT GCC
TCC CGT AGG AGT 3'), 531R, 810R (5' GGC GTG GAC TTC CAG GGT ATC T 3'),
776F (5' AGC AAA CAG GAT TAG ATA CCC TGG 3'), 1087F (5' GGT TAA GTC
CCG CAA CGA 3'), modified-M13F, and the modified-M13R. Each primer was used in
duplicate sequencing reactions. Sequencing was performed at the Michigan State
University Research Technology Support Facility (genomics.msu.edu/index.html).
Sequences were assembled using the DNA Star LaserGene Software (Madison, WI) and
aligned using the ARB Software (31), and compared to acidobacterial 16S rRNA gene
sequences downloaded from GenBank. The phylogeny algorithms in ARB were used for
the generation of the phylogenetic trees; PAUP* Version 4.0b10 was used for

bootstrapping analysis (5 4).

Colony and cell morphology Bacterial colonies on plates of MHM medium were
examined under a Nikon SMZ-2T dissecting microscope at 10-15X for size,
pigmentation, form, elevation, and margin (50)., Gram stain reactions were performed as
described previously (48) and cells were stained with India ink to determine if they were
capsulated. Gross cell morphology and motility were assessed by using phase contrast

microscopy with a Zeiss Axioskop Microscope (Carl Zeiss Inc., Thomwood, NY).

124

Transmission electron microscopy was completed at the Michigan State University

 

Center for Advanced Microscopy (\wrwzceo.msu.edu).

Characterization of genomic DNA The mole percent G + C content of genomic
DNA from strains KBS 83 and KBS 96 was determined as described previously (33).
Briefly, genomic DNA was extracted using a Qiagen Genomic DNA Extraction Kit
(Qiagen, Valencia, CA), and approximately 2 ug of DNA was digested with P1 nuclease
and alkaline phosphatase. The nucleosides were separated and quantified using a
ShimadzuTM High Pressure Liquid Chromatograph fitted with a UV detector and a VP
Series Alltima C18 column (250 x 4.6mm; particle size Sum) (Alltech Associates, Inc.,
Deerfield, IL). Genomic DNA purified from A. capsulatum (ATCC Number 51196) was
used as a positive control.

The number of 16S rRNA-encoding genes was determined by non-radioactive
Southern hybridization after restriction endonuclease digestion of genomic DNA, as

described previously (25, 26) (http://rrndb.cme.msu.edu/midb/scrvlet/controller). A

 

DIG-labeled, acidobacteria-specific 16S rRNA gene probe was made targeting regions
between 31 and 531 (E. coli numbering). Genomic DNA purified from A. capsulatum

was used as a positive control.

RESULTS and DISCUSSION
Isolation of new members of subdivision 1 & 3 The recovery of soil
microorganisms using this new medium base, VL55, along with more native carbon

sources such as plant polymer mixture and xylan was between 0.6 to 1.3 x 107 CFU/gram

125

of soil (dry wt) (Figure 4.1), which was lower then previous recoveries which ranged
between 3.5 to 5 x 107 CFU/gram of soil (dry wt) (16). The major difference in these
cultivation experiments from previous ones was the use of Gellan Gum, a bacterial
exopolysaccharide, as the solidifying agent which was shown previously to be superior to
agar for isolating novel members of the acidobacteria (21, 44). Regardless of the media
additive or pH, the recovery under hypoxic conditions was similar, however under oxic
conditions there was a ca. 1.3-fold higher recovery of microorganisms when xylan and
the plant polymer mixture were used as the source of carbon with the conventional
agriculture soil (treatment 1) as the source of inoculum as compared to the mid-
successional soil (treatment 8) at pH 5.5 (Figure “4.1).

The inclusion of these compounds appears to have led to the successful
cultivation of the new subdivision 1 strain KBS 83 which was isolated from VL55-PP at
a pH of 5.5 fiom the conventional agriculture plot (treatment 1) at the KBS LTER after
incubation at room temperature for ca. 30 days in an oxic atmosphere. Perhaps the use of
these more recalcitrant carbon sources prevented substrate-accelerated death (53) for
strain KBS 83 since these polymers must be hydrolysed before used as a source of carbon
(7, 30), as suggested to be responsible for the increased viable counts of soil bacteria
from Australian soil (44). The new strain in subdivision 3, KBS 96, was isolated on
VL55-R2B under the same conditions described for KBS 83.

Genotypic, physiological, and morphological properties of KBS 83 and KBS 96
will primarily be described in this chapter. Additional strains isolated during these
experiments were strains KBS 5 and 13, isolated on VL55-R2B at pH 5.5 and 6.5,

respectively, and KBS 3 was isolated on VL55-PP, pH 5.5 from the never-tilled,

126

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Figure 4.1. Summary of cultivation experiments. Panels A & B depict total recovery 1
standard error after 30 days of growth fiom plates incubated under oxic conditions with
the media at a pH of 5.5 and 6.5, respectively. Panels C & D depict total recovery alter
30 days for growth from plates incubated under hypoxic conditions with media at a pH of
5.5 and 6.5, respectively.

127

successional community soil (treatment 8) at the KBS LTER after incubation at room
temperature for ca. 30 days in an oxic or hypoxic (KBS 13) atmosphere. Strain KBS 146
was isolated previously from the conventional agricultural soil (treatment 1) on VL55-

R2B at pH 6.0 under a hypoxic atmosphere.

Colony and cellular morphology Colonies of all acidobacteria were small,
approximately 1 mm in diameter (after ca. 14-21 days of incubation), and had a circular
form with a convex elevation and undulate margin when grown on MHM-5 with glucose
under oxic conditions. Colonies of strains KBS 83 were smooth, glutinous white
colonies taking approximately two to three weeks to form visible colonies, whereas the
colonies of strain KBS 96 had smooth, butyrous pale yellow pigmented colonies taking
approximately three weeks to form a visible colony. Neither strain KBS 83 nor KBS 96
were pigmented like strains of the T erriglobus genus (16).

Cells of all strains are short, plump, Gram negative, nonmotile rods measuring ca.
0.5 pm x 0.4 pm (strain KBS 83) and 1 pm x 0.4 pm (strain KBS 96) when grown on
MHM-5 with glucose. Cells possessed exaggerated convoluted outer membranes when
viewed by TEM as is typical of a Grarn-negative type cell wall (Figure 4.2, panels A&B
(strain KBS 83), C&D (strain KBS 96)). Both strains produced an extracellular matrix of
as yet unknown chemical composition, but which apparently caused cells to stick
together tightly in colonies and form visible clumps in liquid culture similar to the
previously isolated strains in the Terriglobus genus (16). Strain KBS 83 typically

appears in clumps of four cells as seen under phase contrast and transmission electron

128

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Figure 4.2. Phase contrast and transmission electron micrograph of KBS 83 (panels A &
B) and KBS 96 (panels C & D). Arrow in panel A indicates the capsular material layer
produced by KBS 83. Scale bar indicates 1 pm (panels B & D) and 200 nm (panels A &
C).

129 .

microscopy (Figure 4.2, panels A&B), and are held together in this tetrad by capsular
material visible in electron micrograph (Figure 4.2, panel A). Capsules or extracellular
polysaccharides has been shown to promote bacterial adhesion (9), prevent desiccation
(39), and soil aggregation (1, 3), all of which would be beneficial for a soil

microorganism.

Capacity to utilize nitrate Strains KBS 83 and KBS 96 were tested for their ability to
use nitrate either via denitrification, nitrate reduction to ammonium, or nitrate reduction.
Under the conditions tested, strain KBS 83 does not appear to be capable of using nitrate.
After approximately 25 days of growth, there were no detectable amounts of nitrite or
higher concentrations of ammonium in the spent medium, and nitrate concentrations were
not different from the control (Table 4.1).

During the course of determining this strain’s ability to use nitrate, it was
discovered that strain KBS 83 was able to utilize ethylene. Growth under anoxic
conditions preliminarily appears to be a result of either ethylene or glucose utilization
(Table 4.1). This strain could not grow anaerobically in this medium without the
presence of ethylene in the headspace; however it could not grow solely on ethylene
under anaerobic conditions. It was interesting to note that growth of this strain under
these conditions was inconsistent, reaching a maximum optical density ranging from 0.02
to 0.102 at 600 nm; perhaps the inconsistent maximum optical density noted during
growth resulted from possible impurities of the ethylene gas. Ethylene is a plant
hormone used to promote fruit ripening of higher plants. This highly unusual substrate

has been seen previously for growth in Nitrosomonas europaea (19) as well as other soil

130

Table 4.1. Summary of growth and end products of KBS 83 & KBS 96 under anoxic

conditions with nitrate.

 

 

 

Strain KBS 83 Strain KBS 96
Concentrations (mM) Blank T25T Blank T13T
Carbon source
Glucose 9.7 i: 0.2 9.0 i 0.3 9.7 i 0.2 7.9 :t 0.8
Ethylene 5.3 i 0.3 2.6 :t 0.4 5.3 i 0.3 4.2 i 0.2
End products
N03' ca. 49.3 49.0 i 0.9 ca. 49.3 47.8 i 1.1
N02' ca.0 0:0 ca.0 1.1i0.1
NH4+ ca. 5.93 6.0 i 0.3 ca. 5.93 5 .5 i 0.2
N20 0 i 0 0 :t 0 0 :1: 0 0 i 0
C02 0 i 0 0.17 :1: 0.9 0 i 0 0.46 i 0.04

 

 

1 Indicates the number of days it took for culture to reach stationary phase.

131

microorganisms (l2), furthermore certain plants and microbes such as Pseudomonas
syringae (59) and Penicillium cyclopium (8) can produce ethylene at sufficiently high
concentrations for growth of other organisms.

Alternatively strain KBS 96 exhibited a substantially different pattern of grth
under these conditions where growth commenced after ca. 3 days instead of ca. 17 days
as seen with strain KBS 83. After approximately 13 days of growth, ca. 3% of the nitrate
was converted into nitrite and ca. 20% of the glucose was utilized (Table 4.1). The
ammonium concentrations were not substantially higher fiom the initial concentrations
and no nitrous oxide was detected in the headspace. Like strain KBS 83, it can also use
ethylene given the ca. 1.2-fold reduction of ethylene in the headspace; there was no
grth without the presence of ethylene in the headspace.

These results preliminarily suggest that nitrate was being reduced to nitrite while
glucose and/or ethylene were either being oxidized through a respiratory pathway or
glucose being oxidized by fermentation. Preliminary there was ca. 2-fold higher growth
rate with ethylene and glucose in the presence of nitrate as compared to the absence of
nitrate. Although very little of the nitrate was reduced to nitrite, it appears that there was
enough energy produced for grth from the partial reduction of nitrate to nitrite for
anaerobic respiration (for both glucose and ethylene) based on fiee energy (AG)
calculations even with scenarios where high free energy products were produced. Since
nitrate was in excess, the low optical density readings of the strain were presumed to be
of result a build-up of an inhibitory compound(s) resulting from growth or limiting
concentration of a cofactor such as molybdenum which is necessary for the

metalloprotein, nitrate reductase.

132

Strain KBS 96 preliminarily appears to be capable of reducing nitrate to nitrite
under anoxic conditions, but not is able to denitrify. This trait is fairly ubiquitous and is
the most widely distributed variant of using nitrogenous oxides for respiratory purposes
(64). Perhaps this ability could provide a selective advantage allowing them to survive
during rainfall events, creating anoxic pockets in the soil especially in fertilized fields
high in nitrate. Additionally, this physiology could be used to develop novel cultivation
methods to isolate additional strains in this poorly represented subdivision. Readily
available strains of subdivision 1, T erriglobus roseus strain KBS 63, T erriglobus sp.
strain TAA 43 (16), and Acidobacteriaceae strain KBS 146 were not able to grow under
anoxic conditions with nitrate and glucose as the carbon source. Strain KBS 96 was
initially cultivated under aerobic conditions, however it was further isolated in pure
culture on the MHM-5 with 10 mM glucose under anaerobic conditions, illustrating its
capacity to form a colony on a plate under these nitrate reducing conditions, typically

taking about three weeks.

Other physiological properties The characteristics of the new strains are
smnmarized in Table 4.2. Similar to the T erriglobus strains (16), KBS 83 and KBS 96
preferred slightly acidic pH conditions. KBS 83 grew at pHs between 4.5 and 6, growing
optimally at 5.0, whereas KBS 96 grew between pHs 4.0 and 6, optimally growing at 6.0
which is consistent with the pH of their native environment. As suggested previously,
members of subdivision 1 have a preference for mildly acidic pH conditions (16, 43).
Preliminary these data suggests that members of subdivision 3 might also follow this

trend given the growth of strain KBS 96 under mildly acidic conditions.

133

Table 4.2. Phenotypic characteristics of Terriglobus, Acidobacteriaceae, and
Acidobacterium strains.

134

 

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Strain KBS 83 grew significantly faster at oxygen concentrations between 4% to
16% (avg. p-value = 0.01) as compared to atmospheric concentrations of oxygen,
growing optimally at 8% oxygen (u = 0.014 hr‘l) (Figure 4.3), suggesting that it was
microaerophilic. Microaerophiles are defined as organisms not capable of growing or
growing poorly under atmospheric concentrations of oxygen presumably due to an
increased sensitivity to toxic forms of oxygen such as H202, 02', and OH-(27). Although
strain KBS 83 was catalase positive, suggesting that cells have the capacity to deal with
peroxides, this enzyme’s effectiveness might be low, similar to certain Campylobacter
species classically described as microaerophiles (41). The preference of this strain for
low levels of oxygen is consistent with their native environment; oxygen can be depleted
in the soil enviromnent, particularly in soil aggregates when soil moisture increases (6,
35, 58). Strain KBS 96 qualitatively grew better under atmospheric concentrations of
oxygen; this strain’s growth requirements demanded the use of a undefined medium
which made optimal growth testing under different oxygen concentrations difficult since
the concentration of oxygen was not in excess relative to the unknown amount of carbon.
Nevertheless, it was clear the strain KBS 96 preferred more oxygen rich environments.

Characterization data from recently isolated strains of the Acidobacteria (16)
along with data obtained from cultivation experiments using PWPCR suggest that some
acidobacteria are microaerophiles. The inclusion of the catalase enzyme as well as
incubation under microoxic conditions increased with frequency of acidobacteria
detection by ca. 3.3 and 1.3-fold, respectively on initial cultivation experiments (Chapter

2). The newly isolated strain KBS 83 grows optimally at low concentrations of oxygen.

137

0.014 ~ .

0.013 ~

0.012 _.

0.011 -

0.010 -

Growth rate (hr'1)+/- standard error

0.009 -

 

 

 

4% 8% 16% 21%
Oxygen concentrations

0% 2%

Figure 4.3. Summary of growth rates for KBS 83 under different concentrations of
oxygen. Note growth rate for an oxygen concentration of 0% was obtained fiom studies
assessing the capacity to use nitrate.

138

Although not following the classic definition of microaerophily, members of the recently
described genus T erriglobus produce a carotenoid(s) whose quantity per cell increased in
response to oxygen, suggestive of an oxidative stress response (16). These data taken
together suggest that oxygen and its damaging effects is a major concern to the lifestyle
of acidobacteria, and strains harbor traits to respond accordingly.

Relatively slow growth rate in acidobacteria strains is becoming a trend with
recent cultivars which is consistent with their low number of genes coding for the 16S
rRNA gene. These new strains contain either one (KBS 83) or two (KBS 96) copies of
the rrs gene (Figure 4.4). Additionally, recently sequenced genomes of Acidobacterium
str. Ellin 345 and Solibacter usitatus strain Ellin 6076 reveal that both harbor one copy of

the rrs gene ( wvnvj gidoe. gov), and in summary no strain in the phylum Acidobacteria

 

to date contains an rrs copy number greater than 2. This is additional support for
previous claims suggesting that members of the phylum Acidobacteria can be described
as oligotrophic given their high abundance in soil environments where nutrients are low
(17), relatively slow growth rate, and possession of either one or two copies of the 16S
rRNA encoding gene — a genomic marker of oligotrophic bacteria (25).

There were major differences in the carbon utilization profiles of these new
strains as compared to previously isolated strains in T erriglobus and Acidobacteriaceae
(16). Strain KBS 83 and KBS 96 had the capacity to oxidize more native soil carbon
such as xylan, cellulose, carboxymethyl cellulose, syringate, pectin, trimethoxybenzoate,
and ferulate (Table 4.2), which are typically subject to slow turnover in the soil
environment (6). I propose that the use of these more recalcitrant carbon compounds was

responsible for its successful isolation of strain KBS 83, presumably because the use of

139 '

 

9 KBS 83 KBS96 9
kb 2 I I fl 2
__P. 1 2 3 4 5 6 7 8
23.] — Q m ..
9.4 — - -
6.6 — C "" -
2.3 — u- .- -

Figure 4.4. Southern hybridization of digested genomic DNA from strains KBS 83 &
KBS 96. Strains were cut with EcoRI (lanes 2 & 5), EcoRVI (lane 3 & 6), and HindIII
(lane 4 & 7). The lambda DNA Hind III marker (lane 1 & 8) provided size estimates.

140

these compounds prevented substrate-accelerated death (37, 53) since these polymers
must be hydrolysed before used as a source of carbon (7, 30) as suggested to be
responsible for the increased viable counts of soil bacteria from Australian soil previously
(44).

Plants in terrestrial ecosystems contain 560 Pg of the total carbon content on the
planet (61). Plant tissues are the dominant source of soil polysaccharides, specifically
cellulose, hemicellulose, and lignin (55). Depending on the type of plant, xylans can
comprise from 7% to 35% (dry weight of the plant) (60), and they are typically oxidized
by nearby microbial communities. Although the soil environment is referred to as carbon
limiting, plant polymers and products of the degradation are major sources of carbon for
indigenous populations of microorganisms. Microorganisms that harbor active enzymes
to degrade these plant polymers are essential for maintaining an active carbon flow in the
soil.

The ability of these acidobacteria to oxidize more recalcitrant carbon compounds
is a practical trait to have in the soil environment. One could predict that strains capable
of oxidizing plant polymers might be prevalent-in environments where carbon pools are
low such as in the conventional agriculture soil (treatment 1) where carbon
concentrations primarily consisting of degradation of plant material from corn, soybean,
and wheat that is plowed into the soil annually and are ca. 2 to 3-fold lower as compared
to the successional treatment (treatment 8). In order to test this prediction, acidobacteria
16S rRNA gene sequences from treatment 8 and l soils were analyzed, specifically

assessing if there was a higher prevalence of sequences from treatment 1 in the clade

141

harboring the acidobacteria strain capable of oxidizing these more recalcitrant carbon
sources based on the assumption that phylogeny follows function. Figure 4.5 depicts a
phylogenetic tree of this clade along with the number of acidobacteria sequences from
treatments 8 and 1. Approximately half of the clones in this clade belonged to treatment
1, suggesting that this trait is not unique to acidobacteria from low carbon environments
and is fairly ubiquitous in both conventional agriculture and mid-successional community

soils.

l6S rRNA gene phylogeny Figure 4.6 depicts a phylogenetic tree of nearly full length
sequences for the new strains. Strains KBS 13, 5, and 3 are ca. 97% similar to T. roseus,
strain KBS 63, therefore preliminary appears to be members of the T erriglobus genus
(16). The closest cultivar to strain KBS 146 is strain TAA 166 with a sequence similarity
of ca. 99%.

Strain KBS 83 is more closely related to A. capsulatum (ca. 96% sequence
similarity) than to members of T erriglobus and is most similar (ca. 99%) to an
environmental clone from a polychlorinated. biphenyl-polluted soil (env. WD277,
AJ292577) (34). The strain belonging to subdivision 3, KBS 96 is ca. 94% similar to an
environmental clone from a reaction system treating monochlorobenzene contaminated
groundwater (env. GOUT8, AY050595) (2), and is only ca. 93% similar to Solibacter
usitatus strain Ellin 6076.

Strain KBS 83 and KBS 96 are distinct from previously isolated acidobacteria to
call for the creation of a new genus and species in subdivisions 1 and 3, respectively.

Strain KBS 83 is a member of subdivision 1 and only 96% similar to A. capsulatum, and

142

  
  
  
  

 

Acbt.capsl(DZ6l7l: 4,,

K8883 Q . l

env.WDZ77 A1292577 2 3
O

Ellin5225 (AY2345'Q6) «- l

.ElhnSO 7 AY23 34) ° «

Ellin5237(A _234588) .. ... g 2
— E

. . ‘.=. —.- a: 3 l

7‘ 1

0.10 q

- 4

O

 

 

 

Figure 4.5. Neighbor-joining phylogenetic tree depicting the number of 16S rRNA gene
clone sequences generated from treatments 1 and 8 (black and gray boxes, respectively)
at the KBS LTER in the clade harboring strain KBS 83.

143

 

 

 

 

ll__.||._._.lL__.l
Ul 8 a

fi env.SHA18 (M249099)
env.SJAl49 (M009495)
Solibacter usitatus, strain Ellin6076 (AY234728)
env.WD205 (A129ZS7 1)
env.AzCOOZ (AF013515) 3
env.WD254 (M292582)
K8896
env.OOUTBS (AY050595)
env.OPB3 (AF 027004)
Acbt.capsl (D261 71 )
K8883
env.WD277 (M292577)
Ellin5017 (AY234434)
Ellin5225 (AY234576)
6 Ellin5237 (AY234588)
Ellin337 (AF498719)
KBSl46 .
TAA166 (AY587230)
Ellin351 (AF498733)
K8889 (AY587227) l
env.TM2 (X97098)
K8813
K883
K885 .
Terriglobus sp. strain TAA43 (AY587228)
Terriglobus sp. strain TAA48 (AY587229)
Terriglobus roseus, strain K88112 (DQ660895)
Terriglobus roseus, strain K8868 (DQ660894)
Terriglobus roseus, strain KBS62 (DQ660893)
Terriglobus roseus, strain K8863 (DQ660892)

 

  
   
   
  

 

 

 

 

   
  

 

 

 

 

0.10

 

Figure 4.6. Maximum-likelihood tree of the Acidobacteria subdivisions 1, 3, 4, 5, and 6
(indicated to the right of the group) based on 16S rRNA gene using sequences obtained
from cultivated representatives and environmental clones. Geothrix fermentans and
Holophaga foetida of subdivision 8 were used as an outgroup (not shown). Strains from
this study are in bold font. Each gray trapezoid represents a different subdivision with
approximately 5 representative sequences. Intemal nodes supported by a bootstrap value
of >95% are indicated with a filled circle ( o ) and of >70 with an open circle ( o ). The
scale bar indicates 0.10 changes per nucleotide.

144

94% similar to the T erriglobus strains. Unlike A. capsulatum, this strain cannot grow at
low pH (pH 3.0-6.0) conditions, it is not motile, and it was isolated from soil, not an
acidic mine drainage. Given the low sequence similarity, it does not appear to be
members of T erriglobus. Strain KBS 96 is a member of subdivision 3 and only ca. 93%
similar to previously isolated strain Solibacter usitatus, strain Ellin 6076. The specific
epithets to strains KBS 83 and KBS 96 could not be determined for my dissertation, but

will be given in the publication of this work.

Description of new genus and species in subdivisions 1 Gram negative,
nonmotile capsulated short rods characteristically in groups of four (ca. 0.5 x 0.4 pm)
when grown on MHM-5 with 10 mM glucose. Moderate acidophile with an optimal pH
of 5. Colonies are non-pigmented, taking approximately two to three weeks to form a
colony. Catalase positive; oxidase negative; one copy of the rrs; isolated from a
conventional agriculture treatment at the K88 LTER; able to grow at 12°C, 25°C and
37°C, but not 4°C; G+C content ca. 62 mol%; microaerophilic growing optimally at
oxygen concentrations between 4 to 16%; and able to oxidize more recalcitrant carbon
sources such as xylan, syringate, pectin, methyl cellulose, ferulate, and cellulose, in

addition to various mono-, di- and trisaccharides. Type strain KBS 83.

Description of new genus and species in subdivisions 3 Gram negative,
nonmotile short rods (ca. 1.0 x 0.5 pm) when grown on MHM-5 with 10 mM glucose.
Moderate acidophile with an optimal pH of 6. Colonies are pale yellow, taking

approximately three weeks to form a colony. Catalase positive; oxidase positive; two

145

copies of the rrs; isolated from a conventional agriculture treatment at the KBS LTER;
able to grow at 12°C, 25°C, and 37°C, but not 4°C; G+C content ca. 60 mol%; reduces
nitrate to nitrite; can growth at low concentrations of oxygen, but prefers atmospheric
concentrations; and able to oxidize more recalcitrant carbon sources such as xylan,
syringate, pectin, trimethylbenzoate, methyl cellulose, ferulate, and cellulose in addition
to various mono-, di- and trisaccharides. Type strain KBS 96.

These strains will be deposited into the USDA Agricultural Research Service
Culture Collection and the Deutsche Sammlung von Mikroorganismen und Zellkulturen

GmbH.

ACKNOWLEDGEMENTS
This work was funded by the USDA (#2001-35107—09939), NSF (#MCB-
0135880), and the NSF Long-Term Ecological Research Program at the Kellogg
Biological Station and the Michigan Agricultural Experiment Station.
I would like to thank Dr. Jon Dahl at the Michigan State University Soil Testing

Laboratory for his assistance with analyzing nitrate, nitrite, and ammonium.

146

10.

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153

CHAPTER 5

THE EXPLORATION OF METHANOTROPHY AND METHYLOTROPHY

AMONG MEMBERS OF THE PHYLUM ACIDOBAC T ERLA

INTRODUCTION

Methane is a plentiful gas in the soil environment. Forests, grasslands, and other
unsaturated soils are major sinks for atmospheric methane (11). Microorganisms are
responsible for methane oxidization in these environments, consuming 20 to 60 Tg of
methane per year (11). Unfortunately, these communities are largely unexplored as
evident by the discrepancy between the rates of known methane oxidizers and soils’
consumption of methane (8, 24).

Microorganisms that are capable of growth on one-carbon compounds such as
methanol, methylated amines, halomethanes, and sulfur-containing methylated
compounds are referred to as methylotrophs, while organisms capable of growing on
methane are designated as methanotrophs. These compounds serve as the
microorganism’s sole source of carbon and energy (11) and this unique group of
microbes are found in soils and sediments throughout the globe (12, 13, 24, 25, 34).
Notably, this unique physiology is not limited to bacteria; it has also been observed in
yeast as well as filamentous fungi (11).

Oxidization of these one-carbon compounds is a simple, yet elegant pathway
converting single carbon compounds into complex, multi-carbon cellular material.

Methane is oxidized to methanol by the methane monooxygenase enzyme and is

154

performed by two phylogenetically distinct groups of methane oxidizers, Type I and II,
which differ by their assimilation pathway and the type of methane monooxygenase
enzyme. Type I methanotrophs are members of the y-Proteobacteria phylum, oxidizing
methane with a membrane-bound or particulate methane monooxygenase (pMMO), and
assimilating carbon into cellular material with the RuMP pathway. Type II
methanotrophs are members of the a-Proteobacteria phylum oxidizing methane with a
cytoplasmic soluble methane monooxygenase (sMMO) and assimilating carbon using the
serine pathway (11). Methanol, resulting from the oxidization of methane or exogenous
sources such as pectin or lignin degradation, is further oxidized to formaldehyde with the
highly conserved periplasmic enzyme, methanol dehydrogenase (MDH) (11, 23).
Remarkably, the presence of methanol promotes the oxidization of methane in pure
cultures and soil enrichments (2). Formaldehyde is then shuttled to one of three
assimilation pathways, ribulose monophosphate (RuMP), serine, or ribulose-1,5-
bisphosphate carboxylase (RuBP), to create multi-carbon compounds for the generation
of cellular material (Figure 5.1) (11).

Previous studies have implicated members of the phylum Acidobacteria with the
capacity to oxidize one-carbon compounds. Radajewski and colleagues (26), after
incubating oak forest soil samples with l3C-methanol for a period of 40 days, revealed
that acidobacterial DNA was present in the heavy fraction of a cesium chloride gradient
(Figure 5.2) (26-28). Heavy-labeled acidobacterial DNA suggests these acidobacteria
used the methanol as a substrate for growth thus incorporating the 13 C into their DNA.

The ubiquity of acidobacteria in the soil along with the scarcity of knowledge correlating

155

Methylated

Methylated sulfur

amines\ / compounds

CH4 —> CH3OH —> HCHO —> HCOOH —> C02

Methane Methanol

Monooxygenase Dehydrogen17 \

 

Type I Methanotrophs
RuMP Pathway

 

 

 

 

 

Type II Methanotrophs
Serine Pathway

l

 

 

 

RuBP
{Pathway

 

\ //

Cell Material

Figure 5.1. Simplified overview of the pathway for the oxidation and assimilation
of one-carbon compounds into cellular material using the RuMP, RuBP, or serine

pathways.

156

a-subclass Proteobacteria

 

 

 

 

 

 

Hyphomicrobium vulgare
Methylobacterium extorquens Methylosinus trichosporium
Paracoccus denitrificans RhOgOpseudomonas
A _ h . . acrdophrla
_ crdomonas met anohca UP2 ( 47%)
Beyerinckia indica
UP1 (49%)
- UP3 (1 96)
Methylomonas methamca eu b a ct eriu m K
Methylococcus
cap sulatus Thermotoga maritima
Methylophilus
methylotrophus _
‘ Holophaga foetida
Arthrobacter globiformis
- - Unidentified
Bacrllus methanolrcus eu bacterium RBO4
Unidentified
eubacterium R816 UA3 (1%)
(1 %) ‘ Acidobacterium capsulatum
”A1 UAZ (1%)
1 0%

Acidobacterium division

 

Figure 5 .2. Acidobacteria clones obtained after incubation of acidic oak forest soil
samples with 350 mM methanol (26).

157

upland soils’ methane fluxes to known, cultivated microorganisms could be two pieces of
the complex soil puzzle, which if fit together, can greatly improve our understanding of
these bacteria in the soil. The goal of this study is to assess the metabolic potential of
acidobacteria to oxide one-carbon compound(s) using growth and molecular-based

approaches.

MATERIALS and METHODS

Pure culture approach

Media composition and growth conditions Strains were tested for the capacity
to oxidize one-carbon compounds on VSB-7 as described previously (Chapter 3) as well
as on a nitrate mineral salts medium (pH 7). Nitrate mineral salts, Whittenbury medium
(NMS) contained the following components (per liter): MgSO4, 0.49g; KNO3, 0.1g;
CaClz-HzO, 0.2g; Whittenbury trace element solution (lmL/L); NazMoO4, 0.0005g;
FeEDTA, 0.0038g; CuSO4, 0.8mg; phosphate stock solution (lOmL); and vitamin stock
solution (10mL). The Whittenbury trace elements solution contained the following (per
liter): FeSO4-7HZO, 0.5g; ZnSO4-7HzO, 0.4g; MnC12-4HZO, 0.02g; CoClz-6HzO, 0.05g;
NiClz-6H20, 0.01g; H3BO3, 0.015g; and NaEDTA, 0.25g. The phosphate stock solution
contained the following (per liter): KHzPO4, 26g, and NazHPO4, 33g. The vitamin stock
solution contains the following (per liter): biotin, 2mg; folic acid, 2mg; thiamin-HCI,
5mg; calcium pantothenate, 5mg; vitamin B12, 0.1mg; riboflavin, 5mg; and
nicotinamide, 5mg. For solid medium, agar was added to a final concentration of 1.5%

(g/vol).

158

Carbon Utilization Testing All strains were grown on replicate agar plates of
VSB-7 and NMS with 10 mM of one of the following one-carbon compounds: methanol;
forrnate; methylated amines containing dimethylamine (2.8 mM), trimethylamine (1.8
mM), and methylamine (5.4 mM); and dimethylsulfoxide. Plates were incubated at room
temperature (ca. 23°C), under low light for a period of 40-60 days, under one of the
following atmospheres: CO2-enriched hypoxia (vol/vol: 2% O2, 5% CO2, and bal N2);
CO2-enriched air (vol/vol: 95% air, 5% CO2), or air. Liquid cultures of the NMS medium
were grown with an oxic headspace supplemented with methane (3 0% vol/vol).

Strains KBS 63, TAA 43, and A. capsulatum were also grown on the VSB-6 with
50 mM and 100 mM methanol in 750 ml sealed, side-arm flasks. Some of these cultures
were supplemented with a combination of 2.5 mM glucose and 1 mM formaldehyde.
Optical density at 600 nm was monitored every 5 to 10 hours with a Thermo Spectronic
model 20D+ spectrophotometer. The headspace was monitored for a period of 30 days
using an Aerograph flame ionization gas chromatograph series 2440 (V arian Instrument
Division, Palo Alto, California) with a 5 foot long, 1/8th inch stainless column with

Porapak Q packing material.

PCR amplification and cloning DNA was extracted from the pure cultures using the
MoBio Fecal DNA Extraction Kit (MoBio Labs. Inc., Carlsbad, California). Genomic
DNA was screened for the presence of the methanol dehydrogenase gene (mxaf) using
primers mxa f1003 and ma r1561 (23), and the different forms of methane
monoxygenase using A189/A682 (active site for pMMO) and ammonium

monooxygenase) (14), A189/mb661 (active subunit for pMMO) (4), mmox536f/898r (9)

159

and mmox206f/886r (15) (active subunit for sMMO), mmox206F/886R (active subunit
for sMMO) (15). Purified genomic DNA from M. capsulatum (ATCC Number 19069-
D) for was used as a positive control. Each 25 ul PCR reaction contained 1x PCR Buffer,
1mM magnesium choloride, 0.02mM of each dNTP, 0.2uM each primer, and 5U Taq
polymerase (Invitrogen, Carlsbad, California). The PCR program consisted of the
following steps: (1) 95°C for 3 minutes, (2) 95°C for 30 seconds, specific annealing
temperature for 30 seconds, 72°C for 45 seconds (step 2, repeated for 30 cycles), (3) 72°C
for 10 minutes, and (4) hold at 4°C. In addition to these reactions, a PCR amplification of
the 16S rRNA gene was performed to ensure the genomic DNA from the strains was
amplifiable. Purified genomic DNA from A. Capsulatum (ATCC Number 51196) was
used as a positive control. PCR reactions were electrophoresed on a 1% agarose gel and
stained with GelStar Nucleic Acid Stain (BioWhittaker Molecular Applications,
Rockland, MA). If reactions yielded a positive amplification of the correct size, PCR
products were cloned into the Invitrogen TOPO TA Cloning Kit for Sequencing
(Invitrogen Carlsbad, California) as per the protocol instructed. The PCR product was
then re-amplified using the following primers: modified-M13F and modified-M13R (19)
with the PCR conditions as described above. PCR products were purified for sequencing
using a 1:8 dilution of ExoSAP-IT (U SB, Cleveland, Ohio) following incubation at 37°C
for 30 minutes and 80°C for 15 minutes, and reactions were sent to the Michigan State
University Genomics Technology Support Facility (RTSF)

(http://genomics.msu.edu/index.html).

 

Environmental molecular-based approach

160

Soil microcosms Soil samples were collected July 10‘“, 2001 from the Michigan
State University WK. Kellogg Biological Station Long Term Ecological Research Site
(KBS LTER). The KBS site is a 48-hectare plot designed to study ecological processes
in agroecosystems, which prior to its establishment in 1989 was uniformly farmed for
more than 50 years. A description of this site can be accessed at

http://\va.kbs.msu.edu. Soil cores (2.5 cm diameter x 10 cm depth) were collected

 

from a mid-successional community with no history of tillage (treatment 8), and stored at
4°C for approximately two weeks prior to establishing the microcosms. Treatment 8 soil
was used for this microcosm because preliminary clone analysis of acidobacterial 16S
rRNA genes revealed a high sequence similarity to Radjewski’s and colleagues (26)
clone sequences.

The soil was homogenized in a sterile 500 mL beaker by gently mixing with a
sterile spatula. Replicate, sealed microcosms consisted of approximately 10 grams in a
125 mL crimp-top vial supplemented with 336 umoles of methanol. The remaining
homogenized soil was frozen at -20°C and later used to extract genomic soil DNA for
time zero in the clone library analysis (below). The water holding capacity was adjusted
to ca. 35% with sterile distilled water (20). The microcosms were incubated at room
temperature (ca. 23°C) under low light conditions.

The headspace was monitored routinely for methanol using the Aerograph flame
ionization gas chromatograph series 2440 (Varian Instrument Division, Palo Alto, Calif).
The headspace was flushed every other week to maintain an oxic atmosphere. Once the
initial addition of methanol was consumed, methanol was re-added at the same initial

concentration.

161

Enrichment plating Upon the consumption of the second addition of methanol, a
portion of the soil was used for DNA extraction (see below) and the remaining soil was
used as a source of inoculum in an attempt to cultivate potential methylotrophic
acidobacteria using a water agar based medium with 25 mM, 50 mM, and 490 mM of
methanol. Plates were incubated at room temperature (ca. 23°C) under CO2-enriched air

for ca. 30 days. Plates were screened for the presences of acidobacteria using PWPCR

(33).

Methanol microcosm clone libraries DNA was extracted using the MoBio Fecal
DNA Extraction Kit (MoBio Labs. Inc., Carlsbad, California) from soil before methanol
addition (T0) and after the consumption of the‘second addition of methanol (T90). T0
and T90 soil DNA was used to amplify the acidobacteria 16S rRNA gene using the
phylum-specific Acidobacteria forward primer, ACD31F (1), and the broadly inclusive
bacterial primer, 1492R (21). Amplicons were cloned into the Invitrogen TOPO TA
vector and prepared for sequencing with conditions described above, and submitted with
the 531R (5’ TAC CGC GGC TGC TGG CAC 3’) primer. Sequences were aligned using
ARB Software (22) along with other sequences were downloaded from Genbank and the
Ribosomal Database Project (RDP) (3) for the generation of the phylogenetic tree. I-
LIBSHUFF was used to determine the similarities of the libraries before and afier

methanol additions (31, 32).

RESULTS

162

Pure culture approach

Strain similarity to acidic forest soil clones Clone sequences of the 16S rRNA
genes obtained from acidic oak forest soil microcosms incubated with methanol (UAl,
UA2, and UA3) (26) had a high sequence similarity to acidobacteria clones obtained
from KBS LTER soil. KBS Acido Clone 29 was 98.8% similar to UA3; KBS Acido
Clone 25 was 96.4% similar to UAl; and KBS Acido Clone 31 was 95% similar to UA2
(Figure 5.3). All clone sequences were associated with subdivision 1 of the phylum

Acidobacteria.

Carbon ultization testing Strains, KBS 62, KBS 63, KBS 68, KBS 112, KBS 89,
TAA 43, TAA 48, and TAA 166 were tested for their ability to oxidize different one-
carbon compounds. These newly isolated strains had a sequence similarity to the putative
methanol-oxidizing clones (26) of ca. 91% ranging from 88 to 94%. No growth was
detected on the plates containing neither one-carbon compounds nor liquid cultures with
30% (vol/vol) methane in the headspace under these conditions. Positive controls of
VSB-7 and NMS with glucose and glucose with one-carbon compound(s) resulted in
growth indicating that the conditions were ample for growth of the strains. Neither
growth nor depletion of methanol in the headspace was detected for KBS 63, TAA 43, or
A. capsulatum in liquid cultures with 50 mM or 100 mM methanol with and without
supplementation of 1 mM formaldehyde. Previous studies indicated that the presence of
formaldehyde acts as an important trigger for the expression of one-carbon compound
genes (6); therefore these cultures were supplemented with 1 mM formaldehyde.

Controls containing glucose I and methanol (

163

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164

with and without 1 mM formaldehyde) exhibited no difference in normal growth patterns,
suggesting that methanol was not inhibitory to the any of the strains. Under all
conditions tested, there was no evidence for these strains harboring the capacity to

oxidize these one-carbon compounds.

PCR amplifaction of ma f and mm _ Purified genomic DNA of these
acidobacteria stains did not yield positive amplification for the methanol dehydrogenase
nor methane monooxygenase gene with readily available primer sets, even though
positive controls yielded amplicons of the correct size. This result was not surprising
since these primers were designed from the Proteobacteria; members of the phylum
Acidobacteria could have a methane monooxygenase and/or methanol dehydrogenase
gene, but these primer sets may not have amplified their genes, most likely due to the

proteobacteria-specific primers not allowing any degeneracy for other diverse phyla.

Environmental molecular-based approach

Soil microcosm Since the KBS acidobacteria clones were similar in sequence to the
putative methylotrophic Radajewski and colleagues’ clones (26), methanol was used as
an enrichment with soil from the KBS LTER. A steady decrease in methanol was
observed for 90 days (Figure 5.4). Abiotic controls consisting of sterile soil
supplemented with 338 umoles of methanol“ had significantly higher (p < 0.006)
concentrations of methanol compared to experimental replicates after 90 days,
demonstrating that the depletion of methanol was due to a biological component, rather

than abiotic degradation. Upon depletion of the second addition of methanol, the soil was

165 .

 

E <-* -—- ———-— methanol re-addition

- ‘__~___,;. - ——- methanol re-addition

 

 

E
V i Q
r: 250 A :l - '
.2 . " I
E; I: . I
“g, 200 A d T
é‘) ‘ ' - o
.5 150 — H L __
g ' m Ii ‘
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a) 100 - 6
2 . E ‘
' 3
50 - :
. l l L
0 I l ”I I I I - l ‘7 I
0 20 40 ' 60 80 100

Days of Incubation

Figure 5.4. Methanol consumption by KBS LTER replicate soil microcosms, replicate l
(gray boxes) and replicate 2 (black boxes). Each point represents the mean i standard
deviation from triplicate measurements of methanol in the headspace as measured by GC-

FID.

166 -

used as a source of inoculum to isolate potential, methylotrophic acidobacteria. After
thirty days of growth, the water agar treatment supplemented with 25 mM methanol
yielded a positive amplification for acidobacteria (Figure 5.5, Sequence ID JAB 971)
using PWPCR (33). Attempts to isolate acidobacteria from replicate plates were

unsuccessful.

Phylogenetic analysis (I - LIBSHUFF) Acidobacteria clone libraries were made
from microcosm soil before (TO) and after methanol addition (T90) to determine if
acidobacteria responded to methanol additions. The composition of acidobacteria clone
libraries following methanol incubation significantly differed from acidobacteria clone
libraries before methanol incubation (p < 0.001). Furthermore, methanol enriched for a
phylogenetically cohesive group of Acidobacteria in subdivision 1 (Figure 5.6). It was

this subdivision where the clones from the Radajewski and colleagues’ study (26) reside.

DISCUSSION and FUTURE DIRECTION
The goal of this study is to assess the capacity of acidobacteria to oxidize one-
carbon compounds using growth and molecular-based approaches. At this time, there is
no evidence to suggest that these newly isolated strains have the capacity to oxidize one-
carbon compounds under the conditions tested. Nevertheless it is interesting to note that
during the soil microcosm experiment there was a shift in the acidobacterial community

after methanol consumption. Methanol clearly had an affect on the acidobacteria

167

Acbt.capsl (D26l7l)
l E env.WD277 (AJZ92577)
Ellin5017 (AY234434)
Ellin5225 (AY234576)
Ellin351 (AF498733)
Ellin337 (AF498719)
TAA166 (AY587230)

env.TM2 (X97098)

 

 

JAB97l

 

KBSB9 (AY587227)
TAA43 (AY587228)
TAA48 (AY587229)
KBsnz (DQ660895)
KBS62 (DQ660893)
KBS63 (DQ660892)

J KBS68 (DQ660894)

 

 

 

 

 

l8 / subdivision 3
i7 / subdivision 5

8 / subdivision 4
——‘l 8 / subdivision 6

 

 

 

 

 

 

 

0.10

Figure 5.5. Neighbor-joining tree of the Acidobacteria subdivisions 1, 3, 4, 5, and 6
based on 16S rRNA gene using sequences obtained from cultivated representatives and
environmental clones. Geothrixfermentans and Holophagafoetida of subdivision 8 were
used as the outgroup (not shown). Strains from this study are in bolded text. JAB 971 in
subdivision 1 is a partial 16S rRNA gene sequence from cultivation experiments using
soil from the methanol microcosms as a source of inoculum. The scale bar indicates 0.10
changes per nucleotide.

168

Figure 5.6. Neighbor-joining tree of partial sequences of 16S rRNA gene clones in the
phylum Acidobacteria obtained before and after incubation of KBS LTER treatment 8
soil with 350 mM methanol. Acidobacteria] clones before and after the addition of
methanol are represented in light gray and dark gray boxes, respectively. White boxes
with black outlines represent Acidobacteria l6S rRNA gene clones obtained from oak
forest soil incubated with 13C-methanol, previously (26).

169 i

 

No. of clones
b)
o
l

 

 

 

 

 

 

 

Before Afier

 

 

 

 

No. of clones
b.’

1
10 j ’ . 69.-s! -
I

“.3.“ ., '_ .
‘ . .

 

 

 

 

 

 

Before ' Afier

 

 

No. of clones
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Before After

Before methanol addition

I Afier methanol addition
[I l3C-methanol microcosm (Nature, Vol. 403, No. 6770, 646-649)

 

 

170

community, significantly selecting for members of subdivision 1; notably, the
acidobacterial subdivision with the most success in cultivating to date (5, 7, 10, 16-18,
29, 30, 33) and harboring the putative, methanol-oxidizing clones (26). It is unclear if
this was a result of acidobacteria oxidizing methanol or an indirect affect, such as a
reduction in the nearby, competing communities thereby allowing the acidobacterial
community to flourish. Alternatively, the acidobacterial community could have been
interacting with or cross-feeding with the nearby methylotrophic community(s). Perhaps
acidobacteria are capable of methylotrophy, but only under a symbiosis with nearby
community(s) of known methylotrophs due to an auxotrophic requirement of some
metabolite which can only be obtained through cross-feeding of nutrients.

Interestingly, during the course of these studies alternative explanations were
realized about Radjewski and colleagues’ claims of methanol oxidization in
acidobacteria. At the time of the paper, stable isotope probing was a new technique and
since then, scientists have revealed potential pitfalls of this technique that were not
addressed at the time of this publication. The possibility exists that the DNA from the so-
called heavy fraction might not have been heavy at all; rather the DNA could have been
G + C rich 12C-DNA. In stable isotope probing, DNA is separated based on its buoyant
density in a cesium chloride gradient; if the non-heavy, 12C DNA has a high proportion of
G + C, it would settle around the area considered to be the heavy, 13C DNA, thereby
introducing unlabeled DNA in the presumed heavy fraction. This scenario is illustrated
in Figure 5.7 (area of ambiguity) where non-labeled, 12C DNA having a G + C content of
ca. 62% has the same buoyant density at labeled, '3 C -DNA with a G + C content of ca.

20%. Recently isolated strains of subdivision 1 in the phylum Acidobacteria have a G +

171 i

1.78
1.77
1.76
1.75
1.74
1.73
1.72
1.71
1.70
1.69
1.68

1.67 T ' l ‘ I 1 l ‘ l ' 1 ' T
20 30 4O _50 60 7O 80

G + C content (%)

 

Buoyant density (g/mL)

 

 

 

 

lllJLlllllllllllllillJ

 

 

Figure 5. 7. G + C content of '2C- and ”0 DNA 1n relation to its buoyant density on a
cesium chloride gradient.

172

C content (%) ranging from 55 to 62% (Chapters 3 & 4), values within the area of
ambiguity (Figure 5.7). In order to address this concern, researchers typically have a
replicate microcosm with the non-labeled substrate, and a subtractive analysis is
performed to truly determine the active community(s) oxidizing the tested substrate.
Another possible explanation of Radajewski and colleagues’ (26) results could be that the
acidobacterial DNA became heavily-labeled not through the oxidization of the methanol,
but by the acidobacteria cross-feeding on the byproducts of methanol oxidization of the
nearby methylotrophic communities.

Fortunately, there is still hope to address this interesting physiology in the phylum
Acidobacteria with the advent of genome analysis. There have been three, recently
sequenced genomes of members of the phylum Acidobacteria: Acidobacterium
capsulatum (subdivision 1), Acidobacterium sp. strain Ellin345 (subdivision 1), and
Solibacter usitatus strain Ellin6076 (formerly, Acidobacteria-3 sp. Ellin6076)
(subdivision 3). The recently annotated Acidobacterium sp. strain Ellin345 and
Solibacter usitatus strain Ellin6076 genomes preliminary indicates the presence of a
methanol dehydrogenase gene as well as the ability to oxidize other one-carbon
compounds such as methane, methylamine, dimethylamine, trimethylamine,

formaldehyde, and forrnate (httpz/Ammnigidoegovn. These data have provided more

 

compelling evidence that members of the Acidobacteria phylum may have the potential
to oxidize one-carbon compounds. The future direction of this research would be to
ascertain if these genes are functional in Acidobacterium sp. strain Ellin345 and
Solibacter usitatus strain Ellin6076 by testing their ability to oxidize these one-carbon

compounds in culture. If these endeavors prove fruitful, the next step would be to

173

determine how ubiquitous this physiology is among members of the phylum by testing
other currently, cultivated strains as well as designing alternative cultivation experiments
to select for this novel physiology in this phylum. Eventually, one could assess if
acidobacteria are truly one of the unexplored communities of methanotmphs responsible

for the discrepancy in affinities between the environment and known methanotrophs.

ACKNOWLEDGEMENTS
This work was funded by the USDA (#2001-35107-09939), NSF (#MCB-
0135880), and the NSF Long-Tenn Ecological Research Program at the Kellogg
Biological Station and the Michigan Agricultural Experiment Station.
I would like to thank Christopher D. Engles for his help with the l-LIBSHUFF

analysis.

174

10.

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Barns, S. M., S. L. Takala, and C. R. Kuske. 1999. Wide distribution and
diversity of members of the bacterial kingdom Acidobacterium in the
environment. Appl. Environ. Microbiol. 65:173 1-1737.

Benstead, J., G. M. King, and H. G. Williams. 1998. Methanol promotes
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178

CHAPTER 6

SUMMARY and SIGNIFICANCE

The soil environment is a rich habitat of life with members of the Bacteria,
Archaea, and Eukarya. However, this environment is largely unexplored since only 0.1
to 1% of the total microbial community has been cultivated. The exploration of this
untapped microbial diversity will allow us to understand the role microorganisms play in
the soil environment. This knowledge will enhance our understanding of nutrient
cycling, biogeochemistry, and agricultural practices. A microbial community that is
highly abundant in soil, but poorly understood is the phylum Acidobacteria. This
research was a union of both cultivation and non-cultivation based approaches, which
generated information pertaining to members of the phylum Acidobacteria in the soil
environment.

We knew previous to this study that members of the phylum Acidobacteria were
prevalent in a diverse collection of soils with varying degrees of relative abundance;
however the distribution of the subdivisions in relation to the various edaphic properties
was unknown. Acidobacteria comprise 1 to 6% of the total microbial community at the
KBS LTER based on their abundance of rRNA. Basic questions were asked about the
diversity of Acidobacteria at the KBS LTER, specifically how the structure of the
community changed in relation to various edaphic properties.

Phylogenetic libraries of the 16S rRNA gene were made from the conventional

agriculture (treatment 1), historically cultivated successional (treatment 7), and never

179

cultivated successional (treatment 8) plots. The practice of agriculture appears to
influence the acidobacterial communities through a shift in the relative abundance of the
different subdivisions. These data reveal that members of subdivision 1 and 6 are
dominant in the never cultivated successional treatment, whereas members of subdivision
4 dominate the conventional agriculture treatment.

The relative percentage of clones of each subdivision varied with depth and
treatment, suggesting that physical and chemical properties of the soil influenced the
distribution of the various subdivisions. Subdivisions responded to soil moisture
(subdivision 3), carbon concentrations (subdivision 4), soil pH (subdivision 1), methane
fluxes (subdivision 1), and nitrous oxide fluxes (subdivision 4). In particular, the relative
abundance of subdivision 1 clones in each treatment appeared to be related to soil pH
which was consistent with the pH optima of recently isolated strain in subdivision 1
(below). Additionally, members of subdivision 4 are more prevalent in the conventional
agriculture soil and respond positively to nitrous oxide fluxes suggesting that members of
this subdivision are capable of denitrification or respond favorably to nearby denitrifing
communities. These trends provide insight into the physiology of members of this
phylum as well as generated information which can be used to develop cultivation
strategies to isolate additional strains.

Successful cultivation attempts were made from the native and cultivated plots
from the KBS LTER. The use of media compositions and incubation conditions that
mimicked their native soil environment such as low nutrient media, atmospheric gas
concentrations, and extended incubation times as well as the development of a novel

screening method called Plate Wash PCR (PWPCR) was instrumental in isolating more

180

than a dozen strains from subdivisions 1 and 3. Using our novel cultivation approach, we
recovered between 3.5 to 5 x 107 CFU/gram of soil (dry wt), which was ca. 2.5 to 6% of
the microbial community based on direct counts. Extended incubation time, low
concentrations of carbon, and soil inoculum with high moisture (significantly) improved
total recovery of aerobic heterotrophs from soil. A facile, high-throughput method called
Plate wash PCR was developed to rapidly screen a multitude of plates for the target
organism(s) using group-specific primers. Additionally, PWPCR revealed that
acidobacteria were more frequently detected with elevated levels of carbon dioxide
(significantly), the presence of the catalase enzyme, low carbon, and low oxygen
concentrations.

Characterization of these strains revealed the nature and importance of members
of the phylum Acidobacteria in the soil environment. Typically, strains of subdivision 1
were isolated from soils using low-nutrient media incubated under elevated levels of
carbon dioxide for extended periods. Such incubation conditions resulted in a slightly
acidified medium that matched the pH optima of the strains (between 5 and 6), which
also was similar to their native soil pH. All strains were Gram negative, aerobic,
chemoorganotrophic, nonmotile rods that produced an extracellular matrix which causes
cells to clump in liquid culture. The production of this extracellular material is believed
to promote adhesion, prevent desiccation, and promote soil aggregation, all of which
would be beneficial to a soil microorganism. Colonies were small, approximately 1 mm
in diameter and either white, pale yellow or pink in color, the latter owing to the presence
of a carotenoid(s) whose formation was much greater under 20% oxygen as opposed to

2%. The presence of a carotenoid(s) in these strains is believed to be an oxidative stress

181

response. Additionally, strain KBS 83 appears to be a microaerophile, growing optimally
at oxygen concentrations between 4 to 16%. The ability to grow and/or grow better
under lower concentrations of oxygen is a valuable trait in the soil environment, since
oxygen can be depleted after rain events and in the inner layers of soil aggregates.

All strains contained either 1 or 2 copies of the 16S ribosomal RNA encoding
gene which along with extended doubling time (10-15 hours at ca. 23°C), is suggestive of
an oligotrophic lifestyle. Major differences in carbon utilization profiles were noted
during routine characterization of the strains. Strains KBS 83 (subdivision 1) and KBS
96 (subdivision 3) were able to use more recalcitrant carbon source such as cellulose,
syringate, ferulate, and pectin whereas members of the newly described T erriglobus
genus (below) could not. The ability to oxidize these more recalcitrant carbon source(s)
is a practical trait in the soil environment. The soil is typically defined as carbon
limiting, however plant polymers and their products of degradation are major source of
carbon for indigenous populations of microorganisms. Preliminarily, strain KBS 96 is
capable of reducing nitrate to nitrite under anoxic conditions, but not able to denitrify.
This strain was isolated from a conventional agriculture soil, high in nitrate
concentrations. Recently isolated strains from subdivision 1 were not able to grow under
these conditions; therefore it appears that this physiology could be used as a means of
selection for isolating additional members in subdivision 3.

Previous work has suggested that members of subdivision 1 in the phylum
Acidobacteria are capable of oxidizing one-carbon compounds such as methanol, a
physiology known as methylotrophy. A molecular, growth, and cultivation-based

approach were taken to explore this interesting physiology. The phylogenetic survey

182

revealed that members of subdivision 1 were positively influenced by methane, a one-
carbon compound. Methanol-enriched soil selected for a phylogenetic clade of
acidobacteria in subdivision 1, however no methanol-oxidizing acidobacteria were
cultivated. Readily available strains were tested for their ability to growth on various
one-carbon compounds, however under the conditions tested, they were not able to grow.
At this time, our strains do not appear to have the capability to growth on one-carbon
compounds under the conditions tested.

Six of the strains in subdivision 1 are sufficiently similar to one another, but
distinct from previously named acidobacteria, to warrant creation of a new genus;
T erriglobus, with T. roseus defined as the type species. The characteristics of members
of the phylum Acidobacteria elucidated in this study are consistent with the widespread
distribution of acidobacteria in soil. Type strain KBS 63 has been deposited into the
Deutsche Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany
and the USDA Agricultural Research Service Culture Collection to allow the our lab and
the scientific community to work with these novel strains and continue to make future
discoveries.

In conclusion, this research addressed some basic questions about acidobacteria in
relation to their native environment using both molecular and growth-based approaches.
Differences in the relative percentages of the different acidobacterial subdivisions in the
various plots at the KBS LTER are suggestive of their niche or ecological role in the soil
environment. The development of novel cultivation strategies including PWPCR along
with the successful isolation more than a dozen members of the phylum Acidobacteria

are major contributions to the scientific community. This research revealed not only

183

interesting properties of the acidobacteria, but also laid the foundation for further

questions and research to continue on this diverse group of microorganisms.

184

APPENDIX A

OVERVIEW
This appendix is a brief description of the VSB-#, the “vitamins and salts base”, and
MHM-#, the “modified hyphomicrobium medium”; # indicates the respective pH of the

basal solution.

1. VSB-#

1. 1X basal medium I

2. SL—lO trace element solution (lmL/liter)

3. Vitamin mixture (lmL/liter)

4. Vitamin B12 (lmL/liter)

5. Buffer (HEPES, MOPS, MES) (10mM final concentration)

6. distilled water (up to 1 liter)
Basal Medium I (2)

Per liter Final Concentration

NaCl 1.0g ca. l7mM
MgC12°6H20 0.62g ca. 3.1mM
CaC12-2H20 0.15g ca.1.0mM
NH4C1 0.25g ca. 4.7mM
KH2P04 0.2g ca. 1.5mM
KCl 0.5g ca. 6.7mM
Na2804 0.28g ca. 2mM

NOTE: When making up the Basal Medium 1, add each component one at a time and
wait until it is completely dissolved before adding the next component. Follow the order
of components exactly for successful preparation. In my experience, the addition of
Na2804 in the Basal Medium I reduced the solubility of the salts. A separate stock of
Na2804 was prepared at a concentration of 1.25M and 1600uL was added per liter.

SL-10 Trace Metals (1 ml/liter) (2)

Per liter
HCl (25%) 10ml
FCC12‘4H20 l .5 g
COC12‘6H20 190mg
MnC12‘4H20 100mg
ZnC 12, 70mg
H3BO3 6mg

185

N32M004‘2H20 36mg
N1C12'6H20 24mg
CuC12-2HzO 2mg

NOTE: When making up the SL10 trace metals solution, add each component one at a
time and wait until it is completely dissolved before adding the next component. Follow
the order of components exactly for successful preparation. In my experience, the
dissolving of FeC12-4H20 was problematic, therefore, first dissolve FeClz-4H20 in the
HCl (25%) before addition of water or the other components.

Vitamin Mixture (lml/liter)(2)

Per 100ml
Sodium phosphate buffer (10mM; pH 7.1) 100ml
4-aminobenzoic acid 4mg
D (+)-biotin 1mg
Nicotinic acid 10mg
Calcium D (+)-pantothenate 5mg
Pyridoxine dihydrochloride 15mg

Vitamin B12 solution (50 mg/I) (lml/liter)

186

2. MHM-#

1X Freshwater base

150mM mix of K2HPO4 & KH2P04 (below) (pH 6) (lml/liter)
SL-10 trace element solution (lmL/liter)

Vitamin mixture (lmL/liter)

Vitamin BIZ (lmL/liter)

Buffer (HEPES, MOPS, MES) (20mM final concentration)
Desired carbon source

distilled water (up to 1 liter)

WNQMPVNT‘

Freshwater Base (1)

Per liter Final Concentration

NaCl 1.0g ca. 17mM
MgClz°6H20 0.4g ca. 2mM

CaC12-2H20 0.1g ca. 0.7mM
NH4C1 0.25g ca. 4.7mM
KH2P04 0.2g ca. 1.5mM
KCl 0.5g ca. 6.7mM
NaZSO4 0.28g ca. 2mM

KNO3 5g ca. 54mM

NOTE: When making up the MHM, add each component one at a time and wait until it
is completely dissolved before adding the next component. Follow the order of
components exactly for successful preparation. In my experience, the addition of Na2$O4
in the Basal Medium I reduced the solubility of the salts. A separate stock of NaZSO4 was
prepared at a concentration of 1.25M and 1600111 was added per liter.

150mM mix of K2HPO4 & KHzPO4 (pH 6)
KZHPO4 6g
KH2P04 19g

SL-l0 Trace Metals (1 ml/liter) (above) (2)
Vitamin Mixture (lmI/liter) (above) (2)
Vitamin B12 solution (50 mg/l) (1 ml/liter)

187

REFERENCES

Poindexter, J. S. Dimorphic prosthecate bacteria: the genera Caulobacter,
Asticcacaulis, Hyphomicrobium, Pedomicrobium, Hyphomonas, and
T hiodendron. In A. Balows, H.G. Trueper, M. Dworkin, W. Harder, K.H.
Schleifer (ed.), The Prokaryotes: a handbook on the biology of bacteria:
ecophysiology, isolation, identification, applications, 2 ed, vol. 3. Springer-
Verlag, New York.

Widdel, F., and F. Bak. 1991. Gram-negative mesophilic sulfate-reducing
bacteria, p. 3352-3378. In A. Balows, H.G. Trfiper, M. Dworkin, W. Harder, and
K.H. Schleifer (ed.), The Prokaryotes, 2 ed, vol. 4. Springer-Verlag, New York,
NY.

188

APPENDIX B

OVERVIEW
This appendix contains all of the raw data for the soil plating experiments from the KBS
LTER including collection date, plating date, treatment & replicate(s), soil moisture (%),

treatments, and average recovery in CFU/gram of soil (dry weight).

189

1. SOIL PLATING EXPERIMENT #1

SOIL COLLECTION DATA:

Collection Date: July 10, 2001

Plating Date: August 13, 2001

Treatment & Replicate: KBS LTER, Treatment 8, Replicate 1, sampling stations 1-5
Soil Moisture: 7.1 :1: 1.2%

 

Average CFU/gram of soil :1: 3.1] (dry wt.) (10f)1r

 

After 24 days
Air

Agarose 0.27 :1: 0.04

Agar 1.06 d: 0.13

Agar, 0.001 R2A 1.09 :1: 0.14

Agarose, 0.001 R2A 0.28 i 0.01
a b c

““40 ’ aHSL ’Feg 0.81 $0.03
catalase

thcr, aHSL”, Fe° 0.58 i 0.12

NH‘CI’Fe; l.18:1:0.13
catalase

thcr, Fe" 1.34 .1: 0.09
a b

““40 ’ aHSL; 0.84 a 0.24
catalase

thcr, aHSLb 0.30 s: 0.13

thcr, catalased 1.22 :t 0.39

NH4Cl“ 0.98 :1: 0.21

aHSLb, Fec, catalased 0.69 s 0.07

aHSL", Fe° 0.55 :1: 0.16

F e°, catalased 1.07 :1: 0.49

Fe° 1.15 :1: 0.04

aHSLb, catalased 0.54 a 0.32

aHSL” 0.61 :1: 0.03

catalased 0.98 :1: 0.24

no additives 1.06 :1: 0.22

0.001R2A 1.18 :1: 0.04

0.001 R2A, PEG‘ 0.01 e 0.01

 

1 Unless otherwise noted, medium was water BactoTM agar (1.5%).

' Ammonium chloride (NH4C1) was at a final concentration of 1mM.

b aHSL cocktail included: ketocaproyl-, octanoyl-, hexanoyl-, butyryl-, heptanoyl-, and
tetradecanoyl-DL- at a 111M concentration each. The HSL cocktail stock was made in acidified
ethyl acetate and added directly to cooled, molten agar medium immediately before pouring.

° Ferric chloride (Fe) was at a final concentration

d Catalase was added at a final concentration of 65U/mL.

° Polyethylene glycol was added at a final concentration of 0.58M.

190

2. SOIL PLATING EXPERIMENT #2

SOIL COLLECTION DATA:

Collection Date: September 21, 2001

Plating Date: October 2, 2001

Treatment & Replicate: KBS LTER, Treatment 8, Replicate 1, sampling stations 1-5

Soil Moisture: 24.8 :1: 0.7%
Average CFU/gram of soil :1: s.d (dry wt.) (107)

 

After 28 days
Elevated CO; Hypoxia Air
Washed agar" 4.57 a 0.76 4.84 i 0.17 3.12 i 0.16
Washed agar‘ w/
VSB-7“ 4.61 :1: 0.39 6.02 :1: 0.78 4.13 s 0.42
Catalaseb w/
VSB-7a 6.67 e 4.15 4.20 d: 0.13 4.84 :1: 1.38
AQDSd w/
VSB-7‘ 4.87 s 0.61 4.04 i 0.25 3.99 3: 0.54
AQDSd, catalaseb
w/ v513.7a 4.68 i 1.17 3.69 d: 0.27 4.11 :1: 0.11
VSB-7“ 5.82 i 2.30 4.38 :1: 0.68 4.66 a 1.67
AEA“ w/ VSB-7"
N0 additives 5.13 :1: 1.24 5.06 s 0.20 2.89 :t 0.11

 

' Unless otherwise noted, the medium consisted of VSB-7 (2) with HEPES (10mM), pH 7 as a
buffering system, and aHSL cocktail. aHSL cocktail included: ketocaproyl-, octanoyl-,
hexanoyl-, butyryl-, heptanoyl-, and tetradecanoyl-DL- at a 111M concentration each. The HSL
cocktail stock was made in acidified ethyl acetate and added directly to cooled, molten agar
medium immediately before pouring:

b Catalase was added at a final concentration of 65U/mL.

° “AEA” includes the addition of acidified ethyl acetate without the aHSL.

d AQDS had a final concentration of 25mM.

° BactoTM agar was washed twice in distilled water.

191

3. SOIL PLATING EXPERIMENT #3

SOIL COLLECTION DATA:

Collection Date: November 27, 2001

Plating Date: November 30, 2001

Treatment & Replicate: KBS LTER, Treatment 8, Replicate 1, sampling stations 1-5
Soil Moisture: 32 d: 1.4%

 

Average CFU/gram of soil :1: s.d (dry wt.) (107) '

 

After 22 days
Elevated CO; Hypoxia Air Knallgas Mix(l)
HEPES? 6.53 a 0.61 6.20 a 0.76 3.22 a 0.38 2.96 a 1.98
catalasec
HEPES" 5.20 a 0.58 4.96 a 0.49 3.19 :1: 0.59 -
HEPES", 4.94 :1: 1.56 5.23 a 1.06 4.11 a 2.84 -
catalasec, R2Ad
HEPES", R2A“ 5.86 a 1.11 5.03 r 1.51 3.07 :t 0.63 -
HEPES", 5.86 a 1.85 6.55 a 1.23 5.06 3; 0.50 -
catalase°, aHSL°
HEPES", 4.29 a 0.74 3.84 r. 0.59 4.36 a 0.56 -
catalase°, AEAf
HEPES", - - 5.06 a 1.33 -
catalase‘,
Bacxell8
HEPES", - - 4.24 a 0.58 -
catalase°,
Bacxellg, R2Ad
HEPESb, - - 5.30 a 0.92 2.84 a: 2.18
catalase°,
Bacxellg, aHSL°
HEPES", aHSL‘, - - 5.80 a 1.10 -
Bacxellg, R2Ad
catalase° 4.94 a 0.68 4.99 a 0.63 3.29 r 0.38 -
Rainwater 8.95 r 6.03 4.81 r 1.56 3.26 a 0.43 2.32 a 1.83

extract“, catalasec

 

" Unless otherwise noted, the medium consisted of VSB-7 (2).

b HEPES (10mM), pH 7 was used as a buffering system.
° Catalase was added at a final concentration of 65U/mL.
d R2A carbon additive included all carbonaceous components of R2A medium except pyruvate.
It was at a 0.1x concentration.

° aHSL cocktail included:

ketocaproyl-, octanoyl-, hexanoyl-, butyryl-, heptanoyl-, and

tetradecanoyl-DL— at a 111M concentration each. The HSL cocktail stock was made in acidified
ethyl acetate and added directly to cooled, molten agar medium immediately before pouring.

' Acidified ethyl acetate was added as a control for the aHSL treatment.
‘ Bacxell was an additive given to us by another lab speculated to increase recovery of

microorganisms.

h Rainwater extract was prepared via a cold extraction of soil from the KBS LTER.

192

4. SOIL PLATING EXPERIMENT #4

SOIL COLLECTION DATA:
Collection Date: June 10, 2002
Plating Date: June 11, 2002

Treatment & Replicate: KBS LTER Treatment 8, Replicate 4, sampling stations 1-5

Soil Moisture: 17.8 :1: 1%
Average CFU/gram of soil :1: s.d (dry wt.) (107) '

 

After 30 days
Elevated CO; Hypoxia Air

aHSL", catalase° 5.21 a 0.66 6.91 :r 0.25 7.02 a 0.95
catalasec 5.8 :1: 1.3 6.63 :1: 0.81 9.67 a 2.79
catalase°, light 4.76 i 050 - 7,75 :1: 0,93
aHSLb’ “‘35:; 5.07 :1: 0.53 6.66 a 0.58 4.00 i 3.63

aHSLb 4.61 a 2.41 6.55 :1: 0.78 -

No additives 6.25 a 0.28 5.65 3: 1.93 -

aHSL”, R2Ad 5.36 a 0.25 5.57 a: 2.33 -

 

' Unless otherwise noted, the medium consisted of VSB-7 (2) and HEPES (10mM), pH 6.8 as a
buffering system.

b aHSL cocktail included: ketocaproyl-, octanoyl-, hexanoyl-, butyryl-, heptanoyl-, and
tetradecanoyl-DL- at a 111M concentration each. The HSL cocktail stock was made in acidified
ethyl acetate and added directly to cooled, molten agar medium immediately before pouring.

° Catalase was at a concentration of 65U/mL.

d R2A carbon additive included all carbonaceous components of R2A medium except pyruvate.
It was at a 0.1x concentration.

193

5. SOIL PLATING EXPERIMENT #5

SOIL COLLECTION DATA:

Collection Date: November 12, 2002

Plating Date: November 22, 2002

Treatment & Replicate: KBS LTER Treatment 8, Replicate 3, sampling stations 1-5;
Treatment 1, Replicate 3, sampling stations 1-3; Treatment 7, Replicate 2, sampling
stations 1-3; Deciduous forest, Replicate 2, sampling stations 1-5

Soil Moisture: T8, 21.1 i 0.7%; T1 & 7 (reps. 1-3 for each), 17.5 :t 0.5%; DF, 17.2 i

 

 

0.4%
Average CFU/gram of soil i s.d (dry wt.) (107) '
20 days of growth
Anoxia Hypoxia
T8 soil
aHSL‘l RZA" 6.26 r 0.21 -
aHSL°, Methyl
cocktail d 0.19 i 0.18 9.51 i 0.76
aHSL“. AQDS ° 2.69 a 0.39 8.0 a 2.08
T1 & 7 soil (pooledb)
aHSL’RzA 39910.53 5.11 a0.41
aHSL“, Methyl
cocktail d 0.44 i 0.20 5.45 d: 1.10
aHSL", AQDS ° 1.62 d: 0.34 5.16 r 1.01
Deciduous forest solil
aHSL°, R2A 0.89 a 0.64 1.53 a 0.31
aHSL°, Methyl
cocktail d 0.03 i 0.03 0.84 i 0.24
aHSL“. AQDS ° 0.38 a 0.23 2.49 a: 1.51

 

" Unless otherwise noted, the medium consisted of VSB-7 (2) catalase (65U/mL), and HEPES
(10mM), pH 6.8 as a buffering system.

b R2A carbon additive included all carbonaceous components of R2A medium except pyruvate.
It was at a 0.1x concentration.

° aHSL cocktail included: ketocaproyl-, octanoyl-, hexanoy1-, butyryl—, heptanoyl-, and
tetradecanoyl-DL- at a 111M concentration each. The HSL cocktail stock was made in acidified
ethyl acetate and added directly to cooled, molten agar medium immediately before pouring.

d Methyl cocktail contained SmM methanol, 3mM methylamine, and 1mM dimethylamine. This
treatment was buffered with bicarbonate.

° AQDS had a final concentration of 25mM.

194

6. SOIL PLATING EXPERIMENT #6

SOIL COLLECTION DATA:
Collection Date: June 6, 2003
Plating Date: June 26, 2003
Treatment & Replicate: KBS LTER, Treatment 8, Replicate 3, sampling station 1-5
Soil Moisture: 10 :t 0.2%

Average CFU/gram of soil :1: s.d (dry wt.) (107)

 

47 days of growth
Elevated CO; Hypoxia Air Anoxia
12°C'
No additives 0.51 a: 0.21 0.98 3: 0.16 0.94 a 0.19 0.90 :1: 0.43
aHSL° 0.61 a 0.34 0.90 a; 0.21 0.84zt 0.23 0.87 a 0.10
AQDSd 0.54 a: 0.30 0.98 a: 0.30 0.99 a 0.20 0.48 a 0.13
R2Ab 0.83 r 0.09 0.84 a: 0.16 0.75 :1: 0.17 0.90 a 0.23
23°C'
No additives 1.27 a 0.17 1.14 a 0.17 1.12 a 0.28 0.58 r 0.32
aHSL° 1.90 a 0.52 1.16 a 0.28 1.46 a 0.35 0.81 a 0.21
AQDSd 1.03 10.17 1.00:0.15 1.10a032 1.08:1:0.36
R2A” 1.12 :1: 0.16 0.89 a 0.49 1.20 a 0.25 0.98 3: 0.37

 

' Unless otherwise noted, the medium consisted of VSB-7 (2) catalase (65U/mL), and HEPES

(10mM), pH 6.8 as a buffering system.

b R2A carbon additive included all carbonaceous components of R2A medium except pyruvate.
It was at a 0.1x concentration.

° aHSL cocktail included:

ketocaproyl-, octanoyl-, hexanoyl-, butyryl-, heptanoyl-, and

tetradecanoyl-DL- at a 111M concentration each. The HSL cocktail stock was made in acidified
ethyl acetate and added directly to cooled, molten agar medium immediately before pouring.

d AQDS had a final concentration of 25mM.

195

7. SOIL PLATING EXPERIMENT #7

SOIL COLLECTION DATA:
Collection Date: July 21, 2006
Plating Date: July 22, 2006
Treatment & Replicate: KBS LTER, Treatment 8, Replicate 1, sampling station 1-5;
Treatment 1, Replicate 1, sampling station 1-5;
Soil Moisture: T8, 15.0 :1: 0.0%; T1, 8.0 d: 0.0%

 

Average CFU/gram of soil 4 s.d (dry wt.) (10’)

 

After 30 days
Hypoxia
pH 5.5 pH 6.5 pH 5.5 pH 6.5
T1 Soil
VL55". plan; 1.07 a 0.27 0.98 :1: 0.40 0.82 :r 0.02 0.87 :1: 0.03
polymers
VLSSb, xylane 0.91 i 0.24 1.16 a 0.21 0.94 :1: 0.04 1.09 :1: 0.01
VLSSb, R2Bc 0.91 i 0.15 1.23 :1: 0.07 1.04 :1: 0.01 1.07 :t 0.04
VSBa, xylan‘ 0.00 i 0.00 0.97 :t 0.11 0.02 :1: 0.001 0.76 :1: 0.03
VSBa , R2B° 0.51 i 0.19 0.36 :t 0.08 0.68 :1: 0.04 0.54 :1: 0.02
Soil extract mediumf - - - -
R2ATM 0.54 :1: 0.09 0.97 :1: 0.02
T8 Soil
VLSS". plan; 0.75 a 0.19 0.55 a 0.32 0.82 a 0.05 0.59 a 0.61
polymers
VLSS", xylan° 0.77 d: 0.36 0.90 i 0.27 1.02 i 0.29 1.05 :1: 0.50
VLSSb, R2Bc 1.29 :1: 0.45 0.80 i 0.23 1.07 i 0.47 0.98 :1: 0.69
VSB“, xylan° 0.00 :1: 0.00 0.69 i 0.17 0.29 d: 0.09 0.57 d: 0.30
VSB“ , RZBc 0.51 i 0.19 0.69 d: 0.23 0.48 :1: 0.23 0.44 :t 0.20
Soil extract mediumf - - - -
R2ATM 0.51 :t 0.15 0.61 i 0.04

 

 

196

' Unless otherwise noted, the medium consisted of VSB-# (2) catalase (65U/mL), and HEPES
(10mM), pH 6.8 as a buffering system.

b VL55 is described previously (3).

° R2A carbon additive included all carbonaceous components of R2A medium except pyruvate.
It was at a 0.1x concentration.

d Plant polymers mix consisted of xylan (oat spels), xanthan, pectin, xylan (Birchwood), methyl
cellulose, and xylan (Larch) each at 0.008% (wt/vol) giving a final concentration of 0.05%
(wt/vol).

° Xylan (Birchwood) was dissolved in water and added to a final concentration of 0.05%
(wt/vol).

f Soil extract medium consisted of a mixture of equal volume of soil (T8 & T1) to water with agar added as
the solidifying agent.

197

REFERENCES

Aragno, M., and H.G. Schlegel. 1992. The mesophilic hydrogen-oxidizing
(Knallgas) bacteria, p. 344-384. In A. Balows, H.G. Trilper, M. Dworkin, W.
Harder, and K.H. Schleifer (ed.), The Prokaryotes, vol. 1. Springer-Verlag, New
York, NY.

Eichorst, S. A., J.A. Breznak, and T.M. Schmidt. 2007. Isolation and
characterization of newly isolated soil bacteria that define Terriglobus gen. nov.,
in the phylum Acidobacteria. Appl. Environ. Microbiol. 73:2708-2717.

Joseph, S. J., P. Hugenholtz, P. Sangwan, C. A. Osborne, and P. H. Janssen.

2003. Laboratory cultivation of widespread and previously uncultured soil
bacteria. Appl. Environ. Microbiol. 69: 72 1 0-72 1 5.

198

This appendix is a list of primers, both 16S rRNA and functional genes, used throughout

APPENDIX C

OVERVIEW

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

my dissertation.

PCR Primers Opick Reference

Name Gene Sequence Ref.
8F 16S rRNA 5’ AGA GTT TGA TCC TGG CTC AG 3’ (9)
ACD 31F l6S rRNA 5’ GAT CCT GGC TCA GAA TC 3’ (1)
1492R 16S rRNA 5’ GGT TAC CT’T GTT ACG ACT T 3’ (9)
338R 16S rRNA 5’ GCT GCC TCC CGT AGG AGT 3’ (8)
338F l6S rRNA 5’ CTC CTA CGG GAG GCA GCA GT 3’ (8)
531R l6S rRNA 5’ TAC CGC GGC TGC TGG CAC 3’ (8)
810R l6S rRNA 5’ GGC GTG GAC TTC CAG GGT ATC T 3’ (8)
776F 168 rRNA 5’ AGC AAA CAG GAT TAG ATA CCC TGG 3’ (8)
1087F 16S rRNA 5' GGT TAA GTC CCG CAA CGA 3’ (8)
mg?“ TOPO TA 5’ CAG TCA CGA cor TGT AAA ACG ACG GC 3’ (7)

vector
"‘Od‘fied TOPO TA 5’ CAG GAA ACA GCT ATG ACC ATG 3’ (7)
M13R vector
mxaf(MDH
active site mxa f1003 5’ GCG GCA CCA ACT GGG GCT GGT 3’ (10)
subunit) mxa f156l 5’ GGG CAG CAT GAA GGG CTC CC 3’
mmoX
(sMMO mmoX 206F 5’ ATC GCB AAR GAA TAY GCS CG 3’ ( 6)
active site mmoX 886R 5’ ACC CAN GGC TCG ACY TTG AA 3’
subunit)
mmoX
(sMMO mmoX F536 5’ CCG CTG TGG AAG GGC ATG AA 3’ (4)
active site mmoX R898 5’ CAC TCG TAG CGC TCC GGC T 3’
subunit)
pmoA
(pMMO A189 5’GGN GAC TGG GAC TTC TGG 3’ (3)
active mb661 5’ CCG GMG CAA CGT CYT TAC C 3’
subunit)
pmoA &
amoA
“’“m‘ive A189 5’GGN GAC TGG GAC TTC TGG 3’
active site (5)
. A682 5’ GAA SGC NGA GAA GAA SGC 3’

subunit of
pMMO and
AMO)
Nitrogenase
gene (2 sets; IGK, 5’ AAR GGN GGN ATH GGN AA 3’
IGK & ni 1H KAD, 5’ TGY GAY CCN AAR GCN GA 3’ (1 l)
YAA and GEM, 5’ AND GCC ATC ATY TCN CC 3’
KAD & YAA, 5’ ATR TTR TTN GCN GCR TA 3’
GEM)

 

 

 

 

199

 

 

PCR Primers Quick Reference cont’d

 

 

 

 

Name Gene Sequence Ref.
Nitrite
reductase nirKlF 5’ GG(A/C) ATG GT(G/T) CC(C/G) TGG CA 3’ (2)
gene nirKSR 5’ GCC TCG ATC AG(A/G) TT(A/G) TGG 3’
Nitrite
reductase nirSlF 5’ CCT A(C/T)T GGC CGC C(A/G)C A(A/G)T 3’ (2)
gene nirS6R 5’ CGT TGA ACT T(A/G)C CGG T 3’

 

 

 

 

200

 

10.

REFERENCES

Barns, S. M., S. L. Takala, and C. R. Kuske. 1999. Wide distribution and
diversity of members of the bacterial kingdom Acidobacterium in the
environment. Appl. Environ. Microbiol. 65: 1731—1737.

Braker, G., A. Fesefeldt, and K.-P. Witzel. 1998. Development of PCR primer
systems for amplification of nitrite reductase genes (nirK and nirS) to detect
denitrifying bacteria in environmental samples. Appl. Environ. Microbiol.
64:3769-3775.

Costello, A. M., and M. E. Lidstrom. 1999. Molecular characterization of
firnctional and phylogenetic genes from natural populations of methanotrophs in
lake sediments. Appl. Environ. Microbiol. 65:5066-5074.

Fuse, H. M., M. Ohta, O. Takimura, K. Murakami, H. Inoue, Y. Yamaoka,
J.M. Oclarit, and T. Omori. 1998. Oxidation of trichloroethylene and dirnethyl
sulfide by a marine Methylomicrobium strain containing soluble methane
monooxygenase. Biosci. Biotechnol. Biochem. 62:1925-1931.

Holmes, A. J., A.M. Costello, M.E. Lidstrom, and J.C. Murrell. 1995.
Evidence that particulate methane monooxygenase and ammonium
monooxygenase may be evolutionarily related. FEMS Microbiol. Lett. 132:203-

208.

Hutchens, E., S. Radajewski, M. G. Dumont, I. R. McDonald, and J. C.
Murrell. 2004. Analysis of methanotrophic bacteria in Movile Cave by stable
isotope probing. Environ. Microbiol. 6:1 1 1-120.

Kim, K. S., T.G. Lilburn, M.J. Renner, and J.A. Breznak. 1998. arfI and arflI,
two genes encoding alpha-L-arabinofuranosidases in Cytophaga xylanolytica.
Appl. Environ. Microbiol. 64: 1919-1923.

Lane, D. J. 1991. Chapter 6 16S/23S rRNA sequencing. In E. S. M. Goodfellow
(ed.), Nucleic acid techniques in bacterial systematics. John Wiley & Sons Ltd.,
New York.

Lane, D. J., B. Pace, G. J. Olsen, D. A. Stahl, M. L. Sogin, and N. R. Pace.
1985. Rapid determination of 16S ribosomal RNA sequences for phylogenetic
analyses. Proc. Natl. Acad. Sci. USA. 82:6955-6959.

McDonald, 1., and J. Murrell. 1997. The methanol dehydrogenase structural
gene mxaF and its use as a functional gene probe for methanotrophs and
methylotrophs. Appl. Environ. Microbiol. 63:3218-3224.

201

11. Ohkuma, M., S. Noda, R. Usami, K. Horikoshi, and T. Kudo. 1996. Diversity
of nitrogen fixation genes in the symbiotic intestinal microflora of the termite
Reticulitermes speratus. Appl. Environ. Microbiol. 62:2747-2752.

202

APPENDIX D

BACTERIAL GROWTH EFFICIENCY: UNDERSTANDING ECOLOGICAL

STRATEGIES OF COPIOTROPHS AND OLIGOTROPHS

INTRODUCTION

Ecologists R.H. MacArthur and BC. Wilson developed a theory of r and K
selection that classifies organisms based on their life history including traits such as
development, growth, reproductive strategies, and life span. R-strategists are defined as
species found in low population densities where growth can be exponential and substrate
availability is high (21), therefore growth is regulated independent of population density.
They are greatly affected by catastrophic events in the physical environment, and their
characteristics include high growth rate, reduced body size, and reduced intraspecific
competition (2, 16). K-strategists are typically found in high population densities at or
near the carrying capacity (K) where there is intense intraspecific competition for scarce
resources (21) and growth is regulated by the population density. Characteristics of a
K-strategists include reduced growth rate, increased body size, and presumed higher
efficiency in the utilization of resources and competition (2, 16). There is a systematic
trade-off when applying this theory: an organism can either be a r-strategist or a K-
strategist, but not both (20, 21, 39, 40) because it contradicts all notions of evolution of a
generalist versus a specialist (39).

It is relatively easy to observe this theory in plants and animals when one

considers traits such as clutch size or parental care. But how can this theory be observed

203

and applied to bacteria? Andrew and Harris (1986) hypothesized that bacteria with high
maximum growth rates and a requirement for abundant resources during growth are r-
strategists, where as bacteria with low maximum growth rate and require few resources
for growth are K-strategists (1). Depending on their metabolic attributes, one can classify
these organisms as being more oligotrophic (K-selected) or copiotrophic (r-selected).
Copiotrophs are microbes that grow and multiply in the presence of an abundance of
carbon, whereas oligotrophs are defined as microbes that are isolated on media
containing 1-15mg organic C/liter (30).

The classic definition of oligotrophy as defined by Poindexter states that the
bacteria must be isolated on medium containing 1-15 mg/L of carbon (30). This
definition might be too stringent. The requirement to grow at low concentrations of
carbon might be a characteristic to some more obligate oligotrophic bacteria, but its
absence does not exclude the possibility of other strains to be considered oligotrophic as
suggested in an article by Arthur Koch (14). Koch suggested there are four categories of
oligotrophy; 1. bacteria that cannot be cultivated; 2. bacteria cultivated on poor medium;
3. bacteria isolated on poor medium, but are able to grow on rich medium; and 4. bacteria
only initially cultivated (14). The third category is consistent with the growth dynamics
of Sphingopyxis alaskensis strain RB2256 (formerly Spingomonas alaskensis), the model
oligotroph, which was isolated on synthetic seawater medium containing less that 1 mg of
carbon per liter, but subsequently adapted in the lab to grow on high nutrient medium
(3 3). Recently, a more inclusive definition has been suggested by Fierer and colleagues
that includes attributes such as relatively slow growth rate, high affinity uptake systems,

and better growth under nutrient-poor conditions (9) which has a genetic marker of the

204

16S rRNA-encoding gene in the rm operon (13, 37). It is this more inclusive definition
of oligotrophy which will be used in this chapter.

The 16S rRNA-encoding gene in the rrn operon appears to be a good genetic
marker to preliminarily categorize a bacterium as being more copitrophic or oligotrophic,
specifically when discussing growth rate (13, 37). The genes encoding the 5S, 16S, and
23S rRNAs in bacteria are typically organized into an operon (Figure DJ), and the
number of 16S-rRNA encoding gene(s), rrs, are determined using Southern hybridization
with specific probes. In general, bacteria with few rrs copies (_<_ 2 copies) are found in
low nutrient environments and are slow-growing such as Terriglobus roseus strain KBS
63 and Sphingopyxis alaskensis strain RB2256, where as bacteria with multiple copies of
the rrs (3 3 copies) are found in nutrient rich environment and are fast-growing such as
E. coli. This correlation between high number of rrs and faster growth rate is clearly
illustrated when growing different species of Mycobacterium and Rhizobium that differ in
rrs copy number (17, 36), presumed to be a result of these strain increased ability to
respond to changes in the environment. Since the rrs appears to be a good genetic
marker for assessing bacteria’s ecological strategy, fast-growing bacteria with a high rrs
copy number (3 3 copies) will be referred to as copiotrophs, and slow-growing bacteria
with a low rrs copy number (5 2 copies) will be referred to as oligotrophs.

There are systematic differences in metabolic efficiency between copiotrophs and
oligotrophs. The idea of efficiency has been discussed in the literature with such terms as
YATp and bacterial growth efficiency. Y”): is a term more fi'equently used by microbial

physiologists, and it is defined as the amount of ATP required for the formation of

205

Internally

Promoters (P1 & P2) transcribed Terminators
spacer
l '—' l

   
   

5' 3.

5S rDNA

Figure D. 1. Classic organization of the rrn operon.

206

cellular material (38), where yield is ultimately dependent on the amount of ATP
produced by the organism. YA—rp can vary dramatically with the type of carbon source
(104 ATP mole/grams cell formed): glucose and pyruvate, 28.8 and 13.5, respectively) as
well as medium supplemented with amino acids (31.9 x 104 ATP mole/grams cell
formed) and nucleic acid bases (28.8 x 104 ATP mole/grams cell formed) (38). Bacterial
growth efficiency or BGE is a term used by microbial ecologists, and it is defined as the
amount of new biomass produced (bacterial production or BP) per unit organic carbon
substrate consumed (BP + bacterial respiration (BR)) (6). BGE has been explored
primarily in natural planktonic ecosystems where it increases along with productivity,
reaching an asymptote at a BGE of 0.5 (5). Hereafier, this concept of efficiency will be
referred to as bacterial growth efficiency (BGE), and it will be defined as moles of carbon
in biomass/moles of carbon consumed.

The goal of this study was to determine if there is a difference in BGE among a
diverse collection of bacteria which differ in the rrs copy number. Previous work has
illustrated that there is a strong, positive correlation between rrs copy number and growth
rate indicating that the rrs copy number is a suitable genetic marker for ecological
strategy (13). In addition, other studies revealed differences in YATP or BGE of
communities and/or pure cultures suggesting systematic differences among strains’
ability to utilize substrate. The next logical question is to determine if there is a
correlation between BGE and rrs copy number.

Copiotrophs’ strategy is to maximize their growth rate, typically having a high
number of offspring per unit time; therefore, it is plausible to assume that when

maximizing growth rate, it would be less efficient (decrease in cells per mole of carbon)

207

or wasteful with their resources. On the other hand, an oligotroph’s strategy is to limit
their population density based on the amount of limited resources available; in order to do
so, it would demand the most efficient (increase in cells per mole of carbon) use of
resources. Oligotrophs focus in on quality then quantity therefore they would be less
likely to be wasteful with their substrate. Based on these strategies, the prediction is that
there is a negative correlation between BGE and rrs copy number.

This model was tested on a collection of soil microorganisms isolated from the
KBS LTER and isolates from rice paddy soils from Tsutomu Hattori (Institute of Genetic
Ecology, Tohoku University, Sendai, Japan). T erriglobus roseus strain KBS 63 is a
recently cultivated and characterized members of the phylum Acidobacteria that harbor
traits suggesting an oligotrophic lifestyle such as low rrs copy number (1 or 2), slow
growth rate, and abundance in low nutrient environments such as soil (8). As a reference,
Escherichia coli REL 607 (y -Proteobacteria), our model copiotroph, and Spingopyxis
alaskensis RB2256 (01- Proteobacteria), our model oligotroph, were used in the analysis.
The rrs copy number of the test strains has previously been determined as well as their
phylogeny. When possible, related pairs of the isolates (i.e. a copiotroph & oligotroph of

the B-Proteobacteria) were used to reduce the threat of shared evolutionary histories.

MATERIALS and METHODS
Description of bacterial strains Strains used in this study were obtained from long
term plating experiments from either soil at the WK. Kellogg Biological Station Long
Term Ecological Research site (KBS LTER) or rice paddy ( 10, 12, 13). Escherichia coli

strain REL 607, a derivative of E. coli B/r was a gift obtained from Richard E. Lenski

208

(19), EC2 & LC9 were gifts obtained from Joel A. Klappenbach isolated from the KBS
LTER (12, 13), HF3 and HF2 were gifts obtained from T. Hattori isolated from rice
paddy soils (10), and Sphingopyxis alaskensis RB2256 (formally Sphingomonas), a
marine ultramicrobacterium, was a gifi obtained from R. Cavicchioli and M. Ostrowski
(7, 25, 34, 35). These phylogenetically diverse strains were divided into two categories,
copiotrophs and oligotrophs: copiotrophs were defined as having a rrs copy number of

three or more, whereas oligotrophs were defined as having a rrs copy number of two or

less (Table D.1).

Medium Composition and Incubation Conditions Replicate 750 mL side-arm
flasks of all the strains were grown at room temperature (ca. 23°C), in 30mL of VSB-6
(Terriglobus roseus KBS 63) or -7 (all other strains) (8) with glucose (2.5mM) as the
primary carbon and energy source under yield-reduced conditions, supplemented with
Sug/mL yeast extract (BD, Franklin Lakes, NJ), and shaken on an orbital shaker (ca. 225
rotations/min). Yield-reduced conditions are defined as the limiting factor of cellular
yield where the amount of growth is proportional to the amount of carbon. This ensured
a decreased chance of a build-up of storage materials such as glycogen and
polyhydroxybutarates and promoted complete utilization of the 14C glucose label. A
magnetic stir bar was present during the growth of all strains in the 750 mL side-arm
flask to ensure successful processing of the sample (below). Typical inocula consisted of
a 1:100 dilution of cultures in mid/late log phase (ca. 1x108 cells/mL) growing on the
VSB-6,7 with 2.5 mM glucose and Sug/mL yeast extract. The optical density was

monitored periodically with a ThermoTM Spectronic 20D+ at 600 nm.

209

 

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Addition of 14C label The 14C labeled-glucose was added to the culture vessel
once the culture reached early-log phase, typically having an optimal density of ca. 0.05
at 600 nm. Uniformly labeled 0.1 umoles of glucose, 14C-UL-glucose (Sigma-Aldrich,
St. Louis, Missouri), was added with a time zero activity of ca. 444,000 counts per
minute (CPM). After three, 100 11L samples from each vessel were taken for the time
zero measurement; the flasks were sealed and allowed to grow until they reached the

transition point between log and stationary phase.

Processing of 14C Glucose The replicate flasks were processed to separate and
quantify the labeled biomass, carbon dioxide, and spent-medium once the culture reached
the transition point between log and stationary phase. The flasks were connected to a
series of hoses to vent the 14C02 in the headspace into three C02 trap in tandem
consisting of phenylethylaminezmethanol (1:1 vol/vol) (Figure D2). The headspace of
each flask was vented with nitrogen gas, briefly, for ca. 5 to 10 minutes to ensure that any
labeled l4C02 in the headspace was trapped. Venting was stopped, and a 5 mL aliquot of
the medium was removed from the flask and distributed into five, 1 mL aliquots in 1.5
mL microcentrifuge tubes. These microcentrifuge tubes were spun at 4°C at 16,000 g for
25 minutes which allowed separation of the biomass from the spent-medium. The
medium was decanted into scintillation vials and the biomass was resuspended in the
scintillate. Simultaneously, venting with nitrogen gas continued and trichloracetic acid
(final concentration of 2% (vol/vol)) was added to the culture to acidify the medium to
convert any dissolved CO; in the medium to gaseous form thereby allowing it to be

trapped in the phenylethylaminezmethanol C02 trap. The replicate flasks were vented for

211

Addition of
trichloroacetic acid
Nitrogen Gas

 

 

Bacterial culture 0' o'
phenylethylaminezmethanol (1 : 1) trap

 

 

 

 

 

Figure D.2. Methodology for trapping l4C-labeled carbon dioxide in the bacterial growth
efficiency experiments. Approximately 5mL ,of a 10% (vol/vol) tricholoracetic acid
solution was added to the 30 mL of culture to acidify, thereby allowing the dissolved
carbon dioxide to convert into a gaseous form. The gaseous carbon dioxide was trapped
with a carbon dioxide trap (phenylethylamine and methanol, 1:1 (vol/vol)) upon venting
the headspace with nitrogen gas.

212 V

 

a period of 3.5 to 4 hours. Triplicate one milliliter aliquots of each of the C02 traps were
measured for activity.

The labeled biomass, carbon dioxide, and spent-medium was quantified using a
Beckman Coulter LS 6000TA Liquid Scintillation System (Fullerton, California),
courtesy of the Dr. Ronald J. Patterson lab using Biosafe 11 (Research Products
International Inc., Mt. Prospect, Illinois) as the scintillate. All solutions in which the 14C
radioactive label was present were assessed for their capacity to quench the signal. The
only solution which indicated a significant affect (p < 0.002) of quenching was the

phenylethylamine:methanol C02 trap, and a correction factor of 3.38% was used.

Data Analysis In order to fully account for the 14C label, the biomass, carbon
dioxide, and spent-medium were summed and no analysis was run unless 2 96% of the
label could be recovered. During the development and use of this technique, attaining
full recovery of the 14C label was problematic. Numerous replicates of the strains were
processed to achieve 3 96% recovery. Typically, incomplete recovery was due to the loss
of carbon dioxide either during incubation of the strains or processing of the l4C-labeled
carbon dioxide, since the biomass and spent-medium counts were consistent.

BGE was determined using the values in the biomass and carbon dioxide
measurements. Briefly, the activity per umole of carbon (CPM/umole C) was
determined by taking the time zero measurement, and dividing it by the total amount of
carbon added in umoles (2.5mM glucose = ca. 75 umoles). It was then divided by six

since there are six carbons in glucose, in order to determine the specific activity of one

213

umole of carbon, which was then used to determine the umoles of carbon in the biomass
and umoles of carbon in the carbon dioxide by dividing the total activity for each by the
specific activity of one umole of carbon. The amount of carbon in ug could then be
determined from umole of carbon, which was the value used to determine BGE. BGE
was determined using the following equation: (ug of carbon in biomass)/ (ug of carbon

in biomass + 11g of carbon in carbon dioxide).

RESULTS
Description of bacterial strains Bacterial strains were obtained from the previous
soil cultivation experiments from the KBS LTER (8, l3) and rice paddy soils from
Tsutomu Hattori (Institute of Genetic Ecology, Tohoku University, Sendai, Japan).
Additional strains including Verrucomicrobium spinosum, Escherichia coli strain REL
607 (y -Proteobacteria) (our model copiotroph) and Spingopyxis alaskensis (formely
Spingomonas alaskensis) RB2256 (01- Proteobacteria) (our model oligotroph) were used
in the analysis. Strains were selected based on their phylogenetic affiliation, rrs copy
number, and ability to grown in VSB-6 or 7. These strains were separated into two
groups: oligotrophic bacteria and copiotrophic bacteria. Oligotrophs were defined as
having an rrs copy number of two or less, whereas copiotrophs were defined as having a
rrs copy number of three or more. The number of rrs operons per genome were
experimentally determined previously (8, 13) using non-radioactive Southern
hybridizations. Specific phylogenetic affiliations included members of the following
phyla: Acidobacteria, Verrucomicrobia, Bacteriodetes, and a, B, and y Proteobacteria

(Figure D.3). When possible, related pairs of the isolates (i.e. a copiotroph & oligotroph

214

rrs copy #

Bur.cepaci (M22518)

__1 Vrv.pardox (D30793) B
. 5
7

 

 

HF2-$21012 (084564)

 

 

 

 

 

 

 

 

 

 

 

 

Escherichia coli (A 3000129) g
Yers. pcstis (M232222) E
— Ps.stutzer (U26262) 7 5
Ps. putida (A8016428) E
HF3-$21027 (AY337597) ‘ 4 i
_{ Caulob. crescentus (AF125194) a
_ Sphingvpyfls alaskensis strain R 8225 6 (2.73631 ) I
KBS-LC9 (AY337604) 2 a
4— Sph.comtns (X91814) Bacteroldetes
KBS-EC2 (AF183150) 6
Pd.heprins (M11657)

 

—:Acbt.capsl (D261 71) Acidobacteria

Terriglobus roseus strain K8363 (1)0660892) 2 4

 

l'c’rrucmnicmbimn spinasum (X59721) l

 

 

Prsb.fusfm (U60015) Verrucomicrobla
env.slT62 (294005)

 

0.10

Figure D.3. Neighbor-joining phylogenetic tree'of the 168 rRNA gene of strains used to
determine bacterial growth efficiency. Strains from this study are in bolded text; bolded
in black represent copiotrophs (n=4) whereas bolded in gray represents oligotrophs (n=4).
The rrs copy number and major phylogenetic affiliations are illustrated to the right of the
phylogenetic tree. The scale bar indicates 0.10 changes per nucleotide.

215

of the B-Proteobacteria) were used to reduce the potential of shared evolutionary
histories; however this was not successful for some genera due to several strains’

inability to grow in the medium.

Bacterial growth efficiency, cell yield, and growth rates Strains were grown
on VSB-6 or 7 with glucose as the sole carbon and energy source under yield-reduced
conditions to determine bacterial growth efficiency, cell yield, and grth rates. The
results are illustrated in Table D1 In general, the oligotrophs had growth rates ranging
from 0.04 — 0.14 hr '1 whereas the copiotrophs growth rates ranged from 0.18 — 0.37 hr '1.
A significant (p < 0.05) positive correlation existed between bacterial grth rates and
the rrs copy number, where 50% of the data could be explained by the rrs copy number
(Figure D4).

The BGE for each organism was determined under saturating conditions of
glucose (final concentration, 2.5mM glucose) by comparing the amount of 14C in the
biomass over the total 14C consumed. BGE among all organisms tested ranged between
0.29 to 0.52. There was no relationship between bacterial growth efficiency and rrs copy
number (Figure D.5). As expected, there was a significant relationship (p < 0.006)
between cellular yield and bacterial growth efficiency where 87% of the differences in

cellular yield could be explained by the organism’s BGE.

Phylogenomic analyses of ATP-dependent genes and response regulators

01i gotrophs harbor traits including, but not limited to, reduced growth rate, increased

216 p

0.40 —

0.35 - E

0.30 _

I;

  

Growth rate (hr'l) +/- standard deviation
.o .o .o
H N N
L11 0 LII
1 m l L l

 

 

f
=0.7x+0.8
010—h y
4 I p<0.05
0.05— I
0.00 ,, ..... ,
1 2 4 8

rrs copy number

Figure D.4. Relationship between bacterial growth rate and rrs copy number. Each point
represents the average growth rate from two replicates grown in the VSB-6,7 with
2.5mM glucose and Sug/mL yeast extract at room temperature.

217

2600 —r

2400 ...

2200 ->

2000 -

   

 

1800-
, y = -350 + 7744x -4980x2
1600- R2 = 0.87
p < 0.006
1400— I
1200 - r - . .

 

0.25 0.30 0.35 0'40 ' 0'45 ' 0'50 ' 0.55
Bacterial growth efficiency

Cell yield (ug of carbon in biomass/flask) +/- stud. dev.

0.55 :E
0.50 5 u i
'1

0.45 7
0.40 ‘
0.35 ‘

0.30 ‘

Bacterial growth efficiency +/- stud. dev.
l-I-I
I

 

 

0.25

171' I fT T I l'

1 2 - 4 8
rrs copy number

n
_

Figure D.5. Panel A depicts the relationship between bacterial growth efficiency and cell
yield. Panel B depicts the relationship between bacterial growth efficiency and rrs copy
number.

218

body size, and presumed higher efficiency in resource consumption. Their presumed
high metabolic efficiency in resource utilization results from adapting to environments
where carbon is scarce. One adaptation to this environment would be their high affinity
uptake systems allowing these oligotrophs to take up carbon when they are scarce; in
order to transport carbon against the gradient into the cell for growth, active transport
systems such as ATP-binding cassette (ABC) transporters, are required. The relationship
between the proportion of ATP-dependent proteins out of the total protein versus rrs
copy number was evaluated. The hypothesis was that transporters in the genomes of
these oligotrophs would have a higher percentage of ATP-binding cassette (ABC)
transporters, and there would be a negative correlation between rrs copy number and the
percentage of ATP-dependent transporters out of the total proteins. The ability of
oligotrophs to transport carbon for growth provides them with the capacity to uptake and
therefore use this carbon, however it comes with an energetic cost which will decrease
efficiency. The trade-off is that copiotrophs with less energy-dependent transporters
would not be able to survive in these low nutrient environments because of the
diminished capacity to uptake carbon.

This hypothesis was tested by analyzing 133 readily available data of sequenced
non-symbiotic, heterotrophic, bacterial genomes from the TransportDB
(http://www.membranetransportorg) (31), specifically calculating the total number of
ATP-dependent proteins out of the total number of proteins. There was a significant
negative correlation with the percent abundance’of ATP-dependent proteins and rrs copy
number (p < 0.0001). However, only 18% of the increase in ATP-dependent proteins can

be explained by a decrease in rrs copy number (Figure D.6). Bacteria harboring a low

219

Percent of ATP-dependent proteins (out of total proteins)

80- I

70-

 

  

60

y = -4.2x + 47.1
- R2 = 0.18
p < 0.0001

50

   

 

40

30

20

10I ......, ......., ........ .......
1 2 4 8 16

rrs copy number

Figure D.6. Percent of ATP-dependent proteins (out of the total proteins) versus rrs copy
number for non-symbiotic heterotrophs. Data obtained from 133 sequenced genomes
from the TransportDB (http://www.membranetransportorg) (31).

220

rrs copy number have a higher portion of their genome dedicated to ATP-dependent
transporters which allow them to survive in low nutrient environments. This relationship
clearly illustrates an intrinsic property, ATP-dependent proteins, which reflect a different
ecological strategy for bacteria.

Another intrinsic property that was assessed to determine any correlation with rrs
copy number was the number of response regulators in two-component systems in
bacterial genomes, not associated with chemotaxis. Two-component systems are
pathways some bacteria use to sense environmental stimuli through the use of a histidine
kinase sensor protein typically in the periplasm that transmits a signal to the carboxyl-
terminal of the response regulator (22). The response regulators typically are DNA-
binding, transcriptional regulators which allow the transcription of the necessary genes in
response to the environmental stimuli (3). Copiotrophs harbor traits including, but not
limited to, high growth rate and abundance in nutrient rich and fluxing environments. It
was hypothesized that these bacteria would have to dedicate a high portion of their
genome to sense changes in these fluxing environments such as response regulator-
encoding genes, which would lend to a positive correlation between rrs copy number and
the percent of the genome devoted to response regulator genes. Response regulators
associated with chemotaxis are excluded in. the analysis because the property of
chemotaxis allows them to physically change their surrounding environment, thereby not
directly linking gene function with environmental stimuli. This hypothesis was tested by
analyzing 187 readily available data of sequenced non-symbiotic, bacterial genomes from

Response Regulator Census on the NCBI website

221

(http://www.ncbi.nlm.nih.gov/Complete Genomes/RRcensus.html), Specifically

 

calculating the percent of response regulator genes out of the total genes in each of the
respective genome. There was a significant positive correlation with the percent
abundance of response regulator genes and rrs copy number (p < 0.0001). Bacteria
harboring a high rrs copy number have a higher portion of their genome dedicated to
sensing changes in the environment. The increase in rrs copy number explains ca. 20%
of the increase in response regulators, which is “as hypothesized, where the bacteria with
high rrs copy number have the capability of sensing a fluctuating environment (Figure

D.7).

DISCUSSION and FUTURE DIRECTIONS

The primary goal of this study was ‘to determine if there was a negative
correlation between bacterial growth efficiency and rrs copy number, the genetic marker
for ecological strategies in bacteria, among a phylogenetically diverse group of bacteria.
Under saturating concentrations of glucose, there was not an observable trend between
bacterial growth efficiency and rrs copy number (Figure D.5). One explanation for this
result could be due to intrinsic differences in the metabolic pathways to oxidize carbon,
synthesize vitamins, enzymes, co-factors, and create reducing power for energy
production (Table D2). There is no available metabolic information on these organisms
to address this concern. For the purposes of these experiments, it was assumed that all
matters pertaining to growth among these strains was equal. Furthermore, the
environmental conditions in which these strains were grown, in particular pH and

temperature, could be metabolically costly for some organisms to grow due to a possible,

222

A

U)

E-i

:1)

OD 2.0 I
76 I

H

0

:3

VJ

5 15

H

E

:3

ea

8

g) 1.0

(D

1:

8-

8

c... 05

o -../g I
H I
5 I

3 .
o I

D- 0.0 I I

....E i.

y = 0.16x + 0.31
R’=0.20
p<0.0001

 

I Hi

4

l I I i ‘ rTTU'

rrs copy number

Figure D.7. Percent of response regulators in two-component systems not associated
with chemotaxis, versus rrs copy number for bacteria and archaea. Data obtained 187

sequenced genomes

obtained

from TIGR

(http://www.ncbi.nlm.nih.gov/Complete Genomes/RRcensus.html).

223 .

Table D2. Intrinsic metabolic features and environmental factors that influence bacterial

 

 

 

 

growth efficiency.
Metabolic Features Proposed Mechanisms Reterencem
. Varying proportions of transporters that consume
ACthC transport (23)
ATP or IMF
_ . Consumption of IMF in flagellar motility or synthesis
Mommy . . . .. (29)
of excreted compounds 1n gliding mot1lrty
rrs copy number Fast (high rrs) growth vs. slow (low rrs) growth (13, 37)
Tradeoff between speed and accuracy of translational
Translational machinery machinery influences amount of functional protein (18)
synthesized per mole of ATP
_ Tradeoff between rate and efficiency of analogous
Pathway utilization (11)
pathways
_ . . Consumption of IMP or ATP to maintain internal pH
Maintenance of ion gradient _ . _ (29)
or intracellular ion concentrations
Synthesis of extracellular Extracellular enzymes or matrix materials are not (24)
macromolecules typically measured as biomass
Half life of proteins and ribosomes dictates demand
Turnover of macromolecules . (32)
for new synthe51s of these macromolecules
Synthesis and utilization of storage Variable energy costs or savings associated with (6)
compounds intracellular storage of C, S, N, and P
ATP-producing pathways Trade-off between rate and yield of ATP (27)
Environmental Factors
Free energy of substrates Limits maximal ATP yield (6)
, . Cost of assimilatory reduction of C, S, N, and P for
0x1datron state of substrates (6)
biosynthesis
Increase ATP demand for transport at low substrate
Substrate concentration concentrations; energy spilling at excess (4)
concentrations
. High ratio demands carbon is oxidized before
Carbon: Nitrogen , (15)
nitrogen can be used for biosyntheSIS
. _ _ Demand for synthesis of compounds not available
Substrate avallability . (6)
from the env1ronment
Synthesis of macromolecules required for
Deviation from a microbe‘s optimal growth/survival in suboptimal conditions; costs (26)

growth conditions

associated with removing or degrading inhibitory

compounds

 

224

deviation from their optimal conditions. The varying degrees of these demands could
negatively influence the bacterial growth efficiency calculation by indicating an
adversely low value since more carbon would be diverted to non-growth related functions
to compensate for these non-ideal conditions. One could speculate that the sample size
would need to be rather large in subsequent experiments to overcome this variability to
reveal true differences reflected by an organism’s rrs copy number.

Another explanation for the lack of correlation between bacterial growth
efficiency and rrs copy number is due to the saturating glucose conditions under which
the efficiency measurements were made. Oligotrophs are typically found in stable
environments where carbon concentrations are low; therefore, one could predict it would
be under these low, but stable nutrient concentrations where an oligotroph would
maximize their efficiency. The efficiency experiments were performed under saturating
concentrations of carbon, therefore masking any meaningful relationship between
efficiency and low rrs copy number.

In order to address this concern, the next step in this research would be to grow
these strains under stable, low nutrient concentrations and measure their bacterial growth
efficiency. Oligotrophs are typically more abundant in low nutrient environments;
therefore, they have evolved to be more fit under these conditions. If oligotrophs are
more efficient at utilizing substrates, it would be plausible to assume that they would do
so under conditions typical of their native environment. Furthermore oligotrophs
typically have a higher portion of ATP-dependent transporters (Figure D6) which would
allow them to uptake carbon in carbon-poor environments. Strains would need to be

grown using a chemostat whereby the population density is controlled by the dilution rate

225

which is a reflection of the nutrient concentration in the system. The hypothesis would
be that oligotrophs reach their maximum growth efficiency at lower dilution rates,
reflective of a low nutrient environment, as compared to copiotrophs reaching their
maximum efficiency at high dilution rates or nutrient rich environments (Figure D.8,
panel A). The model would be that there is a positive correlation between rrs copy
number and resource concentration (as measured by dilution rate in a chemostat) at which
the organism reaches its maximum growth efficiency (Figure D.8, panel B).

To further understand this proposed study, it would be beneficial to predict why
copiotrophs have lower bacterial growth efficiencies at low nutrient concentrations.
When a bacterium oxidizes carbon, the energy obtained is either use for growth or non-
growth. Energy used for non-growth related functions is typically defined as
maintenance energy. It is believed that the demand for maintenance energy increases the
firrther away from optimal conditions. Maintenance energy can be quantified through the
use of Pirt’s double reciprocal plots (l/yield versus l/growth rate) (28, 29).

Under steady state, low nutrient concentrations, the maintenance demands would
have a greater effect in terms of efficiency in copiotrophs than oligotrophs because it is
an environment not conducive to fast growth. The proposed hypothesis is that
maintenance energy is lower in oligotrophs than copiotrophs as measured by a double
reciprocal plot of the inverse of yield and growth rate under steady state, low nutrient
conditions (Figure D.9, panel A) at dilution rates below maximum efficiency.
Copiotrophs would have a larger slope than oligotrophs because the demand for

maintenance energy would be higher under these low nutrient conditions. The model

226

 

 

 

 

Oligotrophs
Copiotrophs

m

O

m

Diultion rate

B.

2

1";

Cl

.9

E

Q

 

 

rrs copy number

Figure D.8. Proposed model for future bacterial growth efficiency experiments. Panel A
depicts the proposed relationship between bacterial growth efficiency (BGE) versus
dilution rate. Panel B depicts the proposed linear relationship between dilution rate, a
proxy for resource concentration indicating rate at which the organism reaches its
maximum growth efficiency (Panel A) measured in a chemostat, versus rrs copy number.

227

Copiotrophs

1/Y Oligotrophs

 

 

I/u

Maintanence energy

 

 

rrs copy number

Figure D.9. Proposed model for future bacterial growth efficiency experiments. Panel A
depicts the proposed relationship for maintenance energy for oligotrophs and copiotrophs
as illustrated by Pirt’s double reciprocal plots of the inverse of yield (l/Y) versus the
inverse of grth rate (1/ u) at steady state.‘ Panel B depicts the proposed linear
relationship between maintenance energy of oligotrophs and copiotrophs grown at steady
state dilution rates versus rrs copy number.

228

would be that there is a positive correlation between maintenance energy, as measured by
the slope of the double reciprocal plot, and rrs copy number (Figure D.9, panel B)
meaning there would be a lower demand in maintenance energy in oligotrophs when
growing under their ideal conditions The juxtaposition of these experiments, as
mentioned above, is that one could speculate that the sample size would need to be rather
large in subsequent experiments to overcome the variability of different physiological as
well as environmental attributes of the strains to reveal true differences in efficiency
reflected by rrs copy number.

The foundation of this research brings an ecological theory into a microbe’s
perspective. It is a unifying step of multiple disciplines: microbiology, ecology, and
evolution. Understanding bacteria’s ecological strategies in the environment is necessary
to appreciate the community’s dynamics especially when only a small fraction of the soil
microbial community has been cultivated, such as the members of the phylum
Acidobacteria. By knowing a bacteriurn’s efficiency as well as the environmental
regime, we will be able to make predictions of the impacts certain communities have on

nutrient cycling, biogeochemistry, and agricultural practices.

ACKNOWLEDGEMENTS
This work was supported by the National Science Foundation (IBN 9875254).
I would like to thank Zarraz May-Ping Lee for her helpful discussions on

ribosomes and bacterial grth efficiency.

229

 

10.

11.

12.

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