Identification and Cultivation of Exopolysaccharide-Degrading Bacteria in Two Soils

University of Calgary PRISM: University of Calgary's Digital Repository

Graduate Studies The Vault: Electronic Theses and Dissertations

2013-09-24 Identification and Cultivation of Exopolysaccharide-Degrading Bacteria in Two Soils

Wang, Xiaoqing

Wang, X. (2013). Identification and Cultivation of Exopolysaccharide-Degrading Bacteria in Two Soils (Unpublished master's thesis). University of Calgary, Calgary, AB. doi:10.11575/PRISM/26419 http://hdl.handle.net/11023/1033 master thesis

University of Calgary graduate students retain copyright ownership and moral rights for their thesis. You may use this material in any way that is permitted by the Copyright Act or through licensing that has been assigned to the document. For uses that are not allowable under copyright legislation or licensing, you are required to seek permission. Downloaded from PRISM: https://prism.ucalgary.ca UNIVERSITY OF CALGARY

Identification and Cultivation of Exopolysaccharide-Degrading Bacteria in Two Soils

Xiaoqing Emily Wang

A THESIS

SUBMITTED TO THE FACULTY OF GRADUATE STUDIES

IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE

DEGREE OF MASTER OF SCIENCE

DEPARTMENT OF BIOLOGICAL SCIENCES

CALGARY, ALBERTA

SEPTEMBER, 2013

In this study, we hypothesized that bacterial exopolysaccharides (EPS) may serve as energy substrates for K-selected bacteria in soils, and we would be able to identify and culture previously uncultured species using various EPS as growth substrates.

The exopolysaccharides gellan, indican_2, and cellulose were produced by the bacteria

Sphingomonas elodea (ATCC 31461), Beijerinckia indica (ATCC 9039) and Gluconacetobacter xylinus (ATCC 53524) respectively. Two experiments were performed using these exopolysaccharides (EPS) as growth substrates. The first experiment involved detecting EPS- degrading bacteria by using a stable isotope probing technique. 13C labeled EPS were used as growth substrates for Big Hill Spring (BHS) and Paint Pots mound (PP) soil samples. EPS- degrading bacteria were identified using DNA density ultracentrifugation coupled with 454- pyrosequencing of the 16S rRNA genes recovered from 13C-labelled DNA fractions. Several uncultured Xanthomonadaceae sp. were enriched on cellulose (produced by G. xylinus) during incubation from BHS soil and several previously uncultured Planctomycetes were highly enriched on indican_2 (produced by B. indica) incubation from PP soil. The second experiment was to isolate novel bacteria by using gellan or indican_2 as sole carbon sources. Based on cultivation results, we found several previously uncultured bacteria could be identified and isolated from our gellan (produced by S. elodea) and indican_2 plates. For instance, we successfully isolated previous uncultured Acidobacteria by using gellan and indican_2 as our carbon source.

All in all, in this study, we found some previously uncultured bacteria belonging to the groups of Acidobacteria, Verrucomicrobia, Planctomycetes, Chloroﬂexi, Actinobacteria,

Cyanobacteria, Proteobacteria, Bacteroidetes, Armatimonadetes and Candidate division OD1,

ii which are readily enriched or cultured using these EPS substrates. These poorly understood groups, especially some members of Planctomycetes, showed preference for our EPS indican_2.

The research results indicate that using novel EPS as carbon sources might be a new way to improve current cultivation techniques.

iii Acknowledgements

I am grateful to the many people who helped and encouraged me to be successful.

Without their support, this thesis would not be possible. In particular, I would like to express my deepest gratitude to my supervisor, Dr. Peter Dunfield, for providing me the opportunity to work on this interesting project. To Dr. Peter Dunfield, thanks for always believing in me and allowing me a chance to grow. Your knowledge and insights guide me through the project. Thanks to the members of my committee Dr. Lisa Gieg, Dr. Raymond Turner, and Dr. Kenneth Sanderson for taking the time to support and encourage my academic efforts. To Dr. Kenneth Sanderson, thanks to encourage me when I am feel lack of confidence at very beginning of my master study, your support and care means a lot to me. To Dr. Allyson Brady and Christine Sharp, without your assisting me with so many details of this research, this research would not have been possible. You were not only taught me many hands-on experimental skills, the knowledge of research, you also helped me to know Canada. You bring a lot of first time to me, first Camping, first ski, first make camp fire...thanks you guys to bring such wonderful experiences to me. I will be forever grateful! Thanks to Gareth Jones for all cellulose supports and all the grammar corrections, without his help this thesis could be done. Thanks all the members in the Dunfield lab Dr. Angela Smirnova, Dr. Joongjae Kim, AliReza Saidi-Mehrabad, Fauziah Rochman and

Roshan Khadka, for sharing their experience and knowledge.

Finally, I wish to show my appreciation to my family and many wonderful friends for their love and support. To my Dad and Mom, without your love and supports, I cannot finish this study. To my fiancé Mark Zhong, thanks for always be with me in those difficult time. This research would not have been possible without you all. Thank you!

iv Table of Contents

Abstract ...... ii Acknowledgements ...... iv Table of Contents ...... v List of Tables ...... viii List of Figures and Illustrations ...... x

CHAPTER ONE: THESIS INTRODUCTION ...... 1 1.1 Introduction ...... 1 1.2 Hypothesis ...... 4 1.3 Research Objectives ...... 4 1.4 Study Sites ...... 4

CHAPTER TWO: LITERATURE REVIEW ...... 8 2.1 Known and Unknown Microbial Diversity ...... 8 2.1.1 Culture-Independent Methods Overview ...... 8 2.1.1.1 Total Nucleic Acid Extraction from Soil ...... 8 2.1.1.2 PCR of the 16S rRNA Gene ...... 9 2.1.1.3 Fingerprinting Method ...... 10 2.1.2 Abundance and Diversity of Soil Bacterial Communities ...... 13 2.2 r- and K-Selection Theory ...... 18 2.3 Microbial cultivation methods ...... 19

CHAPTER THREE: CHARACTERIZATION OF SOIL MICROBIAL COMMUNITIES24 3.1 Introduction ...... 24 3.2 Methods ...... 24 3.2.1 Sampling ...... 24 3.2.1.1 Paint Pots Site ...... 24 3.2.1.2 Big Hill Spring Site ...... 24 3.2.2 Extraction of Total DNA ...... 25 3.2.3 PCR Amplification, Purification and Quantification ...... 27 3.2.4 Microbial Community Analyses ...... 28 3.3 Results and Discussion: ...... 28 3.3.1 Previous Paint Pots Microbial Community Research Review ...... 28 3.3.2 Big Hill Spring Microbial Community...... 29

CHAPTER FOUR: DETECTION OF AS-YET-UNCULTURED BACTERIAL GROUPS IN BHS AND PP SOIL USING STABLE ISOTOPE PROBING ...... 32 4.1 Introduction ...... 32 4.2 Materials and Methods ...... 35 4.2.1 13C and 12C EPS Production ...... 35 4.2.1.1 Cellulose Production ...... 35 4.2.1.2 Indican_2 Production ...... 36 4.2.1.3 Gellan Production ...... 36 4.2.2 EPS Extraction ...... 38

v 4.2.2.1 Cellulose Extraction ...... 38 4.2.2.2 Indican_2 Extraction ...... 40 4.2.2.3 Gellan Extraction ...... 42 4.2.3 SIP Incubation and CO2 Production Measurement ...... 44 4.2.4 DNA Extraction, Fractionation, and Quantification ...... 47 4.2.5 16S rRNA Gene Pyrotag Sequencing ...... 48 4.2.6 Microbial Communities Analyses ...... 49 4.3 Results ...... 51 4.3.1 CO2 Production Rate ...... 51 4.3.1.1 Cellulose incubation CO2 Production ...... 51 4.3.1.2 Indican_2 incubation CO2 Production ...... 51 4.3.1 13C-EPS Incubation Results ...... 52 4.3.1.1 13C-Cellulose incubation DNA gradients ...... 52 4.3.1.2 13C-Indican_2 incubation DNA gradients ...... 52 4.3.1.3 13C-Cellulose incubation sequencing results ...... 56 4.3.1.4 13C-Indican_2 incubation ...... 65 4.4 Discussion ...... 76 4.4.1 Cellulose Incubation ...... 76 4.4.2 Indican_2 Incubation ...... 80 4.4.3 Potential Errors ...... 83

CHAPTER FIVE: ISOLATION OF SOIL BACTERIA ON EPS-SUPPLEMENTED MEDIA...... 86 5.1 Introduction ...... 86 5.2 Materials and Methods ...... 86 5.2.1 12C EPS Production ...... 86 5.2.1.1 Indican_2 Production ...... 86 5.2.2 EPS Extraction ...... 87 5.2.2.1 Indican_2 Extraction ...... 87 5.2.2.2 Gellan Extraction ...... 87 5.2.3 Cultivation Experiments ...... 88 5.2.4 Identiﬁcation of Colonies ...... 89 5.3 Results and Discussion ...... 90 5.3.1 Indican_2 Supplemented Medium ...... 90 5.3.1.1 DNMS5.8+Indican_2 ...... 90 5.3.2 Gellan Supplemented Medium ...... 94 5.3.2.1 DNMS 5.8+gellan ...... 94 5.3.2.2 DNMS 3.8+gellan ...... 98 5.3.3 Culturability Factors ...... 102

CHAPTER SIX: CONCLUSIONS ...... 107

REFERENCE ...... 110

APPENDIX A: MEDIA ...... 129

APPENDIX B: EPS STRUCTURE ...... 134

vi Cellulose structure ...... 134 Indican_2 Structure : ...... 134 Gellan structure: ...... 134

APPENDIX C: PERMISSIONS ...... 135

vii List of Tables

Table 2.1 Summary of common fingerprinting methods (Spiegelman et al. 2005; Liesack and Dunfield 2002; Suzuki et al. 1998)...... 12

Table 2.2 Contribution of 16S rRNA and 16S rRNA genes from members of different phyla and subphylum groups (class, subdivision, or subclass) to soil bacterial communities (Janssen 2006) (see Appendix C for permission)...... 14

Table 2.3 Some attributes of r-and K-selected microorganisms (Langer et al. 2004)...... 19

Table 3.1 Sample description ...... 25

Table 3.2 The top six most abundant phyla across all research soil samples (as a percentage of all reads)...... 30

Table 4.1 Incubation Setting Information ...... 45

Table 4.3 The most abundant (>1%) OTUs in the heavy fractions of PP soil incubated with 13C-Cellulose, compared to their abundances in the original unincubated PP soil (as a percentage of all reads)...... 63

Table 4.4 The most abundant (>1%) OTUs in BHS with 13C-Indican_2 incubation compared to the original unincubated BHS site abundance (as a percentage of all reads)...... 67

Table 4.5 The top ten abundant species in PP soil with 13C-Indican_2 incubation compared to the original unincubated PP site abundance (as a percentage of all reads)...... 72

Table 4.6 Taxonomic descriptions of the top six uncultured Planctomycetes species in PP with 13C-Indican_2 incubation compare to the original PP site abundance (as a percentage of all reads)...... 73

Table 4.7 OTUs related to known cellulose-degrading species list ...... 77

Table 5.1 Analysis of isolates obtained via ContagiExpress software on the basis of partial (~ 600 bp) 16S rRNA gene sequences, including the closest taxonomically validated relatives identiﬁed using the BLAST program in the NCBI databases...... 92

Table 5.2 Analysis of isolates obtained via ContagiExpress software on the basis of partial (~ 600 bp and use only 9f as sequencing primer) 16S rRNA gene sequences, including the closest taxonomically validated relatives identiﬁed using the BLAST program in the NCBI databases...... 96

viii Table 5.3 Analysis of isolates obtained via ContagiExpress software on the basis of partial (~ 600 bp and use only 9f as sequencing primer) 16S rRNA gene sequences, including the closest taxonomically validated relatives identiﬁed using the BLAST program in the NCBI databases...... 100

Table 5.4 Summary of the most taxonomically novel isolates obtained in this study...... 106

ix List of Figures and Illustrations

Figure 1.1 Map showing the location of the Paint Pots springs and the distribution of iron oxide deposits (Ochre beds) precipitated in the outﬂow stream and surrounding marsh (after van Everdingen 1970) (Grasby et al. 2013) (see Appendix C for permission)...... 6

Figure 1.2 Big Hill Springs study site. Photo source: Google (public domain)...... 7

Figure 3.1 The Paint Pots Springs site (A-D) and BHS site (E). A) Downstream of the spring, a bog fed by the spring water B) From a relic spring mound, C) Soil near the source spring, taken just under a pine tree D) Sediment in main spring pool soil, E) Under pine tree, taken after removed top pine leaves. Photos taken by Christine E. Sharp (A-D) and Peter F. Dunfield (E)...... 26

Figure 3.2 Relative abundance of different bacterial and archaeal phyla for each soil library. 16S rRNA gene sequences were classiﬁed according to the nearest neighbour in the SILVA108 database: (A) mound (based on 9919 reads), (B) Ochre bed bog (based on 6788 reads), (C) outflow marsh (based on 4386 reads), (D) soil (based on 6238 reads), (E) BHS Soil (based on 8990 reads)...... 31

Figure 4.1 Flow diagram of experimental procedure...... 35

Figure 4.2 Incubations of EPS producing bacteria A) Gluconacetobacter xylinus B) Beijerinckia indica C) Sphingomonas elodea...... 37

Figure 4.3 Step by step method for G. xylinus EPS cellulose extraction, after Schramm and Hestin (1954)...... 39

Figure 4.4 Step by step method for B.indica EPS (indican_2) extraction, modified from (Liu and Fang 2002)...... 41

Figure 4.5 Step by step method for S.elodea EPS (gellan) extraction, after Liu and Fang (2002) and Goh (2004)...... 43

Figure 4.6 The final EPS products A) Cellulose (the EPS product of G.xylinus strain) B) Indican_2 (the EPS product of B.indica strain) C) Gellan (the EPS product of S.elodea strain)...... 44

Figure 4.7 Bottles set up A) cellulose with soil incubation B) indican_2 with soil incubation.... 46

Figure 4.8 The flow diagram for DNA centrifugation and fractionation. Part of the figure is courtesy of Allyson Brady (Brady et al. 2011)...... 48

Figure 4.9 CO2 production over time in soils incubated with or without EPS. (A) BHS soil incubated with 12C-cellulose compared to BHS Soil control group. (B) PP soil incubated with 12C-cellulose compared to PP Soil control group. (C) BHS soil incubated with 12C- indican_2 compared to BHS Soil control group. (D) PP soil incubated with 12C-

x indican_2 compared to PP Soil control group. Bars=±SEMs.Where the error bars are not seen they are smaller than the symbols...... 53

Figure 4.10 Relative DNA concentrations recovered from CsCl gradient fractions for PP BHS soil (A) and PP soil (B) incubated with 13C-Cellulose or 12C-Cellulose control. Solid lines represent incubations with 12C-Cellulose control and dashed lines represent incubations with 13C-Cellulose. The x-axis indicates the relative DNA concentrations recovered from each gradient fraction, with the highest quantity detected in any gradient fraction set to 1.0. Relative concentration rather than absolute DNA copies counts are used to facilitate comparison across the different experiments. The arrows indicate the fractions sent for sequencing...... 54

Figure 4.11 Relative DNA concentrations recovered from CsCl gradient fractions for BHS soil (A and B) and PP soil (C and D) with 13C-Indican_2 and 12C-Indican_2 control. Solid lines represent incubations with 12C-Indican_2 control and dashed lines represent incubations with 13C-Indican_2. The x-axis indicates the relative DNA concentrations recovered from each gradient fraction, with the highest quantity detected in any gradient fraction set to 1.0. Relative concentration rather than absolute DNA copies counts are used to facilitate comparison across the different experiments. Experiments were performed in duplicate BHS soil (C) and PP soil (D). The arrows indicate the fractions sent for sequencing...... 55

Figure 4.12 Relative abundance of different bacterial phyla and archaeal phyla from BHS soil with 13C-Cellulose incubation in the heavy DNA fractions (density 1.73 g ml−1 and 1.74 g ml−1) compared to original BHS soil. 16S rRNA gene sequences were clustered based on 97% identity and classiﬁed via BLAST (Altschul et al. 1990) based on the SILVA108 database of bacterial and archaeal 16S rRNA gene sequences (Haas et al. 2011) ...... 57

Figure 4.13 16S rRNA gene sequence-based tree showing phylogenetic position of the most enriched cellulose-degrading bacteria identified in BHS soil (designated with prefix ‘Emily’ in bold) assembled as per the Neighbor-Joining method with a Jukes-Cantor correction. Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB-SILVA database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents 0.1 change per nucleotide position...... 60

Figure 4.14 Relative abundance of different bacterial phyla and archaeal phyla from PP soil with 13C-Cellulose incubation in the heavy DNA fractions (density 1.74 g ml−1) compared to original PP soil. 16S rRNA gene sequences were clustered based on 97% identity and classiﬁed via BLAST (Altschul et al. 1990) based on the SILVA108 Database of bacterial and archaeal 16S rRNA gene sequences (Haas et al. 2011)...... 62

xi correction. Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB-SILVA database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents 0.1 change per nucleotide position...... 64

Figure 4.16 Relative abundance of different bacterial phyla and archaeal phyla from BHS soil with 13C-Indican_2 incubation in the heavy DNA fractions (density 1.73 g ml−1 and 1.74 g ml−1) compared to original BHS soil. 16S rRNA gene sequences were clustered based on 97% identity and classiﬁed via BLAST (Altschul et al. 1990) based on the SILVA108 database of bacterial and archaeal 16S rRNA gene sequences (Haas et al. 2011)...... 66

Figure 4.17 16S rRNA gene sequence-based tree showing phylogenetic position of the most enriched indican_2-degrading bacteria identified in BHS soil (designated with prefix ‘Emily’ in bold) assembled as per the Neighbor-Joining method with a Jukes-Cantor correction. Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB-Silva database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents a 0.1 change per nucleotide position...... 69

Figure 4.18 Relative abundance of different bacterial phyla and archaeal phyla from PP soil with 13C-Indican_2 incubation in the heavy DNA fractions (density 1.74 g ml−1). 16S rRNA gene sequences were clustered based on 97% identity and classiﬁed via BLAST (Altschul et al. 1990) based on the SILVA108 database of bacterial and archaeal 16S rRNA gene sequences (Haas et al. 2011)...... 71

Figure 4.19 16S rRNA gene sequence-based tree showing phylogenetic position of the most enriched indican_2-degrading bacteria identified in PP soil (designated with prefix ‘Emily’ in bold) assembled as per the Neighbor-Joining method with a Jukes-Cantor correction. Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB-SILVA database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents a 0.1 change per nucleotide position...... 74

Figure 4.20 16S rRNA gene sequence-based tree showing phylogenetic position of the most enriched indican_2-degrading Planctomycetes bacteria identified in PP soil (designated with prefix ‘Emily’ in bold) assembled as per the Neighbor-Joining method with a Jukes-Cantor correction. Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB-Silva database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents a 0.1 change per nucleotide position...... 75

Figure 5.1 Flow diagram of the experimental procedure...... 87

xii Figure 5.2 Bacterial isolates obtained from gellan-containing media...... 89

Figure 5.3 BHS soil 10-4 dilution spread on indican_2 -containing DNMS 5.8 Plates from Day 1 to Day 27...... 93

Figure 5.4 Incubation of BHS soil 10-4 to 10-7 dilution after 2 weeks on indican_2 -containing DNMS 5.8 Plates...... 94

Figure 5.5 BHS soil 10-7 dilution spread on gellan-containing DNMS 5.8 Plates from Day 1 to Day 25...... 97

Figure 5.6 Incubation of BHS soil 10-6 dilution after 1 month on gellan-containing DNMS 5.8 Plates. Some colonies start to spread out and couple similar colonies appearing around a mother colony...... 97

Figure 5.7 PP soil 10-7 dilution spread on gellan-containing DNMS 3.8 Plates from Day 1 to Day 25...... 101

Figure 5.8 Incubation of PP soil 10-6 dilution after 1 month on gellan-containing DNMS 3.8 plates. As colonies spread out, smaller colonies sometimes appear around a mother colony...... 101

Figure 5.9 Picture was taken (from below) from a 10-7 PP soil + gellan plate after 3 month incubation (10× Magnification by SZ61 zoom stereomicroscope) Note the depressions in the surface under some colonies, presumably caused by the degradation of the gellan gel...... 103

xiii List of Symbols, Abbreviations and Nomenclature

Abbreviation Definition

ARDRA amplified ribosomal DNA restriction analysis

B. indica Beijerinckia indica

BHS Big Hill Spring bp base pair

C carbon

CFU colony-forming unit

CO2 carbon dioxide

DES DNase/Pyrogen-free water

DGGE/TGGE denaturing/temperature gradient gel electrophoresis

DNMS dilute nitrate mineral salts

EPS Exopolysaccharides

EPS* Extracellular polymeric substances

FID ﬂame ionization detector

G. xylinus Gluconacetobacter xylinus

GC gas chromatograph

GTC guanidine thiocyanate

HTC high-throughput culturing

LH-PCR length heterogeneity polymerase chain reaction

OTU Operational Taxonomic Unit

PCR polymerase chain reaction

PP Paint Pots mound

xiv QIIME quantitative insights into microbial ecology rRNA ribosomal ribonucleic acid

S. elodea Sphingomonas elodea

SDS sodium dodecyl sulfate

SIP Stable isotope probing sp. species

SSCP single-strand conformation polymorphism

T-RFLP terminal-restriction fragment length polymorphism

Chapter One: THESIS INTRODUCTION

1.1 Introduction

At one time, microbiologists believed that they could grow and identify all different bacterial species present in soil based on culturing methods (Conn 1909). However, several years later they realized that the methods that relied on cultivation retrieved only 1.5 to 10% of the bacterial cells in soil (Conn 1918). Fifty years later, Vagn Jensen (1968) also concluded that the bacterial colonies which were growing on agar plates were not the total bacterial community. In the early 1990s, based on PCR amplifications and sequencing of 16S rRNA genes from soil, microbiologists were able to identify bacterial species without the need for cultivation. The cultivation-independent methods demonstrated that only about 10% of all 16S rRNA clones recovered from soils can be assigned to a cultured genus (Janssen 2006). Although cultivation- independent methods have become more popular, cultured isolates still play a very important role in studying bacterial physiology, genetics, and ecology (Janssen 2006). Also because most bacteria are uncultured, they may be a source of antibiotics and other useful compounds (Keller and Zengler 2004).

At present, the Domain Bacteria is divided into 29 main groups or phyla based on 16S rRNA gene sequence similarity. Some phyla like Proteobacteria and Actinobacteria are well- known from previous cultivation studies. However, others like Acidobacteria, Verrucomicrobia and Planctomycetes are poorly studied and have few cultured species (Janssen 2006). Janssen

(2006) pointed out that most bacteria in nature are uncultured, and will not grow on standard rich nutrient media used in the laboratory. For example, based on analyses of 16S rRNA gene clone libraries, members of the phylum Acidobacteria typically represent about 20% of soil bacterial communities (Janssen 2006). This means they are widely distributed and abundant in soils. 1

However, they are currently poorly understood because they are difficult to grow in the lab

(Eichorst et al. 2007). This phylum has been proposed to consist of 26 subdivisions, and most of these subdivisions do not contain cultured representatives (Losey et al. 2013). There are only about 14 genera cultivated, and they belong to Acidobacteria subdivisions 1,3,4,8 and 10 (Losey et al. 2013). Similarly to Acidobacteria, the phylum Planctomycetes is also abundant in soil but poorly known and has few cultured species. Planctomycetes makes up an average of 2% of soil bacterial communities. However, until now, there are only 3 orders (Planctomycetales,

Phycisphaerales, and “Candidatus Brocadiales”), 11 described genera and 14 species, and 5

Candidatus genera with 14 Candidatus species (Lage and Bondoso 2010) have been reported.

One theory proposed to explain the inability to culture certain bacteria is that different species are r-selected or K-selected bacteria. Typical r-selected bacteria are organisms that prefer to grow in uncrowded, nutrient-rich environments (De Leij et al. 1994). In general, these organisms are capable of rapid growth in most nutrient-rich microbiological media and should be favored with simple substrates like glucose and amino acids. K-selected bacteria, on the other hand, are efficient competitors. They have a more efficient cell metabolism than r-selected bacteria and they prefer complex substances (Langer et al. 2004) such as lignin and xylan. Davis et al. (2005) were able to isolate many members of the poorly-uncultured bacteria, like

Acidobacteria, Verrucomicrobia and Chloroflexi, by using some polysaccharides such as xylan as the sole carbon sources in laboratory media. In their studies, they tested the effect of a range of growth substances (pectin, alginate, carboxymethylcellulose, xylan, xanthan, N-acetyl- glucosamine, amino acid mix, mix of D-galacturonate, D-glucuronate acid, L-ascorbate, and D- gluconate, mix of D-glucose, D-galactose, D-xylose, and L-arabinose and mix of acetate,

benzoate, L-lactate, and methanol) in low concentration of mineral salt VL55 medium.

Compared to the other substances, they found that xylan plates showed the highest average viable counts and the most of the colonies belonged to the previously uncultured K-selected bacteria. Xylan was also used successfully in other studies as a growth substrate for isolation of representatives of poorly studied groups of bacteria (Sait et al. 2002; Davis et al. 2011; Sait et al.

2006; Janssen 2002). These studies support the idea that as-yet-uncultured bacteria may belong to K-selected species, and they may prefer complex polysaccharides. The most common complex polysaccharides in nature are plant compounds like cellulose. However, another source, like those found in bacterial exopolysaccharides, is also a common source in nature. In this study, we used cellulose, gellan and another unusual EPS, indican_2, as growth subtracts to enrich and isolate EPS degrading bacterial communities (EPS. Note that throughout this thesis, I will use

EPS as the abbreviation for exopolysaccharides and EPS* as the abbreviation for extracellular polymeric substances).

Extracellular polymeric substances (EPS*) are high-molecular weight secretions produced by microorganisms that accumulate on the bacterial cell surface (Staudt et al. 2004;

Morgan et al. 1990). EPS* are comprised of polysaccharides, proteins, lipids, humiclike substances and even nucleic acids (e.g. DNA) and are involved in biofilm formation (Flemming and Wingender 2001; Allensen-Holm et al. 2006; Eboigbodin and Biggs 2008). EPS* form a protective layer that shields cells against a hostile external environment and also serve as carbon and energy reserves during time periods when nutrients are lacking (Liu and Fang 2002). High molecular weight polymer exopolysaccharides (EPS) as a key component of EPS* are composed of sugar residues (Gutierrez et al. 2013).

The purpose of this study is to contribute to efforts for culturing some previous uncultured K-selected bacteria. Some of them may have great potentially for study their metabolism in general.

1.2 Hypothesis

We hypothesized that bacterial exopolysaccharides (EPS) may serve as energy substrates for K-selected bacteria in soils, and that we would be able to identify and culture previously uncultured species using various EPS as growth substrates.

1.3 Research Objectives

There are three objectives for this research.

1. Develop culturing techniques to enhance EPS* production in selected organisms

Sphingomonas elodea (ATCC 31461), Beijerinckia indica (ATCC 9039) and

Gluconacetobacter xylinus (ATCC 53524).

2. Use 13C glucose to produce 13C EPS* exopolysaccharides. Then, use 13C-labeled EPS

in DNA-Stable Isotope Probing experiments to identify uncultured K-selected

bacteria in two soil communities (Big Hill Spring Provincial Park, Calgary, AB, pH

~6.0; and Paint Pots Spring Soils, Kootenay National Park, BC, pH ~3.5).

3. Use EPS as the sole energy substrate in culture media to cultivate novel soil bacteria.

1.4 Study Sites

Two study sites were selected, one with acidic soil and one with neutral soil. The acidic soil (pH ~3.5) study area is a group of cold natural acid mineral springs, known as the Paint Pots. 4

It is located just west of Vermilion Pass along Highway 93, in Kootenay National Park, British

Columbia (Grasby et al. 2013) (Figure 1.1). The springs have been known for a long time as an important cultural site. The Ktunaxa people used the iron oxide from the spring deposits as pigments. In 1858, this site was first formally described by Dr. James Hector of the Palliser

Expedition (Grasby et al. 2013). The spring sediment at the outflow, soil near the outflow, soil from atop a relic spring mound, and soil from the Ochre beds were sampled by Christine E.

Sharp (Figure 1.1). In previous Paint Pots studies, Everdingen (1970) indicate that this site water is acidic and has high heavy-metal content. The acidic water comes from the underground spring water on Ochre Hill which flows though the sulphide minerals deposits. When oxygen reacts to exposed sulphide minerals, the minerals form sulphuric acid with high dissolved metal content

(Grasby et al. 2013). The discharge from the spring acidic water flows down hill to form this acidic site.

Compared to the acidic PP soil, the neutral soil (pH ~6) area is Big Hill Springs

Provincial Park, Alberta, Canada. It is located northwest of Calgary, 10 km north of Cochrane, 6 km from Highway 22 on Highway 567 (Figure 1.2). The main attraction is a series of small waterfalls that flow year-round over rocky terraces covered with a lush growth of shrubs and grasses. The soil sample used for study was sampled from under a pine tree. Big Hill Springs water passes through calcareous sandstones and large calcium carbonate deposits resulting in a pH around 8 to 8.2 (Turner and Jones 2005), which keeps the soil quite neutral.

Figure 1.2 Big Hill Springs study site. Photo source: Google (public domain).

Chapter Two: LITERATURE REVIEW

2.1 Known and Unknown Microbial Diversity

2.1.1 Culture-Independent Methods Overview

The traditional cultivation methods used to describe microbial communities are limited and biased. Previous research showed less than 1% of the viable species present in a community can grow on traditional media (Liesack and Dunfield 2002). On the other hand, a different methodological tool, molecular analysis of a microbial community directly based on DNA or

RNA extracted from the environment, has increased microbiologists’ knowledge of microbial diversity (Spiegelman et al. 2005).

2.1.1.1 Total Nucleic Acid Extraction from Soil

Compared to labile RNA, DNA is more common to use in both extraction and subsequent molecular analysis. DNA extraction can be direct or indirect. The difference between these two procedures is whether cells are lysed within soil or not (Liesack and Dunfield 2002). Bead mill grinding and enzymatic lysis are two common protocols to use for cell lysis. Bead mill grinding usually extracts DNA fragments up to 25 kb in size which is suitable for PCR- based assessment of microbial diversity study (Kozdrój and van Elsas 2000). In contrast, enzymatic lysis which allowed extracting about 40-100 kb DNA fragments is more suitable for metagenomic analysis

(Krsek and Wellington 1999). For the subsequent analysis, DNA purification is required. Some research reported that humic acid which is tightly associated with the soil DNA extractions, complexes with proteins and inhibits PCR through interaction with Taq DNA polymerase

(Watson and Blackwell 2000). Many purification protocols have been applied to separate DNA

from humic acid, such as cesium chloride density gradient centrifugation and agarose gel electrophoresis (Liesack and Dunfield 2002) or washes using 5.5 M guanidine thiocyanate (GTC)

(Knief et al. 2003). The most commonly used are commercial kits which include purification steps.

2.1.1.2 PCR of the 16S rRNA Gene

Polymerase chain reaction (PCR) is used to amplify previously extracted DNA. In the process, the double-stranded DNA is denaturanted into single strands at high temperature. Then, two unique primers anneal to matching targets of the denatured target DNA sequence. The unique primers allow PCR to target 16S rRNA or other interesting genes. The 16S, 23S, and 5S rRNAs are three types of rRNA found in the ribosomes of prokaryotes. Based on the size difference, they are separated as small-subunit (SSU) known as 16S rRNA and large-subunit

(LSU) known as 23S rRNA and 5S rRNA (Spiegelman et al. 2005). They are ideal bio-molecule markers because they encode a highly conserved structure, universal to prokaryotes, rarely transfered from one species to another and have different regions of different mutation rates which allow inference of phylogenetic relationships (Liesack and Dunfield 2002; Spiegelman et al. 2005; Gürtler and Stanisich 1996). The most widely used gene to characterize microbial communities is the 16S rRNA gene. 16S rRNA genes contain highly conserved regions and have similar length (about 1.5 kb). Sequencing of the 16S rRNA allowed classiﬁcation of all prokaryotes into a Tree of Life (Woese et al. 1990). After years of 16S rRNA data collection, many public databases, such as the Ribosomal Database Project (Cole et al. 2009), Greengenes

(DeSantis et al. 2006) and ARB-Silva (Pruesse et al. 2007), have become available for comparative sequence alignments and analysis. However, we also need to keep in mind that 16S

rRNA PCR has some limitations. Primer bias is an important one. There are many 16S universal primers that can be used to target 16S rRNA genes. However, these universal premiers are not really universal (Baker et al. 2003). These “universal primers” are only short sequences (~20 bp) which can bind only to the small regions of 16S rRNA genes. Therefore, these primers will not attach to some prokaryotes which do not have these particular sequences in their 16S rRNA genes. Due to the primer bias, the primer selections affect the community analysis results

(Narzisi and Mishra 2011).

2.1.1.3 Fingerprinting Method

There are many molecular ﬁngerprinting methods available for us to analyze microbial communities. The most popular methods are denaturing/temperature gradient gel electrophoresis

(DGGE/TGGE), terminal-restriction fragment length polymorphism (T-RFLP), length heterogeneity polymerase chain reaction (LH-PCR), single-strand conformation polymorphism

(SSCP), amplified ribosomal DNA restriction analysis (ARDRA) (Spiegelman et al. 2005) and

16S rRNA gene pyrosequencing (Margulies et al. 2005). However, there is no single method listed above that is perfect and suitable for all research purposes. When we face all these choices, we should consider the purpose of study and choose a proper ﬁngerprinting method to use (Table

2.1). For example, T-RFLP is a highly quantitative analysis method compared to other techniques, but it is not suitable for phylogenetic deduction studies as many T-RFs are not specific for species (Liesack and Dunfield 2002). 454 pyrosequencing of 16S rRNA genes has been the most widely used fingerprinting technique recently. 454 pyrosequencing, invented by

454 Life Sciences, has been adapted as a high-throughput sequencing technology to discover the diversity of bacterial communities (Margulies et al. 2005). This method can provide hundreds of

thousands of sequences in a single run, and by adding a unique barcode to each sample allows as many as 120 samples to be sequenced at the same time on the Roche 454 Genome Sequencer

FLX System. Since 16S rRNA clone libraries were often done before pyrosequencing was available, 16S rRNA gene pyrosequencing is more convenient and efficient compared to other fingerprinting methods.This efficient fingerprinting method provides the most complete overview of the bacterial community structure that makes discovering a true picture of bacterial diversity possible (Cleary et al. 2012).

Table 2.1 Summary of common fingerprinting methods (Spiegelman et al. 2005; Liesack and Dunfield 2002; Suzuki et al. 1998).

Methods DGGE /TGGE T-RFLP LH-PCR SSCP ARDRA Pyrosequencing

Denaturing Terminal- Length Amplified Single-strand The Roche 454/GS /Temperature restriction heterogeneity ribosomal Full name conformation FLX sequencing gradient gel fragment length polymerase chain DNA restriction polymorphism. technology. electrophoresis. polymorphism. reaction. analysis. Separated PCR products by Separates mixed Separates mixed Separates amplified detecting Separates mix PCR PCR products by Separates mixed PCR products by 16S sDNA pyrophosphate Description products by %G-C using fluorescent PCR products by using restriction by 3D release on content. PCR primers and length. enzyme. structure nucleotide restriction enzyme. incorporation.

Low cost; Simple and rapid; Extracted band can Best with simple Simple procedure; Rapid; Efficient; Computer analysis; Rapid; be sequenced; communities; Can separate Can process Reproducible; Reproducible; Pros Difficult to Rapid and closely related complex Requires small Low cost. reproduce a gel. reproducible; fragment. environmental sample sizes. Computer analysis. samples.

High re-annealing Hard to work on Required clone Overlapping size rate; Doesn’t allow for complex library construction classes can leave Only short Unstable phylogenetic group environmental to assign peaks; ambiguities; sequence can be Cons sensitivity; identification; samples; Cannot recover and Technical problem obtained (~400 bp); No specific Limited by enzyme Often poor obtain sequence for accuracy peak Too much data. database for this bias. sequence recovery. data. detection. method.

2.1.2 Abundance and Diversity of Soil Bacterial Communities

Almost 30 years ago, Carl Woese and colleagues used a 16S rRNA phylogeny to divide the tree of life into three Domains. The Domain Bacteria originally contained 11 main phyla

(Woese et al. 1985). After molecular ecology techniques were widely used, over 100 bacterial phyla have been suggested (Achtman and Wagner 2008). So far only a small part of the bacteria in nature have been cultivated (Keller and Zengler 2004), and for most phyla (> 70%), we do not have a single isolate (Achtman and Wagner 2008).

Different soil bacterial communities may vary in the relative contributions of different phyla (Table 2.2) (Janssen 2006). However, nine dominant phyla (Proteobacteria,

Acidobacteria, Actinobacteria, Verrucomicrobia, Bacteroidetes, Chloroﬂexi, Planctomycetes,

Gemmatimonadetes and Firmicutes) make up an average of 92% of soil 16S rRNA clone libraries (Table 2.2) (Janssen 2006). Some of these phyla were quite well studied, for example

Proteobacteria and Bacteroidetes, but others have few or no known pure culture representatives from soils (Keller and Zengler 2004). Below I briefly describe the most important soil phyla that relate to the research results described later.

Proteobacteria and Acidobacteria are the most abundant phyla in soil bacterial communities (Table 2.2) (Janssen 2006). Proteobacteria comprised weighted average of 39% of total soil bacterial communities. Proteobacteria now can be classiﬁed within 6 classes:

Alphaproteobacteria, Betaproteobacteria, Gammaproteobacteria, Deltaproteobacteria,

Epsilonproteobacteria and Acidithiobacillia (Williams and Kelly 2013). The phylum

Proteobacteria currently contains 972 named and described genera (Euzeby 2013), but the number of proteobacterial sequences that can be assigned to known genera is relatively low.

Acidobacteria comprise on average 20% of all soil bacterial communities (Table 2.2).

Acidobacteria was recognized in 1997 as a novel phylum division (Kuske et al. 1997). Quaiser et al. (2003) based on years of cultivation- independent molecular ecology studies, as well as

Janssen (2006) pointed out that this phylum has a high phylogenetic diversity, and is particularly ubiquitous and abundant in soil habitats. The phylum Acidobacteria is divided by means of phylogenetic analyses. It was originally described as having 4 to 5 subdivisions, with subsequent expansion to having between 8 and 11 subdivisions (Kielak et al. 2010). However, genetic and physiological information about Acidobacteria are rare. Until now only three complete genome sequences of isolates of this phylum have been determined (“Solibacter usitatus”, “Korebacter versatilis” Ellin345 and Acidobacterium capsulatum) ( Kielak et al. 2010). Based on the report on List of Bacterial Names with Standing in Nomenclature (LPSN) website (Euzeby 2013), there are 11 described genera for Acidobacteria. Eight of them (Acidicapsa, Acidobacterium,

Bryobacter, Bryocella, Edaphobacter, Granulicella, Terriglobus and Telmatobacter) belong to subdivision 1. The other three (Acanthopleuribacter, Geothrix and Holophaga) belong to subdivision 8 (Losey et al. 2013). The previous research on cultured species showed the physiological versatility of cultured Acidobacteria species. In the recent Bergey's Manual of

Systematic Bacteriology (2012), Acidobacterium capsulatum, Terriglobus roseus and

Edaphobacter modestus are moderately acidophilic aerobic heterotrophs (Thrash and Coates

2010). Holophaga foetida and Geothrix fermentans are restricted anaerobes and have obligately fermentative metabolism (Thrash and Coates 2010).

Members of the phylum Planctomycetes make up an average of 2% of soil bacterial communities (Table 2.2). The first planctomycete observed was Planctomyces bekeﬁi (Gimesi

1924). Strangely, it was never isolated in pure culture. James T. Staley was the first to report the isolation of a planctomycete (Staley 1973). This organism was renamed several times, ﬁnally as

Pirellula staleyi (Schlesner and Hirsch 1987). In 1986, based on 16S rRNA gene sequence analysis, several members of a new order and family Planctomycetales and Planctomycetaceae were described (Schlesner and Stackebrandt 1986). At present, the Planctomycetes consists of 3 orders (Planctomycetales, Phycisphaerales, and “Candidatus Brocadiales”), 11 described genera and 14 species, and 5 candidatus genera with 14 candidatus species (Lage and Bondoso 2010).

Although the diversity is limited, these numbers are increasing. In the last 4 years, one new order, five new genera, and six species have been described (Lage and Bondoso 2010). Many researchers are trying to isolate novel species from this phylum. After years of studies,

Planctomycetes are recognized as slow growing organisms which have relatively long generation times. They also have low demand for carbon and nitrogen sources (Lage and Bondoso 2010).

However, many researchers also pointed out that most Planctomycetes are not strictly oligotrophic, they are also found in nutrient- rich environments (Jenkins and Staley 2013). Many cultivation strategies were used to enrich and isolate planctomycetes strains. However, due to their long incubation time and low demand of nutrient source, Planctomycetes are hard to isolate from nutrient-rich media. Because the nutrient-rich media can result in other rapid growing bacteria overgrowth, low nutrient media was used to isolate planctomycetes strains.

Planctomycetes has only three classes, but their lifestyles are variable. Some planctomycetes species, especially class candidatus “Brocadiae”, are anaerobic ammonium oxidation (anammox) bacteria. These bacteria are lithoautotrophs which are able to reduce nitrite and oxidise ammonium to produce dinitrogen gas under anaerobic conditions (Strous et al. 1999). The interesting part that is during the reaction, these planctomycetes produce a toxic metabolic

intermediate, hydrazine, which is a compound used in rocket fuel (Strous et al. 1999). These anammox planctomycetes play an important role in the global nitrogen cycle (Jenkins and Staley

2013). Other Planctomycetes even in the same genus showed different lifestyles. For instance,

Planctomyces maris, Planctomyces limnophilus and Planctomyces brasiliensis all belong to the

Planctomyces genus, and they are the only three named species that have been isolated in pure culture. Planctomyces maris are obligate aerobes, while Planctomyces limnophilus and

Planctomyces brasiliensis are facultative anaerobes (Thrash and Coates 2010). And they have different salt tolerance. Planctomyces limnophilus is stenohaline (low salt tolerance) while

Planctomyces maris and Planctomyces brasiliensis are euryhaline (high salt tolerance) (Jenkins and Staley 2013). There are some special strains in this phylum that are chemoheterotrophic, such as Isosphaera pallida. This strain was initially isolated by Giovannoni et al. (1987) with no organic carbon source added to the media. This strain can use only the organic contaminants within agar and was inhibited by 0.05% glucose (Giovannoni et al. 1987). Rhodopirellula baltiva, the best studied of all Planctomycetes, has been analysed by genome sequencing. Based on the genome, this strain was indicated to have abilities to degrade complex polysaccharides such as fucoidan (Jenkins and Staley 2013). This discovery might indicate other Planctomycetes may prefer complex polysaccharides as their energy source as well.

According to Bergey’s Manual of Systematic Bacteriology (2012) the Actinobacteria phylum contains five classes and 221 genera. However, many new taxa continue to be discovered (Goodfellow 2012). Actinobacteria make up an average of 13% of soil bacterial communities (Table 2.2). However, the understanding of this phylum is still limited, especially the subclass Acidimicrobidae. Until now, no validly named and described members of this

subclass from soil exist, and only ﬁve isolates from soil have been reported (Davis et al. 2005;

Joseph et al. 2003).

Similarly to Acidobacteria, Verrucomicrobia, Bacteroidetes, and Chloroﬂexi are all abundant in the soil, but limited on cultivation studies. For example, there are only 15 genera of

Verrucomicrobia that have currently been described (Euzeby 2013). Therefore, information is scant on these phyla, and cultivation research needs to be developed.

2.2 r- and K-Selection Theory

Why are so few groups of bacteria routinely cultivated? One theory behind this is that there are r-selected and K-selected species. r-selected and K-selected species are defined on the basis of their growth and ecological characteristics (Bottomley 1999). Typical r-selected bacteria are organisms that prefer to grow in uncrowded, nutrient-rich environments (De Leij et al. 1994). In general, these organisms are capable of rapid growth in most nutrient-rich microbiological media and should be favored by simple substrates like glucose. K-selected bacteria, on the other hand, are efficient competitors. They have a more efficient cell metabolism than r-selected and they prefer complex substances (Langer et al. 2004) such as lignin and xylan. The following table

(Table 2.3) compares the basic difference between r-selected and K-selected.

Table 2.3 Some attributes of r-and K-selected microorganisms (Langer et al. 2004). r-selected species K-selected species Population growth rate High Low Competitive ability Low High Substrate demand rate High Low Stress tolerance Low High Life span Short Long Nutrient uptake kinetics Low-affinity, high specificity High-affinity, low specificity Substrate diversity Simple and readily available Diverse and complex materials

2.3 Microbial cultivation methods

Until now, only about 30% of all bacterial phyla contain cultivated microorganisms

(Achtman and Wagner 2008). To study soil bacteria, cultivation plays a very important role in order to study physiology, genetics, and ecology (Janssen 2006). More than 50% of all bacterial phyla have been described based only on the 16S rRNA gene sequences recovered from the environment and therefore consist of as-yet-uncultured bacteria (Dojka et al. 2000). The phyla

Actinobacteria, Bacteroidetes, Cyanobacteria, Firmicutes and Proteobacteria represent about

95% of all cultivated and published species (Table 2.2). Others represent only 5% of all published species (Table 2.2) (Janssen 2006). The traditional enrichment and cultivation techniques are selective and biased (Eiler et al. 2000). Most of the species present in the environment do not grow on standard rich nutrient media in the lab like R2A medium and generally do not form visible colonies on plates (Eiler et al. 2000). One particular reason is the natural environments of the microorganisms are very different from traditional cultivation

conditions (Eiler et al. 2000). Also, the minimum visualization of colonies by unaided eyes need

105 cells in a colony. Therefore, slowly growing species are often overlooked because of short incubation times (Keller and Zengler 2004). Long incubation time, slow growing and complex substrate preference might indicate these species are considered as K-selected bacteria. This might explain why those slow- growing bacteria are hard to culture.

In recent decades, scientists have developed culturing methods for bacteria yet to be successfully grown on typical media (as-yet-uncultured bacteria) (Vartoukian et al. 2010).

Janssen et al. (2002) used simple modifications to traditional cultivation strategies, such as lowering the nutrient concentrations and increasing incubation times. They found that over 10% of the cells of the mean microscopically determined total cell count were potentially viable after careful observations based on plating, and most of them might belong to some of the as-yet- uncultured groups. They hypothesised that it was possible to discover and isolate previously uncultured bacteria by increasing the culturable fraction of soil bacteria. In their experiment, they compared the mean viable counts obtained using different media. They found the media that supported the highest mean viable counts also contained the most unclassified isolates (Janssen et al. 2002). They also noted that extended incubation times (>12 weeks) were needed to allow many mini-colonies to grow, and some of the bacteria isolated by this method were previously poorly represented by cultivated representatives such as members of the Acidobacteria and

Verrucomicrobia. Stevenson et al. (2004) also found similar results when they obtained pure cultures of previously uncultivated members of the divisions Acidobacteria from agricultural soil. They also used simple modifications to traditional cultivation strategies: the use of agar media with little or no added nutrients, longer incubation periods (more than 30 days) and

incubation under high concentrations of CO2 (5%). After the modification, they found that the abundance of Acidobacteria incubated with 5% CO2 on isolation plates was significantly higher than on non-treated plates. Dedysh (2011) also concluded that by improved cultivation strategies like dilute cultivation media and extended incubation time, previously uncultured bacteria from northern wetlands could be grown in the laboratory. Staley (1973) and Schmidt (1978) successfully isolated Planctomycetes strains by using low nutrient media from fresh water. Some marine strains such as Phycisphaera mikurensis have also been successfully isolated by using diluted artificial seawater and marine agar (Fukunaga et al. 2009).

Janssen et al. (2002) also amended media with non-traditional sources of nutrients like complex polysaccharides in order to increase culturability. In one experiment (Davis et al. 2005), xylan added as the growth substance showed the highest average viable counts compared to other simple substrates such as glucose. The use of gellan gum as a gelling agent instead of agar is also a strategy being widely used. Tamaki et al. (2009) indicated that growth inhibition for some bacteria was caused by using agar as a gelling agent. Gellan, also known as phytagel, is a bacterial polysaccharide produced by a Sphingomonas sp. (Sa-Correia et al. 2002). The molecular structure is a linear heteropolysaccharide consisting of repeating units of beta-D- glucose, beta-D-glucuronic acid, beta-D-glucose and alpha-L-rhamnose (Tamaki et al. 2009).

Several experiments done by other groups also indicate that using gellan gum as a gelling agent had better cultivation results than agar (Janssen et al. 2002; Sait et al. 2002; Davis et al. 2005).

Recently, Stott et al. (2008) and Dunfield et al. (2012) suggested that in addition to acting as a gelling agent, gellan gum was also used as an energy source for growth.

In addition to the improvement of traditional cultivation strategies, many other new cultivation strategies have also been used. One method called high-throughput culturing (HTC) utilized the concept of extinction culturing to isolate individual cells in small volumes of low- nutrient media (Zengler et al. 2002). Connon and Giovannoni (2002) also isolated and cultivated marine bacteria by using HTC method. This experiment used nutrient concentrations 3 orders of magnitude less than common laboratory media. This method successfully cultivated marine bacterioplankton that were previously uncultured (Connon and Giovannoni 2002). Another method is in situ cultivation. Kaeberlein et al. (2002) successfully grew previously uncultured bacteria from marine sediment in situ. They constructed a diffusion chamber that allowed natural environment substances across a membrane which simulated environmental conditions.

Some other strategies are also quite popular currently, such as combining cultivation with culture-independent methods. Giovannoni et al. (2007) indicate that culture-independent studies can deliver important information on the metabolic requirements of uncultured bacteria and that will lead to successful culturing of new species. Stable isotope probing (SIP) techniques can identify the growth substrate preference for uncultured bacteria in soil (Sharp et al. 2012;

Haicher et al. 2007; Eichorst and Kuske 2012), and is also a new strategy which can be

13 13 combined with cultivation methods. This strategy uses a C labeled substrate such as C-CO2,

13 13 C-CH4 (Sharp et al. 2012), C- cellulose (Haicher et al. 2007; Eichorst and Kuske 2012) as sole carbon source to detect labeled substrate-consuming bacteria and then uses a fingerprinting method to identify the taxonomy of them. Haicher et al. (2007) and Eichorst and Kuske (2012) by using 13C- cellulose successfully identified several cellulose-degrading bacteria, and also found some cellulose- responding bacteria which may use cellobiose and other secondary

products as their energy source. These results can help future researchers customize the bacteria energy source preference and lead to more cultivation success in the future studies.

Although so many strategies have been developed, the simplest modifications to traditional media recipes lead to the most successful isolation of previously uncultured bacteria (Davis et al.

2005; Sait et al. 2002). Simulating the natural conditions as much as possible should lead to a higher probability of isolation (Kaeberlein et al. 2002). Furthermore, a good understanding of metabolic pathways is also useful for developing bacterial cultivation strategies (Lee et al. 2009).

To sum up, cultivation-independent methods are widely used and microbiologists are able to identify bacterial species without the need for cultivation. This greatly increased the whole picture of diversity of microbial communities. Cultivation-independent methods are important for studying soil bacterial communities, but not for studying bacterial physiology, genetics, and ecology (Janssen 2006). To better study and understand the unknown bacteria species, cultured isolates still play a very important role. The K-selected and r-selected theory has helped explain the reason so few groups of bacteria being routinely cultivated. Many poorly represented phyla such as Acidobacteria, Planctomycetes, and Verrucomicrobia are reported as extremely difficult to maintain in pure culture (Janssen 2008). Improving the cultivation strategies to bring more as- yet-uncultured bacteria to pure culture has been a hot topic in microbiology (Eiler et al. 2000;

Kaeberlein et al. 2002; Sait et al. 2002; Davis et al. 2005; Haicher et al. 2007; Janssen 2008;

Sharp et al. 2012; Eichorst and Kuske 2012).

Chapter Three: CHARACTERIZATION OF SOIL MICROBIAL COMMUNITIES

3.1 Introduction

Different soils have different bacterial compositions. The objective for this chapter was to preselect soil samples that have higher potential of as-yet-uncultured bacteria by using the 454 pyrosequencing technique.

3.2 Methods

3.2.1 Sampling

3.2.1.1 Paint Pots Site

Samples for incubation and microbial community analysis were obtained from four locations around the Paint Pots Springs (Table 3.1). The spring sediment at the outﬂow, soil near the outﬂow, soil from atop a relic spring mound, and soil from the ochre beds were sampled by

Christine E. Sharp (Figure 3.1 A,B,C,D). All samples were taken 0-5 cm below the soil surface.

Temperature was measured in the field using a handheld temperature probe model HI9060

(Hanna Instruments). The pH was measured upon return to the laboratory using an Accumet

Basic AB15 pH meter (Fisher Scientiﬁc) by mixing soil with distilled water at a 1:1 ratio.

Samples were stored at 4 °C in the dark for less than 24 h before subsampling for DNA extraction and 16S rRNA PCR.

3.2.1.2 Big Hill Spring Site

Big Hill Spring (BHS) soil (Figure 3.1 E) sample for study and microbial community analysis was sampled under a pine tree (Table 3.1).Temperature was measured in the field using

a handheld temperature probe model HI9060 (Hanna Instruments). The pH for microbial samples was measured upon return to the laboratory using an Accumet Basic AB15 pH meter (Fisher

Scientiﬁc) by dissolving soil with distilled water at a 1:1 ratio. The sample was taken 2-5 cm from the soil surface. Samples were stored at 4 °C in the dark for less than 24 h before subsampling for DNA extraction and 16S rRNA PCR.

3.2.2 Extraction of Total DNA

Bacterial and archaeal diversity was analyzed based on recovery of 16S rRNA genes.

DNA was extracted from 500 mg of sampled soil for BHS soil (Table 3.1) using the FastDNA®

SPIN Kit for soil following the manufacturer's instructions (MP Biomedicals). Additional washes using 5.5 M guanidine thiocyanate (GTC) were used to remove excess humic acids and to increase DNA recovery (Knief et al. 2003). DNA was suspended in a final volume of 50 μl

DNase/Pyrogen-free water (DES) and stored at -20 °C for future use.

Table 3.1 Sample description Sample name Description T(°C) pH

Ochre bed bog Downstream of the spring, a bog fed by the spring water 21 3.29

Mound Soil from a relic spring mound 23 4.19

Soil Soil near the source spring, taken just under a pine tree 23 2.88

Outﬂow marsh Sediment in main spring pool 24 3.36

BHS Soil under pine tree, taken after removed top pine leaves 23 5.62

3.2.3 PCR Amplification, Purification and Quantification

The 16S rRNA gene was targeted using FLX Titanium amplicon primers 454T_RA_X and 454T_F containing the target primers 926f (5′-AAA CTY AAA KGA ATT GAC GG-3′) and

1392r (5′-ACG GGC GGT GTG TRC-3′), which will attached the V8 regions of 16S rRNA gene

(Engelbrekson et al. 2010), at their 3’-ends, along with the adaptors necessary for the Roche

Titanium chemistry (Ramos-Padrón et al. 2011). The reverse primer was different for each sample and contained a unique 10- nucleotide identifier barcode sequence that allowed for sequences to be binned according to the particular sample (Sharp et al. 2012). The PCR reaction mixtures contained 0.04 μM of the forward primer, 25 μl of 2 × Premix F (Interscience), 1.25 U of Taq DNA polymerase (Fermentas), 0.04 μM of the reverse primer with its unique barcode sequence for each sample, 1 μl of template DNA and nuclease-free water (Qiagen) to make up the total volume of 50 μl (Sharp et al. 2012). PCR amplification was performed with a thermal cycler with a temperature profile as follows: initial denaturation at 95 ° C for 3 min followed by

35 cycles of denaturation at 95 ° C for 30 s, annealing at 55 ° C for 45 s and extension at 72 ° C for 90 s, and a final elongation at 72 ° C for 10 min. The PCR products were checked on a 1% agarose gel and then purified using an EZ-10 Spin Column PCR Purification Kit (BioBasic Inc.).

For quality control purposes, the purified PCR products were quantified via a Qubit Fluorometer using a Qubit-iT dsDNA HS Assay Kit (Invitrogen) to ensure a sufficient amount of DNA was available for the pyrosequencing. Purified PCR products (typically 150 ng total DNA) were sequenced at the Genome Quebec and McGill University Innovation Centre, Montreal, Quebec, with a Genome Sequencer FLX Instrument, using a GS FLX Titanium Series Kit XLR70 (Roche

Diagnostics Corporation.).

3.2.4 Microbial Community Analyses

The QIIME (pronounced “chime”) open-source software platform (Caporaso et al. 2010) standing for Quantitative Insights into Microbial Ecology was used to analyze the pyrosequenced

16S rRNA genes. In the process, we first input and combined fasta data files from the 454 sequencing results with a metadata mapping file to run pre-processing analysis. In this step,

QIIME removed primers and barcode reads from all samples, and performed quality filtering.

QIIME then removed low-quality sequences by using PyroNoise based on a minimum quality score of 25. Then, QIIME clustered Operational Taxonomic Units (OTU) based on 97% identity, and classiﬁed the sequences via BLAST using a reference database. Classiﬁcation was based on the SILVA108 database for bacterial and archaeal 16S rRNA gene amplicons (Haas et al. 2011).

Once the OTUs were picked, QIIME automatically chose the representative sequences, assigned taxonomy, aligned the sequences, and output a table with the counts of each OTU in each sample.

All the eukaryotic sequences were recognized by QIIME, marked and named these sequences end with _ eukaryotic.txt listed on eukaryotic sequences OTUs table (DeSantis et al. 2006). By order QIIME directly deleted the _ eukaryotic.txt file, all the eukaryotic sequences (including chloroplasts) were removed from the analysis (Caporaso et al. 2010).

3.3 Results and Discussion:

3.3.1 Previous Paint Pots Microbial Community Research Review

Christine E. Sharp in our lab collected and did analysis on all Paint Pots samples (Grasby et al. 2013). Based on previous 16S rRNA gene pyrotag sequencing results, Proteobacteria was found to be the dominant phylum in the Ochre bed, outflow marsh and soil samples (Figure 3.2

B, C, D), comprising 30% of sequence reads. However, the highest proportions of target phyla

Acidobacteria, Chloroﬂexi, Planctomycetes, and Actinobacteria were found in the relic mound

(Table 3.2). So, based on 454 pyrotag sequencing results, the relic mound sample was selected as the future acidic incubation soil sample.

3.3.2 Big Hill Spring Microbial Community

Based on 16S rRNA gene pyrosequencing results, 20.5% Proteobacteria was found in

Big Hill Springs soil sample (Table 3.2). Phylum Acidobacteria (11.35%), Chloroﬂexi (3.6%),

Planctomycetes (3.5%), and Actinobacteria (32%) were over 50% in total in this soil (Figure 3.2

E).

Based on 454 pyrosequencing analysis results and previous results, Paint Pots relic mound (PP) soil sample and Big Hill Springs under pine tree soil (BHS) were selected as the soil samples for incubation studies. Janssen (2006) did research on different phyla contributions in bacteria soil communities based on 16S rRNA survey from 2920 clones in 21 libraries from different soils. The dominant phyla in soils were Proteobacteria, Acidobacteria, Actinobacteria,

Verrucomicrobia, Bacteroidetes, Chloroﬂexi, Planctomycetes, Gemmatimonadetes and

Firmicutes. Compared to his results, our Paint Pots mound sample which contained unusual by high abundance of Candidate division WPS2 and Chloroﬂexi should not be considered as a typical soil sample. However, for the purpose of detecting and isolating previously uncultured bacteria, and due to the high contributions of the as-yet-uncultured groups in the original Paint

Pots relic mound (PP) soil sample and Big Hill Springs under pine tree soil (BHS), these two soil samples were finally selected for the following studies.

Table 3.2 The top six most abundant phyla across all research soil samples (as a percentage of all reads).

Mound Ochre Bed Outflow Marsh Soil BHS Proteobacteria 17.1 34.0 37.3 47.7 20.5

Chloroflexi 19.8 23.1 11.1 13.6 3.6

Actinobacteria 7.8 8.5 8.3 12.8 32.0

Acidobacteria 13.7 9.2 9.5 12.7 11.3

Planctomycetes 6.5 3.1 2.5 6.7 3.5

Candidate division WPS2 23.0 4.1 0.4 2.3 0.1

120

100

Percentage% 40

0 A B C D E

Figure 3.2 Relative abundance of different bacterial and archaeal phyla for each soil library. 16S rRNA gene sequences were Chloroflexi classiﬁed accordingCyanobacteria to the nearest neighbour in the SILVA108 database: (A) mound (based on 9919 reads), (B) Ochre bed bog Proteobacteria (based on 6788 reads),Acidobacteria (C) outflow marsh (based on 4386 reads), (D) soil (based on 6238 reads), (E) BHS Soil (based on 8990 reads). Actinobacteria Planctomycetes Crenarchaeota 31 Thermotogae Bacteroidetes Gemmatimonadetes Candidate division OP11 Firmicutes Verrucomicrobia Others

Chapter Four: DETECTION OF AS-YET-UNCULTURED BACTERIAL GROUPS IN BHS AND PP SOIL USING STABLE ISOTOPE PROBING

4.1 Introduction

The use of stable isotope probing to identify the growth substrate preference for uncultured bacteria in soil is a new and popular strategy which combines enrichment with culture-independent methods to discover as-yet-uncultured bacteria. It allows us, without the need for cultivation on the plates, to identify species that incorporate particular substrates into specific biomarkers. This strategy successfully used 13C labeled substrate such as labeled 13C-

13 13 CO2, C- CH4 (Sharp et al. 2012), C- cellulose (Haicher et al. 2007; Eichorst and Kuske 2012) to detect labeled-substrate consuming bacteria. 13C labeled sources are added to environmental samples and certain microbes will incorporate it into their DNA. The heavy, 13C labeled DNA is separated from the light 12C DNA by using a cesium-chloride (CsCl) density gradient in an ultracentrifuge. Microbial community fingerprinting methods are then used to analyze the labeled nucleic acids to identify the microorganisms that metabolize the particular labeled substrate (Neufeld et al. 2007).

Many studies support the idea that as-yet-uncultured bacteria may belong to K-selected species, and they may prefer complex polysaccharides (Kaeberlein et al. 2002; Davis et al. 2005;

Sait et al. 2002; Zengler et al. 2002). The most important complex polysaccharide in nature is cellulose (structure see appendix B). Most cellulose is produced by plants and depends on photosynthesis. However, not only plants but also some bacteria can produce cellulose as well.

Gluconacetobacter xylinus has been recognized as a strong cellulose producer for a long time

(Hestrin and Schramm 1954). Highly pure 13C- labeled bacterial cellulose combined with the

stable isotope probing technique has been used to identify the cellulose degrading bacteria communities (Haichar et al. 2006; Eichorst and Kuske 2012). In this study, we used 13C-labelled bacterial cellulose to identify cellulose degrading bacterial communities in our two soil samples.

Besides cellulose, some other 13C-labelled bacterial exopolysaccharides were also used. Gellan

(structure see appendix B), also known as the commercial product phytagel, is a bacterial polysaccharide produced by Sphingomonas elodea (Sa-Correia et al. 2002). Gellan is used as a solidifying agent and can show better cultivation results than agar (Tamaki et al. 2009; Janssen et al. 2002; Sait et al. 2002; Davis et al. 2005). Stott et al. (2008) and Dunfield et al. (2012) suggested that rather than a gelling agent, gellan was an energy source for growth.

Finally an unusual EPS, indican_2, a heteropolysaccharide comprised by glucuronic acid, glucose and glycero mannoheptose (structure see appendix B), produced by Beijerinckia indica was also used to detect EPS degrading bacterial communities in our studies. The genus

Beijerinckia has suffered from taxonomic confusion for a long time. Some strains of Beijerinckia were misnamed as Sphingomonas and Azotobacter in early literature (Tamas et al. 2010). For example, our Beijerinckia indica (ATCC 9039) strain was earlier named as “Azotobacter indicum”, “Azotobacter lacticogenes” and“Azotobacter acida” (Euzeby 2013), The EPS product of B.indica was also misnamed. In 1980, Lawson and Symes (1980), first named the

EPS production of “Beijerinckia indica” ATCC 21423 as indican. However, ATCC 21423 was later reidentified. It is not Beijerinckia indica but a Sphingomonas sp. After that, Symes (1982) used the name “indican” again to name EPS produced by the bacterium Beijerinckia indica subsp. lacticogenes (ATCC 19361). However, “indican” already had its own definition. It is a colourless organic compound, naturally produced by plant Polygonum sp. (Indigofera tinctoria)

(Maier et al. 1990). Therefore, for clarity, “indican_2” named for the EPS extraction from strain

Beijerinckia indica subsp. indica (ATCC 9039) will be used in this thesis.

In this experiment, 13C labeled glucose or unlabelled 12C-glucose was used as the sole carbon source to feed three EPS producing strains, Gluconacetobacter xylinus (ATCC 53524),

Sphingomonas elodea (ATCC 31461) and Beijerinckia indica (ATCC 9039). Then the produced

13C-labeled and unlabelled 12C- EPS (cellulose, gellan and indican_2) were extracted. Before using 13C glucose to produce labeled EPS, all initial growth and EPS extraction used unlabelled glucose on both solid and liquid media. The method which produced the most EPS was chosen as the final method for the production of EPS from 13C-glucose. The 13C-labeled EPS was used as the sole carbon source provided to soil samples, and unlabelled EPS was also used for incubation of a control group. The EPS consumption was estimated by CO2 production as measured by Gas Chromatography (GC). After a certain time of incubation, DNA was extracted from incubated soil. By using CsCl gradient ultracentrifugate method, heavy labeled DNA was separated from unlabelled light DNA. The heavy DNA was amplified by 454 16S rRNA PCR.

The heavy DNA PCR products were finally sent for pyrosequencing analysis to identify the EPS- degrading communities (Figure 4.1).

12C-labelled

13C-labelled

Figure 4.1 Flow diagram of experimental procedure.

4.2 Materials and Methods

4.2.1 13C and 12C EPS Production

4.2.1.1 Cellulose Production

Gluconacetobacter xylinus (ATCC 53524) was previously kept in lab by routine transfer with 12C-glucose as the sole carbon source. To produced 13C-cellulose G. xylinus strain was grown in buffered S & H (Hestrin and Schramm 1954) medium (appendix A ,Table B)

supplemented with 2% 13C-glucose (> 99% purity, LC Scientific Inc.). Incubations were made aerobically for up to one week at 30 °C under static conditions (Figure 4.2 A).

4.2.1.2 Indican_2 Production

Beijerinckia indica (ATCC 9039) was used to produce 13C-labeled indican_2 and unlabelled 12C-indican_2 from 13C-labeled glucose. B.indica strain was previously maintained by routine transfer and substrate amendment with 12C-glucose as the sole carbon source. B.indica was grown on nitrogen-free mineral medium (Becking, 1984) containing 1% 13C labeled glucose

(see appendix A, Table D). Incubations were made aerobically for up to one week at room temperature (Figure 4.2 B).

4.2.1.3 Gellan Production

Sphingomonas elodea (ATCC 31461) was used to produce 13C-labeled gellan and unlabelled 12C-gellan from 13C-labeled glucose. S.elodea strain was previously maintained in lab by routine transfer and substrate amendment with 12C-glucose as the sole carbon.The S.elodea strain was grown in dilute nitrate mineral salts pH7 liquid medium (DNMS 7) (Hanson et al.

1991) containing 1% 13C labeled glucose (see appendix A, Table A). Incubations were stored aerobically for up to one week at 30 °C under static conditions (Figure 4.2 C).

Figure 4.2 Incubations of EPS producing bacteria A) Gluconacetobacter xylinus B) Beijerinckia indica C) Sphingomonas elodea.

4.2.2 EPS Extraction

4.2.2.1 Cellulose Extraction

G. xylinus produced glycan chains into the growth medium. These chains aggregate into ribbons and finally produce a mesh like pellicle (Ross et al. 1991), and the pellicle is floating on the top of the liquid culture. Raw cellulose pellicles (13C-labeled and unlabelled) (~10 g) were collected from liquid culture by using tweezers, and were put into 500-ml serum vials. The pellicles were boiled two times by 30 min baths in a 4% NaOH solution (Schramm and Hestrin

1954). And then, the pellicles were purified by washes in boiling dH2O after each bath until the pH was neutral. Pellicles were then broken down by bead beating. All pellicles were placed into

Lysing Matrix E tubes which contained one large bead and several small beads in each tube.

Then pellicle was then milled in the MPBio FastPrep Instrument for 3 cycles of 30 s at a speed setting of 6.0 m/s. The purified cellulose was autoclaved and then was frozen at -80 ℃ for 24 h.

Then the cellulose samples (Figure 4.6 A) were freeze dried at -80 °C for 72 h. The entire procedure is summarized in Figure 4.3. This procedure was modified from a previous cellulose extraction strategies study described by Schramm and Hestrin (1954). Haichar et al. (2006) also used this strategy to extract 13C cellulose, and they determined that the extractions have 99% purity.

Raw Cellulose Pellicle (~10 g)

2 × 30 min Baths in Boiling 4% NaOH Solution

Washes in boiling dH2O until the pH to Neutral

Bead Beating 3 Cycles of 30 s at 6.0 m/s Speed

Autoclaved

Lyophilization, -80 °C, 72 h

Figure 4.3 Step by step method for G. xylinus EPS cellulose extraction, after Schramm and Hestin (1954).

4.2.2.2 Indican_2 Extraction

EPS and cells (13C-labelled and unlabelled) (~15ml) were collected from plates of B. indica by scraping, and were put into 50-ml Falcon tubes. An equal volume of 10% salt solution

(NaCl) was added into the tube. The tube was vortexed at room temperature at 2500 times/m to mix the solution. The solution was centrifuged at 20,000 ×g at 4 ℃ for 30 min. The supernatant was then filtered through a 0.2-µm membrane (VWRTM International Inc.) at 25 ℃. This process separates the bacteria and other large particle from the EPS salt solution. Sometimes extra dilution steps were needed if the EPS formed a sticky mass. The EPS solution was then purified with a dialysis membrane (Spectrum Laboratories Inc.) (3500 Daltons) at 4 ℃ for 48 h. The purified EPS solution was frozen at -80 ℃ for 24 h, then, freeze-dried at -80 ℃ for 72 h. The entire procedure is summarized in Fig. 4.4. This procedure was modified from a previous EPS extraction strategy described by Liu and Fang (2002). These authors compared different EPS extraction strategies. To increase the purity of our EPS, we adapted most of their physical procedures and used salt (NaCl) instead of complex chemicals like EDTA. The DNA concentration and protein concentration of freeze-dried indican_2 were detected with a Qubit

Fluorometer using Quant-iT™ dsDNA HS Assay Kit (Invitrogen) and Quant -iT™ protein

Assay Kit (Invitrogen) to determine the EPS purity. Our Indican_2 used in the following incubations contained only 0.04% protein and 0.00074% DNA.

15 ml B.indica EPS and Cell Mixture

Mix with 15 ml 10% Salt (NaCl) Solution

Vortex at Highest Speed 25 °C, 10 min

20000×g Centrifugation 4 °C 30 min

Filtration through 0.2-µm Membrane 25 °C

Purified with Dialysis Membrane (3500 Dalton), 4 °C 48 h

Lyophilization, -80 °C, 72 h

Figure 4.4 Step by step method for B.indica EPS (indican_2) extraction, modified from (Liu and Fang 2002).

4.2.2.3 Gellan Extraction

Cells and EPS (13C-labelled and unlabelled) (~100ml) were collected from S. elodea liquid culture. The solution was centrifuged at 31000×g at 20 ℃ for 30 min. The supernatant was then filtered through a 0.2-µm membrane (VWRTM International Inc.) at 25 ℃. The pretreated

EPS solution was used in two alternate extraction methods. Method one purified the pretreated

EPS solution with a dialysis membrane (Spectrum Laboratories Inc.) (3500 Dalton) at 4 ℃ for 48 h. The purified EPS solution was frozen at -80 ℃ for 24 h, and then freeze dried at -80 ℃ for 72 h. The second method used the pretreated EPS (1 part) precipitated in 3 volumes of chilled absolute ethanol and left overnight at 4 ℃. The clear precipitate formed was separated from the solvent by sieve (1-mm) and allowed to drain for 10 min. The precipitate was redissolved in DES, and then the EPS solution was purified with a dialysis membrane (3500 Dalton) at 4 ℃ for 48 h.

The purified EPS solution was frozen at -80 ℃ freezer for 24 h then freeze dried at -80℃ for 72 h. The entire procedure is summarized in Figure 4.5. This procedure was modified from a previous gellan extraction study described by Goh (2004) and sludge EPS extraction study described by Liu and Fang (2002). After 7 times ethanol purification, Goh (2004) indicated their product was about 98% purity. Unfortunately, the 13C-gellan extraction not succeeded in this research.

100 ml S.elodea Liquid Culture

31000×g Centrifugation 4 °C 30 min

Filtration through 0.2-µm membrane 25 °C

Purified with Dialysis Membrane Precipitated in chilled 100% (3500 Dalton), 4 °C 48 h ethanol 3:1(v/v)

Lyophilization, -80 °C, 72 h Precipitated overnight at 4 °C

Drain for 10 min by 1-mm sieve

Purified with Dialysis Membrane (3500 Dalton), 4 °C 48 h

Lyophilisation, -80 °C, 72 h

Figure 4.5 Step by step method for S.elodea EPS (gellan) extraction, after Liu and Fang (2002) and Goh (2004).

Figure 4.6 The final EPS products A) Cellulose (the EPS product of G.xylinus strain) B) Indican_2 (the EPS product of B.indica strain) C) Gellan (the EPS product of S.elodea strain).

4.2.3 SIP Incubation and CO2 Production Measurement

Five-gram (wet weight) amounts of soil from each sample (BHS and PP) and 0.5% (w/w)

EPS (13C-labelled and unlabelled) were put into 120-ml serum bottles, which were sealed gas- tight with butyl rubber stoppers. Five-gram (wet weight) amounts of soil from each sample (BHS

and PP) with no added EPS were also set up as control groups (Table 4.1). Since the bacteria

13 13 13 taking C EPS, they will produce C-lablled CO2. To prevent some bacteria taking CO2 as

12 13 their carbon source, 10% (v/v) CO2 was added to the headspace of C EPS incubation bottles

13 13 12 to dilute to CO2 for preventing CO2 crossfeeding. Therefore, only C EPS incubation bottles and control bottles’ CO2 production rates were monitored. All samples were set up in triplicate

12 (Figure 4.7 A, B) and incubated at 22 °C in dark. Headspace CO2 mixing ratios of C unlabelled

EPS incubation bottles and control bottles were monitored using a Varian 450 gas chromatograph (GC) equipped with Hayesep N (0.5 m × 1/16×1 mm) and Molsieve 13X (1.2 m

× 1/16×1 mm) columns in series (70 °C) (Sharp et al. 2012), and a thermal conductivity detector (TCD) (detector temperature 190 °C). Mixing ratios were calculated by comparison with a known reference standard. The mixing ratio of the standard gas was 52 ppm CH4 and 500 ppm

CO2 with balance N2 supplied by Praxair Canada. Once the CO2 production radio of incubation samples clear higher (~ 1 time) than control samples for more than 14 d, the incubations were stopped, and the DNA were extracted for next steps.

Table 4.1 Incubation Setting Information

12 13 EPS C Cellulose C Cellulose+10% CO2 Control Sample BHS 1 PP1 BHS 1 PP1 BHS 1 PP1 Label BHS 2 PP2 BHS 2 PP2 BHS 2 PP2 BHS 3 PP3 BHS 3 PP3 BHS 3 PP3

12 13 EPS C indican_2 C indican_2+10% CO2 Control Sample BHS 1 PP1 BHS 1 PP1 BHS 1 PP1 Label BHS 2 PP2 BHS 2 PP2 BHS 2 PP2 BHS 3 PP3 BHS 3 PP3 BHS 3 PP3

Figure 4.7 Bottles set up A) cellulose with soil incubation B) indican_2 with soil incubation.

4.2.4 DNA Extraction, Fractionation, and Quantification

DNA was extracted from 500 mg of each soil using the FastDNA Extraction Kit for Soil

(MP Biomedicals). Additional washes using 5.5 M (GTC) were used to remove excess humic acids and to increase DNA recovery (Knief et al. 2003). DNA was suspended in a final volume of 50 μl DES and stored at -20 °C for next step. Unlabelled incubation soil DNA from both

BHS+12C-indican_2, PP+12C-Indican_2, BHS+ 12C-Cellulose and PP+ 12C-Cellulose were also prepared and used as controls to determine the expected position of unlabelled DNA in the cesium chloride (CsCl) density gradients. Each DNA extract (13C-labeled and control) was about

500 ng. All the DNA centrifugation and fractionation were performed following the procedure described by Neufeld et al. (2007). The steps are also summarized in Figure 4.8. The DNA was combined with CsCl (1.87 g/ml) and gradient buffer into ultracentrifugation tubes (13×51 mm).

Ultracentrifugation was done at the maximum allowable speed of 50,000 rmp (299065 ×g) in the

NVT90 rotor (Optima L-100K, Beckman Coulter Inc.) at 20 °C with vacuum over 65 h. DNA was retrieved by gradient fractionation resulting in 12 fractions of approximately 400 μl each, where fraction tube 1 was the heaviest and fraction tube 12 was the lightest. The density of each fraction was measured with a refractometer (AR200, Reichert) to confirm gradient formation.

DNA was precipitated from the CsCl with polyethylene glycol (PEG) and glycogen, washed with

70% ethanol and eluted in 30 ul of Tris-EDTA buffer (Sharp et al. 2012). The DNA concentration of each fraction was determined with a Qubit Fluorometer using a Quant-iT™ dsDNA HS Assay Kit (Invitrogen).

Figure 4.8 The flow diagram for DNA centrifugation and fractionation. Part of the figure is courtesy of Allyson Brady (Brady et al. 2011).

4.2.5 16S rRNA Gene Pyrotag Sequencing

The 16S rRNA gene was targeted using FLX Titanium amplicon primers 454T_RA_X and 454T_F containing the target primers 926f (5′-AAA CTY AAA KGA ATT GAC GG-3′) and 1392r (5′-ACG GGC GGT GTG TRC-3′), which will attached the V8 regions of 16S rRNA gene (Engelbrekson et al. 2010), at and their 3’-ends, along with the adaptors necessary for the Roche Titanium chemistry (Ramos-Padrón et al. 2011). The reverse primer was different for each sample and contained a unique 10- nucleotide identifier barcode sequence that allowed for sequences to be binned according to the particular sample (Sharp et al. 2012). The PCR

reaction mixtures contained 0.04 μM of the forward primer, 25 μl of 2 × Premix F (Interscience),

1.25 U of Taq DNA polymerase (Fermentas), 0.04 μM of the reverse primer with its unique barcode sequence for each sample, 1 μl of template DNA and nuclease-free water (Qiagen) to make up the total volume of 50 μl (Sharp et al. 2012). PCR amplification was performed with a thermal cycler with a temperature profile as follows: initial denaturation at 95 °C for 3min, followed by 35 cycles of 30 s at 95 °C, 45 s at 55 °C and 90 s at 72°C, and a 10 min ﬁnal elongation at 72 °C. PCR products were visualized on a 1% agarose gel and puriﬁed with an EZ-

10 Spin Column PCR Puriﬁcation Kit (BioBasic Inc.) For quality control purposes, the purified

PCR products were quantified via a Qubit Fluorometer using a Qubit-iT dsDNA HS Assay Kit

(Invitrogen) to ensure a sufficient amount of DNA was available for the pyrosequencing. In some cases, the purification step reduced the concentration of DNA necessary for pyrosequencing and additional PCR products had to be combined and added to the sample.

Purified PCR products (~150 ng total DNA) were sent to Genome Quebec and McGill

University Innovation Centre, Montreal, Quebec. They use a GS FLX Titanium Series Kit

XLR70 (Roche Diagnostics Corporation) and run samples with Genome Sequencer FLX

Instrument.

4.2.6 Microbial Communities Analyses

The results of the pyrotag sequencing revealed an overall amount of 2,443 OTUs from

26,552 reads across 4 samples. The average number of reads per sample was 6,638 with a maximum of 7,018 and minimum of 5,729. The QIIME software platform version 1.3 (Caporaso et al. 2010) was used to analyze the pyrosequenced 16S rRNA genes. In the process, we first input and combine fasta data file from the 454 sequencing results with metadata mapping file to

run pre-processing analysis. In this step, QIIME removed primers and barcode reads from all samples, and performed quality filtering. QIIME removed low-quality sequences based on a minimum quality score of 25. For the Roche Genome Sequencer FLX System, to decrease error in our data, reads with an average quality score above 25 had very few errors (less than 2 %).

Then, QIIME clustered Operational Taxonomic Units (OTUs) based on 98%, 95% and 90% sequence similarity. And then, the QIIME classiﬁed the sequences via BLAST against

SILVA108 database for bacterial and archaeal 16S rRNA gene amplicons (Haas et al. 2011).

QIIME takes the closest BLAST hit reference sequence to assign the taxonomy string (Caporaso et al. 2010). If there is no reference sequence hit, QIIME showed the results based on the least common ancestor method (Quast et al. 2013). For example, if a sequence had >90% similarity to a cultured relative, the family will show; >95% similarity to a cultured relative, the family will show; and > 98% similarity to a cultured relative, a genus will show (Pruesse et al. 2007; Quast et al. 2013). SILVA databases are quality-controlled databases of aligned 16S and 18S rRNA gene sequences from the Bacteria, Archaea and Eukaryota domains (Quast et al. 2013). They are official databases for the software ARB. SILVA108 database is the version released in 2011.

Eukaryotic sequences (including chloroplasts) were removed from the analysis. All the eukaryotic sequences were recognized by QIIME, marked and named these sequences end with _ eukaryotic.txt listed on eukaryotic sequences OTUs table (DeSantis et al. 2006). By order

QIIME directly deleted the _ eukaryotic.txt file, all the eukaryotic sequences (including chloroplasts) were removed from the analysis (Caporaso et al. 2010). Phylogenetic trees were built for visualization of sequence-based phylogenetic analyses. The tool we used for analysis of

16S rRNA sequence data was the ARB (Ludwig et al. 2004) software package. It is available online at http://www.arb-home.de. There are three major approaches for tree reconstruction:

distance matrix, maximum parsimony, and maximum likelihood methods (Ludwig et al. 1998).

In this study, we used maximum likelihood methods to build trees. In a phylogenetic tree, the path of evolution is indicated by branching pattern and the phylogenetic distances are indicated by branch lengths (Ludwig et al. 1998).

4.3 Results

4.3.1 CO2 Production Rate

4.3.1.1 Cellulose incubation CO2 Production

The cumulative carbon dioxide increased over a period of 20 d or 30 d with or without

(Control) supplemented cellulose in triplicate serum bottles of each soil. The results of the BHS soil with 12C cellulose incubation (Figure 4.9 A) suggest that during the 20-d incubation, the

12 BHS+ C Cellulose group showed a higher mean (n=3) CO2 production rate than the control group. At day 10, compared to the control group (1.68±0.05%), the mean production rate of

BHS+12C Cellulose (4.97±0.07%) is about 3-fold higher. The standard errors of the mean (SEM) bars were small and did not overlap. The results of the PP soil with 12C cellulose incubation

(Figure 4.9 B) suggest that during 30-d incubation, the control group showed a higher mean (n=3)

12 CO2 production rate than the PP+ C Cellulose incubation group. However, because the SEM bars overlapped, there was no difference between the PP+12C Cellulose incubation group and the control groups.

4.3.1.2 Indican_2 incubation CO2 Production

The cumulative carbon dioxide increased over a period of 90 d with or without (Control) supplemented indican_2 in triplicate serum bottles of each soil. The results of the BHS soil with

12C indican_2 incubation (Figure 4.9 C) show that during 90-d incubation, there was no difference between the BHS+12C indican_2 incubation group and the control group. The SEM bars also overlapped. The results of the PP soil with 12C indican_2 incubation (Figure 4.9 D) suggest that after 90-d incubation, the PP soil+12C Indican_2 group showed a higher mean (n=3) of CO2 production rate than the control group. However, since their SEM bars overlapped before day 60, there was no significant between PP soil+12C Indican_2 group and control group until day 60. After Day 60, PP soil+12C Indican_2 group showed more increase than control group. At

12 day 90 compared to the control group (0.71±0.03%), the CO2 production rate of PP soil+ C

Indican_2 (0.78±0.02%) was slightly (1.2-fold) higher than control group. The SEM bars were small and did not overlap.

4.3.1 13C-EPS Incubation Results

4.3.1.1 13C-Cellulose incubation DNA gradients

The incorporation of 13C into DNA of two soils was measured by relative DNA concentrations in the individual SIP fractions after 30 d incubation. BHS soils incubated in the presence of 13C-Cellulose incubation showed little shift in density of the DNA compared to unlabelled DNA (Figure 4.10 A). However, PP soil with 13C-Cellulose incubation showed a more obvious increase in DNA density as compared to the unlabelled DNA (Figure 4.10 B).

4.3.1.2 13C-Indican_2 incubation DNA gradients

The incorporation of 13C into DNA of two soils was measured by relative DNA concentrations in the individual SIP fractions after 80 d incubation. BHS soils incubated in the presence of 13C-Indican_2 showed a clear shift in density of the DNA compared to unlabelled

DNA (Figure 4.11 A, B). PP soil with 13C-Indican_2 also showed an increase in DNA density as compared to the unlabelled DNA. (Figure 4.11 C, D)

A B

C D

Figure 4.10 Relative DNA concentrations recovered from CsCl gradient fractions for BHS soil (A) and PP soil (B) incubated with 13C-Cellulose or 12C-Cellulose control. Solid lines represent incubations with 12C-Cellulose control and dashed lines represent incubations with 13C-Cellulose. The x-axis indicates the relative DNA concentrations recovered from each gradient fraction, with the highest quantity detected in any gradient fraction set to 1.0. Relative concentration rather than absolute DNA copies counts are used to facilitate comparison across the different experiments. The arrows indicate the fractions sent for sequencing. 54

4.3.1.3 13C-Cellulose incubation sequencing results

4.3.1.3.1 BHS soil

The dominant taxonomic phyla (Figure 4. 12) in the heavy fraction (Figure 4.10 A) of

BHS incubation sample (>1% of all sequences) were 45.49% Proteobacteria, 23.67%

Actinobacteria, 9.51% Bacteroidetes, 7.47% Acidobacteria, 3.13% Chloroflexi, 2,79%

Planctomycetes, 2.53% Crenarchaeota, 1.22% Gemmatimonadetes 1.10% Verrucomicrobia and

1.04% Firmicutes. The most abundant (>1%) species represented as a percentage of all reads are listed in Table 4.2. All the most abundant species in the heavy DNA fractions were also detected in the original unincubated BHS soil (Table 4.2). Except for one uncultured bacterium in the phylum Chloroflexi, these species were highly enriched in the heavy DNA compared to the original soil. Most of the enriched species were closet related to previous cultured cellulose degrader on taxonomic species level (identity ≥ 98%).

The sequences from the most abundant enriched uncultured species were imported into

ARB (Ludwig et al. 2004) and compared to currently known species. The phylogenetic position of the most abundant bacteria found in this study is shown in the 16S rRNA sequence-based tree

(Figure 4.13). Phylogenetic analysis of the 16S rRNA gene sequences of the top enriched species,

Emily OTU_4474, Emily OTU_4170, Emily OTU_2270, Emily OTU_3756, Emily OTU_4926,

Emily OTU_5740, Emily OTU_1589, Emily OTU_4762 and Emily OTU_663, supported their classiﬁcation as uncultured species of the phylum Proteobacteria. Emily OTU_408 was classified as an uncultured Bacteridetes. Emily OTU_2428 was classified as an uncultured

Actinobacteria (Figure 4.13).

Table 4.2 The most abundant (>1%) species in the heavy fractions of BHS soil incubated with 13C-Cellulose, compared to their abundances in the original unincubated BHS soil (as a percentage of all reads). Abundance Abundance in heavy Sequence identity Enrichment Taxonomic Description (Phylum; Class; Closet relative Rank OTUs in BHS fraction of BHS+13C- to the closest Radio Order; Family;Genus) based on BLAST soil Cellulose incubation relative % Proteobacteria;Alphaproteobacteria; 1 5740 0.01 12.34 1234.0 Caulobacterales;Caulobacteraceae; Caulobacter henricii 98.9 Caulobacter; Proteobacteria;Deltaproteobacteria;Myxococc 2 663 0.01 3.19 319.0 Sorangium cellulosum 93.3 ales; Sorangiineae; Proteobacteria;Betaproteobacteria;Burkholde 3 4926 0.12 2.94 24.5 Rubrivivax gelatinosus 99.5 riales; Comamonadaceae; Leptothrix; Proteobacteria;Gammaproteobacteria;Xantho Pseudoxanthomonas 4 4474 0.18 2.61 14.5 monadales; Xanthomonadaceae; 100.0 yeongjuensis Pseudoxanthomonas; Actinobacteria;Actinobacteria;Micromonospo Catenuloplanes 5 2428 0.10 2.56 25.6 rales; Micromonosporaceae; 100.0 crispus Catelliglobosispora; Bacteroidetes;Cytophagia;Cytophagales;Cyto Sporocytophaga 6 408 0.01 1.55 155.0 99.7 phagaceae; Sporocytophaga; myxococcoides Proteobacteria; Gammaproteobacteria; Ectothiorhodospira 7 3756 0.32 1.45 4.5 92.5 Chromatiales; Ectothiorhodospiraceae; haloalkaliphila Proteobacteria;Alphaproteobacteria; Brevundimonas 8 1589 0.03 1.33 44.3 Caulobacterales;Caulobacteraceae; 99.8 viscosa Brevundimonas; Proteobacteria;Alphaproteobacteria;Rhodospi Magnetospirillum 9 4762 0.01 1.22 122.0 94.4 rillales; Rhodospirillaceae; gryphiswaldense Proteobacteria;Gammaproteobacteria;Xantho Pseudoxanthomonas 10 4170 0.23 1.18 5.1 97.6 monadales Xanthomonadaceae; daejeonensis Sphaerobacter 11 2492 1.19 1.13 0.9 Chloroflexi;KD4-96; 84.6 thermophilus Proteobacteria;Gammaproteobacteria;Xantho 12 2770 0.01 1.07 107.0 Lysobacter niabensis 98.0 monadales; Xanthomonadaceae; Lysobacter;

Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB-

SILVA database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents 0.1 change per nucleotide position.

4.3.1.3.1 PP soil

The dominant taxonomic phyla (Figure 4. 14) in the heavy fraction of PP incubation

(Figure 4.10 B) sample (>1% of all sequences) were 55.71% Actinobacteria, 31.6%

Proteobacteria, 7.12% Bacteroidetes, 2.13% Acidobacteria, 1.22% Chloroflexi, and 1.04%

Cyanobacteria. The most abundant species (>1%) represented as a percentage of all reads are listed in Table 4.3. All the most abundant (>1%) species in heavy DNA fractions were also detected in the original unincubated PP soil (Table 4.3). All the abundant species in the heavy

DNA showed clear enrichment (>32 times) during the incubation. About 50% of the enriched species were uncultured at the species level (identity ≤ 98%). The sequences from the most abundant enriched species were imported into ARB (Ludwig et al. 2004) and compared to the currently known species. The phylogenetic position of the most abundant bacteria found in this study is shown in the 16S rRNA sequence-based tree (Figure 4.15). Phylogenetic analysis of the

16S rRNA gene sequences of the most abundant enriched species, Emily OTU_3404, Emily

OTU_2593 and Emily OTU_3460, supported their classiﬁcations all as members of uncultured species of the phylum Proteobacteria. The most enriched OTU (increased over 1000 times)

Emily OTU_372 belonged to the Actinobacteria and was closely related to the known species

Catenulispora acidiphila (Figure 4.15).

Table 4.3 The most abundant (>1%) OTUs in the heavy fractions of PP soil incubated with 13C-Cellulose, compared to their abundances in the original unincubated PP soil (as a percentage of all reads).

Abundance in Sequence Abundance heavy fraction Enrichment Taxonomic Description (Phylum; Closet relative identity to the Rank OTUs in PP of PP+13C- Radio Class; Order; Family;Genus) based on BLAST closest soil Cellulose relative % incubation Actinobacteria;Actinobacteria; Catenulispora 1 372 0.05 53.53 1070.6 Catenulisporales;Catenulisporaceae; 99.8 acidiphila Catenulispora; Proteobacteria;Alphaproteobacteria; Telmatospirillum 2 3404 0.01 19.13 1913.0 94.3 Rhodospirillales;Rhodospirillaceae; siberiense Proteobacteria;Alphaproteobacteria; Inquilinus 3 2593 0.01 8.00 800.0 92.5 Rhodospirillales; ginsengisoli Bacteroidetes;Sphingobacteria; Bradyrhizobium 4 3460 0.21 6.88 32.8 Sphingobacteriales;Chitinophagaceae; 100.0 elkanii Ferruginibacter;

Figure 4.15 16S rRNA gene sequence-based tree showing phylogenetic position of the most enriched cellulose-degrading bacteria identified in PP soil (designated with prefix ‘Emily’ in bold) assembled as per the Neighbor-Joining method with a Jukes-Cantor correction. Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB- SILVA database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents 0.1 change per nucleotide position.

4.3.1.4 13C-Indican_2 incubation

4.3.1.4.1 BHS soil

The dominant taxonomic phyla (Figure 4. 16) in the heavy fraction of the BHS incubation

(Figure 4.11 A) sample (>1% of all sequences) were 32.66% Actinobacteria, 17.43%

Proteobacteria, 15.36% Planctomycetes, 9.48% Acidobacteria, 9.28% Bacteroidetes, 5.4 %

Chloroflexi, 4.35% Verrucomicrobia, 2.88% Gemmatimonadetes and 1.44% Crenarchaeota. The most abundant (> 1%) species represented as a percentage of all reads are listed in Table 4.4. All the most abundant species in heavy DNA fractions were also detected in the original unincubated

BHS soil (Table 4.4). Other abundant species all showed enrichment during the incubation, although some showed only about 1 time enrichment. Most of the enriched species were uncultured species (identity ≤ 98%).

These sequences from the most abundant enriched uncultured species were imported into

ARB (Ludwig et al. 2004) and compared to the currently known species. The phylogenetic positions of the most abundant bacteria found in this study are shown in the 16S rRNA sequence- based tree (Figure 4.17). Phylogenetic analysis of the 16S rRNA gene sequences of the top 11 enriched species, Emily OTU_6297, Emily OTU_5443, Emily OTU_879 Emily OTU_4746 and

Emily OTU_665 supported their classiﬁcation as members of uncultured species of the phylum

Actinobacteria. Emily OTU_2492 was classified as an uncultured Chloroflexi. Emily OTU_6349 was classified as an uncultured Acidobacteria. Emily OTU_5492 and Emily OTU_2571 were classified as members of uncultured Bacteroidetes, Emily OTU_1151 was classified as an uncultured Planctomycetes and Emily OTU_2372 was classified as an uncultured

Verrucomicrobia (Figure 4.17).

Table 4.4 The most abundant (>1%) OTUs in BHS with 13C-Indican_2 incubation compared to the original unincubated BHS site abundance (as a percentage of all reads).

Sequence Abundance in Abundance in heavy Enrichment Taxonomic Description (Phylum; Class; Closet relative identity to the Rank OTUs BHS fraction of BHS+13C- Radio Order; Family;Genus) based on BLAST closest soil Indican_2 incubation relative % 1 1151 0.01 7.18 718.0 Planctomycetes;Phycisphaerae; Planctomyces maris 84.3 Actinobacteria;Actinobacteria;Propionibac Friedmanniella 2 6297 2.64 2.98 1.1 teriales; Propionibacteriaceae; 99.1 capsulata Microlunatus Acidothermus 3 879 1.20 2.70 2.3 Actinobacteria; MB-A2-108; 89.7 cellulolyticus Verrucomicrobia;Spartobacteria; Roseimicrobium 4 2372 0.01 2.57 257.0 88.4 Chthoniobacterales; gellanilyticum Bacteroidetes;Cytophagia; Nafulsella 5 5492 0.39 2.37 6.1 91.9 Cytophagales;Cytophagaceae; Flexibacter; turpanensis Sphaerobacter 6 2492 1.19 2.26 1.9 Chloroflexi; 84.6 thermophilus Bacteroidetes;Cytophagia;Cytophagales; Nafulsella 7 2571 0.38 1.93 5.1 93.1 Cytophagaceae; Flexibacter; turpanensis Actinobacteria;Thermoleophilia; 8 665 1.49 1.85 1.2 Conexibacter arvalis 95.0 Solirubrobacterales; Actinobacteria;Actinobacteria; Kribbella 9 2926 2.50 1.63 0.7 Propionibacteriales;Nocardioidaceae; 99.8 catacumbae Kribbella; Proteobacteria;Alphaproteobacteria; Pseudolabrys 10 2335 1.28 1.19 0.9 95.8 Rhizobiales;Xanthobacteraceae; taiwanensis Geobacter 11 6349 0.66 1.18 1.8 Acidobacteria;Acidobacteria; 87.3 psychrophilus Actinobacteria;Actinobacteria; Thermasporomyces 12 5443 0.86 1.15 1.3 92.9 Micrococcales;Dermatophilaceae; composti 13 4746 0.58 1.08 1.9 Actinobacteria;Thermoleophilia Gaiella occulta 94.8

4.3.1.4.2 PP soil

The dominant taxonomic phyla (Figure 4. 18) in the heavy fraction of the PP incubation

(Figure 4.11 C) sample (>1% of all sequences) were 68.18% Planctomycetes, 11.91%

Proteobacteria, 4.77% Cyanobacteria, 4.70% Chloroflexi, 3.14% Acidobacteria, 2.38%

Armatimonadetes, 2.20% Actinobacteria, and 1.46% Verrucomicrobia. The most abundant (>1%) species represented as a percentage of all reads are listed in Table 4.5. All the most abundant species in heavy DNA fractions were also detected in the original unincubated PP soil (Table

4.5). The most abundant species all showed enrichment during the incubation. Especially, the top enriched uncultured Plancomycetes, increased from only 0.04% to 59.13%. One species belonging to the Acidobacteria also showed about 2 times enrichment. All of the enriched species were uncultured species.

The sequences from the most abundant enriched species were imported into ARB

(Ludwig et al. 2004) and compared to the currently known species. The phylogenetic position of the most abundant uncultured bacteria found in this study is shown in the 16S rRNA sequence-

based tree (Figure 4.19). Phylogenetic analysis of the 16S rRNA gene sequences of three enriched uncultured species, Emily OTU_1253, Emily OTU_5016 and Emily OTU_4846 strongly supported their classiﬁcation as members of uncultured species of the phylum

Planctomycetes. Emily OTU_4106 was classified as an uncultured Acidobacteria. Emily

OTU_353 was classified as an uncultured Cyanobacteria, and Emily OTU_1270 was classified as an uncultured Armatimonadetes (Figure 4.19).

Due to the high abundance of Planctomycetes in the results, the top six abundant enriched

Planctomycetes species represented as a percentage of all reads are listed in Table 4.6. All five of them showed enrichment during the incubation and the minimum increased over 44-fold. All of them belonged to uncultured Planctomycetes species. These sequences were also imported into

ARB (Ludwig et al. 2004) and compared to the currently known species (Figure 4.20).

Table 4.5 The top ten abundant species in PP soil with 13C-Indican_2 incubation compared to the original unincubated PP site abundance (as a percentage of all reads).

Sequence Abundance Abundance in heavy Enrichment Taxonomic Description (Phylum; Class; Closet relative identity to the Rank OTUs in PP fraction of PP+13C- Radio Order; Family;Genus) based on BLAST closest soil Indican_2 incubation relative % 1 4846 0.04 59.13 1478.3 Planctomycetes;Phycisphaerae; Planctomyces maris 85.0 Proteobacteria;Gammaproteobacteria; Caldalkalibacillus 2 1355 0.01 7.12 712.0 78.3 Legionellales;Coxiellaceae;Aquicella; thermarum 3 353 0.01 2.78 278.0 Cyanobacteria; Microcystis aeruginosa 88.0 4 5016 0.01 1.67 167.0 Planctomycetes;Phycisphaerae; Zavarzinella formosa 88.8 5 1720 0.01 1.40 140.0 Armatimonadetes; Frankia alni 86.8 Planctomycetes;Planctomycetacia; 6 1253 0.04 1.08 27.0 Planctomycetales;Planctomycetaceae; Singulisphaera rosea 95.6 Singulisphaera; 7 4106 0.78 1.02 1.3 Acidobacteria;Holophagae; Nitrospina gracilis 88.7

Table 4.6 Taxonomic descriptions of the top six uncultured Planctomycetes species in PP with 13C-Indican_2 incubation compare to the original PP site abundance (as a percentage of all reads).

Abundance in Sequence Abundance heavy fraction of Enrichment Taxonomic Description (Phylum; Class; Order; Closet relative identity to Rank OTUs in PP PP+13C-Indican_2 Radio Family;Genus) based on BLAST the closest soil incubation relative % Planctomyces 1 4846 0.04 59.13 1478.3 Planctomycetes;Phycisphaerae 85.0 maris Zavarzinella 2 5016 0.01 1.67 167.0 Planctomycetes;Phycisphaerae; 88.8 formosa Planctomycetes;Planctomycetacia;Planctomycet Singulisphaera 3 1253 0.04 1.08 27.0 95.6 ales; Planctomycetaceae; Singulisphaera rosea Zavarzinella 4 3450 0.01 0.51 51.0 Planctomycetes;Phycisphaerae; 88.3 formosa Planctomyces 5 341 0.01 0.44 44.0 Planctomycetes;Phycisphaerae; 85.1 maris Planctomycetes;Planctomycetacia;Planctomycet Gemmata 6 1097 0.01 0.43 43.0 90.7 ales;Planctomycetaceae; obscuriglobus

Figure 4.19 16S rRNA gene sequence-based tree showing phylogenetic position of the most enriched indican_2-degrading bacteria identified in PP soil (designated with prefix ‘Emily’ in bold) assembled as per the Neighbor-Joining method with a Jukes-Cantor correction. Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB- SILVA database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents a 0.1 change per nucleotide position. 74

Figure 4.20 16S rRNA gene sequence-based tree showing phylogenetic position of the most enriched indican_2-degrading Planctomycetes bacteria identified in PP soil (designated with prefix ‘Emily’ in bold) assembled as per the Neighbor-Joining method with a Jukes- Cantor correction. Bootstrap support values (% of 1,000 replicates) are shown at the nodes. A skeleton tree was made from nearly full-length sequences (>1400 bp) and the shorter pyrosequences (450 bp) were added to this tree via parsimony. The sequence alignments were from the ARB-Silva database and calculations were made using ARB (Ludwig et al. 2004). The scale bar represents a 0.1 change per nucleotide position.

4.4 Discussion

4.4.1 Cellulose Incubation

After 20 days incubation with cellulose a CO2 production difference was apparent in BHS soil (Table 4.9 A) but not in PP soil (Table 4.9 B), and there was tiny shift apparent in community DNA density for BHS incubation studies but a clear peak in PP incubation. There have been some similar experiments done by other research groups examining bacteria involved in cellulose degradation in a complex soil environment (Haichar et al. 2006; Eichorst and Kuske

2012). These all indicated that moistening the soil is an important step during the incubation, especially for aggregated and clumped soil. Also, in Eichorst and Kuske’s (2012) research, they indicated increased incubation time to 30 to 35 days allowed more 13C incorporation into DNA, especially for some slow growing organisms. In our experiment the soil may have been too dry, and only 20 days incubation might not given enough time to allow sufficient 13C incorporation into some slow-growing organisms. In the future, increased incubation times might help us find more cellulose degrading bacteria. We originally did this experiment for longer (over 300 days) incubation time (data not shown) for cellulose incubation. However, in this round of experiments, our SIP peaks showed no shift at all. The possible explanation could be the cellulose and soil were dry and not mixed well.

In our cellulose incubation results with BHS soil (Table 4.2; Table 4.3), the most enriched groups belonged to previously discovered cellulose degraders. OTU_5740, OTU_408,

OTU_4474, OTU_4170 and OTU_663 were related to known cellulose-degrading species (Table

4.7). We also found some members of Family Rhodospirillaceae (OTU_4762, OTU_3404 and

OTU_2593), Micromonosporaceae (OTU_2428), Caulobacteraceae (OTU_1589) and family

Xanthomonadaceae (OTU_2770), were enriched in other cellulose incubation research (Eichorst and Kuske 2012; Kanokratanae et al. 2012; Schrempf and Walter 1999).

Table 4.7 OTUs related to known cellulose-degrading species list

OTUs Related cellulose-degrading species Reference showing cellulose degradation

OTU_5740 Caulobacter henricii Godden and Penninckx 1984

OTU_408 Sporocytophaga myxococcoides

OTU_4474 Pseudoxanthomonas yeongjuensis Trujillo-Cabrera et al. 2013

OTU_4170 Pseudoxanthomonas daejeonensis

OTU_663 Sorangium cellulosum Eichorst and Kuske 2012

OTU_5740, the most enriched bacterium in the heavy DNA fractions of BHS soil incubations, has 98.9% identity to Caulobacter henricii (Table 4.2). In microbial ecology, ≥98%

16S rRNA gene sequence identity was used as a guideline to indicate distinct species

(Stackebrandt and Goebel 1994), ≥96% indicating genera (Sait et al. 2002), with ≥ 90% identity indicating a family (Everett et al. 1999). OTU_5740 has 98.9% identity to Caulobacter henricii which means OTU_5740 and Caulobacter henricii are the same species. Caulobacter henricii consists of primarily aerobic species that are well known for their ability to degrade cellulose

(Godden and Penninckx 1984). The Caulobacteraceae family (Phylum: Proteobacteria; Class:

Alphaproteobacteria; Order: Caulobacterales) is typically found in low-nutrient soil and aquatic environments (Eichorst and Kuske 2012). The recently sequenced Caulobacter crescentus has genes to degrade plant-derived biopolymers like cellulases, xylanases and xylosidases (Hottes et

al. 2004). OTU_1589 also belonged to the family Caulobacterales (Table 4.2). Until now its role in cellulose degradation is not well understood. However, the enrichment in our experiment might suggest this strain may have some have ability related to cellulose degradation. More studies are needed in the future.

OTU_408 was also enriched in the heavy DNA fractions of BHS soil incubations studies and has 99.7% identity to Sporocytophaga myxococcoides. Sporocytophaga myxococcoides consists of primarily aerobic species that are well known for their ability to degrade cellulose

(Godden and Penninckx 1984). They belong to the Cytophagacea family (Phylum:Bacteroidetes;

Class: Cytophagia) (Table 4.2) which have been shown to contain cellulolytic and chitinolytic species (Eichorst and Kuske 2012; Kanokratanae et al. 2012).

OTU_4474, OTU_4170 and OTU_2770 all belonged to family Xanthomonadaceae

(Phylum: Proteobacteria; Class Gammaproteobacteria). This family is exclusively plant- associated bacteria (Pieretti et al. 2012). Many of their genomes encode putative cell wall degrading enzymes (e.g. cellulases, polygalacturonases, rhamnogalacturonases, beta- glucosidases and xylanases), showing their strong ability to degrade cellulose (Pieretti et al.

2012). OTU_4474 had 100.0% identity to Pseudoxanthomonas yeongjuensis and OTU_4170 has

97.6% identity to Pseudoxanthomonas daejeonensis. These two species both belong to the

Pseudoxanthomonas genus which was reported to have cellulose-degrading ability recently

(Trujillo-Cabrera et al. 2013).

OTU_663 was most closely related (93.3% identity) to the cellulose-degrading strain

Sorangium cellulosum (Figure 4.13) Sorangium cellulosum belongs to the Myxococcales order

(Phylum: Proteobacteria; Class Deltaproteobacteria) and was also enriched by Eichorst and

Kuske (2012) using cellulose as the incubation substrate. There is also a recently sequenced

genome of the Myxococcales bacterium Byssovorax cruenta that suggested it has the potential genes to degrade cellulose (Reichenbach et al. 2006)

OTU_2428 belongs to family Micromonosporaceae (Phylum: Actinobacteria; Class

Actinobacteria) and was also enriched in cellulose BHS soil incubation sample (Table 4.2).

Although its role in cellulose degradation is not well understood, some related species

(Actinoplanes aurantiacea and Streptomyces reticuli) have been identified as classic cellulose- degrading bacteria (Schrempf and Walter 1999). OTU_4762, OTU_3404 and OTU_2593 belong to Rhodospirillaceae family Rhodospirillales (Phylum: Proteobacteria; Class

Alphaproteobacteria), were also enriched in cellulose incubation sample (Table 4.2 and Table

4.3). Although its role in cellulose degradation is not well understood, it was also found be enriched during cellulose incubation in other research (Eichorst and Kuske 2012; DeAngelis et al. 2011).

OTU_372 has 99.8% identity to Catenulispora acidiphila (Phylum: Actinobacteria;

Class Actinobacteria) (Table 4.3) and was highly enriched in PP soil with cellulose incubation in our experiment. This strain has a gene belonging to Cellulose-binding family II. However, it was not found to be able to utilize cellulose by itself (Busti 2006). Some research showed it was also able to use xylose as energy source (Busti 2006). These results indicate this strain might degrade cellulose in collaboration with other strains in soil. OTU_3756 and OTU_3460 are not reported as cellulose-degrader in previous research, but our research results might indicate their might have cellulose degrading ability.

4.4.2 Indican_2 Incubation

Our Indican_2 incubation results (Table 4.4; Table 4.5; Table 4.6) provided strong evidence that our most enriched groups belong to uncultured species (≤ 98% identity). 15 OTUs were affiliated with 9 phylum-level groups and could be assigned to a total of 14 different family-level groups. Of these 14 family-level groups, only 4 (29%) OTUs belonged to a known family (Dermatophilaceae; Cytophagaceae; Planctomycetaceae; Coxiellaceae ). The rest did not correspond validly named or described species (or families). All OTUs therefore fall into new genera or new species in the Planctomycetes, Verrucomicrobia, Acidobacteria, Cyanobacteria,

Chloroflexi and Armatimonadetes (formerly Candidate phylum OP10).

OTU_4846, OTU_5016, OTU_1253 and OTU_1151 were affiliated to the phylum

Planctomycetes. These are the most enriched phylum in the heavy fractions of the indican_2 enrichments. Especially, OTU_4846 increased about 1466 times during the incubation. That strongly indicates that this strain uses our indican_2 as its carbon source. OTU_4846, OTU_5016 and OTU_1151 all belong to unknown order of Phycisphaerae Class. The unknown order they belonged to was named as WPS-1 (Nogales et al. 2001) which are mainly detected in marine and soil habitats (Fakunaga et al. 2009). OTU_1253 belongs to the Singulisphaera genus which was previous described by Kulichevskaya et al. in 2008. Singulisphaera acidiphila is the closest related known species to OTU_1253. Singulisphaera acidiphila is capable to hydrolysing laminarin, pectin, chondroitin sulfate, aesculin, gelatin,pullulan, lichenan and xylan

(Kulichevskaya et al. 2008). Several cultivation methods and media formulations for the isolation of Planctomycetes were tried by many researchers in past few decades (Lage and

Bondoso 2012). However, the isolation in pure culture has rarely succeeded (Lage and Bondoso

2012). One main difficulty is due to their slow growth rate. In our incubations, we incubated over 80 days, which seems to give them enough time to enrich.

OTU_1720 from the PP soil incubation belonged to the plylum Armatimonadetes formally called the candidate phylum OP10 (Dunfield et al. 2012). Candidate phylum OP10 was originally detected in Obsidian Pool in Yellowstone National Park (Hugenholtz et al. 1998). In

2011, Tamaki’s group proposed to use the candidate phylum OP10 as the new phylum name for

Armatimonadetes (Tamaki et al. 2011). OP10 usually was found in extremely environment, like hot springs and acidic soil (Dunfield et al. 2012). Stott et al. (2008) reported the first cultivation of an OP10 bacterium from Te Manaroa Spring in New Zealand. They indicate the keys to cultivation of this strain were to use dilute, low nutrient media, extended incubation time, use gellan as energy source and also maintain the low pH to copy the in situ conditions (Dunfield et al. 2012). Similar to gellan our indican_2 is also a complex polysaccharide served as energy source for incubated acidic PP soil. OTU_1720 enrichment might indicate this strain is also able to degrade other complex carbon sources like indican_2.

OTU_6349 from the BHS soil incubation affiliated to subdivision 1 of Acidobacteria

(Figure 4.17). The closed related known species (Figure 4.17) is called Granulicella rosea (87.3% identify) which was described as a species capable of degrading laminarin, pectin, lichenan, starch and xylan species (Pankratov and Dedysh 2010). Other research also indicates that using complex growth substrate helps isolate strains of Acidobacteria (Sait et al. 2002).

OTU_353 from the PP soil incubation belonged to the Cyanobacteria. Cyanobacteria are diverse in soil OTU_353 belonged to an unnamed class and was only 88.0% identical to known

Cyanobacteria species Microcystis aeruginosa. Many Cyanobacteria are photosynthetic;

however, we do not believe OTU_353 is photosynthetic. Because all the incubations were processed in dark, we do not have evidence to say this strain is photosynthetic.

OTU_5443, OTU_879 and OTU_4746 all from the BHS soil incubation were affiliated to the phylum Actinobacteria (Figure 4.17). OTU_5443 is 92.9% identical with species belong to the genus Dermatophilus. OTU_4746 belongs to the subclasses Thermoleophilia. This subclass contains only very few known isolates in the genera Rubrobacter and Thermoleophilum (Joseph et al. 2003). Most were isolated with complex substrates like gellan and xylan (Joseph et al.

2003). OTU_879 is more unusual and belongs to an unknown class of Actinobacteria that has no isolates.

OTU_5492 and OTU_2571 from the BHS soil incubation all affiliated to different species of Cytophagaceae in Bacteroidetes. Cytophagaceae are well studied and known as cellulose degrading bacteria (Schellenberger et al. 2011). They are able to utilize cellulose and cellulose-derived saccharides (e.g., cellobiose). We hypothesize that some strains are also able to degrade other complex carbon sources like indican_2.

OTU_2492 from the BHS soil incubation belonged to the phylum Chloroﬂexi.

Chloroﬂexi comprises on average of 3% of soil bacterial communities (Janssen 2006). Based on the list on LPSN website, only 19 described genera are known, and these are not evenly distributed within the classes (Euzeby 2012). OTU_1355 from the PP soil incubation belong to the Aquicella genus of Gammaproteobacteria. There is evidence to show they can grow in protozoa (Santos et al. 2003). OTU_2372 from the BHS soil incubation belonged to the order

Chthoniobacterales of Verrucomicrobia. Chthoniobacterales are free-living microorganisms common in soil, and they are use mainly mono-, di-, and polysaccharides and their derivatives

(Janssen and Hedlund 2011). This might explain the reason of these three OTUs enrichment in our research.

The most striking and interesting finding was that several Planctomycetes were highly enriched in our EPS indican_2 incubation results. Until now, there are only a few cultivable

Planctomycetes available and the knowledge for Planctomycetes is limited. Although in last few decades microbiologists have increased the success of Planctomycetes isolation (Schlesner 1994;

Zengler et al. 2002), their isolation in pure culture is difficult (Lage and Bondoso 2010). Due to the difficulty of growing planctomycetes, their enrichment in our incubations is exciting. We predict based on previous research (Schlesner 1994; Zengler et al. 2002) planctomycetes may like some complex polysaccharides. Therefore, our EPS could be very useful in future cultivation studies.

4.4.3 Potential Errors

Some cellulolytic species can completely degrade cellulose using their own multiple enzymes. Other species may use cellobiose and other secondary products as their energy source.

Cellulose degradation and degradation of cellulose breakdown products will occur at the same time in the soil. It is therefore difficult to clearly distinguish the primary cellulose-degrading community just by using SIP experiments. Eichorst and Kuske (2012) also indicated the similar results and limitation in their experiment.

Density gradient centrifugation was employed to separate 12C and 13C but this can also lead to fractionation through differences in nucleic acid GC-content (Holben et al. 2004). In our figure 4.10 A, there was an overlap between 12C and 13C patterns at 1.73 DNA density. For 12C

DNA this might represent DNA with high GC-content. This might confuse our 13C results. We

compared the cellulose-responsive bacterial communities in incubated soil to the initial unincubated soil community for distinct composition (Table 4.2). If one OTU only abundant in our 13C incubation results not in the original incubated soil, we are more confident to say this

OTU is truly enriched by incubating with our EPS.

All of this research was based on 16S rRNA gene molecular method. However, there are shortcomings for this method. First, there may be biases in the contributions of the various bacterial groups to libraries (Frostegard et al. 1999). Since the 16S rRNA gene has different regions targeted by different primers and different primers have different biases to binding on the sequence, the selection of primers affects the community analysis result (Narzisi and Mishra

2011). In addition, the libraries of sequences may only present an incomplete sampling. In the meantime, these available libraries of 16S rRNA and 16S rRNA genes permit an initial survey of the global soil bacterial community structure. We should also keep in mind that the libraries of

PCR-ampliﬁed 16S rRNA and 16S rRNA genes may not represent a true picture of the bacterial community. It is also possible that members of some groups may be underrepresented in our result, because the cells or spores may be difficult to lyse and so were not detected in PCR-based analyses that rely on DNA extraction from soil (Janssen 2006).

When we look at the results of indican_2 incubation enrichment, we found a couple of

OTUs only enriched about 1%. This slight increase may be due random errors in sampling. To decrease the sources of error, we only considered OTUs that were very strongly enriched in the heavy DNA fractions, not ones that are only a little higher.

Current QIIME BLAST result is limited by the SILVA108 database, which contains only a subset of high-quality reference sequences. Therefore, sometimes it brings errors. For example, in our cellulose with PP soil incubation, the QIIME BLAST result on OTU_3460 against

SILVA108 database, it is classified as uncultured Ferruginibacter bacteria. But if we BLAST the sequencing on the NCBI website which combined more than one database, this OTU is 100.0% identity to Bradyrhizobium elkanii. To decrease the error, we also BLASTed and identified the each individual abundant sequencing on NCBI website to find the relative closed strain manually.

The 13C gellan incubations were not successful because we were unable to develop a good system for 13C EPS extraction. Future studies need to improve gellan extraction.

Chapter Five: ISOLATION OF SOIL BACTERIA ON EPS-SUPPLEMENTED MEDIA.

5.1 Introduction

In recent decades, scientists have developed improved culturing methods for as-yet- uncultured bacteria. Many previous researchers (Janssen et al. 2002; Dedysh et al. 2000;

Kulichevskaya et al. 2009; Davis et al. 2011; Giovannoni et al. 2007; Overmann 2010; Zengler et al. 2002), by lowering the nutrient concentrations, using non-traditional energy sources and increasing incubation time had great success in isolating previously uncultured bacteria.

In this experiment, unlabelled EPS 12C-Indican_2 produced by B.indica and 12C-gellan produced by S. elodea were used as the sole carbon sources added to dilute nitrate mineral salts

(DNMS) plates. These plates containing EPS were inoculated with the same two dilute soil samples (PP soil and BHS soil) as used in the stable isotope probing study. The purpose of this study is trying to directly isolate previously uncultured bacteria by using our EPS. Pictures were taken daily or weekly to record the growth. To identify the individual species, individual colonies were selected for direct colony PCR and Sanger 16S rRNA gene sequencing. Figure 5.1 shows a flow diagram of the experiment.

5.2 Materials and Methods

5.2.1 12C EPS Production

5.2.1.1 Indican_2 Production

Was performed as described in the last chapter (Section 4.2.1.2).

5.2.2 EPS Extraction

5.2.2.1 Indican_2 Extraction

Was performed as described in the last chapter (Section 4.2.1.3).

5.2.2.2 Gellan Extraction

In this experiment gellan, or phytagel (Sigma), was used. This commercial product was produced from S. elodea.

Figure 5.1 Flow diagram of the experimental procedure.

5.2.3 Cultivation Experiments

1.0 g of wet freshly sieved soil was dispersed in 100 ml of sterile distilled water. These

10-2 diluted aliquots were serially diluted to 10-3, 10-4, 10-5, 10-6, and 10-7, and 200 µl of each dilution was used to inoculate each of 2 replicate plates. The dilutions were spread over the surface of the Indican_2 - or gellan-containing medium (Appendix A) by the use of sterile glass spreading rods. For BHS soil, two different media (pH 5.8) were used (DNMS 5.8+gellan and

DNMS 5.8+indican_2). For PP soil, one medium (pH 3.8) was used (DNMS 3.8+gellan). A total of 42 plates were incubated at room temperature in the dark for over 12 weeks in sealed polyethylene bags to prevent desiccation. Colony formation was monitored by camera every 3 d.

Due to frequent contamination by fungi all experiments were repeated 3 times. All plates were observed under a SZ61 zoom stereomicroscope (Olympus, Tokyo Japan) at 10× magnification for observing the colony morphology and the selection of the colonies of interest. Colonies were picked and transferred to new Indican_2 -or gellan-containing plates for isolation (Figure 5.2).

Figure 5.2 Bacterial isolates obtained from gellan-containing media.

5.2.4 Identiﬁcation of Colonies

Morphologically different colonies were picked with autoclaved toothpicks, and put into a DNA- free PCR tube and frozen at -80◦C overnight to lyse the cells. The next day they were thawed and

16S rRNA genes from colonies were ampliﬁed using the primer pair 9f (5’ GTT GTT GGG AAT

GGT TAC GG 3’) and 1492b (5’ GGT TAC CTT GTT ACG ACTT 3’) which will attached the whole variable regions (V1-V9) of 16S rRNA gene. PCR mixtures contained 25 μl of 2× Premix

F (Interscience), 1.25 U Taq DNA polymerase (Fermentas), 0.5 μM of each primer and the DNA, made up to 50 μl total with nuclease-free water (Qiagen). PCR reaction conditions were: initial denaturation at 94 °C for 5 min, followed by 35 cycles of 45 s at 94 °C, 1 min at 48 °C and 2 min

at 72 °C, and a 5-min final elongation at 72 ◦C. PCR products were visualized on a 1% agarose gel and purified with an EZ-10 Spin Column PCR Purification Kit (BioBasic Inc.). DNA concentration was determined via a Qubit Fluorometer using a Quant-iT™ dsDNA HS Assay Kit

(Invitrogen). Puriﬁed PCR products (typcally~ 120 ng total DNA) with 9f primer were sent for capillary-based automated Sanger sequencing at University of Calgary Core DNA sequencing facility. Sequencing results were visually checked for any background noise and mismatches via

ContigExpress version 11.5.1 (component of Vector NTI Advanced, Invitrogen part of Life

Science 2011). Trimmed sequences (removed low-quality sequences) about 600 bp long were compared to the reference sequences in NCBI database via nucleotide BLAST tool

(http://blast.ncbi.nlm.nih.gov/Blast.cgi).

5.3 Results and Discussion

5.3.1 Indican_2 Supplemented Medium

5.3.1.1 DNMS5.8+Indican_2

During the incubation colonies continued to become visible on the plates to the naked eye over the entire 12 weeks incubation period. Figure 5.3 shows a representative plate in which colonies gradually showed up during the incubation. After two weeks incubation, the most concentrated inocula grew on plates compared to other dilute inocula that had generally few, small (~1 mm in diameter) colonies (Figure 5.4). A total of 70 colonies from a number of dilution series (10-4, 10-5, 10-6 and 10-7) on agar supplemented with indican_2 were subcultured onto the same medium. 60% of these colonies did not grow after transfer, even after 2 months of incubation. About 20 subcultures were lost to fungal contamination. The remaining 8 isolates

yielded bacterial cultures, 2 of which were initially mixures for two species and were subsequently separated, which gave a total of 10 isolates. All isolates were identiﬁed by analysis of partial 16S rRNA gene sequences (Table 5.1). The isolates were assigned to the phyla

Proteobacteria, Actinobacteria, Acidobacteria, and Bacteroidetes.

In the following discussion, ≥98% 16S rRNA gene sequence identity was used as a guideline to indicate distinct species (Stackebrandt and Goebel 1994), ≥96% indicating genera

(Sait et al. 2002), with ≥ 90% identity indicating a family (Everett et al. 1999). Based on the results of the BLAST analyses many of the isolates were closely related (> 98% sequence identity) to members of characterized and named bacterial species, such as Microbacterium ginsengisoli and Bradyrhizobium elkanii (Table 5.1). However, other isolates were less related to previously cultivated bacteria, and some belonged to genera or even families with few or no known cultivated representatives. Isolate E120517_Brown_8 belonged to the Proteobacteria.

This isolate was 88.8% (a family level difference) identical to the previously cultured strain

Steroidobacter denitrificans which belongs to the Gammaproteobacteria. BLAST also showed this isolate had 98% identity to a bacterial isolate “Ellin5280” (Joseph et al. 2003), which was isolated using xylan as the carbon source. One isolate E120517_Brown_6 belonged to the

Acidobacteria. This isolate was 96.9% identical to Edaphobacter modestus which belongs to subdivision 1 of the phylum Acidobacteria (Koch et al. 2008). The BLAST result also showed that this isolate had 98% related to bacterium isolate “Ellin5236” (Joseph et al. 2003), which was isolated from soil using xylan as carbon source. Isolate E120517_Brown_2 belonged to the

Bacteroidetes. This isolate had 92.0% (a genus level difference) identity to Sediminibacterium salmoneum. Sediminibacterium salmoneum belongs to the class Sphingobacteria which is common in soils (Janssen 2006).

Nucleotide Isolates Top match to names cultured sp. Phylum Identity %

E120517_Brown_1 Microbacterium ginsengisoli Actinobacteria 100.0

E120517_Brown_2 Sediminibacterium salmoneum Bacteroidetes 92.0

E120517_Brown_3 Bradyrhizobium elkanii Proteobacteria 100.0

E120517_Brown_5 Microbacterium ginsengisoli Actinobacteria 100.0

E120517_Brown_6 Edaphobacter modestus Acidobacteria 96.9

E120517_Brown_8 Steroidobacter denitrificans Proteobacteria 88.8

E120517_Brown_9 Microbacterium ginsengisoli Actinobacteria 100.0

E120517_Brown_10 Microbacterium ginsengisoli Actinobacteria 100.0

E130506_1-6 Microbacterium ginsengisoli Actinobacteria 100.0

Figure 5.3 BHS soil 10-4 dilution spread on indican_2 -containing DNMS 5.8 Plates from Day 1 to Day 27.

Figure 5.4 Incubation of BHS soil 10-4 to 10-7 dilution after 2 weeks on indican_2 - containing DNMS 5.8 Plates.

5.3.2 Gellan Supplemented Medium

5.3.2.1 DNMS 5.8+gellan

During the incubation colonies continued to become visible on the plates to the naked eye over the entire 12 weeks incubation period. Figure 5.5 shows a representative plate in which colonies gradually showed up during the incubation. After 2 weeks incubation, there were some small colonies on the plate appearing around or covered by a large colony. This situation yielded new small colonies that were difficult to separate from the large colony (Figure 5.6). Possibly the small colonies could not grow alone on our medium but as the large colony covers them it somehow provides something for them to grow on. A total of 100 colonies from a number of dilution series (10-4, 10-5, 10-6 and 10-7) on gellan solidiﬁed DNMS plates were subcultured onto the same medium. 50% of these colonies did not grow after transfer, even after 2 months of 94

incubation. About 15 subcultures were lost to fungal contamination. The remaining 35 isolates were used for colony PCR and the PCR products sent to University of Calgary Core DNA sequencing facility. 19 isolates were not successfully sequenced, perhaps due to contamination.

Other non-contaminated isolates were identiﬁed by analysis of partial 16S rRNA gene sequences

(Table 5.2). The isolates were assigned to the phyla Proteobacteria, Actinobacteria, Firmicutes, and Bacteroidetes.

Based on the results of the BLAST analyses many of the isolates were closely related

(>98% sequence identity) to members of characterized and named bacterial species, such as

Streptomyces galilaeus, Ralstonia pickettii, Caulobacter vibriodes, Paenibacillus alginolyricus,

Sphingomonas panni, Ralstonia solanacearum, Pelomonas puraquae and Pelomonas aquatica

(Table 5.2). However, three isolates were less related to previously cultivated bacteria, and some belonged to groups with few or no known cultivated representatives. E130212_4-5 and

E130506_1-2 belonged to the phylum Bacteroidetes. Isolate E130212_4-5 had 95.3% (a genus level difference) identity to Niastella populi. Isolate E130506_1-2 had 92.1% (a genus level difference) identity to Sediminibacterium salmoneum. Sediminibacterium salmoneum and

Niastella populi all belong to the class Sphingobacteria which is common in soils (Janssen

2006). Isolate E130212_5-6 belonged to the phylum Actinobacteria. This isolate had 86.8 % (a family level difference) identity to Streptomyces aurantiacus. Streptomyces aurantiacus comes from Actinobacteridae which is the most well known and well studied subclass for this phylum

(Janssen 2006).

E120106_13 Streptomyces galilaeus Actinobacteria 98.8

E120106_15 Ralstonia pickettii Proteobacteria 99.4

E120306_Blue_4 Caulobacter vibrioides Proteobacteria 98.9

E120306_Blue_6 Ralstonia pickettii Proteobacteria 99.8

E120410_Black_3 Paenibacillus alginolyticus Firmicutes 99.4

E120410_Black_4 Sphingomonas panni Proteobacteria 99.6

E130212_1-2 Pelomonas puraquae Proteobacteria 99.8

E130212_1-3 Ralstonia solanacearum Proteobacteria 100.0

E130212_1-8 Ralstonia solanacearum Proteobacteria 99.6

E130212_2-8 Pelomonas aquatica Proteobacteria 100.0

E130212_4-5 Niastella populi Bacteroidetes 95.3

E130212_5-1 Ralstonia pickettii Proteobacteria 99.9

E130212_5-2 Ralstonia solanacearum Proteobacteria 99.5

E130212_5-6 Streptomyces aurantiacus Actinobacteria 86.8

Sediminibacterium E130506_1-2 Bacteroidetes 92.1 salmoneum

Figure 5.5 BHS soil 10-7 dilution spread on gellan-containing DNMS 5.8 Plates from Day 1 to Day 25.

Figure 5.6 Incubation of BHS soil 10-6 dilution after 1 month on gellan-containing DNMS 5.8 Plates. Some colonies start to spread out and couple similar colonies appearing around a mother colony.

5.3.2.2 DNMS 3.8+gellan

During the incubation colonies continued to become visible on the plates to the naked eye over the entire 12 weeks incubation period. Figure 5.7 shows a representative plate in which colonies gradually showed up during the incubation. After 2 weeks incubation, some small colonies on the plate appeared around or covered by a large colony. This situation yielded new small colonies that were difficult to separate from the large colony (Figure 5.8). The small colonies possibly could not grow alone on our medium but as the large colony covers them it somehow provides something for them to grow on. A total of 100 colonies from a number (10-4,

10-5, 10-6 and 10-7) on gellan solidiﬁed DNMS plates were subcultured onto the same medium.

55% of these colonies did not grow after transfer, even after 2 months of incubation. About 15 subcultures were lost to fungal contamination. The remaining 35 isolates used for 16S rRNA colony PCR, and PCR products were sent to University of Calgary Core DNA sequencing facility. 8 isolates were contaminated by Ralstonia sp. DNA. 22 non-contaminated isolates were identiﬁed by analysis of partial 16S rRNA gene sequences (Table 5.3). The isolates were assigned to the phyla Proteobacteria, Actinobacteria, Acidobacteria, and Bacteroidetes.

Based on the results of the BLAST analyses 14 isolates are closely related (>98% sequence identity) to members of characterized and named bacterial species, such as

Streptomyces prunicolor, Ralstonia pickettii, Caulobacter vibrioides, Microbacterium ginsengisoli, Ralstonia solanacearum and Leifsona kribbensis (Table 5.2). However, the other seven isolates were less related to previously cultivated bacteria, and some belonged to genera or even families with few or no known cultivated representatives. Isolates E120417_Purple_4,

E120628_1 and E120628_4 all belonged to Bacteroidetes. Isolate E120417_Purple_4 have

95.3% (a genus level difference) identity to Niastella populi. Isolate E120628_4 had 91.4% (a

genus level difference) identity to Sediminibacterium salmoneum. Sediminibacterium salmoneum belong to the class Sphingobacteria which was common in soils (Janssen 2006). Isolate

E120628_1 had 84.5% (a family level difference) identity to Algoriphagus aquimarinus.

Algoriphagus aquimarinus belongs to the Cytophagia class and Cyclobacteriaceae family. Due to the family level difference (85%), this isolated might represent a new Bacteroidetes family.

Isolate E120416_Orange_8 belonged to Proteobacteria. This isolate had 87.2% (a family level difference) identity to Steroidobacter denitrificans and could be reproducibly assigned to

Gammaproteobacteria of Proteobacteria. The BLAST result also showed this isolate had 99.5% related to bacterium isolates “Ellin6101” (Joseph et al. 2003). Ellin6101 is also an unclassified

Gammaproteobacteria (Joseph et al. 2003), and was isolated by using xylan as carbon source.

Isolates E120106_1 and E130506_1-4 belonged to Actinobacteria. Isolates E120106_1 had 96.3 %

(a species level difference) identity to Streptomyces galilaeus. Isolate E130506_1-4 had 97.6 %

(a species level difference) identity to Leifsonia kribbensis. These two strains all came from

Actinobacteridae which is the most well known and well studied subclass (Janssen 2006). One isolate from the incubation E120628_6 belonged to the Acidobacteria. This isolate had 97.3% identity to Edaphobacter modestus and could be reproducibly assigned to subdivision 1 of

Acidobacteria (Koch et al, 2008).

Table 5.3 Analysis of isolates obtained via ContagiExpress software on the basis of partial (~ 600 bp and use only 9f as sequencing primer) 16S rRNA gene sequences, including the closest taxonomically validated relatives identiﬁed using the BLAST program in the NCBI databases.

Nucleotide Isolates Top match to cultured sp. Phylum Identity % E120106_1 Streptomyces galilaeus Actinobacteria 96.3 E120106_3 Streptomyces prunicolor Actinobacteria 99.6 E120106_4 Ralstonia pickettii Proteobacteria 100.0 E120106_6 Streptomyces prunicolor Actinobacteria 99.8 E120106_7 Ralstonia pickettii Proteobacteria 100.0 E120106_8 Caulobacter vibrioides Proteobacteria 100.0 E120106_9 Caulobacter vibrioides Proteobacteria 99.6 E120106_10 Caulobacter vibrioides Proteobacteria 100.0 E120416_Orange_8 Steroidobacter denitrificans Proteobacteria 87.2 E120416_Purple_5 Microbacterium ginsengisoli Actinobacteria 100.0 E120416_Purple_8 Microbacterium ginsengisoli Actinobacteria 100.0 E120417_Purple_3 Microbacterium ginsengisoli Actinobacteria 100.0 E120417_Purple_4 Niastella populi Bacteroidetes 95.3 E120417_Purple_5 Microbacterium ginsengisoli Actinobacteria 100.0 E120417_Purple_6 Microbacterium ginsengisoli Actinobacteria 100.0 E120417_Purple_8 Microbacterium ginsengisoli Actinobacteria 100.0 E120628_1 Algoriphagus aquimarinus Bacteroidetes 84.5 E120628_4 Sediminibacterium salmoneum Bacteroidetes 91.4 E120628_6 Edaphobacter modestus Acidobacteria 97.3 E130212_6-1 Ralstonia solanacearum Proteobacteria 99.5 E130506_1-4 Leifsonia kribbensis Actinobacteria 97.6 E130506_1-5 Leifsonia kribbensis Actinobacteria 98.5

100

Figure 5.7 PP soil 10-7 dilution spread on gellan-containing DNMS 3.8 Plates from Day 1 to Day 25.

Figure 5.8 Incubation of PP soil 10-6 dilution after 1 month on gellan-containing DNMS 3.8 plates. As colonies spread out, smaller colonies sometimes appear around a mother colony.

101

5.3.3 Culturability Factors

During our study we noted that some colonies were large and fast-growing, but other small colonies took extremely long incubation times (>3 months) to become visible. In r versus

K-selection theory, r-selected species like to use simple substrates such as glucose and amino acids, usually form large colonies, and grow rapidly. On the other hand, K-selected species prefer complex substrates like polysaccharides, usually take long time to grow, and form small colonies (Langer et al. 2004). This results indicate these small and slow-growing colonies might be the K-selected bacteria we are looking for. However, even after such long incubation, some colonies were still visible to the naked eye. Davis et al. (2010) also mentioned in their research that some Acidobacteria, Actindobacteria or Chloroﬂexi normally formed late-appearing colonies (visible > 12 weeks) or developed only mini-colonies. We also observed many mini- colonies after about 3 month incubation. Gellan degrading colonies appeared like small volcanos

(Figure 5.9). In our research, we found that increasing the incubation time resulted in increased small colony visibility. Janssen et al. (2003) also observed that increasing the incubation time results in increased viable counts, particularly on media with low nutrient concentrations like ours. However, incubation times on the order of months are only rarely used, due to the limited research or project time. In future studies extended incubation times will be important. Davis et al. (2010) suggested that some rarely isolated K-selected groups might need more than 2 months incubation on suitable media, and pure cultures from these groups took an extremely long time to produce visible colonies.

102

103

Based on 16S rRNA sequence libraries in previous studies, Proteobacteria,

Acidobacteria, Actinobacteria, Verrucomicrobia, Bacteroidetes, Chloroﬂexi, Planctomycetes,

Gemmatimonadetes and Firmicutes are the most abundant groups in soils (Janssen 2006).

Certain phyla such as Acidobacteria, Planctomycetes and Verrucomicrobia are quite common in soils based on 16S sequencing, but rarely cultured (Janssen 2006; Keller and Zengler 2004).

Besides the effect of incubation time, growth media composition and inoculum size (Schoenborn et al. 2004) also are considered as important conditions determining culturability. Davis et al.

(2005) mention these three factors in their paper. In their research, many different media were used to compare isolation ability. After long incubations, they found that using non-traditional media helped them to isolate some previously uncultured groups (Davis et al. 2005; Janssen et al.

2002; Sait et al. 2002; Sait et al. 2006). Their non-traditional media mimicked the low concentrations of inorganic ions in soils, adjusted media pH to soil pH at the site being studied, used gellan instead of agar as the solidifying agent and also used different growth substrates like xylan. Xylan was also used successfully as a growth substrate for the isolation of representatives of poorly studied groups of bacteria in earlier studies (Joseph et al. 2003; Sait et al. 2002; Joseph et al. 2003; Schoenborn et al. 2004). Later on, some researchers indicate gellan may not be only a gelling agent, but also a substrate for growth (Stott et al. 2008; Dunfield et al. 2012). Stott et al.

(2008) successfully cultivated an OP10 bacterium by using gellan as the growth substrate. The previous studies and this research also indicated that, rather than simple substrates such as glucose or amino acids, complex substrates like xylan, gellan and other EPS as cultivation energy source lead may help isolate previously uncultured K-selected bacteria (Davis et al. 2005;

Joseph et al. 2003; Sait et al. 2002; Joseph et al. 2003; Janssen 2008; Stott et al. 2008; Dunfield et al. 2012; Schoenborn et al. 2004).

104

In our study, we also adjusted our media pH (3.8 or 5.8) and used complex growth substrates (EPS) as our energy source. However, PP soil contained high concentrations of iron in soils (Grasby et al. 2013), therefore, increased iron level in our media might be a good way in the future to increase the isolation of the poorly studied groups. Besides the overgrowth of rapid growing bacteria, another problem is the rapid and invasive growth of fungi. To inhibit fungal growth, cyclohexamide was added to our plates. However, these antifungal compounds have not always proven to be effective in our case. To prevent culture loss we need to do more research to solve this problem.

Use of our EPS as carbon sources did show some evidence that we can isolate some previous uncultured species in the phyla Actinobacteria, Proteobacteria, Bacterioidetes and

Acidobacteria (Table 5.4). The members of Acidobacteria (E120517_Brown_6 and E120628_6) were isolated from both indican_2 and gellan plates (Table 5.4). Our SIP experiment also found some members of Acidobacteria enriched by using indican_2 as a carbon source (Table 4.4;

Table 4.5). In SIP studies, more than 50% of the enriched OTUs belonged to Actinobacteria

(Chapter 4). Our cultivation studies also had similar results. Most isolates belonged to the

Actinobacteria. However, the similarity stopped at the phylum level, until now we have not detected any of the same bacterial species in both SIP and cultivation studies. Since agar cannot solidify at low pH, our PP soil was not processed using indican_2 as sole carbon for source cultivation studies. In SIP enrichment studies, we found in PP soil with indican_2 incubations, some members of Planctomycetes were highly enriched. Because of the limitation of agar, which will not solid on low pH, we do not have any isolate of Planctomycetes on the plates. In the future studies, we could use gellan as gelling agent and Indican_2 as carbon source to try to cultivate some Planctomycetes.

105

Table 5.4 Summary of the most taxonomically novel isolates obtained in this study.

Energy Nucleotide Soil Isolates Top match to cultured sp. source Identity %

E120517_Brown_2 Indican_2 Sediminibacterium salmoneum 92.0

E120517_Brown_6 Indican_2 Edaphobacter modestus 96.9

E120517_Brown_8 Indican_2 Steroidobacter denitrificans 88.8 BHS soil E130212_4-5 Gellan Niastella populi 95.3

E130212_5-6 Gellan Streptomyces aurantiacus 86.8

E130506_1-2 Gellan Sediminibacterium salmoneum 92.1

E120106_1 Gellan Streptomyces galilaeus 96.3

E120416_Orange_8 Gellan Steroidobacter denitrificans 87.2

E120417_Purple_4 Gellan Niastella populi 95.3

PP soil E120628_1 Gellan Algoriphagus aquimarinus 84.5

E120628_4 Gellan Sediminibacterium salmoneum 91.4

E120628_6 Gellan Edaphobacter modestus 97.3

E130506_1-4 Gellan Leifsonia kribbensis 97.6

106

Chapter Six: CONCLUSIONS

Culturing the total diversity of species currently seems an unreachable goal. But if we could figure out what the bacteria require, such as key carbon and energy sources, we will get a basic idea how to culture more uncultured species.

Some progress in culturing previously uncultured soil bacteria has been made in the last few years (Eiler et al. 2000; Janssen 2008; Kaeberlein et al. 2002; Davis et al. 2005; Sait et al.

2002; Sharp et al. 2012; Haicher et al. 2007; Eichorst and Kuske 2012; Lee et al. 2009; Connon and Giovannoni 2002; Zengler et al. 2002; Stevenson et al. 2004; Schoenborn et al. 2004). These recent advances mean that it is probably wrong to say the majority of bacterial species in soil are unculturable. However, many rarely isolated groups are very slow-growing and are difficult to maintain in the laboratory and the colonies require weeks or months rather than hours or days to become visible (Davis et al. 2005; Janssen 2008; Stevenson et al. 2004; Janssen et al. 2002).

Therefore for isolation or enrichment, we should be patient and carefully select appropriate strategies. Based on our research we found some members of Acidobacteria, Verrucomicrobia,

Planctomycetes, Chloroﬂexi, Actinobacteria, Cyanobacteria, Proteobacteria, Bacteroidetes, and

Armatimonadetes (OP10), are actually readily enriched on complex substrates like some members of the Acidobacteria were isolated on both indican_2 and gellan plates.

Although our cultivation studies did not succeed in cultivating many new species (Table

5.1, 5.2 and 5.3), our SIP studies showed that some poorly-cultured groups (Acidobacteria,

Verrucomicrobia, Planctomycetes, Chloroﬂexi, Cyanobacteria and Armatimonadetes) may consume complex substrates that are not often used in laboratory media. Especially, Some

Planctomycetes which are not related to any cultured Planctomycetes species, but highly

107

enriched by our Indican_2. That gives us an idea, that in the future many of these poorly-cultured bacteria will be readily culturable. By using complex substrate as cultivation supplement, we can develop culture-based approaches that can become more powerful qualitative tools to investigate more many as-yet-uncultivated bacteria.

Comparing our two experiments, the results of the SIP studies were more interesting taxnomically. SIP studies mimic the in situ environment, and also maintained the connections between individual bacteria. By SIP studies, we can learn and understand what the bacteria demand, and that might lead to more success on culturing them in the future.

Some work needs to be done in the future on this project. First, enriched soil communities identified via SIP should be isolated to pure cultures, especially the members of

Planctomycetes. In this study, to use indican_2 as sole carbon source, we highly enriched some

Planctomycetes as evidenced by SIP studies. However, due to their acidic soil environment, we were not able to process this in the agar plate cultivation studies. In the future, using indican_2 as carbon source and using gellan as solid agent should be processed to isolate novel

Planctomycetes. Second, soil might be pretreated before cultivation studies. Because most

Planctomycetes have a large cell size (Jenkins and Staley 2013), we could use flow cytometry machine (Kvist et al. 2007) to separate and collect the Planctomycetes cells, and then enrich the cell collection on media. After previous enrichment and separation, the cultivation and isolation will become easier to process. Third, 13C EPS extraction techniques on strain S.elodea can be improved. Until now, we only have some gellan plate cultivation experiment results. These results only showed us a tiny part of the whole gellan-degrading communities in the soil. If we can successfully extract 13C labeled gellan in the future, we believe that there will be more novel gellan-degrading species can be found in our 13C gellan SIP studies. Last but not least, we could

108

use the novel culture-independent method-combined SIP with metagenomics (Chen and Murrell

2010) to reconstruct metabolic pathways for EPS-degrading bacteria communities, thus helping to predict the cultivation requirements for isolating these EPS-enriched uncultured bacteria.

Unfortunately, I cannot finish all the improvement for this research in my master study period, but I believe if we continue doing this project in the future we will discover more of the unknown world of bacteria.

109

REFERENCE

Achtman, M. and Wagner, M. 2008. Microbial diversity and the genetic nature of microbial

species. Nature Reviews Microbiology. 6(6): 431-440.

Altschul, S. F., Gish, W., Miller, W., Myers, E.W. and Lipman, D. J. 1990. Basic local alignment

search tool. Journal of Molecular Biology. 215(3): 403–410.

Allensen-Holm, M., Barken, K., Yang, L., Klausen, M., Webb, J., Kjelleberg, S., Molin, S.,

Givskov, M. and Tolker, N. T. 2006. A characterization of DNA release in Pseudomonas

aeruginosa cultures and biofilms. Molecular Microbiology. 59: 1114–1128.

Baker, G. C., Smith, J. J. and Cowan, D. A. 2003. Review and re-analysis of domain-specific

16S primers. Journal of Microbiological Methods. 55(3): 541-555.

Brady, A., Sharp, C. E., Grasby, S. E. and Dunfield, P. F. 2011.Exploring microbial diversity at

geothermal hot springs in British Columbia and potential applications to biotechnology.

From 61st Annual Conference of the Canadian Society of Microbiologists Memorial

University of Newfoundland, St. John’s, Newfoundland.

Becking, J. H. 1984. Genus Beijerinckia Derx 1950, 145AL. In Bergey’s Manual of Systematic

Bacteriology.1: 311–321.

Borneman, J., Skroch, P. W., O’Sullivan, K. M., Palus, J. A., Rumjanek, N. G., Jansen, J. L.,

Nienhuis, J. and Triplett, E. W. 1996. Molecular microbial diversity of an agricultural

soil in Wisconsin. Applied and Environmental Microbiology. 62: 1935–1943.

Bottomley, P. J. 1999. From Principles and applications of soil microbiology. Fuhrmann, J. J.,

Hartel, P., and Zuberer, D. A. (Eds.). New Jersey: Pearson Prentice Hall. pp. 417-420.

Busti, E., Cavaletti, L., Monciardini, P., Schumann, P., Rohde, M., Sosio, M., and Donadio, S.

110

2006. Catenulispora acidiphila gen. nov., sp. nov., a novel, mycelium-forming

actinomycete, and proposal of Catenulisporaceae fam. nov. International Journal of

Systematic and Evolutionary Microbiology. 56(8): 1741-1746.

Caporaso, J. G., Kuczynski, J., Stombaugh, J., Bittinger, K., Bushman, F. D., Costello, E. K.,

Fierer, N., Pena, A. G., Goodrich, J. K., Gordon, J. I., Huttley, G. A., Kelley, S. T.,

Knights, D., Koenig, J. E., Ley, R. E., Lozupone, C. A., McDonald, D., Muegge, B. D.,

Pirrung, M., Reeder, J., Sevinsky, J. R., Turnbaugh, P. J., Walters, W. A., Widmann, J.,

Yatsuneko, T., Zaneveld, J. and Knight, R. 2010. QIIME allows analysis of high-

throughput community sequencing data. Nature Methods. 7(5): 335-336.

Chen, Y. and Murrell, J. C. 2010. When metagenomics meets stable-isotope probing: progress

and perspectives. Trends in Microbiology. 18(4): 157-163.

Cleary, D. F., Smalla, K., Mendonça-Hagler, L. C. and Gomes, N. C. 2012. Assessment of

variation in bacterial composition among microhabitats in a mangrove environment using

DGGE fingerprints and barcoded pyrosequencing. PloS One. 7(1): e29380.

Cronan, C. S. and Grigal, D. F. 1995. Use of calcium/aluminum ratios as indicators of stress in

forest ecosystems. Journal of Environmental Quality. 24(2): 209-226.

Cole, J. R., Wang, Q., Cardenas, E., Fish, J., Chai, B., Farris, R. J., Kulam-Syed-Mohideen A.

S., McGarrell, D. M., Marsh, T., Garrity, G. M. and Tiedje, J. M. 2009. The Ribosomal

Database Project: improved alignments and new tools for rRNA analysis. Nucleic Acids

Research. 37: 141-145.

Conn, H. J. 1909. Future methods of soil bacteriological investigations. Zentbl Bakteriol

Parasitenkd Infektkrankh. 2 (25): 454-457.

Conn, H. J. 1918. The microscopic study of bacteria and fungi in soil. Agriculture Technical

111

Bulletin. 64: 3-20.

Connon, S. A. and Giovannoni, S. J. 2002. High-throughput methods for culturing

microorganisms in very-low-nutrient media yield diverse new marine isolates. Applied

and Environmental Microbiology. 68(8): 3878-3885.

Davis, E. R. K., Joseph, J. S. and Janssen, H. P. 2005. Effects of growth medium, inoculum size,

and incubation time on culturability and isolation of soil bacteria. Applied and

Environmental Microbiology. 71(2): 826-834.

Davis, K. E., Sangwan, P. and Janssen, P. H. 2011. Acidobacteria, Rubrobacteridae and

Chloroflexi are abundant among very slow‐growing and mini‐colony‐forming soil

bacteria. Environmental Microbiology. 13(3): 798-805.

DeAngelis, K. M., Allgaier, M., Chavarria, Y., Fortney, J. L., Hugenholtz, P., Simmons, B.

Sublette, K., Silver, W. L. and Hazen, T. C. 2011. Characterization of trapped lignin-

degrading microbes in tropical forest soil. PloS One. 6(4): e19306.

Dedysh, S. N. 2011. Cultivating uncultured bacteria from northern wetlands: knowledge gained

and remaining gaps. Frontiers in Microbiology. 2(184): 1-15.

Dedysh, S. N., Liesack, W., Khmelenina, V. N., Suzina, N. E., Trotsenko, Y. A., Semrau, J. D.,

Bares, A. M., Panikov, N. S. and Tiedje, J. M. 2000. Methylocella palustris gen. nov., sp.

nov., a new methane-oxidizing acidophilic bacterium from peat bogs, representing a

novel subtype of serine-pathway methanotrophs. International Journal of Systematic and

Evolutionary Microbiology. 50(3): 955-969.

DeSantis, T. Z., Hugenholtz, P., Larsen, N., Rojas, M., Brodie, E. L., Keller, K., Fournier, J.,

112

Voordouw, G. and Gieg, M.L. 2006.Greengenes, a chimera-checked 16s rRNA gene

database and workbench compatible with ARB. Applied and Environmental

Microbiology. 72(7): 5069-5072.

Desantis, T. Z., Hugenholtz, P., Larsen, N., Rojas, M., Brodie, E. L., Keller, K., Huber, T.,

Dalevi, D., Hu, P. and Andersen, G. L. 2006. Greengenes, a chimera-checked 16S rRNA

gene databaseand workbench compatible with ARB. Applied and Environmental

Microbiology. 72(7): 5069-5072.

De Leij, F. A. A. M., Whipps, J. M. and Lynch, J. M. 1994. The use of colony development for

the characterization of bacterial communities in soil and on roots. Microbial Ecology.

27(1): 81-97.

Dojka, M. A., Harris, J. K. and Pace, N. R. 2000. Expanding the known diversity and

environmental distribution of an uncultured phylogenetic division of bacteria. Applied

and Environmental Microbiology. 66(4): 1617-1621.

Dunfield, P. F., Tamas, I., Lee, K. C., Morgan, X. C., McDonald, I. R. and Stott, M. B. 2012.

Electing a candidate: a speculative history of the bacterial phylum OP10. Environmental

Microbiology. 14(12): 3069-3080.

Eboigbodin, K. E. and Biggs, C. A. 2008. Characterization of the extracellular polymeric

substances produced by Escherichia coli using infrared spectroscopic, proteomic, and

aggregation studies. Biomacromolecules. 9: 686- 695.

Eichorst, S. A., Breznak, J. A. and Schmidt, T. M. 2007. Isolation and characterization of soil

bacteria that deﬁne Terriglobus gen. nov., in the phylum Acidobacteria. Applied and

Environmental Microbiology. 73(8): 2708-2717.

Eichorst, S. A. and Kuske, C. R. 2012. Identification of cellulose-responsive bacterial and

113

fungal communities in geographically and edaphically different soils by using stable

isotope probing. Applied and environmental microbiology. 78(7): 2316-2327.

Eilers, H., Pernthaler, J., Glöckner, F. O. and Amann, R. 2000. Culturability and in situ

abundance of pelagic bacteria from the North Sea. Applied and Environmental

Microbiology. 66(7): 3044-3051.

Engelbrektson, A., Kunin, V., Wrighton, K. C., Zvenigorodsky, N., Chen, F., Ochman, H. and

Hugenholtz, P. 2010. Experimental factors affecting PCR-based estimates of microbial

species richness and evenness. The ISME Journal. 4(5):642-647.

Euzeby, J. P. 2013. List of Prokaryotic names with Standing in Nomenclature (LPSN). Online

at: http://www. bacterio. net/ (accessed July 2013).

Everett, K. D., Bush, R. M. and Andersen, A. A. 1999. Emended description of the order

Chlamydiales, proposal of Parachlamydiaceae fam. nov. and Simkaniaceae fam. nov.,

each containing one monotypic genus, revised taxonomy of the family Chlamydiaceae,

including a new genus and five new species, and standards for the identification of

organisms. International Journal of Systematic Bacteriology. 49(2): 415-440.

Flemming, H. and Wingender, J. 2001. Relevance of microbial extracellular polymeric

substances (EPSs): part I. Structural and ecological aspects. Water Science and

Technology. 43: 1-8.

Frostegard, A., Courtois, S., Ramisse,V., Clerc, S., Bernillon, D., Gall, F. Le, Jeannin, P.,

Nesme, X. and Simonet, P. 1999. Quantiﬁcation of bias related to the extraction of DNA

directly from soils. Applied and Environmental Microbiology. 65: 5409-5420.

Fukunaga, Y., Kurahashi, M., Sakiyama, Y., Ohuchi, M., Yokota, A. and Harayama, S. 2009.

114

Phycisphaera mikurensis gen. nov., sp. nov., isolated from a marine alga, and proposal of

Phycisphaeraceae fam. nov., Phycisphaerales ord. nov. and Phycisphaerae classis nov.

in the phylum Planctomycetes. The Journal of General and Applied Microbiology. 55(4):

267-275.

Garrity, G. M., Bell, J. A. and Lilburn, T. G. 2004. Taxonomic outline of the prokaryotes, release

5.0. Springer-Verlag. New York, N.Y.

Gimesi, N. 1924. Hydrobiologiai Tanulmanyok (Hydrobiologische Studien). I: Planctomyces

bekeﬁi Gim. nov. gen. et sp. [in Hungarian, with German translation]. Budapest: Kiadja a

Magyar Ciszterci Rend.

Giovannoni, S. J., Foster, R. A., Rappé, M. S. and Epstein, S. 2007. New cultivation strategies

bring more microbial plankton species into the laboratory. Oceanograph. 20(2): 62-69.

Giovannoni, S. J., Schabtach, E. and Castenholz, R. W. 1987. Isosphaera pallida, gen. and

comb. nov., a gliding, budding eubacterium from hot springs. Archives of Microbiology.

147(3): 276-284.

Gray, N. D. and Head, I. M. 2001. Linking genetic identity and function in communities of

uncultured bacteria. Environment Microbiology. 3(8): 481-492.

Grasby, S. E., Richards, B. C., Sharp, C. E., Brady, A. L., Jones, G. M. and Dunfield, P. F.

2013. The Paint Pots, Kootenay National Park, Canada-a natural acid spring analogue for

Mars 1, 2. Canadian Journal of Earth Sciences. 50(1): 94-108.

Gürtler, V. and Stanisich, V. A. 1996. New approaches to typing and identification of bacteria

using the16S-23S rDNA spacer region. Microbiology. 142(1): 3-16.

Godden, B. and Penninckx, M. J. 1984. Identification and evolution of the cellulolytic

115

microflora present during composting of cattle manure: on the role of actinomycetes sp.

Annales de l'Institut Pasteur/Microbiologie. 135(1): 69-78.

Goh, K. K. T. 2004. Isolation and characterisation of bacterial exopolysaccharides produced by

Lactobacillus delbrueckii subsp. bulgaricus NCFB 2483 and Sphingomonas elodea

ATCC 31461: a thesis presented in partial fulfilment of the requirements for the degree of

Doctor of Philosophy in Food Technology at Massey University, New Zealand. pp. 115-

116.

Goodfellow, M. 2012. Phylum XXVI. Actinobacteria phyl. nov. In Bergey’s Manual® of

Systematic Bacteriology pp. 33-2028. Springer New York.

Gutierrez, T., Berry, D., Yang, T., Mishamandani, S., McKay, L., Teske, A. and Aitken, M. D.

2013. Role of Bacterial Exopolysaccharides (EPS) in the Fate of the Oil Released during

the Deepwater Horizon Oil Spill. PloS One. 8(6): e67717.

Haichar, F. Z., Achouak, W., Christen, R., Heulin, T., Marol, C., Marais, M., Mougel, C.,

Ranjard, L., Balesdent, J. and Berge, O. 2007. Identification of cellulolytic bacteria in

soil by stable isotope probing. Environmental Microbiology. 9(3): 625-634.

Hanson, R. S., Netrusov, A. I. and Tsuji, K. 1991. The obligate methanotrophic bacteria

Methylococcus,Methylomonas, and Methylosinus. In The Prokaryotes. pp. 2350-2365.

Haas, B. J., Gevers, D., Earl, A. M., Feldgarden, M., Ward, D.V., Giannoukos, G., Ciulla, D.,

Tabba, D., Highlander, S. K., Sodergren, E., Methe, B., DeSantis, T. Z., Petrosino, J. F.,

Knight, R. and Birren, B. W. 2011. Chimeric The Human Microbiome Consortium 16S

rRNA sequence formation and detection in Sanger 454-pyrosequenced PCR amplicons.

Genome Research. 21: 494-504.

Hestrin, S. and Schramm, M. 1954. Synthesis of Cellulose by Acetobacter xylinum 2.

116

Preparation of Freeze-Dried Cells Capable of Polymerizing Glucose to Cellulose.

Biochemical Journal. 58: 345-352.

Hottes, A. K., Meewan, M., Yang, D., Arana, N., Romero, P., McAdams, H. H. and Stephens,

C. 2004. Transcriptional profiling of Caulobacter crescentus during growth on complex

and minimal media. Journal of Bacteriology. 186(5): 1448-1461.

Holben, W. E., Feris, K. P., Kettunen, A. and Apajalahti, J. H. 2004. GC fractionation enhances

microbial community diversity assessment and detection of minority populations of

bacteria by denaturing gradient gel electrophoresis. Applied and Environmental

Microbiology. 70(4): 2263-2270.

Hugenholtz , P., Goebel, M. and Pace, N. R. 1998. Impact of culture-independent studies on the

emerging phylogenetic view of bacterial diversity. Journal of Bacteriology. 180: 4765 -

4774.

Janssen, P. H. 2006. Minireviews: Identifying the dominant soil bacterial taxa in libraries of 16S

rRNA and 16S rRNA genes. Applied and Environmental Microbiology. 72(3): 1719-

1728.

Janssen, P. H. 2008. New cultivation strategies for terrestrial microorganisms. In Accessing

Uncultivated Microorganisms: From the Environment to Organisms and Genomes and

Back. Zengler, K. (ed.). Washington, DC, USA. pp. 173–192.

Janssen, P. H. and Hedlund, B. P. 2011. Order I. Chthoniobacterales ord. nov. In Bergey’s

Manual® of Systematic Bacteriology. Springer New York. pp.836.

Janssen, P. H., Yates, P. S., Grinton, B. E., Taylor, P. M. and Sait, M. 2002. Improved

117

culturability of soil bacteria and isolation in pure culture of novel members of the

divisions Acidobacteria, Actinobacteria, Proteobacteria, and Verrucomicrobia. Applied

and Environmental Microbiology. 68(5): 2391-2396.

Jenkins, C. and Staley, J. T. 2013. History, Classification and Cultivation of the Planctomycetes.

In Planctomycetes: Cell Structure, Origins and Biology. pp. 1-38.

Jensen, V. 1968. The plate count technique. In Gray, T.R.G. and Parkinson D. editors. The

ecology of soil bacteria. Liverpool University Press, Liverpool (UK). pp. 158-170.

Joseph, S. J., Hugenholtz, P., Sangwan, P., Osborne, C. A. and Janssen, P. H. 2003. Laboratory

cultivation of widespread and previously uncultured soil bacteria. Applied and

Environmental Microbiology. 69(12): 7210-7215.

Kaeberlein, T., Lewis, K. and Epstein, S. S. 2002. Isolating" uncultivable" microorganisms in

pure culture in a simulated natural environment. Science. 296(5570): 1127-1129.

Kanokratana, P., Mhuantong, W., Laothanachareon, T., Tangphatsornruang, S., Eurwilaichitr, L.,

Pootanakit, K. and Champreda, V. 2013. Phylogenetic Analysis and Metabolic Potential

of Microbial Communities in an Industrial Bagasse Collection Site. Microbial Ecology.

pp. 1-13.

Keller, M. and Zengler, K. 2004. Tapping into microbial diversity. Nature Reviews

Microbiology. 2(2): 141-105.

Kerr, T. J., Kerr, R. D. and Benner, R. 1983. Isolation of a bacterium capable of degrading

peanut hull lignin. Applied and Environmental Microbiology. 46(5):1201-1206.

Kishimoto, N., Kosako, Y. and Tano, T. 1991. Acidobacterium capsulatum gen. nov., sp. nov.

an acidophilic chemoorganotrophic bacterium containing menaquinone from an acidic

mineral environment. Current Microbiology. 22: 1–7.

118

Knief, C., Lipski, A. and Dunﬁeld, P. F. 2003. Diversity and activity of methanotrophic bacteria

in different upland soils. Applied and Environmental Microbiology. 69: 6703-6714.

Koch, I. H., Gich, F., Dunfield, P. F. and Overmann, J. 2008. Edaphobacter modestus gen. nov.,

sp. nov., and Edaphobacter aggregans sp. nov., acidobacteria isolated from alpine and

forest soils. International Journal of Systematic and Evolutionary Microbiology. 58(5):

1114-1122.

Kozdrój, J. and van Elsas, J. D. 2000. Application of polymerase chain reaction-denaturing

gradient gel electrophoresis for comparison of direct and indirect extraction methods of

soil DNA used for microbial community fingerprinting. Biology and Fertility of Soils.

31(5): 372-378.

Krsek, M. and Wellington, E. M. H. 1999. Comparison of different methods for the isolation

and purification of total community DNA from soil. Journal of Microbiological Methods.

39(1): 1-16.

Kulichevskaya, I. S., Ivanova, A. O., Baulina, O. I., Bodelier, P. L., Damsté, J. S. S. and

Dedysh, S. N. 2008. Singulisphaera acidiphila gen. nov., sp. nov., a non-filamentous,

Isosphaera-like planctomycete from acidic northern wetlands. International Journal of

Systematic and Evolutionary Microbiology. 58(5): 1186-1193.

Kulichevskaya, I. S., Baulina, O. I., Bodelier, P. L., Rijpstra, W. I. C., Damsté, J. S. S. and

Dedysh, S. N. 2009. Zavarzinella formosa gen. nov., sp. nov., a novel stalked, Gemmata-

like planctomycete from a Siberian peat bog. International Journal of Systematic and

Evolutionary Microbiology. 59(2): 357-364.

Kvist, T., Ahring, B. K., Lasken, R. S. and Westermann, P. 2007. Specific single-cell isolation

119

and genomic amplification of uncultured microorganisms. Applied Microbiology and

Biotechnology. 74(4): 926-935.

Lage, O. M. and Bondoso, J. 2012. Bringing Planctomycetes into pure culture. Frontiers in

Microbiology. (405): 1-6.

Langer, U., Böhme, L. and Böhme, F. 2004. Classification of soil microorganisms based on

growth properties: a critical view of some commonly used terms. Journal of Plant

Nutrition and Soil Science. 167(3): 267-269.

Lawson, C. J. and Symes, K. C. 1980. Microbial polysaccharide, its preparation and

use. European Patent Application 19800300523.

Lee, K. C., Webb, R. I., Janssen, P. H., Sangwan, P., Romeo, T., Staley, J. T. and Fuerst, J. A.

2009. Phylum Verrucomicrobia representatives share a compartmentalized cell plan with

members of bacterial phylum Planctomycetes. BMC Microbiology. 9(1): 5.

Liesack, W. and Dunfield, P. F. 2002. Biodiversity in soils: use of molecular methods for its

characterization. Encyclopedia of Environmental Microbiology.

Liesack, W. and Stackebrandt, E. 1992. Occurrence of novel groups of the domain Bacteria as

revealed by analysis of genetic material isolated from an Australian terrestrial

environment. Journal of Biotechnology. 174:5072-5078.

Liu, H. and Fang, H. H. P. 2002. Extraction of extracellular polymeric substances (EPS) of

sludges. Journal of Biotechnology. 95(3): 249-256.

Losey, N. A., Stevenson, B. S., Busse, H. J., Damsté, J. S. S., Rijpstra, W. I. C., Rudd, S. and

Lawson, P. A. 2013. Thermoanaerobaculum aquaticus, gen. nov., sp nov., the first

cultivated member of Acidobacteria subdivision 23, isolated from a hot spring.

International Journal of Systematic and Evolutionary Microbiology. 67(7).

120

Ludwig, W., Strunk, O., Westram, R., Richter, L., Meier, H., Buchner, Y. A., Lai, T., Steppi, S.,

Jobb, G., Forster, W., Brettske, I., Gerber, S., Ginhart, A.W., Gross, O., Grumann, S.,

Hermann, S., Jost, R., Konig, A., Liss, T., Lubmann, R., May, M., Nonhoff, B., Reichel,

B., Strehlow, R., Stamatakis, A., Stuckmann, N., Vilbig, A., Lenke, M., Ludwig, T.,

Bode, A. and Schleifer, K. H. 2004. ARB: A software environment for sequence data.

Nucleic Acids Research. 32(4): 1363-1371.

Ludwig, W., Strunk, O., Klugbauer, S., Klugbauer, N., Weizenegger, M., Neumaier, J.,

Bachleitner, M. and Schleifer, K. H. 1998. Bacterial phylogeny based on comparative

sequence analysis (review). Electrophoresis. 19(4), 554-568.

Maier, W., Schumann, B. and Gröger, D. 1990. Biosynthesis of indoxyl derivatives in Isatis

tinctoria and Polygonum tinctorium. Phytochemistry. 29(3): 817-819.

Margulies, M., Egholm, M., Altman, W. E., Attiya, S., Bader, J. S., Bemben, L. A., Berka, J.,

Braverman, M. S., Chen, Y. J., Chen, Z., Dewell, S. B., Du, L., Fierro, J. M., Gomes, X.

V., Godwin, B. C., He, W., Helgesen, S., Ho, C. H., Irzyk, G. P., Jando, S. C., Alenquer,

M. L., Jarvie, T. P., Jirage, K. B., Kim, J. B., Knight, J. R., Lanza, J. R., Leamon, J. H.,

Lefkowitz, S. M., Lei, M., Li, J., Lohman, K. L., Lu, H., Makhijani, V. B., McDade, K.

E., McKenna, M. P., Myers, E. W., Nickerson, E., Nobile, J. R., Plant, R., Puc, B. P.,

Ronan, M. T., Roth, G. T., Sarkis, G. J., Simons, J. F., Simpson, J. W., Srinivasan, M.,

Tartaro, K. R., Tomasz, A., Vogt, K. A., Volkmer, G. A., Wang, S. H., Wang, Y., Weiner,

M. P., Yu, P., Begley, R. F. and Rothberg, J. M. 2005. Genome sequencing in

microfabricated high-density picolitre reactors. Nature. 437(7057): 376-380.

Markowitz, V. M., Szeto, E., Palaniappan, K., Grechkin, Y., Chu, K., Chen, I. M. A, Dubchak,

121

I., Anderson, I., Lykidids,A.,Mavromatis, K., Ivanova, N.N. and Kyrpides, N. C. 2008.

The integrated microbial genomes (IMG) system in 2007: data content and analysis tool

extensions. Nucleic Acids Research. 36(1): D528-D533.

Metting Jr, F. B. 1992. Soil microbial ecology: applications in agricultural and environmental

management. Marcel Dekker, Inc. pp.11-17.

Monciardini, P., Cavaletti, L., Schumann, P., Rohde, M. and Donadio, S. 2003. Conexibacter

woesii gen. nov., sp. nov., a novel representative of a deep evolutionary line of descent

within the class Actinobacteria. International Journal of Systematic and Evolutionary

Microbiology. 53: 569-576.

Morgan, J. W., Forster, C. F. and Evison, L. M. 1990. A comparative study of the nature of

biopolymers extracted from anaerobic and activated sludges. Water Research. 6: 743-

750.

Narzisi, G. and Mishra, B. 2011. Comparing de novo genome assembly: the long and short of

it. PLoS One. 6(4):e19175.

Neufeld, J. D., Vohra, J., Dumont, M. G., Lueders, T., Maneﬁeld, M., Friedrich, M. W. and

Murrell, J. C. 2007. DNA stable-isotope probing. Nature Protocols. 2(4): 860-866.

Nogales, B., Moore, E. R., Llobet-Brossa, E., Rossello-Mora, R., Amann, R. and Timmis, K. N.

2001. Combined use of 16S ribosomal DNA and 16S rRNA to study the bacterial

community of polychlorinated biphenyl-polluted soil. Applied and Environmental

Microbiology. 67(4):1874-1884.

Overmann, J. 2010. Novel cultivation strategies for environmentally important microorganisms.

In Geomicrobiology: Molecular and Environmental Perspective pp. 69-89.

Pankratov, T. A. and Dedysh, S. N. 2010. Granulicella paludicola gen. nov., sp. nov.,

122

Granulicella pectinivorans sp. nov., Granulicella aggregans sp. nov. and Granulicella

rosea sp. nov., acidophilic, polymer-degrading acidobacteria from Sphagnum peat bogs.

International journal of systematic and evolutionary microbiology. 60(12): 2951-2959.

Parikh, V. M. and Jones, J. K. N. 1963. The structure of the extracellular polysaccharide of

Azotobacter indicum. Canadian Journal of Chemistry. 41(11): 2826-2835.

Pieretti, I., Royer, M., Barbe, V., Carrere, S., Koebnik, R., Couloux, A., Darrasse, A.,Gouzy, J.,

Jacques, M. A., Lauber, E., Manceau, C., Mangenot, C., Poussier, S., Segurens, B.,

Szurek, B., Verdier, V., Arlat, M., Gabriel, D. W. Rott, P. and Cociancich, S. 2012.

Genomic insights into strategies used by Xanthomonas albilineans with its reduced

artillery to spread within sugarcane xylem vessels. BMC Genomics. 13(1):658.

Pruesse, E., Quast, C., Knittel, K., Fuchs, B. M., Ludwig, W., Peplies, J. and Glöckner, F. O.

2007. SILVA: a comprehensive online resource for quality checked and aligned

ribosomal RNA sequence data compatible with ARB. Nucleic Acids Research.

35(21):7188-7196.

Quast, C., Pruesse, E., Yilmaz, P., Gerken, J., Schweer, T., Yarza, P., Pepline, J. and Glöckner,

F. O. 2013. The SILVA ribosomal RNA gene database project: improved data processing

and web-based tools. Nucleic Acids Research. 41(D1): D590-D596.

Ramos-Padrón, E., Bordenave, S., Lin, S., Bhaskar, I. M., Dong, X., Sensen, C.W., Fournier,J.,

Vorrdouw, G. and Gieg, L. M. 2011.Carbon and Sulfur Cycling by Microbial

Communities in a Gypsum-Treated Oil Sands Tailings Pond. Environmental Science and

Technology. 45(2): 439-446.

Rappe´, M. S. and Giovannoni, S. J. 2003. The uncultured microbial majority. Annual Review of

Microbiology.57:369-394.

123

Reichenbach, H., Lang, E., Schumann, P. and Spröer, C. 2006. Byssovorax cruenta gen. nov.,

sp. nov., nom. rev., a cellulose-degrading myxobacterium: rediscovery of ‘Myxococcus

cruentus’ Thaxter 1897. International Journal of Systematic and Evolutionary

Microbiology. 56(10): 2357-2363.

Rezaei, F., Xing, D., Wagner, R., Regan, J. M., Richard, T. L. and Logan, B. E. 2009.

Simultaneous cellulose degradation and electricity production by Enterobacter cloacae in

a microbial fuel cell. Applied and Environmental Microbiology. 75(11): 3673-3678.

Ross, P., Mayer, R. and Benziman, M. 1991. Cellulose biosynthesis and function in

bacteria. Microbiological Reviews. 55(1): 35-58.

Ronaghi M. 2001. Pyrosequencing sheds light on DNA sequencing. Genome Research. 11: 3-11.

Sá-Correia, I., Fialho, A. M., Videira, P., Moreira, L. M., Marques, A. R. and Albano, H. 2002.

Gellan gum biosynthesis in Sphingomonas paucimobilis ATCC 31461: genes, enzymes

and exopolysaccharide production engineering. Journal of Industrial Microbiology and

Biotechnology. 29(4): 170-176.

Sait, M., Hugenholtz, P. and Janssen, H. P. 2002. Cultivation of globally distributed soil bacteria

from phylogenetic lineages previously only detected in cultivation-independent surveys.

Applied and Environmental Microbiology. 4: 654-666.

Sait, M., Davis, K. E., and Janssen, P. H. 2006. Effect of pH on isolation and distribution of

members of subdivision 1 of the phylum Acidobacteria occurring in soil. Applied and

Environmental Microbiology. 72(3):1852-1857.

Schramm M and Hestrin S. 1954. Synthesis of Cellulose by Acetobacter xylinum 1.

Micromethod for the Determination of Cellulose. Biochemical Journal. 56:163-166.

Schoenborn, L., Yates, P. S., Grinton, B. E., Hugenholtz, P. and Janssen, P. H. 2004. Liquid

124

serial dilution is inferior to solid media for isolation of cultures representative of the

phylum-level diversity of soil bacteria. Applied and Environmental Microbiology. 70(7):

4363-4366.

Santos, P., Pinhal, I., Rainey, F. A., Empadinhas, N., Costa, J., Fields, B., Banson, R.

Verissimon, A. and da Costa, M. S. 2003. GammaProteobacteria Aquicella lusitana gen.

nov., sp. nov., and Aquicella siphonis sp. nov. Infect Protozoa and Require Activated

Charcoal for Growth in Laboratory Media. Applied and Environmental Microbiology.

69(11): 6533-6540.

Sharp C. E., Stott, M. B. and Dunfield, P. F. 2012. Detection of autotrophic verrucomicrobial

methanotrophs in a geothermal environment using stable isotope probing. Frontiers in

Microbiology. 3(303): 1-9.

Schellenberger, S., Drake, H. L. and Kolb, S. 2011. Functionally redundant cellobiose

degrading soil bacteria respond differentially to oxygen. Applied and Environmental

Microbiology. 77(17): 6043-6048.

Schlesner, H. and Hirsch, P. 1987. Rejection of the genus name Pirella for pear-shaped budding

bacteria and proposal to create the genus Pirellula gen. nov. International Journal of

Systematic and Evolutionary Microbiology. 37: 441.

Schmidt, J. M. 1978. Isolation and ultrastructure of freshwater strains of Planctomyces. Current

Microbiology. 1(2): 65-70.

Schlesner, H. and Stackebrandt, E. 1986. Assignment of the genera Planctomyces and Pirella

toanew family Planctomycetaceae fam. nov. and description of the order

Planctomycetales ord. nov. Systematic and Applied Microbiology. 8: 174-176.

Schrempf, H. and Walter, S. 1995. The cellulolytic system of Streptomyces reticuli.

125

International Journal of Biological Macromolecules. 17(6): 353-355.

Singleton, D. R., Furlong, M. A., Peacock, A. D., White, D. C., Coleman, D. C. and Whitman,

W. B. 2003. Solirubrobacter pauli gen. nov., sp. nov., a mesophilic bacterium within the

Rubrobacteridae related to common soil clones. International Journal of Systematic and

Evolutionary. 53:485-490.

Spiegelman, D., Whissell, G. and Greer, C. W. 2005. A survey of the methods for the

characterization of microbial consortia and communities. Canadian Journal of

Microbiology. 51(5): 355-386.

Staley, J. T. 1973. Budding bacteria of the Pasteuria-Blastobacter group. Canadian Journal of

Microbiology. 19: 609-614.

Staley, J. T. 2006. The bacterial species dilemma and the genomic–phylogenetic species

concept. Philosophical Transactions of the Royal Society B: Biological Sciences.

361(1475):1899-1909.

Staudt, C., Horn, H., Hempel, D. C, Neu, T. R. 2004. Volumetric measurements of bacterial cells

and extracellular polymeric substance glycoconjugates in biofilms. Biotechnology and

Bioengineering. 88(5): 585-592.

Stevenson ,B. S., Eichorst, S. A., Wertz, J. T., Schmidt, T. M. and Breznak, J. A. 2004. New

strategies for cultivation and detection of previously uncultured microbes. Applied and

Environmental Microbiology. 70: 4748-4755.

Stott, M. B., Crowe, M. A., Mountain, B. W., Smirnova, A. V., Hou, S., Alam, M. and Dunfield,

P. F. 2008. Isolation of novel bacteria, including a candidate division, from geothermal

soils in New Zealand. Environmental Microbiology. 10(8): 2030-2041.

Strous, M., Fuerst, J. A., Kramer, E. H., Logemann, S., Muyzer, G., van de Pas-Schoonen, K. T

126

Webb R., Kuenen J. G. and Jetten, M. S. 1999. Missing lithotroph identified as new

planctomycete. Nature. 400(6743): 446-449.

Suzuki, M., Rappé, M. S. and Giovannoni, S. J. 1998. Kinetic bias in estimates of coastal

picoplankton community structure obtained by measurements of small-subunit rRNA

gene PCR amplicon length heterogeneity. Applied and Environmental Microbiology.

64(11): 4522-4529.

Tamaki, H., Hanada, S., Sekiguchi, Y., Tanaka, Y. and Kamagata, Y. 2009. Effect of gelling

agent on colony formation in solid cultivation of microbial community in lake sediment.

Environmental Microbiology. 11(7): 1827-1834.

Tamaki, H., Tanaka, Y., Matsuzawa, H., Muramatsu, M., Meng, X. Y., Hanada, S., Mori, K. and

Kamagata, Y. 2011. Armatimonas rosea gen. nov., sp. nov., of a novel bacterial phylum,

Armatimonadetes phyl. nov., formally called the candidate phylum OP10. International

Journal of Systematic and Evolutionary Microbiology. 61(6):1442-1447.

Tamas, I., Dedysh, S. N., Liesack, W., Stott, M. B., Alam, M., Murrell, J. C. and Dunfield, P. F.

2010. Complete Genome Sequence of Beijerinckia indica subsp. indica. Journal of

Bacteriology. 192(17): 4532-4533.

Thrash, J. C. and Coates, J. D. 2010. Phylum XVII. Acidobacteria phyl. nov. InBergey’s

Manual® of Systematic Bacteriology. pp. 725-735. Springer New York.

Torsvik, V., Goksøyr, J. and Daae, F. L. 1990. High diversity in DNA of soilbacteria. Applied

and Environmental Microbiology. 56: 782-787.

Trujillo-Cabrera, Y., Ponce-Mendoza, A., Vásquez-Murrieta, M. S., Rivera-Orduña, F. N. and

Wang, E. T. 2013. Diverse cellulolytic bacteria isolated from the high humus, alkaline-

saline chinampa soils. Annals of Microbiology. pp. 1-14.

127

Turner, E. C. and Jones, B. 2005. Microscopic calcite dendrites in cold‐water tufa:

Implications for nucleation of micrite and cement. Sedimentology. 52(5):1043-1066.

Vartoukian, S. R., Palmer, R. M. and Wade, W. G. 2010. Strategies for culture of

‘unculturable’bacteria. FEMS Microbiology Letters. 309(1): 1-7.

Watson, R. J. and Blackwell, B. 2000. Purification and characterization of a common soil

component which inhibits the polymerase chain reaction. Canadian Journal of

Microbiology. 46(7):633-642.

Williams, K. P. and Kelly, D. P. 2013. Proposal for a new Class within the Proteobacteria, the

Acidithiobacillia, with the Acidithiobacillales as the type Order. International Journal of

Systematic and Evolutionary Microbiology.

Woese, C.R. 1987. Bacterial evolution. Microbiological Reviews.51: 221–271.

Woese, C. R., Kandler, O. and Wheelis, M. L. 1990. Towards a natural system of organisms:

proposal for the domains Archaea, Bacteria, and Eucarya. Proceedings of the National

Academy of Sciences. 87(12): 4576-4579.

Woese, C. R., Stackebrandt, E., Macke, T. J. and Fox, G. E. 1985. A phylogenetic definition of

the major eubacterial taxa. Systematic and Applied Microbiology. 6(2): 143-151.

Zengler, K., Toledo, G., Rappé, M., Elkins, J., Mathur, E. J., Short, J. M. and Keller, M. 2002.

Cultivating the uncultured. Proceedings of the National Academy of Science. 99(24):

15681-15686.

128

APPENDIX A: MEDIA

Table A: Composition of DNMS 7 Component (per L)

MgSO4·7H2O 0.2 g

CaCl2 0.04 g

Sequestrene Fe 0.001 g

Trace elements 0.5 ml

NH4Cl (1.85mM) 0.1 g

Adjust to required pH before autoclaving (S.elodea incubation pH ~7).

-1 after autoclaving add 10 ml L of 100 mM PO4 buffer pH 7 with sterile syringe and filter make

100 mM Na2HPO4 and 100mM KH2PO4 mix together until the pH is 7.

* Composition of trace element solution is shown in Table C.

Table A_1: Composition of DNMS5.8 + Indican_2 medium. Component (per L)

Indican _2 0.5 g

MgSO4·7H2O 0.2 g

CaCl2 0.04 g

Sequestrene Fe 0.001 g

Trace elements 0.5 ml

NH4Cl (1.85mM) 0.1 g

Agar 15 g

Adjust to pH 5.8 before autoclaving.

129

-1 after autoclaving add 10 ml L of 100 mM PO4 buffer pH 5.8with sterile syringe and filter make

100 mM Na2HPO4 and 100mM KH2PO4 mix together until the pH is 5.8.

* Composition of trace element solution is shown in Table C.

Table A_2: Composition of DNMS 3.5 + gellan medium. Solution 1

Component (per L)

CaCl2 0.04 g

Sequestrene Fe 0.001 g

Trace elements 0.5 ml

NH4Cl (1.85 mM) 0.1 g

Adjust pH to 3.5

Solution 2

Component (per L) phytagel 15 g

MgSO4·7H2O 0.2 g

Add phytagel slowly with vigorous stirring to avoid clumping.

Adjust pH to 3.5

-1 after autoclaving add 10 ml L of 100 mM PO4 buffer pH 3.5 with sterile syringe and filter make

100 mM Na2HPO4 and 100mM KH2PO4 mix together until the pH is 3.5.

* Composition of trace element solution is shown in Table C.

130

Table A_3: Composition of DNMS 5.8 + gellan medium. Solution 1

Component (per L)

CaCl2 0.04 g

Sequestrene Fe 0.001 g

Trace elements 0.5 ml

NH4Cl (1.85 mM) 0.1 g

Adjust pH to 3.5

Solution 2

Component (per L) phytagel 15 g

MgSO4·7H2O 0.2 g

Add phytagel slowly with vigorous stirring to avoid clumping.

Adjust pH to 5.8

-1 after autoclaving add 10 ml L of 100 mM PO4 buffer pH 5.8 with sterile syringe and filter make

100 mM Na2HPO4 and 100mM KH2PO4 mix together until the pH is 5.8.

* Composition of trace element solution is shown in Table C.

Table B: Composition of buffered S&H medium

Component (per L)

Peptone 5.0 g

Yeast extract 5.0 g

131

Na2HPO4 2.7 g

Citric acid ·H2O 1.15 g

50% Glucose (filter-sterilized) 40.0 ml

Agar (if necessary) 15.0 g

Distilled water 960.0 ml

Adjust medium for final pH 5.0 with HCl. For solid medium, autoclave agar separately.

Aseptically add filter-sterilized glucose solution.

Table C: Composition of trace element solution Component (per L)

. ZnSO4 7 H2O 440 mg

. CuSO4 5 H2O 200 mg

. MnSO4 2 H2O 170 mg

. Na2MoO4 2 H2O 60 mg

H3BO3 100 mg

. CoCl2 6 H2O 80 mg dH2O 980ml

132

Table D: B.indica incubation medium Component (per L) glucose (13C or 12C) 10.0 g

K2HPO4 0.8 g

KH2PO4 0.2 g

MgSO4× 7H2O 0.1 g

FeSO4× 7H2O 20.0 mg

MnSO4× 6H2O 2.0 mg

Zn SO4× 6H2O 5.0 mg

Cu SO4× 6H2O 4.0 mg

Na2MoO4 × 2H2O 5.0 mg

Agar 15.0 g

Distilled water 950 ml

Adjust PH to 6.5. Sterilize glucose separately (10 g in 50 ml H2O) and mix after cooling

133

APPENDIX B: EPS STRUCTURE

Cellulose structure

β(1→4) linked D-glucose units

This image was taken from (http://en.wikipedia.org/wiki/File:Microbialcellulosestructure.jpg)

Indican_2 Structure :

[D-glucuronic acid β(1→3) D-glucose β(1→4) D-glycero-D-mannoheptose β(1→4)]n

(Parikh and Jones 1963)

This image was taken from Parikh and Jones (1963) permission attracted in Appendix C.

Gellan structure:

[D-glucose (β1→4) D-glucuronic acid (β1→4) D-glucose (β1→4) L-rhamnose (α1→3)]n

(Symes 1982)

This image was taken from (www.kelco.com)

134

APPENDIX C: PERMISSIONS

License Number 3202691471966 License date Aug 05, 2013 Licensed content NRC Research Press publisher Licensed content Canadian Journal of Chemistry publication Licensed content title THE STRUCTURE OF THE EXTRACELLULAR POLYSACCHARIDE OF AZOTOBACTER INDICUM Licensed content author V. M. Parikh, J. K. N. Jones Licensed content date Nov 1, 1963 Type of Use Thesis/Dissertation Volume number 41 Issue number 11 Requestor type Academic Format Print and electronic Portion Figure/table Number of figures/tables 1 Order reference number Title of your thesis / Identification and Cultivation of Exopolysaccharide-Degrading dissertation Bacteria in Two Soils Expected completion Aug 2013 date Estimated size(pages) 100 Total 0.00 USD

License Number 3185560250117 License date Jul 10, 2013 Licensed content NRC Research Press publisher Licensed content Canadian Journal of Earth Sciences publication Licensed content title The Paint Pots, Kootenay National Park, Canada — a natural acid spring analogue for Mars1,2 Licensed content Stephen E. Grasby, Barry C. Richards, Christine E. Sharp, Allyson author L. Brady, Gareth M. Jones, Peter F. Dunfield Licensed content date Jan 1, 2013 Type of Use Thesis/Dissertation Volume number 50 Issue number 1

135

Requestor type Academic Format Print and electronic Portion Figure/table Number of 2 figures/tables Order reference None number Title of your thesis / Identification and Cultivation of Exopolysaccharide-Degrading dissertation Bacteria in Two Soils Expected completion Aug 2013 date Estimated size(pages) 100 Total 0.00 USD

Dear Emily,

Thank you for contacting Copyright Clearance Center's RightsLink service. RightsLink provides permission for the reproduction and distribution of copyrighted materials in print and electronic formats on behalf of copyright owners who list their titles with us.

As per the ASM:

ASM authorizes an advanced degree candidate to republish the requested material in his/her doctoral thesis or dissertation. If your thesis, or dissertation, is to be published commercially, then you must reapply for permission.

If you have any further questions, please contact a Customer Service Representative at 877-622-5543, Monday-Friday, 8:00 AM to 6:00 PM Eastern Time, or by e-mail at [email protected].

Sincerely,

Leon Dequito Customer Service Representative Copyright Clearance Center 222 Rosewood Drive Danvers, MA 01923 +1.855.239.3415 Toll Free +1.978.646.8600 Fax www.copyright.com

Please click here to participate in our online customer service survey: http://www.cvent.com/d/bdqgxd

136