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IDENTIFICATION OF ACTIVE SOIL RDX BIODEGRADING

MICROORGANISMS USING 15N STABLE ISOTOPE
PROBING

presented by

INDUMATHY JAYAMANI

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5108 K:lProj/Aoc&Pres/ClRC/DateDue.indd

IDENTIFICATION OF ACTIVE SOIL RDX BIODEGRADING
MICROORGANISMS USING 15N STABLE ISOTOPE PROBING

By

Indumathy J ayamani

A THESIS

Submitted to
Michigan State University
in partial fulfilhnent of the requirements
for the degree of

MASTER OF SCIENCE
Environmental Engineering

2009

ABSTRACT

IDENTIFICATION OF ACTIVE SOIL RDX BIODEGRADING MICROORGANISMS
USING 15N STABLE ISOTOPE PROBING

By
Indumathy J ayamani
15N DNA stable isotope probing was used to identify microorganisms responsible for
degradation of hexahydro-l,3,5-trinitro-1,3,5-triazine (RDX) from soil microcosms. An
agricultural soil was amended with either unlabeled or labeled (13C315N3) RDX along

with added mineral salts medium and glucose. Following RDX degradation monitored
through HPLC analysis, DNA was extracted from both sets of microcosms. The DNA
samples were then subject to isopycnic density gradient ultracentrifugation, fractionation,

followed by terminal restriction fragment length polymorphism on ‘heavy’ fractions. One

‘ . . . 13 15 .
fragment was dominant In heavy fractlons of C N-RDX amended samples, but not In

the unlabeled controls indicating label uptake by this organism from RDX. Sequencing of
the total DNA indicated the organisms involved in RDX transformation belonged to the
class of Sphingobacteria and Acidobacteria. These organisms have not been associated
with RDX degradation so far, but have been noted for their ability to degrade many other
xenobiotic compounds such as MTBE, BTEX, tetracyclines and PCBs. This study
indicates the potential for identifying more non-indigenous RDX degrading

microorganism from uncontaminated soils.

ACKNOWLEDGEMENTS

I am thankful to my research advisor Dr. Cupples for giving me the opportunity to work
on this project. She also needs to be thanked for her continuous support and guidance
without which this thesis would not be possible. I also would like to thank Dr. Marsh, for
his guidance and for generously allowing me to use the GeneScan Software at his
laboratory. I am also thankfiIl to Dr. Hashsham for being on my committee and as well a

mentor.

Many thanks to Strategic Environmental Research and Developmental Program (SERDP)

for funding this work through the 2008 SEED grant (ER1606).

Thanks to Michael Manzella a doctoral student from Microbiology and Molecular
Genetics, for his contribution to this project. My extended thanks to Melissa Knapp and
Weimin Sun for their valuable advice, guidance and troubleshooting techniques

whenever I needed them. Thanks to Yanlyang Pan, Daniel Williams and Dr. Cha for their
help with the analytical instruments. My special thanks to faculty and staff at the

Department of Civil and Environmental for their continuous support.

The success of this study is due to the support of the faculty, staff and students at

Michigan State University and I am thankful to all who contributed either directly or

indirectly.

TABLE OF CONTENTS

LIST OF TABLES .......................................................................................................................... vi
LIST OF FIGURES ........................................................................................................................ vii
ABBREVIATIONS ........................................................................................................................ xii
1.0 INTRODUCTION ...................................................................................................................... 1
1.1 General ................................................................................................................................... 1
1.2 RDX ....................................................................................................................................... 2
1.3 Toxicity of RDX ..................................................................................................................... 2
1.4 RDX as an environmental contaminant .................................................................................. 3
1.5 Study objectives ..................................................................................................................... 4
2.0 BIOREMEDIATION OF RDX — A REVIEW .......................................................................... 5
2.1 General ................................................................................................................................... 5
2.2 Biodegradation pathways and products .................................................................................. 6
2.3 Anaerobic biodegradation .................................................................................................... 12
2.4 Aerobic degradation ............................................................................................................. 14
2.5 Fungal biodegradation .......................................................................................................... 15
2.6 Degradation of metabolites .................................................................................................. 16
3.0 STABLE ISOTOPE PROBING — A REVIEW ....................................................................... 17
3.1 General ................................................................................................................................. 17
3.2 Markers used in SIP ............................................................................................................. 20
3.3 DNA based SIP .................................................................................................................... 22
3.3.1 General methodology .................................................................................................... 22
3.3.2 DNA SIP in microbial ecological studies ...................................................................... 23
3.3.3 DNA SIP in bioremediation studies .............................................................................. 25
3.3.4 Isotopes and methodological considerations ................................................................. 26

4.0 IDENTIFICATION OF HEXAHYDRO-l,3,5-TRINITRO-l,3,5-TRIAZINE DEGRADING

MICROORGANISMS USING 15N STABLE ISOTOPE PROBING .......................................... 28
4.1 Introduction .......................................................................................................................... 28
4. 2 Materials and methods ........................................................................................................ 31

4.2.1 Chemicals ...................................................................................................................... 31
4.2.2 Soil incubations ............................................................................................................. 31
4.2.3 RDX extraction and HPLC analysis ............................................................. . ................. 33
4.2.4 DNA extraction and ultracentrifilgation ........................................................................ 33
4.2.5 TRFLP and sequencing ................................................................................................. 34
4.3 Results .................................................................................................................................. 37
4.3.1 RDX biodegradation ..................................................................................................... 37
4.3.2 TRFLP results of SIP .................................................................................................... 40
4.3.3 Sequencing .................................................................................................................... 44
4.4 Discussion ............................................................................................................................ 45
4.5 Conclusion and future studies .............................................................................................. 47

iv

Appendix A .................................................................................................................................... 49

Appendix B .................................................................................................................................... 50
Appendix C .................................................................................................................................... 56
REFERENCES ............................................................................................................................... 87

LIST OF TABLES

Table 2.1 - RDX degrading strains and possible biodegradation pathways ..................... 11
Table 2.1 — RDX degrading strains...continued................................................-. .............. 12
Table 2.2 - RDX degrading fungi and possible biodegradation pathways ....................... 12
Table 2.3 - Microorganisms that degrade RDX metabolites ............................................ 16
Table A.1 - List of soils tested and experimental details .................................................. 49

vi

LIST OF FIGURES

Figure 2.1 - Molecular structures of hexahydro-l,3,5-trinitro-1,3,5-triazine (RDX), -1,3-
dinitroso-5-nitro-1,3,5-tn'azine (DNX), hexahydro-l,3,5-t1initroso-1,3,5-t1iazine (TNX),
methylenedinitramine (MDNA), bis-(hydroxymethyl)nitramine (BHNA), 4-nitro-2,4-

diazabutanal (N DAB) and triamino-RDX .......................................................................... 8
Figure 2.2 - Proposed biodegradation pathways for RDX .................................................. 9
Figure 3.1 - Overview of DNA-SIP using fractionation and TRFLP ............................... 19

Figure 4.1 - RDX concentration in SIP microcosms (triplicate live controls, killed
controls and labeled samples) set up with soil 3 (Intial RDX concentration — 45 uM) and
with acetonitrile ....................................................................................................... 39

Figure 4.2 - RDX concentration in microcosms (triplicate live controls, killed controls
and labeled samples) set up with soil 3 (Intial RDX concentration — 45 uM) and with
acetonitrile ......................................................................................................................... 39

Figure 4.3 - The relative abundance of the fragment of length 260 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90uM of labeled and unlabeled .......................................... 41

Figure 4.4 - TRF LP profiles of first two heavy fractions fiom soil amended with labeled
RDX (”CISN) showing 260-bp fragment and its abundance ............................................ 42

Figure 4.5 - Dominance of 260-bp fragment in heavy fiactions from labeled samples in
comparison to abundance in heavy fractions (of corresponding BD) from unlabeled
samples .............................................................................................................................. 43

Figure B] - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 1 and was allowed to go to Oz-depleted conditions ........................................... 50

Figure B.2 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 2 .......................................................................................................................... 50

Figure 8.3 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 3 and was allowed to go to 02-depleted conditions (160 mL bottles) ............... 51

Figure 3.4 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 3 and was aerated daily (160 mL bottles) .......................................................... 52

Figure B.5 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 4 and was allowed to go to Oz depleted conditions (160 mL bottles). .............. 53

vii

Figure 3.6 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 4 and was aerated daily (160 mL bottles) .......................................................... 53

Figure B.7 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 5 and was aerated daily (160 mL bottles) .......................................................... 54

Figure B.8 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 6 and was allowed to go to 02-depleted conditions ........................................... 54

Figure 3.9 - RDX concentration in microcosms (duplicate live) set up with soil 7, 8, 9, 10
and no acetonitrile (Oz-depleted conditions) .................................................................... 55

Figure 3.10 - RDX concentration in microcosms (triplicate live and killed controls) set
up with soil 8, no acetonitrile and was aerated daily ........................................................ 55

Figure C.1 - The relative abundance of the fragment of length 63 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 56

Figure C.2 - The relative abundance of the fragment of length 70.5 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 57

Figure C.3 - The relative abundance of the fragment of length 72 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 58

Figure C.4 - The relative abundance of the fragment of length 126 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 59

Figure C.5 - The relative abundance of the fragment of length 172 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 60

Figure C.6 - The relative abundance of the fragment of length 196 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 61

Figure C.7 - The relative abundance of the fragment of length 198 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 pM of labeled and unlabeled RDX ................................ 62

viii

Figure C.8 - The relative abundance of the fragment of length 204 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 63

Figure C.9 - The relative abundance of the fragment of length 208 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 64

Figure C.10 - The relative abundance of the fragment of length 215 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 65

Figure C.11 - The relative abundance of the fragment of length 226 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 66

Figure C.12 - The relative abundance of the fragment of length 228 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 67

Figure C.13 - The relative abundance of the fragment of length 236 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 68

Figure C. 14 - The relative abundance of the fragment of length 251 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 69

Figure C. 1 5 - The relative abundance of the fragment of length 258 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 70

Figure C.16 - The relative abundance of the fragment of length 263 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 71

Figure C.17 - The relative abundance of the fragment of length 271 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 72

Figure C. l 8 - The relative abundance of the fragment of length 272 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 73

Figure C.19 - The relative abundance of the fragment of length 274 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 74

Figure C.20 - The relative abundance of the fragment of length 281 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 75

Figure C.2l - The relative abundance of the fiagrnent of length 292 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 pM of labeled and unlabeled RDX ................................ 76

Figure C.22 - The relative abundance of the fragment of length 302 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 77

Figure C.23 - The relative abundance of the fragment of length 306 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 78

Figure C.24 - The relative abundance of the fragment of length 316 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 79

Figure C.25 - The relative abundance of the fragment of length 328 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 80

Figure C.26 - The relative abundance of the fragment of length 402 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 81

Figure C.27 - The relative abundance of the fragment of length 404 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 82

Figure C.28 - The relative abundance of the fragment of length 448 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 83

Figure C.29 - The relative abundance of the fragment of length 462 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 84

Figure C.30 - The relative abundance of the fragment of length 885 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 85

Figure C.3l - The relative abundance of the fragment of length 920 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX ................................ 86

xi

bp
CL20

CsCl

CSTFA

DGGE

DNX

DNA

dNTP

EDTA

EPA

GTN

HMX

HPLC

kb

LB

MeBr

MeCl

PAH

PCB

PCR

PETN

ABBREVIATIONS
base pair(s)
2, 4, 6, 8, 10, lZ-hexonitrohexazaisowurtzitane
cesium chloride
cesium trifluoroacetate
denaturing gradient gel electrophoresis
hexahydro-l ,3-dinitroso-5-nitro-l ,3 ,S-triazine
deoxyribose nucleic acid
dinucleotide triphosphate
ethylenedinitraminetetraacetic acid
Environmental Protection agency
glycerol trinitrate, nitroglycerine
high melting explosive, octahydro-l,3,5,7-tetranitro-1,3,5.7-tetrazocine
high performance liquid chromatography
kilobase pair(s)
Luria Bertani broth
methyl bromide
methyl chloride
hexahydro-l -nitroso-3 ,5-dinitro- l ,3 ,5-triazine
polyaromatic hydrocarbon
polychlorinated biphenyls
polymerase chain reaction

pentaerithritol tetranitrate

xii

rDNA
rRNA

SIP

TRFLP

X-gal

ribosomal DNA

ribosomal RNA

stable isotope probing

royal demolition explosive, hexahydro-l,3,5-trinitro-1,3,5-triazine
ribose nucleic acid

total DNA

2,4,6-trinitrotoluene

hexahydro-l ,3,5-trinitroso -1 ,3,5-triazine

terminal restriction fragment length polymorphism

5-bromo-4-chloro-3-indolyl-®-D-galactopyranoside

xiii

1.0 INTRODUCTION

This thesis is divided into four chapters and three appendices. Chapter one provides an
overview and introduces RDX as a problem contaminant. Chapter two provides a review
of the biological degradation pathways and strains isolated for transformation of RDX.
This is followed by a review of the molecular technique stable isotope probing (SIP) and
its applications in bioremediation and microbial ecology. Chapter four presents the
experimental methods, materials, key results, discussion and conclusions from the present

study.

1.1 General

Xenobiotic compounds are chemicals that are human made many of which are present in
the environment at a relatively high concentration. Examples are pesticides, herbicides,
pharmaceuticals, explosives, benzene, toluene, ethylbenzene & xylenes (collectively
known as BTEX), methyl tert-butyl ether (MTBE), polycyclic aromatic compounds
(PAH’S) and polychlorinated biphenyls (PCB’s). Environmental contamination by
organic xenobiotic compounds is a growing global problem (1). These contaminants
persist and accumulate in the environment depending upon their fate and transport
mechanisms. One family of xenobiotics that has been of concern since the early 1900’s is

the explosives.

Explosives are materials which when suitably initiated, undergoes very rapid self-
propagating decomposition resulting in the release of energy. Detonation of explosives

produces more stable products, release of heat and a sudden pressure effect by the action

1

of heat. Explosive chemicals are in general high in nitrogen and oxygen content. Widely
used explosives include nitrate esters (Example: glycerol trinitrate, nitroglycerine (GTN)
and pentaerithritol tetranitrate (PETN)), nitroaromatics (Example: picric acid and 2,4,6-
trinitrotoluene (TNT)) and nitramines (Example: hexahydro-l,3,5-trinitro-1,3,5-triazine
(RDX), octahydro-1,3,5,7-tetranitro-l,3,5.7-tetrazocine (HMX), 2, 4, 6, 8, 10, 12-

hexonitrohexazaisowurtzitane (CL-20)).

1.2 RDX

The nitramine explosive RDX is the most widely used explosive after TNT and is the
most important military high explosive in the United States (82). RDX was also the first
nitramine explosive to be developed. It was discovered in the 1890’s by Hans Hemmings,
when he introduced it as a medicine. It was patented and produced in 1920’s by direct
nitration of hexamine. The US. Bachmann Process for continuous generation of RDX
was developed in the 1940’s (23). It is aspowerful as the nitrate esters but less sensitive
and hence preferred (7). It is primarily used in combination with other plastic explosives

and detonators and is rarely used alone.

1.3 Toxicity of RDX

The USEPA has classified RDX as a class C human carcinogen (90) and has
recommended a life time health advisory of 2ug RDX/L. RDX exerts its primary toxic
effects on the central nervous system in addition to affecting the gastrointestinal and renal
tracts (35) in humans and laboratory animals. However, experiments with rats (25), zebra
fish (71) and earthworms (79) have revealed that RDX also affects the reproductive
system or produces smaller offspring in exposed animals. RDX has also been shown to

2

bio-accumulate in plants and animals (45, 81) denoting a potential danger through food
chain transfer to higher level organisms. RDX was once used as a rat poison and has been
found to cause weight loss and affect offspring size in rats (58). RDX was also shown to
be toxic to humans. RDX processing plant employees who were possibly exposed to
powdered RDX through inhalation, suffered convulsions and unconsciousness (51). A 3
year old child who had ingested RDX pellets become epileptic while all her other body
functions were found normal (95). All of these observed effects on humans and other
animals were short term and were reversible. However, the toxic and carcinogenic nature

of RDX is still questionable and the dangers associated with it should not be overlooked.

1.4 RDX as an environmental contaminant

RDX was extensively used both during and after World War 11. Although it is not
commercially manufactured in the United States, it is still produced, handled, packaged
and deployed at various army ammunition sites in United States (82). These activities
have caused contamination of land, water and air. In addition, past practices of improper
disposal of wastewater from RDX production plants has also caused land and water

pollution (87).

RDX contamination is a significant problem in several countries, including United States,
United Kingdom, Canada, Germany and Australia (82). In the United States alone,
explosives have been found in 115 sites at 25 installations amounting to 45,000 tonnes of

contaminated soil (82). Maximum levels of RDX contamination (e.g. 27 g RDX/kg soil)

at some sites are Significantly above the US. Environmental Protection Agency’s

recommended clean up level of 5.8 mg RDX per kg soil (47, 82).

Contamination of ground water due to leaching of explosives from soil is also a problem.

Alhough RDX has low water solubility (maximum 38 mg/L at 20°C, (82)), its low

sorption coefficient (Kd =0.8 L/kg,(82)) makes it a potential ground water pollutant. Thus

RDX’s potential to migrate quickly in soil and pollute potential water sources makes it a
contaminant of concern. Also, the natural attenuation of RDX in water is shown to

produce nitrate, a known contaminant (19).

1.5 Study objectives

1. To screen environmental samples for RDX biodegradation.

2. To identify the microorganisms responsible for RDX biodegradation in these

complex communities using 15N stable isotope labeling.

2.0 BIOREMEDIATION OF RDX - A REVIEW

A summary of RDX biodegradation pathways and the degradation products are presented
in this chapter. This chapter also provides a list of known RDX degraders and their
corresponding proposed biodegradation pathways. The information is presented by
categorizing these pathways and degraders as aerobic, anaerobic or fungal degraders. A
brief note on the microorganisms that degrades RDX metabolites is provided at the end of

this chapter.

2.1 General

RDX has been considered recalcitrant but a review of the literature reveals the potential
for biodegradation. Microbial mediated degradation of RDX was first illustrated in 1981
by McCormick et. a1. (66). Since then, many bacterial strains and fungi have shown to be
capable of degrading RDX under various conditions. RDX biodegradation takes place
under aerobic, anaerobic and microaerobic (46), nitrate reducing (39), sulfate reducing
(l8), methanogenic (3), manganese reducing (22), iron-reducing (54, 75) and acetogenic
conditions (4). RDX biotransforrnation has also been reported to occur in different types
of environments: surface, subsurface, vadose zone, marine (12, 99), aquifer (10), fresh

water and sewage sludges (9).

In many cases, RDX degradation is achieved by adding a carbon source and.or another
nitrogen source resulting in a diauxic grth (17). In contrast to this, RDX degradation
by other organisms is inhibited by the presence of other organic or inorganic nitrogenous

compounds (72).

2.2 Biodegradation pathways and products

Degradation of RDX occurs through different pathways depending upon the conditions
and the microorganism(s). The molecular structures of RDX and a number of the reported
degradation products are illustrated in Figure 2.1. As summarized by Crocker et. a1.
(2006) (32) three mechanisms have been proposed for RDX degradation: two-electron

reduction, denitration and direct enzymatic cleavage. The two electron reduction

mechanism involves the addition of redox equivalents (2e-/2H+) to RDX, to form the

reduced nitroso-derivatives and the hypothesized hydroxylamino—derivatives and
triamino—derivatives. Denitration of RDX occurs by the addition of a single electron
forming the anion radical RDX' followed by ring cleavage. Enzymatic cleavage of RDX
refers to the breaking of OH bonds or C-N bonds or N-N bonds in the RDX molecule by

direct enzymatic attack.

There are seven proposed RDX degradation pathways based on these three basic
degradation mechanisms(32) as summarized by Crocker et. a1. (2006) (32) (Figure 2.2).
They include (a) the reduction of RDX to nitroso derivatives before ring cleavage first
observed and proposed by McCormick et. a1. (1981) (66); (b) the reduction of RDX to
1,3,5-t1iamino-l,3,5-triazine first observed and proposed by Zhang and Hughes (2003)
(98); (c) the reduction of RDX via Aspergillus niger nitrate oxidoreductase enzyme first
observed and proposed by Bhushan et. a1. (2000) (14); (d) the direct enzymatic cleavage
of RDX first observed and proposed by Hawaii et. a1. (2000) (46); (e) the anaerobic
denitration of RDX first observed and proposed by Zhao et. a1. (2002) (100); (f) the intial

reduction to MNX followed by denitration of RDX first observed and proposed by Zhao

et. a1. (2003) (103); and (g) aerobic denitration of RDX first observed and proposed by

Fournier et. a1. (2002) (37).

The reported degradation products (intermediate and end products) of RDX are
hexahydro-l—nitroso-3,5-dinitro-l,3,5-triazine (MNX), hexahydro-l,3-dinitroso-5-nitro-
1,3,5-triazine (DNX), hexahydro-l,3,5-trinitroso-1,3,5-triazine (TNX), 4-nitro-2,4-
diazabutanal (N DAB), methylenedinitramine (MDNA), carbon dioxide, methanol,
formaldehyde, nitrite, nitrate, nitrous oxide and triamino-RDX. In addition, several
hydroxylamino-derivatives are postulated to form as intermediates during RDX

biodegradation. The enzymatic ring cleavage products are still unknown. The degradation

products also differ based upon the electron acceptor (02) conditions. The ring nitroso

degradation products (reduced forms of RDX) are reported to form only during anaerobic
transformation of RDX and have not been observed during aerobic degradation of RDX.
Formaldehyde, when formed as a result of RDX degradation, is believed to be
transformed to methanol and formic acid and fiirther to carbon dioxide and methane
when acetogenic and methonogenic bacteria are present (32). To date, the majority of
studies have used traditional laboratory culture techniques to isolate RDX degrading
bacteria. A summary of known RDX degrading bacteria has been provided (Tables 2.1.,

2.2., 2.3).

s I ‘N’ ‘N’
r’H (NW r’H
. , ,N N, ’0 ,N N~
O~N’N\/N‘Nvo N V N' N \/ N
O O O O O O
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.0. o. ,o
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r W c)"N'N\/N‘N”O H0.C/N\C.OH
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TNX BHNA
NH2
H H N
o‘ .N. .N. 20 I/ j
‘N C C N N
'6 H2 H HZN' V ‘NH2
NDAB TRIAMINO-RDX

Figure 2.1 - Molecular structures of hexahydro-l,3,5-trinitro-1,3,5-triazine (RDX), -1,3-
dinitroso-S-nitro-l ,3,5-triazine (DNX), hexahydro-l,3,5-trinitroso—1 ,3,5-triazine (TNX),
methylenedinitramine (MDNA), bis-(hydroxymethyl)nitramine (BHNA), 4-nitro-2,4-
diazabutanal (N DAB) and triamino-RDX

Figure 2.2 - Proposed biodegradation pathways for RDX (reproduced from Crocker et. al.
(2006) (32)). Path a Reduction of RDX to nitroso derivatives followed by ring cleavage.
Path b Reduction of RDX to triamino-RDX. Path c Reduction of RDX via Aspergillus
niger nitrate oxidoreductase enzyme. Path d Direct enzymatic cleavage of RDX. Path e
Anaerobic denitration of RDX. Path f Denitration of RDX via the reductive intermediate
MNX. Path g Aerobic denitration of RDX

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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10

Table 2.1 - RDX degrading strains and possible biodegradation pathways

 

T

 

Strain Proposed pathway Reference
Morganella morganii B2 A1 (52, 53)
Providencia rettgeri Bl A (52)
Citrobacterfi'eundii N82 A (52)
Stenotrophomonas maltophila PBl ND (17)
Serratia marcescens A (97)
Rhodococcus Sp. Strain DN22 G (31, 37)
Enterobacter cloacae strain 96-3 A (53)
Rhodococcus rhodochrous sp. llY G (83)
Klebsiella pneumoniae Strain SCZ-l A, E, F (100)
Clostridium bifermentans HAW-1 A, F (103, 104)
Clostridium bifermentans HAW-G3. A, F (104)
Clostridium acetobutylicum C (98)
Desulfovibrio desulfuricans HAW-ESZ A, F (104)
Acetobacterium malicum HAAP-l E, F (3)
Shewanella halifaxensis sp. HAW-EB4 A, F (102, 105)
Shewanella sp. HAW-EBl A, F (105)
Shewanella sp. HAW-EB2 A, F (105)
Shewanella sp. HAW-BBS A, F (105)
Clostridium bifermentans HAW-G4 A, F (104)
Clostridium bifermentans HAW-E3 A, F (104)
Clostridium bifermentans HAW-HCl A, F (104)
Desulfovibrio sp. HAW-EB18 A, F (105)
Clostridium sp. HAW-E317 A, F (105)
Fusobacteria isolate HAW-E821 A, F (105)
Shewanella sediminis sp. HAW-EB3 A, F (101, 105)
Methylobacterium sp. strain B] 001 A (92)
Methylobacterium organophilum A (92)
Methylobacterium extorquens A (92)
Methylobacterium rhodesianum A (92)
Clostridium sp. EDB2 F (13, 15)
Williamsia sp. KTR4 (3* (s 8)
Gordonia sp. KTR9 G (88)
Acetobacterium paludosum ND (84)
Halomonas sp. HAW-0C4 C (12)
Marinobacter sp. HAW-0C1 C (12)
Pseudoalteromonas sp. HAW-0C2 C (12)

 

T Figure 2.2 explains the 7 different proposed pathways using the same alphabet codes.

1 ND Not enough data to determine the pathway

Table 2.1 — RDX degrading strains. . .continued.

 

1.

 

Strain Proposed pathway Reference
Pseudoalteromonas sp. HAW-0C5 CI (1 2)
Bacillus sp. HAW-0C6 C (12)
Rhizobr'um rhizogenes BL ND (55)
Burkholderia sp. ND (55)
Pseudomonas putida ND (29)
Fifteen strains belonging to phyla Actinobacteria, ND (80)

-Pr0teobacteria and y-Proteobacteria

 

T Figure 2.2 explains the 7 different proposed pathways using the same alphabet codes.
1 ND Not enough data to determine the pathway

Table 2.2 - RDX degrading fiingi and possible biodegradation pathways

 

 

Fungi Proposed pathwayI Reference
Phanerochaete chrysosporium ND1 (8, 36, 85)
Aspergillus niger C (14)
Cladosporium resinae ND (8)
Cunninghamella echinulata ND (8)
varelegans

Cyathus pallidus ND (8)
Rhodotorula HAW-OCF 1 ND (1 1)
Bullera HAW-OCF2 ND (1 l)
Acremom'um HAW-OCF3 C, E (1 l)
Penicillium HAW-OCFS ND (11)
Cladosporium cladosporioides ND (55)

 

1' Figure 2.2 explains the 7 different proposed pathways using the same alphabet codes.
1 ND Not enough data to determine the pathway

2.3 Anaerobic biodegradation

RDX biodegradation was first studied under anaerobic condition in mixed cultures

obtained from contaminated soil or industrial sludge (46, 66). McCormick et. al. (1981)

12

(66) reported the first confirmed anaerobic microbial degradation of RDX. They reported
the formation of formaldehyde and methanol indicating ring cleavage. The authors also
reported the detection of hydrazines, and postulated a two-electron pathway for RDX
degradation (Figure. 2.2., Path 3). This pathway also was shown to be followed by many
bacterial strains that were later isolated including Klebsiella pneumoniae strain SCZ—l,
Clostridium bzfermentans strain HAW-l, Shewanella halifaxensis strain HAW-EB4, and
Shewanella sp., Methylobacterium sp., Enterobacteria, Shewanella sp. HAW-E32, as a
major or minor pathway (38, 52, 92, 100, 102, 103, 105). A type I nitroreductase that was
cloned and sequenced from Enterobacter cloacae strain 96-3 degraded RDX through

nitroreductase activity (53) and also oxidized NADPH in the presence of RDX.

An alternate pathway that begins with two electron reduction was observed in a study

with the cell-free extracts of Clostridium acetobutylium (98) (Figure 2.2., path b). This

strain utilized H2 as electron donor and transformed RDX to mono-,di, tri-nitroso

derivatives and mono-, di-, tri-amino derivatives. These compounds did not disappear
during the study and hence RDX was not completely mineralized in this pathway. A
nitrate oxidoreductase was extracted from the fungi Aspergillus niger (l4) and was
shown to transform RDX in the presence of NADPH as electron donor. Though MNX
and MDNA were observed as intermediate degradation products, they were fiirther
transformed to nitrous oxide, formaldehyde and ammonium ion (Figure 2.2., Path c).

Another anaerobic pathway significantly different from that postulated by McCormick et
al was postulated by Hawari et. al. (2000) (46) (Figure 2.2., path (1). In this pathway, the

RDX ring was degraded by enzymatic attack on the ON bonds, leading to the generation

13

of the hydroxylamine intermediates, MDNA and BHNA. These intermediates further

decomposed in water to nitrous oxide, methanol, formic acid and formaldehyde.

Denitration of RDX appears to be a major route of RDX biotransformation (32).
Anaerobic denitration of RDX could occur directly or after a reduction step in which
RDX is transformed to MNX first and then denitrified (Figure 2.2., path e, i). Anaerobic
denitration involves a single electron transfer to form the anion RDX' and subsequent
loss of a nitro group. This destabilizes the molecule leading to ring cleavage and
formation of MDNA. Alternatively, RDX could be first reduced to MNX followed by
denitration. These routes can either be a major or minor pathway in different organisms

(100,103)

2.4 Aerobic degradation

The first report on aerobic biodegradation of RDX identified three pure strains of
Corynebacterium capable of utilizing RDX as a sole source of nitrogen (96). Later
several strains including Stenotrophomonas maltophia PBl, Rhodococcus sp. strain
DN22 and llY, Williamsia sp. KTR4 and Gardenia sp. KTR9 were isolated with the
capability to degrade RDX aerobically (17, 31, 83, 88). Although aerobic and anaerobic
denitration involves the loss of nitro group, they are both slightly different mechanisms.
Aerobic denitration of RDX involves two one-electron transfers leading to the loss of two
nitro groups resulting in ring cleavage (37). As a result NDAB, nitrous oxide,
ammonium, formaldehyde and carbon dioxide was formed (Figure 2.2., path g). The

aerobic bacteria Williamsia sp. and Gardenia sp. isolated by Thompson et. al. (2005)

14

(88), degrade RDX using this pathway and utilize it as a carbon, nitrogen and energy
source suggesting that they are able to link the catabolic pathway for RDX with the

anabolic pathway for carbon and nitrogen assimilation (32).

Although RDX biodegradation has been studied from the early 1980’s, only recently has
a gene corresponding to RDX degradation been identified. The prA gene encoding a
constitutively expressed, fused flavodoxin-cytochrome P450 enzyme was successfully
identified fi'om Rhodococcus rhodochrous Sp. NY (83). Bhushan et .al. (2003) (16) later
provided evidence that RDX degradation catalyzed by a rabbit liver cytochrome P450
enzyme and the strain Rhodococcus DN22 produced the same degradation products
(N DAB). They also found that RDX biotransformation by the prA protein and the rabbit
liver cytochrome P450 enzyme was three times faster under anaerobic conditions when
compared to aerobic conditions (16). Roh et. a1. (2009) (80) recently used this gene to
design specific primers and identify RDX degrading strains by combining it with a

powerful molecular probing technique called stable isotope probing (SIP, chapter 3).

2.5 Fungal biodegradation

A white rot fungi Phanerochaete chrysosporium was found to transform both RDX and
TNT to carbondioxide and nitrous oxide (36, 85). However the studies conducted with P.
chrysosporium did not observe any other biodegradation products at detectable
concentrations, therefore pathways has not been completely elucidated. Further studies

have shown several other fungi are capable of degrading RDX (8, l 1, 55).

15

2.6 Degradation of metabolites

Although many bacteria mineralize RDX to harmless gaseous product, some metabolites
such as NDAB, MNX are recalcitrant in certain biodegradation systems (37, 88). Thus
there is a concern over producing more toxic intermediates during RDX bioremediation.
However, growing evidence indicates a number of degradation products can be degraded.
by other organisms (Table 2.3) therefore biodegradation of RDX continues to be a

valuable cleanup technique.

Table 2.3 - Microorganisms that degrade RDX metabolites

 

 

Strain/Fungi RDX metabolite Reference
Methylobacterium sp. J S178 NDAB (38)
Phanerochaete chrysosporium NDAB (37)
Klebsiella pneumoniae Strain SCZ-I MNX (100)

 

16

3.0 STABLE ISOTOPE PROBING - A REVIEW

This chapter provides a brief summary of stable isotope probing methodology. It also
outlines the different markers that are used in SIP studies and their advantages and
disadvantages. Following this a review of SIP studies, involving microbial ecology is
presented. Subsequently, a review of a select number of studies utilizing DNA-SIP for
bioremediation of environmental contaminants is provided. Finally, a brief note on the
various isotopes used in SIP experiments along with their methodological consideration

is presented.

3.1 General

In addition to an understanding of the contaminants physico-chemical properties, having
an accurate insight of microbial community and function is necessary to assess microbial
degradation of xenobiotic contaminants. Because less than 1% of the total microbial
population is cultivable (89), culture independent techniques that use molecular
biomarkers facilitate a more holistic study of microbial community . Although 16S rRNA
genes are useful to determine the phylogeny of organisms present in a sample, such
information does not always provide insight into function in situ. Metagenomics, an
approach to develop gene libraries of the environmental genome, has offered a new way

to assess the microbial community and functions of even uncultivable microorganisms.

Metagenomics, may fail to identify low-abundance species, and may not completely
represent the diversity. The introduction of a method called stable isotope probing (SIP)

has changed this limitation. Stable isotope probing involves the use of an isotopically

17

labeled compound and aims at linking function to identity while attempting to maintain
experimental conditions closer to in situ. The method is based on the uptake of a label by
microorganisms in mixed cultures and analysis of the biomarkers obtained from the
labeled microorganisms to reveal identity of organisms that were involved. Figure 3.1
provides an overview of the method. In the first step, the environmental sample is
incubated with labeled or unlabeled substrate (killed and live) in microcosms. After the
DNA has been extracted from the labeled sample and live control, it is subjected to
ultracentrifugation and fractionation, followed by terminal restriction fragment length
polymorphism (TRFLP) to identify the phylotypes that are quantitatively dominant in the

‘heavy’ fractions of samples but not controls.

18

     

Sample microcosms Control (live) Control (killed)

with labeled RDX microcosms with microcosms with
(‘3C‘5N) unlabeled RDX unlabeled RDX
(12C14N) (12C14N)
DNA extraction

w

11

Isop ycnic density
gradient centrifugation

A II A.
Fractioning to separate the

U light (background) from \J'
o heavy DNA (label 0

° incorporated) °

 

 

 

 

9%"

\

Identification of p hylotyp es that are
quantitatively dominant in heavy fractions of
samples but not controls (live) using TRFLP

Figure 3.1 - Overview of DNA-SIP using fractionation and TRFLP

19

3.2 Markers used in SIP

Phospholipid fatty acids (PLFA) and nucleic acids have been extensively used as
biomarkers for SIP. Phospholipid fatty acids were the first biomarkers used, in a study by
Boschker et. al. (1998) (21) . This study was focused on organisms noted for their

biogeochemical processes, the sulfate-reducing bacteria that utilized acetate as carbon
source and methane reducing bacteria in aquatic sediments, using 13C labeled acetate and
methane respectively. Their findings showed that a gram-positive Desulfotomaculum
acetoxidans were dominant in 13C-labeled acetate uptake and not the most widely studied

Desulfobacter spp. A type I methanotrophic bacteria possibly belonging to the genera
Methylobacter or Methylomicrobium was identified as the dominant methane oxidizing

organism in this study.

Although attempts to use stable isotopes to study phylogeny and firnctionality were used
during the 20th century, the term ‘stable isotope probing’ came into existence after its
first use by Radejewski et. al. (2000) (78). These researchers grew microorganisms on

13C labeled CH3OH and separated the 13C (heavy) DNA from 12C (light) DNA by

isopycnic density gradient centrifugation in a CsCl/EtBr gradient. The heavy DNA was
used to construct 16S rRNA clone libraries. The dominant methylotrophs were identified

as bacteria from the a-Proteobacteria and Acidobacterium.

The third type of biomarker that followed PLFA and DNA was RNA. RNA stable isotope

probing was first performed and proved practical by Manefield et. al. (2002) (65). These

20

researchers used the approach to study phenol degrading communities in an industrial
bioreactor. Total community DNA and RNA were extracted at various time points and

the researchers showed the level of label enrichment in RNA was much higher than in

DNA during the same time period. The 13C labeled RNA was separated from unlabeled

RNA by equilibrium (isopycnic) density gradient centrifugation in CsTFA, fractionation
and was then analyzed using reverse transcriptase-PCR (RT-PCR) and denaturing
gradient gel electrophoresis (DGGE). They identified the organism that dominated

carbon acquisition from phenol belonging to the genus T hauera.

All three biomarkers (PLFA, DNA, RNA) have been widely used under a variety of
experimental conditions, substrates, time of exposure, label incorporation etc and each
has its own advantages and disadvantages. As summarized by Neufeld JD et. al. (2007)
(74), PLFA-SIP is the most sensitive of these biomarkers and DNA-SIP is the least
sensitive as the label incorporation depends upon DNA replication, which is slow.
However, the highly sensitive PLFA is not useful in studying uncultured bacteria as the
unique PLFA patterns of all microorganisms are not known (34). The need for longer
incubation time or increased substrate concentration (that does not mirror the in situ
conditions) to result in better separation of the heavy and light background nucleic acid
material in DNA/RNA-based SIP, can be overcome by fractionation in CsCl or CsTFA
(73). The shortcomings of nucleic acid SIP include cross feeding, if the study uses longer
incubation periods and requirement of higher than typical or in situ substrate conditions.
Nonetheless, nucleic acid SIP, particularly DNA-SIP is still considered a unique

approach to link function to identity for microorganism in complex samples. In addition,

21

functional genes can be targeted (41). Though DNA-SIP is ideal to identify organisms
that use a particular compound as their sole source of carbon or nitrogen, the spectra of
its use in recent times has widened to study organisms that use other environmentally

important compounds as described below.

3.3 DNA based SIP

3.3.1 General methodology

Stable isotope probing requires the use of a labeled substrate (preferably highly labeled)
supplied to a mixed community sample. Such experiments have been conducted in
microcosms or directly in situ. Following label uptake, the total DNA is extracted and
subject to equilibrium (isopycnic) density gradient centrifugation to separate labeled and
unlabeled DNA. The gradient is then either fractioned or the labeled (heavy) DNA is
extracted using a needle and syringe for downstream analysis. The DNA in the heavy
fractions (or “heavy DNA”) is PCR amplified and can be fingerprinted using tools such
as TRF LP, DGGE, cloning and sequencing. Alternatively, real-time PCR with specific
primers can be used to quantify the targeted organisms at various times points to identify
dominant species. Besides using the SSU rRNA genes for phylogenetic analysis,
functional marker genes that encode enzymes for specific activity can be used to target

microorganisms of geochemical importance.

22

3.3.2 DNA SIP in microbial ecological studies

DNA SIP has been extensively used to study microbial communities of environmental

importance. In particular, it has been utilized in studies of various C1 compounds,

denitrifiers and rhizosphere-microbial interactions. The pioneering DNA-SIP study (78)
illustrated the dominant methylotrophs in an oak forest soil belonged to Acidobacterium
and a-Proteobacteria, which were previously not associated with methanol assimilation.
They utilized primers specific to the mxaF gene that codes (ll-subunit of the methanol
dehydrogenase to target the dominant methylotrophs in the ‘heavy’ DNA. Many studies
that followed also utilized DNA SIP with both 16S rRNA and functional genes to identify

active methylotrophs in various natural environments(67).

Methanotrophs have been widely studied using DNA-SIP. Morris SA et. al. 2002 (70)

characterized the active methanotrophic population in a peat soil using 16S rRNA genes

and three other fiinctional genes encoding CH4 oxidation pathway. They identified a

novel methanotrophic organism closely related to Methylocella palustris along with

organisms from a subclass of Proteobacteria as active organisms in methane

assimilation. Another conducted DNA-SIP using l3CH4 in microcosms that closely relate

to environmental conditions, in microcosms amended with water and microcosms
amended with mineral salts medium (27). While treatments best reflecting the
environmental conditions had cross feeding and slow label uptake, the samples amended

with mineral salts medium labeled a less diverse population of methanotrophs. They

23

observed the diversity of the treatments amended with moisture, with 80% type I

methanotrophs and 20% type II methanotrophs best reflected the actual in situ diversity.

In a study focused on ammonia oxidizing bacteria, although the Nitrosomonas sp.

dominated in fresh water and a Nitrosospira sp. dominated in brackish water, further

investigation with 13C DNA SIP revealed that Nitrosomonas sp. were still the dominant

active ammonium oxidizers in marine environment (40). Similarly in a recent study, Jia
et. al. 2009 (50) found that Archaea are dominant by population, even though bacteria
dominate ammonia oxidation in agricultural soils. DNA-SIP was also applied to study

acetate or methanol assimilating bacteria under nitrate reducing conditions in sludge (43,

76) using 13C acetate. Both studies identified the dominant degraders as closely related to

Comamonadaceae and Rhodocyclaceae in the subclass of B-Proteobacteria.

Miller et. al. (2004) (69) identified methyl chloride and methyl bromide degrading
organisms in soils where these gases are naturally released. While the organisms from
Burkholderia dominated MeBr degradation, organisms related to Rhodobacter,
Lysobacter and Nocardioides were found to degrade MeCl. In a later study using DNA-
SIP to investigate MeCl utilizing bacteria, it was reported that DNA-SIP can identify a
more diverse group of organisms involved in biodegradation compared to enrichment and

isolation techniques (20).

24

3.3.3 DNA SIP in bioremediation studies

Bioremediation of persistent xenobiotics has proved to be effective and ecologically
friendly (91). A wide variety of microorganisms capable of utilizing many pollutants
have been isolated under laboratory settings. However, those isolated and characterized
in a laboratory may not be able to degrade the pollutant under typical environmental
conditions. Thus identifying active degraders (whether dominant or not) using DNA-SIP

is important for the ultimate application of bioremediation technologies.

There have been many pollutants studied using DNA. Most are carbon rich organic

compounds studied using 13C isotope labeling. A field based study conducted by DeRito

et. al. (2005) (33) utilized DNA-SIP under three different 12C and 13C phenol doses and

enrichment conditions to identify both the primary active phenol degraders and the

organisms that assimilate the 13C02 respired from phenol degradation. While a diverse

group of phenol degraders (a-,B-,y-Prote0bacteria) were found in the single dosed
unenriched setup, the primary phenol degraders in the enriched setup were found to be

members of the genera Kocuria and Staphylococcus. They also identified a Pseudomonas

as the dominant cross-feeders of the 13C from phenol degradation.

Many organisms have been isolated with the ability to utilize a variety of the PAH

compounds as a carbon source. Singleton et. al. (2005) (86) studied organisms that
degrade naphthalene, phenanthrene and salicylate in a bioreactor. Combining 13C DNA
with DGGE enabled these researchers to determine that salicylate and naphthalene were

25

transformed by Pseudomonas and Ralstonia species, while phenantharene was primarily

utilized by a bacterium related to the genus Acidovorax.

In addition PCB degradation has been studied using SIP. While cultivation studies
indicated the Rhodococcus sp. dominated in biphenyl degradation, DNA-SIP combined
with T-RFLP analysis identified that the Pseudonocardia sp. dominated in biphenyl

utilization (56, 57). BTEX compounds have also been a focus of study (2, 5). Luo et. al.

(2009) (62) identified a novel TM7 strain capable of degrading toluene using 13’C DNA-

SIP.

3.3.4 Isotopes and methodological considerations

Ideally an isotope of any element present in the DNA (C, H, N or 0) could be used in
SIP. However, the higher the proportion of the element in the biomarker the greater the
signal will be. This is because SIP depends on the increase in buoyant density of the

DNA and the higher the label in the DNA the higher the increase in DNA buoyant

density. Thus the 13C label is extensively used as it provides results in high label

incorporation and therefore a clear separation of the labeled and unlabeled DNA.

However, in recent times (24, 80) there have been attempts to use 15N label to study

several nitrogen rich compounds. Interestingly, 15N isotope labeling was used as early as

1958 by Meselson and Stahl (68) to study DNA replication.

26

Laboratory culture based techniques have isolated many strains degrading nitrogen rich
compounds. However there is great interest in identifying organisms that are responsible
for degradation in situ, or under experimental conditions representing in situ conditions.
The low percentage of nitrogen in DNA (average C/N ratio in DNA, 21:1) (26) and the
risk of toxicity of these compounds if present in higher concentration restrict the amount

of label incorporation and are important limitations to this approach. The required 40 -50

atom% 15N-DNA (variable based on the G+C content) enrichment for a clear separation
of labeled and unlabeled DNA is high compared to the required 20% atom 13C-DNA

enrichment or the 10% atom 13C-RNA enrichment (26).

Thus nitrogen-labeling SIP might not yield a strong signal, and hence methodological
changes have been proposed (26). These include, centrifuging the DNA-CsCl mixture at
a low speed for longer duration (140000 X g for 69 hours) and employing 100% labeled

and unlabeled DNA from pure culture to use as boundaries. In addition to these

considerations, the amount of 15N labeled DNA obtained and used for downstream

analysis is also vital in a successful SIP study.

Thus with the application of these methodological changes and understanding its

limitations 15N stable isotope labeling can be applied to study biological uptake of

environmentally important compounds.

27

4.0 IDENTIFICATION OF HEXAHYDRO-l,3,5-TRINITRO-l,3,5-
TRIAZINE DEGRADING MICROORGANISMS USING 15N
STABLE ISOTOPE PROBIN G

In the current study, 15N DNA stable isotope probing was used to identify

microorganisms responsible for degradation of hexahydro-l,3,5-trinitro-1,3,5-triazine
(RDX) fiom soil microcosms. Following label uptake, the extracted DNA was subject to
ultracentrifugation and fractionation. Terminal restriction fragment length polymorphism
(TRF LP) was utilized to identify phylotypes that were quantitatively dominant in heavy
fractions in samples but not in the controls. The results indicate the organisms involved in
RDX transformation belonged to the class of Sphingobacteria and Acidobacteria. This
chapter details the methodology used including both analytical and molecular methods.
This chapter also provides the TRFLP, cloning and sequencing results, followed by the

discussion and conclusions. Finally, a brief note on future studies is provided.

4.1 Introduction

Hexahydro-l,3,5-tlinitro-1,3,5-trazine (RDX) is a nitramine explosive that has been
widely used since World War II and has caused significant contamination of land and
water in and around military ranges at United States. In particular, the physical-chemical
properties of RDX, including low water solubility, low sorption to soil and nonvolatile
nature, have resulted in significant groundwater contamination. For example, RDX has
been found at concentrations as high as 36 mg/L in groundwater in the Iowa Army

Ammunition Plant (87). RDX has shown to be toxic to humans, terrestrial animals and

28

aquatic system. The US. EPA has classified RDX as a type C carcinogen and estimated a

life time exposure health advisory of 2 u g of RDX/L.

Two common methods for contaminated site cleanup are incineration and composting
(82). While incineration does not remove RDX completely (48) it also potentially creates
more toxic products. In contrast, composting (94) reduces the toxicity and mutagenicity
of the contaminants potentially resulting in successful engineered bioremediation.
Numerous microorganisms have been isolated with the potential to transform RDX to
less harmful end products under either anaerobic or aerobic conditions (Table 2.1, 2.2).
However, successful in situ bioremediation requires knowledge on the microorganisms
able to transform the contaminant under conditions typical of the contaminated site, i.e.
mixed culture, complex samples. This represents a knowledge gap for effective
bioremediation of RDX, because the majority of information on RDX degrading

microorganisms has originated from pure culture or enrichments experiments.

To address this knowledge gap, one study has recently (2009) attempted to identify the

microorganisms responsible for RDX degradation in mixed culture, complex samples
using stable isotope probing (SIP) (80). These researchers used l5N - ring labeled RDX
to enrich the DNA of RDX degrading microorganisms from explosive contaminated
groundwater. The extracted heavy DNA was PCR amplified using both 16S rRNA gene
and the prA functional gene (83) specific to RDX degradation and the active RDX

degraders were identified through 16Sr RNA gene sequencing as belonging to

Actinobacteria, a-Proteobacteria and y-Proteobacteria.

29

Here we present a study utilizing 15N DNA stable isotope probing to identify potential

RDX degraders from a soil previously unexposed to the contaminant but likely to contain

a diverse microbial community.

30

4. 2 Materials and methods
4.2.1 Chemicals

Unlabeled RDX and ring labeled RDX (15N3, 13C3; 50% N Labeled) (>99%) dissolved

in acetonitrile were purchased from Cambridge Isotope Laboratories (Andover, MA,
USA). Reagents were either purchased form Sigma-Aldrich (St.Louis, MO, USA), Fisher
BioReagent (New Jersey, USA), Invitrogen (Carlsbad, CA, USA) unless otherwise
stated. Acetonitrile (HPLC grade; 29.8% purity) was purchased from EMD Chemicals

Inc (New Jersey, USA).

4.2.2 Soil incubations

Soil samples used were either collected from agricultural sites (previously unexposed to
RDX) or BTEX contaminated sites in Michigan. The agricultural sites had been
previously amended with biosolids from a wastewater treatment plant, with the last
application being within 1 to 4 years before sample collection. Soils were manually
sorted, homogenized, air dried and sieved through a 4 mm screen after collection and

stored at 4 °C until use (<1 .5 years). In total, ten different soils were tested for RDX

degradation under 02 rich or depleted conditions (Appendix A). Test microcosms

(triplicate killed controls and live samples) were constructed with unlabeled RDX to
determine RDX degradation potential. Stable isotope probing was conducted only on one

soil (referred to as Soil 3).

Microcosms were constructed as previously described (88). Briefly, microcosms were
assembled with soil (2 g; wet weight), a mineral salts medium (MSM), glucose (5.6 mM)

31

and RDX (45 11M or 90 uM) and were incubated in the dark on a shaker. The MSM was

prepared as previously described (88). Final masses (per liter) in each microcosm were as

follows: KH2P04, 0.218 g; K2HPO4, 0.278 g; MgSO4.7H20, 0.16 mg; FeSO4.7H20, 1.6
mg; CaC12.2HZO, 0.024 mg; MnC12.4HZO, 0.4 mg; H3BO3, 0.04 mg; ZnClz, 0.04 mg;
CuClz, 0.024 mg; NazMoO4.2H20, 0.008 mg; CoC12.6HzO, 0.4 mg; NiC12.6HzO 0.04
mg; NazMO4.2HzO, 0.008 mg; CoC12.6H20, 0.4 mg; NiClz.6H20, 0.04 mg; and

NaZSeO3, 0.4 mg.

The SIP study involved microcosm samples amended with labeled or unlabeled RDX as
well as autoclaved controls. Although all microcosms (50 mL or 160 mL) were closed
with a rubber seal and aluminum crimp, a select number were also aerated between
sampling days. All microcosms were briefly exposed to air during sampling, performed
on day one and when RDX was expected to be removed completely (based on
preliminary studies). Microcosms were prepared in duplicates or triplicates, covered in
heavy-duty aluminum foil (to prevent RDX photo degradation) and were shaken at room

temperature (~20 °C).
Microcosms for testing the extraction efficiency of RDX were constructed as described

above (only live unlabeled samples). The SIP study involved two different RDX

concentrations (45 uM and 90 uM) dissolved in acetonitrile.

32

4.2.3 RDX extraction and HPLC analysis

RDX extraction and analysis were as previously described (88). Briefly, sampling for
RDX involved mixing and removal of 1 mL using a wide tip sterile serological pipette
into a Nalgene Oak Ridge High-Speed F EP Centrifuge Tubes with Tefzel ETFE screw
caps. RDX was extracted by adding equal volumes of acetonitrile and sonicating for 18
hours at 15 °C. The ultrasonic bath (Fischer Scienctific) was coil cooled by circulating
cooled deionized water. At the end of 18 hours, the tubes were centrifuged, at 2900 rpm
for 20 minutes. The supernatant (600 uL) was filtered using acetonitrile wetted filters
(PVDF, 0.22 pm, Whattman). All samples were analyzed on the same day as extraction

to minimize potential for RDX degradation.

HPLC analysis involved the following conditions and instrumentation: injector volume:

20 uL for samples and 10 uL for standard; isocratic 40% acetonitrile and 60% 0.1%

H3PO4 acidified deionized water; mobile phase flow rate: 1 mL/min; Perkin Elmer series

200 autosampler; PE binary LC Pump 250; PB diode array detector 235C, wavelength

255 mm; column: Supelco Reverse Phase PAH C18 (25 cm X 4.6 mm, 5 pm).

4.2.4 DNA extraction and ultracentrifugation

Following the complete removal of RDX, genomic soil DNA from the live labeled and
unlabeled microcosms were extracted using the PowerSoil DNA extraction kit (MO BIO
Laboratories, Inc., Carlsbad, CA) as per manufacturer’s instructions. Ultracentrifugation
was performed in Quick —Seal Polyallomer tubes (Beckrnan Coulter) in a Therrno Sorvall

WX ultra series centrifuge equipped with a step saver rotor system (70V6) for 46 hours at

33

178127 X g and 20 °C. All extracted DNA from a single microcosm (approximately 100
ng or more) was added to a Beckrnan Centrifirge tube along with a TE/CsCl solution.
Buoyant densities (BD) were calculated by measuring the refractive index with a model
AR200 digital hand-held refractometer (Leica Microsystems Inc.) before the tubes were

sealed (Quick-Seal tube topper, Beckrnan Coulter). The initial buoyant density of the

TE/CsCl solution was adjusted to 1.7828 g mL-l, and that of the DNA and TE/CsCl

solution to 1.7276 to 1.7285 g mL’l.

Following isopycnic gradient centrifugation, the DNA was divided into fractions (20-26
fractions) using a fractioning system (Beckman Coulter) and a syringe pump (Kd
scientific). Deionized water was pumped into the top of the ultracentrifugation tubes and
DNA-TE/CsCl mixture was collected from the bottom (heaviest DNA collected first) in
volumes of 150 uL. The BD of each fraction was determined by measuring the refractive
index with a model AR200 digital refractometer (Leica Microsystems, Inc.). The DNA
was separated from the CsCl in each of the fractions by overnight glycogen-ethanol

precipitation. The purified DNA was stored at -20 °C until further analysis.

4.2.5 TRFLP and sequencing
Heavy fractions (first 10 ~12 fractions that had detectable DNA on 1% agarose gel) were

analyzed by 16S rDNA terminal restriction fragment length polymorphism (TRFLP)

9

using standard procedures (60). Universal primers 27F-FAM (5 -

AGAGTI‘TGATCMTGGCTCAG, 5’ end-labeled with carboxyfluorescein) and l492R

34

(5,-GGTTACCTTGTTACGACTT) (Operon Biotechnologies) were utilized for PCR of

all fractions. The PCR reaction mix included the following: 10 uL of template (varying
weight based on fraction DNA concentration); 10 uL of 10x PCR buffer; 0.2mM of
dNTP mix; 50 pmols of 27F-FAM; 50 pmols of 1492R; 2.5 units of Taq; and molecular
biology grade water to a final volume of 100uL. The PCR program was: 94 °C (5 min);
94 °C (30 sees), 55 C (30 secs), 72 °C (1.5 min) (30 cycles); 72 °C (5 min). 15 uL of the
PCR products were run on a 1% agarose gel and the first 10 to 12 heavy fractions that

had a band on the gel were chosen for further analysis.

The PCR products were purified using a Qiagen PCR Purification kit following the
manufacturer’s instruction and concentrated in a 30 uL volume of elution buffer. 13 uL
of the purified product (200 to 800 ng) was digested in a 15 uL digestion volume using
15 units of Hae III restriction enzyme (restriction site: CCGG). The digested DNA
samples were analyzed in duplicates using Capillary Electrophoresis (ABi 3730 Genetic
Analyzer, Research Technology Support Facility, Michigan State University). The

percent abundance of fragments was analyzed using Genescan software.

Total DNA was PCR amplified (as described above with a 30 minutes extension step)

and cloned into Escherichia coli TOPO 10 cells using TOPO TA cloning kit (Invitrogen

Corporation). The E. coli cells were grown on LB broth (25 g L-l) solidified with 15 g

agar L.1 in the presence of 50 pg ampicillin mL‘1 for 16 hours at 37°C. The combined

DNA from first four heavy fractions of one of the triplicates was also used to construct

35

clone libraries. Individual colonies were isolated and grown in LB broth with ampicillin

(50 ug mL-l) for upto 16 hours and checked for growth. The clones with inserts were
verified by PCR using M13 forward (5’-TGTAAAACGACGGCCAGT-3’) and M13

reverse (5,-AACAGCTATGACCATG-3’) primers and the plasmids were extracted

using QIAPrep miniprep system (Qiagen, Inc.) and sequenced (using M13 forward and
M13 reverse primers) at the Research Technology Support Facility at Michigan State
University The Ribosomal Database Project’s (Center for Microbial Ecology, Michigan
State University) analysis tool called “Classifier” was used to assign taxonomic identity.
The clustalW2 web tool (European Bioinforrnatics Institute, European Molecular Biology

Laboratory, United Kingdom) was utilized to align sequences.

36

4.3 Results

4.3.1 RDX biodegradation

The extraction efficiency of RDX was 121.90 (i 6.38) % (non autoclaved samples only)
tested with soil 6, (Appendix A). The RDX concentrations, measured at various time
points for all soils, under various conditions, are summarized in a tabular form Appendix
A and are illustrated in bar charts in Appendix B. Among the study soils, RDX was
degraded over a span of 2 to 3 weeks only in microcosms setup with soils 3, 4 (in the
presence of acetonitrile) and 7, 8, 9 and 10 (in the absence of acetonitrile), whereas little
or no degradation was observed in the corresponding autoclaved controls. However all
degradations occurred only when the microcosms were unopened (no oxygen diffusion)

between day l and the last sampling day (varied between 11-49 days).

In soils 3 and 4 (with acetonitrile), 45 uM RDX was degraded in ~1 1-14 days and 90 uM
RDX was degraded in $6 days. Soil 5 showed only slight degradation (significantly
different from the controls as per ANOVA test). Soils 1, 2 and 6 showed no degradation
under conditions tested. Soils 7, 8, 9 and 10 degraded ~45 uM RDX (no acetonitrile) in

16 days.

Following these preliminary RDX degradation experiments, microcosms were

constructed for SIP, using soil 3 with both 45 llM and 90 uM ring labeled RDX

(3N153C13) dissolved in acetonitrile. After 11 and 16 days respectively, the RDX

concentration was below detection level (<500 ppb) in all labeled samples and unlabeled

37

live control microcosms, while no or little degradation was observed in the killed control

samples (Figure 4.1 and Figure 4.2).

38

 

l Control
Unlabeled
I Labeled

 

Measured RDX Concentration (ppm)

 

Day 0 Day 1 Day 1 l

 

 

 

Figure 4.1 - RDX concentration in SIP microcosms (triplicate live controls, killed
controls and labeled samples) set up with soil 3 (Intial RDX concentration — 45 uM) and
with acetonitrile

All microcosms were allowed to go to 02-depleted conditions.

 

25 _ ----------------------------------------------- I contrOI

L Unlabeled
l Labeled

 

Measured RDX Concentration (ppm)

 

 

Day 0 Day 1 Day 15

 

 

 

Figure 4.2 - RDX concentration in microcosms (triplicate live controls, killed controls
and labeled samples) set up with soil 3 (Intial RDX concentration — 45 EM) and with
acetonitrile

All microcosms were allowed to go to 02-depleted conditions.

39

4.3.2 TRFLP results of SIP

DNA extracts from the labeled and unlabeled RDX amended soil samples were subject to
ultracentrifugation, fractionation of ultracentrifuged samples, followed by TRFLP
analysis on the first 10 fractions that had detectable amplified DNA. The TRF LP data
were used to assess the relative abundance of each fragment in fractions of varying
buoyant density. The TRF LP trends from the microcosms amended with 45 uM did not
Show any significant difference between labeled and unlabeled fractions. However, in the
fractions from microcosms amended with a higher concentration of RDX (90 uM), one
TRFLP fiagrnent (260 bp) showed a trend of label uptake in two of the three triplicates.
In other words, this fragment was of higher relative abundance in the heavier fractions
from labeled samples when compared to the heavier fractions (of comparable BD) from
the unlabeled samples (triplicates) (Figure 4.3). TRFLP profiles of the first few heavy
fractions from both labeled and unlabeled samples are presented in figure 4.4 and 4.5,

showing the dominance of 260-bp fragment.

While many fragments were present in the heavier fractions of both the labeled and
unlabeled samples, only fragments of size 260 bp showed a trend of increased relative
abundance, in heavier fractions from labeled samples when compared to heavier fractions
from unlabeled treatments. The relative abundance of the other peaks were similar in both
treatments. Some terminal fragments (data not shown) had variable trends in each of the

triplicates and were not considered, as the trend was not consistent.

4O

+15N,13C -0-l4N, 12C

 

 

6“A
4.
. 1......
\n,
0 \0 0 0:0 1

1.74 1.75 1.77 1.78

 

 

 

 

 

e\°
E B
S 4 - 9’6
'5
:l
:l
2
o 2 -
:5
.5.
0
n: 0 . T , ...g
1.74 1.75 1.76 1.77 1.78 1.79 1.80
10 ‘
C
8 ..
6 ..
4 - ewae-“ebfiww-e—e
2 ..
0 n r I
1.74 1.76 1.78 1.80
Buoyant Density (g ml-‘)

Figure 4.3 - The relative abundance of the fragment of length 260 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90uM of labeled and unlabeled

41

First two heavy fractions (TRF LP technical
duplicates) from soil amended with labeled RDX
(”C 15N). No fractions from soil with unlabeled RDX
(”C 14N) had corresponding high BD

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Labeled Fractions
60 160 260 360
A L
I l L l l I l I I L l l l l I l J l
I ' . I . Duplicate 1
I -BD: 1.775 g ml 1
mi. L-fihhw J}- -..... Mwibc Jutluil II‘h. I'I Ifiixq ”Ute/III. L48 JI LIJU. 'HJMMWLJ
b I I
'5 . .
5 . . Duplicate 2
ES . III .. ,I II .BD:1.775gml"
'03 MI 1 “Mi ....WA jllnil‘ll‘ IiIluI bill .H’L ' WM..-
5 . .
8 ' .
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..E.’ . , . . ' BD: 1.772 g 1111'1
LL. AAJ- J ll ....H-wm-_._AW-L‘..AIlt.)LJRA .r [LDdJLNAA'rLWL 4- —4
I ' Duplicate 2
. I .- I BD:1.772 g ml'1
... III. Lu...” WILL-lid .h Li LMJJ'LNMI'JIMWW
Fragment length

Figure14. 4- 5TRFLP profiles of first two heavy fractions from soil amended with labeled
RDX (131C5N) showing 260-bp fragment and its abundance

42

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43

4.3.3 Sequencing

Partial l6S rRNA gene sequences obtained from total soil extracted DNA were virtually
digested (restrictionmapperorg) with HaeIII enzyme, to identify clones that corresponded
to the fragments of interest. The sequences, when classified using the Classifier
(Ribosomal Database Project, Michigan State University), belonged to Actinobacteria,
Acidobacteria, a-Proteobacteria, y-Proteobacteria, 6—Proter0bacteria,
Verrucomicrobz’ae, Gemmatimonadetes and Sphingobacteria. Of the 155 clones, 19 had
terminal fragment lengths of 258-264 bp when virtually digested with HaeIII restriction
enzyme. This slight difference in the measured length of fragments and that predicted by
sequence data has observed in other studies (30, 60). The analysis of these partial 16$
rRNA sequences indicated the organism responsible for RDX degradation might belong

to either of the following: Sphingobacteria (18 clones), Acidobacteria (1 clone).

4.4 Discussion

The extraction efficiency of RDX has been found to be high in previous studies and
correlates well with the value observed here when employing an acetonitrile and

sonication extraction procedure (6). Since the microcosms were setup and allowed to go

to 02 depleted conditions the time when RDX degradation begins in not known. Notably,

no degradation was observed in microcosms that were aerated. The initial amount of 02

varied based on the volume of the bottle used. The limited amount of data generated did
not allow the calculation of RDX half lives, however others have reported values between

94 to 154 days (49).

The trend of increased relative abundance of fragment 260 bp, in heavier fractions from
labeled samples when compared to heavier fractions from unlabeled treatments, indicates
label uptake by the organism represented by this fragment. Unfortunately, there was no

opportunity to control for label cross feeding in these experiments.

Although many strains have been isolated with RDX biodegradation potential there has
ben only one previous study that has employed SIP to identify RDX biodegraders active
in situ (80). In that study, RDX degradation was examined in microcosms constructed

with material from RDX contaminated aquifer and groundwater. The microcosms were

constructed with unlabeled or ring-lsN-labeled RDX. Following RDX uptake, the DNA

from the labeled RDX amended microcosms were extracted and ultracentrifuged in CsCl-

EtBr solution. The 15N labeled heavy DNA was extracted using a needle and syringe, as

45

opposed to fractioning as employed in the current study. The ‘heavy DNA’ was amplified
PCR amplification employing specific primers targeting the prA gene. This gene has
been associated with RDX biodegadation. They derived 5 xplA-like genes and showed
that the 168 rRNA gene sequences of the clones from the ‘heavy DNA’ containing RDX

biodegraders belonged to Actinobacteria, a-Proteobacteria and y—Proteobacteria.

In the current study, the SIP data indicate an organism belonging to the phylum
Bacteriodetes and class Sphingobacteria. These are Gram negative bacteria found in
grow in aerobic or anaerobic conditions (44). Sphingobacteria have been isolated from a
variety of environments and enrichments including, oil-contaminated sediments,
consortium of polycyclic aromatic hydrocarbon (PAH) degraders, community of
trichloroethylene (TCE) degraders and community of dentrifiers (28, 61, 64, 77, 93).
Though Sphingobacteria have not been previously linked with RDX biodegradation they
have been noted for their ability to biotransform a number of xenobiotic compounds such
as methyl tert-butyl ether (MTBE) (59), tetracycline (42) and polychlorinated biphenyls
(PCBs) (63). Many strains of Acidobacteria have been enriched in the past with ability to
degrade MTBE (59) and BTEX. Previous studies have primarily isolated RDX degrading
microorganisms from explosive contaminated soil or water. In this study RDX degraders
were studied in an agricultural soil. Since organisms from neither of these classes of
bacteria have been associated with RDX degradation before, our results suggest that
RDX biodegradation might be possible by phylogenetically diverse microbial

populations.

46

4.5 Conclusion and future studies

Bioremediation has the potential to be an effective tool for cleanup of sites contaminated
with RDX. However, the approach requires knowledge of microorganisms that are
capable of metabolizing the compound, the necessary site conditions for successful
remediation, the biode gradation pathway, and the end products produced. In addition,
insight into the enzymes responsible for RDX biodegradation provides the ability to
probe for other organisms that possess similar functionality. This study aimed at

identifying microorganisms capable of utilizing RDX by employing SIP. To our

knowledge this is the first study to use 15N DNA-SIP with fractioning and TRF LP to

study RDX biodegradation. The partial 16S rRNA sequences of the RDX degrading
bacteria were classified as Sphingobacteria or Acidobacteria. As suggested by Binks et.
al. (1995) (17) using non-indigenous microorganism and linking their survival with
contaminant provides for a reliable bioremediation strategy and requires identification of
non-indigenous strains capable of degrading the pollutant. All RDX degrading strains
isolated so far have been isolated from explosive contaminated soil or water and this is
the first study that identified RDX degrading microorganisms from an agricultural soil

amended with biosolids.

However it is not known whether the organism was indigenous to the agricultural soil or
to the biosolids. Further studies to identify the biodegradation products of RDX
biodegradation would clearly be useful. As mentioned earlier (Chapter 2) confirming that
RDX mineralization occurs is crucial to ensure no toxic byproducts accumulate in the

system (66). Employing mass spectrometry to identify intermediates and biodegradation

47

end products will also facilitate the identification of the biodegradation pathway. Specific
functional genes to RDX degradation (xplA) could be used for PCR amplification to
compliment the current study. Real-time PCR using specific flmctional genes helps in
studying the relative abundance of these genes in fiactions (labeled and unlabeled) of
varying buoyant density profile. Further conducting SIP over time will help in identifying

and avoiding any cross feeding issues if any.

48

 

 

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49

 

 

- - I Control

10 - : p : ------- '7 -------------- El Sample

  

 

Measured RDX Concentration (ppm)

Day 0 Day 8 Day 43

 

 

 

Figure B.1 - RDX concentration in microcosms (triplicate live and killed controls) set up

with soil 1 and was allowed to go to 02-depleted conditions
- No significant degradation on both day 8 and day 43 as verified by ANOVA test.

 

I Control

N
LII
I

______________________ ____-_____-_______________ISample

N
O
I

 

Measured RDX Concentration (ppm)

 

 

0

Day 0 Day 8 Day 43

 

 

 

Figure B.2 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 2
- No significant degradation on both day 8 and day 43 as given by AN OVA test.

50

(A)

 

1 6 _ -------------------------------------- - CODtI'OI

Measured RDX Concentration (ppm)

 

 

 

 

 

@

 

I Control

'5? Sample

 

 

Measured RDX Concentration (ppm)

 

Day 0 Day 1 Day 13

 

 

 

Figure B.3 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 3 and was allowed to go to 02-depleted conditions (160 mL bottles)

- Figure (A) and (B) are results of similar but repeated experiments setup. 45 pM
RDX was degraded in 13 to 14 days.

51

(A)

 

 

I Control

5 Sample

Measured RDX Concentration (ppm)

 

 

Day 0 Day 1 Day 5 Day 8 Day 10 Day 14

 

 

 

(B)

 

 

l Control

Ti Sample

 

Measured RDX Concentration (ppm)

 

 

Day 0 Day 1 Day 15

 

 

 

Figure B.4 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 3 and was aerated daily (160 mL bottles)
- Figure (A) and (B) are results of similar but repeated experiments, (A)
continuously sampled (B) sampled on day 1 and day 15 only. No significant
degradation was observed as verified by AN OVA test.

52

 

 

—-
N

I Control

I Sample

  

....
O
l

00
1

Measured RDX Concentration (ppm)
Ch

 

DayO Day 1 Day 14

 

 

 

Figure B.5 - RDX concentration in microcosms (triplicate live and killed controls) set up

with soil 4 and was allowed to go to 02 depleted conditions (160 mL bottles).
- 45pM RDX was degraded in 14 days.

 

 

I Control

5?- Sample

 

 

Measured RDX Concentration (ppm)

 

 

Day 0 Day 1 Day 4 Day 7 Day 14 Day 21

 

 

 

Figure B.6 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 4 and was aerated daily (160 mL bottles)
- No significant degradation until day 14 as verified by ANOVA test.

53

 

 

I Control

5! Sample

 

Measured RDX Concentration (ppm)

 

Day 0 Day 1 Day 15

 

 

 

Figure B.7 - RDX concentration in microcosms (triplicate live and killed controls) set up
with soil 5 and was aerated daily (160 mL bottles)
- No significant degradation until day 15 as verified by ANOVA test.

 

12 — -------------------------------------------------------
I Control

_______________________________________ *1 Sample

.—
o
l

  

00
I

O‘\
l

A
1

Measured RDX Concentration (ppm)
N

 

O
I

Day 0 Day 26 Day 57

 

 

 

Figure B.8 - RDX concentration in microcosms (triplicate live and killed controls) set up

with soil 6 and was allowed to go to 02-depleted conditions
- No significant degradation until day 57 as verified by AN OVA test.

54

 

—s
A

T ---------------------------------------------- I Soil 7
................... -I Soil 8

   

j—l
N
I
T
I
l
I
I
I
I
I
I
I
I
I
I
I
I
I
I
I
I
I

._.

O
l
I

00
l

O\
1

A
r

N
1

Measured RDX Concentration (ppm)

 

O
1

Day 0 Day 1 Day 16

 

 

 

Figure B.9 - RDX concentration in microcosms (duplicate live) set up with soil 7, 8, 9, 10

and no acetonitrile (Oz-depleted conditions)
- 45 uM RDX was degraded in 16 days.

 

I Control

Sample

Measured RDX Concentration (ppm)

 

 

Day 0 Day 1 Day 23

 

 

 

Figure B. 10 - RDX concentration in microcosms (triplicate live and killed controls) set
up with soil 8, no acetonitrile and was aerated daily
- No significant degradation until day 23 as verified by AN OVA test.

55

APPENDIX C

This section presents the relative abundance plots of T-RF’s from SIP study setup with 90
pM RDX.

2.5 -
A -0-15N13C‘=<>r 14N12C
2.0 - <>

1.5 "
1.0 -
0.5 r
0.0

 

 

 

1.72

4.2 ‘
3.6 7
3.0 ‘
2.4 -
1.8 '
1.2 ‘
(ml
0.0

 

Relative Abundance (%)

 

 

 

1.74

 

 

 

0.0 I v u v I
1.74 1.76 1.78 1.80
Buoyant Density (g ml'l)
Figure 0.1 - The relative abundance of the fragment of length 63 bp over a range of

buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

56

-o—15N13c so =-14N12C
0.6 -

0.4 -
0.2 -

 

 

p
o

 

1.72 1.78

—i

 

Relative Abundance (%)
'o 'N J:- O\ 00 'o

0°99?

-———¢—-¢o—o—--<->e+ -9 ecu-<3: ere . ~<>~ -<>=--o——.
1.74 1.76 1.78 1.80

9.0.69
#UIQQ
l I

99.6
Ht)!»

 

 

.0
o

 

l V V l V I

1.74 1.76 1.78 1.80
Buoyant Density (g ml")
Figure C.2 - The relative abundance of the fragment of length 70.5 bp over a range of

buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 11M of labeled and unlabeled RDX

57

06 +15N13C~0~14N12C

0.4

0.2

 

 

N

 

 

Relative Abundance (%)

7‘0 '0 O 0%
1.74 1.76 1.78 1.80

O

0.4' C

 

 

1.74 1.76 1.78 1.80
Buoyant Density (g ml'l)
Figure C.3 - The relative abundance of the fragment of length 72 bp over a range of

buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

58

 

 

 

 

A +15N13Cr-0:=14N12C
1.5 —
1.0 —
0.5 —
0.0
1.72
§ 1.2 _ B
8
5 0.8 —
E
a
.4:
E 0.4 -
e
.3
d)
M 0.0 J——-O—-H-«>-¢- ~m¢¢+~+>r one—fl

 

 

 

1.74 1.76 1.78 1.80
08 — C
0.6 —
0.4 —
0.2 -
0.0 . v .
1.74 1.76 1.78 1.80

Buoyant Density (g ml'l)
Figure C.4 - The relative abundance of the fragment of length 126 bp over a range of

buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

59

A +15N13C -0-14N12C

 

 

 

 

 

  

 

 

2.4 -
1.8 r
1.2 -
0.6 “
0.0
1.72
A 1.6 "
é B
3 12 -
E "N.
i“ J 3°"
31 08 ‘ e
7.5
E 0.4 r
.2
$2 0.0 - —<>—<>———4
1.74 1.76 1.78 1.80
1.8 ' C
930‘
0 ‘ow—G f\0
1.2 r \9
“'6‘ W
0.0 . . Q .
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

Figure C.5 - The relative abundance of the fragment of length 172 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

60

3.6 -

3.0 - A f

2.4 - *
1.8 - 8’0
1.2 -

+15N13C -0-14N12C

  

 

 

0.0 I
1.72 . 1.74 1.76 1.78

:‘3
N
I

3.6 -
3.0 r
2.4 -

_ o
1.8 \e

0.6 - [/9

0.0 "G I
1.74 1.76 1.78 1.80

 

 

 

Relative Abundance (%)

1.6 ' C
1.4 ‘

1.2 - M
1.0 w
0.8 4
0.6 -
0.4 -
0.2 ~
0.0 . I v .

1.74 1.76 ' 1.78 1.80
Buoyant Density (g ml")

 

 

 

Figure C.6 - The relative abundance of the fragment of length 196 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

61

+15N13C —0= 14N12C

 

0
8
6
4- \o
2
0

r V: I Q l

 

 

 

 

 

1.72 1.74 1.76 1.78
I; 12 -
c}, B
o 10 4
C)
s w
3 4 i
3 2 *
é o . a
1 74 1.76 1.78 1.80
18 - C
_ ”a0

 

 

T

1.74 1.76 1.78 1.80
Buoyant Density (g ml“)

OWONC
11

Figure C.7 - The relative abundance of the fragment of length 198 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 nM of labeled and unlabeled RDX

62

+15N13C -0-14N12C
72.0 -

54.0 r /9

36.0 ‘ ’0

- /

,0
’9 W
0.0 9. . .

1.72 1.74 1.76 1.78

fl’f‘e

 

 

36.0
30.0 ‘
24.0 —
18.0
12.0

6.0

I

J

2

 

 

Relative Abundance (%)

1.74 1.76 1.78 1.80

50.0
40.0 ‘
30.0
20.0 -
10.0 ‘

0.0 v u 1
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

0

I

 

 

 

Figure C.8 - The relative abundance of the fragment of length 204 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 pM of labeled and unlabeled RDX

63

+15N13C +14N12C

3"
M
1

i

i—5 I—I N
O U- C
i

I

 

9 9
O Ur
o
l

I V

1.72 1.74 1.76 1.78

p—a

 

Relative Abundance (%)
o N A O\ on 'o

99999

A—o—H—eew-eww-e-nemo—e—fi
1.74 1.76 1.78 1.80

0.8 ' C

0.6 ‘

0.4 ‘
0‘0

0.2 - .

 

 

0.0 1 m x} l
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

Figure C.9 - The relative abundance of the fragment of length 208 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

64

10.00 ~

+15N13C m0*“14N12C

5.00 -

 

 

 

 

 

1.74 1.80

A

o

o
O

MOMOU’IO
COOOOO
llllll

 

 

 

I V I

1.74 1.76 1.78 1.80
Buoyant Density (g ml")
Figure C.10 - The relative abundance of the fragment of length 215 bp over a range of

buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 [4M of labeled and unlabeled RDX

65

6.00 -

 

 

 

 

 

 

 

A -0-ISN13C'=*<>~‘14N12C
4.00 _ W
2.00 —
0.00 . <> :0 oo—%
1.72 1.74 1.76 1.78
A 4 ‘
°\° B
E
=
a
'U
E 2 -
..Q
<
0
E
%
a: 0 x
1.74 1.76 1.78 1.80
2.50 . C
2.00 - \
0’0 5.42
1.50 - “0‘, 9
1.00 -
l.
0.50 ~
0.00 | v I NI.) I
1.74 1.76 1.78 1.80

Buoyant Density (g ml")

Figure 011 - The relative abundance of the fragment of length 226 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

66

+15N13C-o—14N12C
2.0 -

 

 

 

A

1.5 -
«0’0
1.0 -
0.5 -
0.0 r
1.72 1.74 1.76 1.78

1.6 -

B

I—I
N
I

«3

 

 

Relative Abundance (%)
c
00

 

 

 

0.0 .
1.74 1.76 1.78 1.80

2.0 ' C

1.6 . /W

1.2 '

0.8 -

0.4 ‘

0.0 . T 8 1
1.74 1.76 1.78 1.80

Buoyant Density (g ml")

Figure C.12 - The relative abundance of the fragment of length 228 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

67

3.0 '
2.4 -
1.8 -
1.2 r
0.6 '

 

+15N13C -0-14N12C

 

 

 

0.0
1.72

 

1.74 1.76 1.78

 

 

Relative Abundance (%)
E3

 

 

1.76 1.78 1.80

 

1.74

1.76 1.78 1.80
Buoyant Density (g ml")

Figure C. l 3 - The relative abundance of the fragment of length 236 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 pM of labeled and unlabeled RDX

68

A +15N13C-0-14N12C
25.00

20.00
15.00
10.00

5.00

0.00 v . I
1.72 1.74 1.76 1.78

“x.
i.

I

 

 

30.0 -
24.0 i
18.0 '
12.0 ‘

 

6.0 ‘-

0.0 1 .
1.74 1.76 1.78 1.80

/w4\§/

 

 

Relative Abundance (%)

25.0
20.0
15.0 -
10.0 -

5.0

0.0 u u u
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

I

J

 

 

Figure C.14 - The relative abundance of the fragment of length 251 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 pM of labeled and unlabeled RDX

69

+15N13CM<>14N12C
12.0 -

10.0 ‘

 

 

0.0 34> '0'6-6' '6' ~00 .934 0
1.72 1.74 1.76 1.78

 

 

Relative Abundance (%)

 

0.30
0.25
0.20
0.15
0.10
0.05
0.00

 

 

06¢ wMamie-<3 -~o————4
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

Figure C.15 - The relative abundance of the fragment of length 258 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

70

1.50 , -o-15N13c-o-14N12c

1.00 r

0.50 - 8¢\

 

 

 

 

 

0.00 . \e—e
1.72 1.74 1.76 1.78
A 1.0 ‘
3: B
3 0.8 -
5
1a 0.6 ~
I:
3 O4 4
< .
0
.5 0.2 a
% ,
a: 0.0 +——-¢—H-ee+——eveo-ew—9—o—e-e»——.
1.74 1.76 1.78 1.80
1.0 i C
0.8 ‘
0.6 "
0.4 r
0.2 ‘
0.0 ‘WWWH u
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

Figure C.16 - The relative abundance of the fragment of length 263 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

71

 

A .0-15N13C“0 ‘l4N12C

 

 

 

  

 

 

g B
8 12 - - '8
a <> ‘5,
E 08 - \
< "i
g 0.4 -
% W
m 0.0 — - atone—fl
174 1.76 178 180
0.6 —
C
0.5
0.4 -
0.3
0.2
0.1 - .0
0.0 0! I v VA \bO“
1.74 1.76 1.78 1.80

Buoyant Density (g ml")

Figure C.17 - The relative abundance of the fragment of length 271 bp over a range of
buoyant density from DNA extracted fi'om triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

72

+15N13C 4-14N12C
1.50 -

1.00 r

0.50 -

 

 

0.00 . -<>
1.72 1.74 1.76 1.78

H
O‘
I

1.2 ‘

0.8

0.4

 

 

Relative Abundance (%)

 

0.0
1.74 1.76 1.78 1.80

1.6 - C
1.4 -

1.2 '

0.8 "

0.6 - 0* \x"
0.4 -
0.2 —

0.0 r
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

 

 

Figure C.18 - The relative abundance of the fiagment of length 272 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
fiom samples amended with 90 11M of labeled and unlabeled RDX

73

40 +15N13C -0-14N12C

3.0 — 4/6 /°
2.0 - I

 

 

1.72 1.74 1.76 1.78

I—I

.U'

C
I

I

12.0

 

Relative Abundance (%)

 

 

1.74 1.76 1.78 1.80

10.0 -
8.0 -
6.0 -
4.0 -
2.0 - O‘O‘OANM /

\8
0.0 . .

1.74 1.76 1.78 1.80
Buoyant Density (g ml")

 

 

Figure C.19 - The relative abundance of the fragment of length 274 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 11M of labeled and unlabeled RDX

74

4.0 -

3.0 -

2.0 -

1.0 -

+15Nl3C-0-14N12C

”’\/

 

0.0

1.72

4.0 i

3.0 '

2.0 ‘

1.0 .

 

 

Relative Abundance (%)

0.0

1.74

4.0 -

3.0 r

2.0 ‘

1.0 r

1.74 1.76 1.78

ewMe-e?

 

1.76 1.78 1.80

 

0.0

1.74

 

, “V

1.76 1.78 1.80
Buoyant Density (g ml")

Figure C.20 - The relative abundance of the fragment of length 281 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

75

-’o—15N13c +14N12c

18:1“

0.00 r
1.72 1.74 1.76 1.78

1.50 -

I

1.00

l

0.50

 

 

 

$3993.92"
NAQOOO

 

Relative Abundance (%)

 

o—oo—o—eev—«mo-e-e—o—e—o—w-e—fi
1.74 1.76 1.78 1.80

P
o

1.07 C
0.8 i

0.6 -
0.4 '
0.2 ‘

0.0 ‘WW—fi .
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

 

 

Figure C.21 - The relative abundance of the fragment of length 292 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 pM of labeled and unlabeled RDX

76

 

+15N13C +14N12C

3.0 -
2.4 -

0.6 - \
0.0 T ' <> .

 

 

 

 

 

 

 

    

 

 

1.72 1.74 1.76 1.78
r-
»; 3.0 " E
e B .
3 2.4 -
g r
a: 1.8 r
I:
.2 _
< 1.2
0
3 0.6 -
8
a: 0.0 .
1.74 l.76 1.78 1.80
3.0 -
C
2.4 -
1.8 —
1.2 -
0.6 -
0.0 I 1 {z .
1.74 1.76 1.78 1.80

Buoyant Density (g ml")

Figure C.22 - The relative abundance of the fiagment of length 302 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

77

 

 

 

 

  

 

 

 

 

+15N13C-6'14N12C
4.0 -
A
32 -
2.. I
16 - ,9
0.8 -
0.0 . «e——.
1.72 1.74 1.76 1.78
"24 -
e\° ' B
3.7
g 1.8 ‘
"U
.5 12 f8
‘5.
, 0.6
g \«9
g 0.0 —-<>—e——1
1.74 1.76 1.78 1.80
3.0 -
C
2.4 -
1.8 -
’-
1.2 -
0.6 ~
0.0 . . é .
1.74 1.76 1.78 1.80

Buoyant Density (g ml")

Figure C.23 - The relative abundance of the fragment of length 306 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 11M of labeled and unlabeled RDX

78

*15N13C "0-14N12C
1.50

1.00 f
1 *3»

 

 

 

0.50
0.00 I I V ~lo '39 v I
1.72 1.74 1.76 1.78
A 1.0 -
8 B
3 0.8 -
E
e 0.6 -
fl
3 4 _
< 0.
0
.3 0.2 —
a
g 0.0 ————+—~—¢><x>a . 3W¢¢39~W<>m<> :o—.
1.74 1.76 1.78 1.80
1.0 7 C
0.8 -
0.6 -
0.4 —
0.2 —
0.0 -———H - e «e. . 6 acre :. cache-core e—é
1.74 1.76 1.78 1.80

Buoyant Density (g ml")
Figure C.24 - The relative abundance of the fragment of length 316 bp over a range of

buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 11M of labeled and unlabeled RDX

79

+15N13C “-0-14N12C
1.2 ‘

0.9 -

0.6 -

0.3 '

 

0.0 6‘0‘0-“6-0
1.72 1.74 1.76 1.78

 

1.0 '
0.8 -

 

0.6 r
0.4 -
0.2 -

0.0 -r—¢—-o-ee+-¢W~o-e——.
1.74 1.76 1.78 1.80

 

Relative Abundance (%)

0.6 —

0.3 -

 

 

 

0.0 u v .

1.74 1.76 1.78 1.80
Buoyant Density (g ml")

Figure C.25 - The relative abundance of the fiagment of length 328 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

80

 

1.0
0.8
0.6
0.4
0.2
0.0

.0-15N13C '0‘14N12C

 

 

1.72 1.74 1.76 1.78

Relative Abundance (%)

0.0

4.0 ‘
3.2 .
2.4 -
1.6 '
0.8 -

 

.‘u.

 

1 “—0-9—-—I

 

1.74 1.76 1.78 1.80

2.4
2.0
1.6
1.2
0.8
0.4
0.0

 

 

 

1.74 1.76 1.78 1.80

Buoyant Density (g ml")

Figure C.26 - The relative abundance of the fragment of length 402 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 pM of labeled and unlabeled RDX

81

+15N13C -0-14N12C

 

0.0 6%wafiq-«3-MH-O—fi
1.72 1.74 1.76 1.78

 

5"
A
I

 

 

Relative Abundance (%)

 

 

1.74 1.76 1.78 1.80

1.2 r

0.8 '
0.6 '
0.4 ‘
0.2 J
0.0 ‘ '6 O u
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

 

 

Figure C.27 - The relative abundance of the fragment of length 404 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 uM of labeled and unlabeled RDX

82

1.2 ~ -o-15N13C -0-14N12C
0.9 - N
0.6 ~

0.0 G%-0—-0-<>'
1.72 1.74 1.76 1.78

 

 

 

1.0 r
0.8 -
0.6 r
0.4 i
0.2 -

0.0 -——o—oo—o—eo~e—-e~ee-o¢-¢—e—-—e—e-<~>———.
1.74 1.76 1.78 1.80

 

Relative Abundance (%)

4.0 -
3.2
2.4
1.6

0‘04 ”\e
V

0.0 u
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

1

l

 

 

Figure C.28 - The relative abundance of the fi‘agment of length 448 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 pM of labeled and unlabeled RDX

83

 

 

-o—15N13C-~<>u14N12C
0.8

0.6 f

0.4 330
3
0.2

0.0 | I v 3:3 706 V l

1.72 1.74 1.76 1.78
1.0 a
0.8 r
0.6 -

 

0.4 '
0.2 -

 

Relative Abundance (%)

0.0 r—W'O“ “OWO'LW‘V '0“=6———1
1.74 1.76 1.78 1.80

1.07 C
0.8 4
0.6 ~
0.4 -
0.2 -

0.0 -—o—¢w¢ «6.3 «ace-Joe eee»=ee—>=~<>———-——-fi
1.74 1.76 1.78 1.80
Buoyant Density (g ml")

 

Figure C.29 - The relative abundance of the fragment of length 462 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 11M of labeled and unlabeled RDX

84

1.5 -

12 _ ? +15Nl3C‘0-14N12C
'1
i
0

0.9 ‘

 

 

 

Relative Abundance (%)

 

 

 

1.74 1.76 1.78 1.80
Buoyant Density (g ml")

Figure C.30 - The relative abundance of the fragment of length 885 bp over a range of
buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 11M of labeled and unlabeled RDX

85

+15N13C m<3"“*14N12C

 

 

 

 

Relative Abundance (%)
N
'o

 

0,3090

 

0.0 l V T V I
1.74 1.76 1.78 1.80
Buoyant Density (g ml")
Figure C.3l - The relative abundance of the fragment of length 920 bp over a range of

buoyant density from DNA extracted from triplicate microcosms (A, B, C) on day 16
from samples amended with 90 11M of labeled and unlabeled RDX

10.

11.

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Adrian, N. R., and C. M. Arnett. 2004. Anaerobic biodegradation of hexahydro-
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Adrian, N. R., C. M. Arnett, and R. F. Hickey. 2003. Stimulating the anaerobic
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Andreoni, V., and L. Gianfreda. 2007. Bioremediation and monitoring of
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Bailey, A. M., S. G. 2000. Explosives, Propellants & Pyrotechnics. Redwood
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Bayman, P., S. D. Ritchey, and J. W. Bennett. 1995. Fungal interactions with
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Microbiology 15:418-423.

Bell, B. A., and G. J. Hardcastle. 1984. Treatment of a high-strength industrial-
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Beller, H. R. 2002. Anaerobic biotransformation of RDX (hexahydro-l ,3,5-
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