genomic dna extraction and detection of bacteria ... · immobilized in polyvinyl alcohol host...

This thesis was elaborated and defended at the Institute of Chemical Technology Prague within the framework of the

European Erasmus Mundus Programme “Erasmus Mundus International Master of Science in Environmental Technology

and Engineering " (Course N° 2011-0172)

Erasmus Mundus Master Course: IMETE

Thesis submitted in partial fulfilment of the requirements for the joint academic degree of:

International Master of Science in Environmental Technology and Engineering

an Erasmus Mundus Master Course from Ghent University (Belgium), ICTP (Czech Republic), UNESCO-IHE (the Netherlands)

Genomic DNA extraction and detection of bacteria immobilized in polyvinyl alcohol

Host University:

Department of Water Technology and Environmental Engineering

Probyn, Rhys Edward Promoter: Co-promoter:

Prof. Ing. Jiří Wanner. DrSc. Ing. Jan Bartáček, Ph.D.

Tutor:

Hana Stryjová, MSc.

2011 - 2013

http://www.unesco-ihe.org/

http://www.vscht.cz/homepage/english

http://www.vscht.cz/homepage/english

iii

iii

iv

iv

Acknowledgements

VI

Very special thanks to Hana Stryjová, MSc. for going above and beyond the call of duty

in her role as my mentor and for becoming a cherished friend.

Very special thank you to Serena Fraraccio, MSc. for helping us with DGGE and

demonstrating unbelievable kindness and patience while working with us to overcome

technical issues and obtain positive results.

Thank you to Ing. Ondrej Vopicka, Ph.D. for helping us develop and execute the liquid

nitrogen treatment for Lentikat’s Biocatalysts

Thank you to Petr Kelbich, MSc. for operating the reactors and performing the chemical

analyses in Ch. 4 of this thesis and for being a great friend.

Thank you to Prof. Ing. Jiří Wanner, DrSc. for hosting me in his laboratory for this

project.

Very special thanks to ir. Maja Šimpraga, PhD. in the IMETE Coordination unit for

tirelessly working to make this program run smoothly.

Thank you the IMETE Management Board for conceiving of and implementing this

program.

Thank you to Ing. Jana Bartáčková, Ph.D. of ICTP and Ineke Melis of UNESCO-IHE for

handling all arrangements in Prague and Delft.

Thank you to Isabel Del Agua Lopez for helping me prepare for all of the toughest exams

in this masters and for being a great friend.

Very special thanks to the following people without whom I would not have gotten this

far in life: Mom and Dad, Richard F. Commenzo esq., Jonathan Todd, Meredyth Ramsay,

Thomas Brew, William Mebane, Inna V. Grishkan MD. Ph.D., Vansa Chatikavanij MSc.,

David C. Gadsby Ph.D, Stephen M. Highstein MD. Ph.D., Scott Lindell MSc., Ed Enos,

Captain Bill Klimm, William Grossman, Gene Tassinari, Alexi Shalapyonok Ph.D., and

Greg Salamida.

This project was supported by a grant of Ministry of Industry and Trade of Czech

Republic FR-TI4/254 and by Research Plan grant MSM 6046137308.

Abstract

VII

In the first part of this investigation, the ability to extract pure high quality DNA from

Lentikat’s Biocatalysts and activated sludge for downstream PCR based applications was

examined with four different commercial DNA isolation kits. DNA extractions were carried out

in triplicate using the Powersoil® DNA Isolation kit, the QIAmp® DNA Stool kit, the Chemagic

DNA Bacteria Kit, and the MasterPure™ DNA Purification Kit. All kits were found to be

compatible with all Lentikat’s Biocatalyst and activated sludge samples and isolated DNA readily

amplified via Touchdown Polymerase Chain Reaction (PCR). Subsequent denaturing gradient gel

electrophoresis (DGGE) showed insignificant extraction bias between isolation kits applied to the

same samples. The Powersoil® DNA Isolation Kit performed the best in terms of processing time

and DNA extract purity. The MasterPure™ DNA purification kit performed the best in terms of

yield, cost, waste generation, and was second best in DNA extract purity. Results also indicated

that flash freezing Lentikat’s Biocatalysts with liquid nitrogen and grinding them prior to DNA

extraction increased the DNA yield and phylogenetic richness of the isolate, thus further

investigation into enhanced lysis methods is recommended.

In the second part of this investigation the effects of a known NOB inhibitor,

hydroxylamine, on Lentikat’s Nitrifying Biocatalysts (PVA biocarriers) was examined in

laboratory scale partial nitrification reactors. Two separate dosing regimes were tested on

individual reactors. The activity of nitrifying bacteria was determined throughout the experiment

with water chemistry analyses while the presence of AOB and NOB within biocarriers was

determined with fluorescent in situ hybridization (FISH). Water chemistry analyses revealed that

daily dosing to 484 µM hydroxylamine for 10 days followed by 49 days at 121 µM achieved

nearly complete nitritation during peak dosing but was not sufficient to achieve long term

sustained NOB inhibition. On the other hand hydroxylamine dosing to 1,211 µM for 14 days

followed by 4 days at 302 µM achieved nearly complete nitritation that was sustained for nearly

78 days and greater than nitratation for 182 days. FISH analyses revealed significant populations

of NOB remained in biocarriers after 14, 54, and 116 days of the onset of severe nitratation

inhibition. This implies a greater resistance of hydroxylamine inhibited NOB immobilized in

PVA biocarriers to washout in comparison with those in RBC biofilms or aerobic granules. NOB

detected by FISH are suspected to include: a small fraction performing limited nitratation, a

significant proportion of dead biomass, an unknown fraction that switched to alternative organic

substrates for survival, and in one case an unknown fraction that may have been seeded from

anaerobic digester effluent.

Table of Contents

VIII

Assignment of Diploma Thesis………………………………………………………………………iii

Certification…………………….…………………………………………….……………………….V

Acknowledgements...……………….……………………………………….……………………….VI

Abstract……….…………………………….………………………………..……………………...VII

List of Figures……………………………………….…………………….…………………………..X

List of Tables……………………………………………………………..…………………………..XI

Abbreviations…………………………………………………………..……………………………XII

Chapter 1 Introduction 1.1 Introduction ....................................................................................................................................... 1

Chapter 2 Literature Review

2.1 The Importance of Nitrogen Removal in Wastewater Treatment ..................................................... 1

2.2 Biological Nitrogen Removal (BNR) Pathways ............................................................................... 2

2.2.1 Nitrogen Speciation ................................................................................................................... 2

2.2.2 Nitrification ................................................................................................................................ 3

2.2.2.1 Ammonia Oxidation (Nitritation) ....................................................................................... 3

2.2.2.2 Nitrite Oxidation (Nitratation) ............................................................................................ 4

2.2.3 Denitrification ............................................................................................................................ 5

2.2.4 Anaerobic Ammonia Oxidation (Anammox) ............................................................................ 6

2.3 Applications of Biological Nitrogen Removal (BNR) Pathways in Wastewater Treatment and the

Case for Partial Nitrification ................................................................................................................... 7

2.4 Lentikat Biocatalysts ......................................................................................................................... 9

2.5 Molecular Methods for Assessing Microbial Communities in WWTP .......................................... 10

2.5.1 DNA Isolation .......................................................................................................................... 11

2.5.2 Polymerase Chain Reaction (PCR) .......................................................................................... 12

2.5.3 Denaturing Gradient Gel Electrophoresis (DGGE) ................................................................. 13

2.5.4 Fluorescence in Situ Hybridization (FISH) .............................................................................. 14

Chapter 3 Genomic DNA Isolation and Amplification from Bacteria Immobilized in Poly Vinyl

Alcohol Biocarriers

3.1 Objectives ....................................................................................................................................... 18

3.2 Materials and Methods .................................................................................................................... 19

3.2.1 Poly Vinyl Alcohol Pellets and Activated Sludge Samples ..................................................... 19

3.2.2 Activated Sludge Characterization ........................................................................................... 20

3.2.3 DNA Isolation .......................................................................................................................... 21

3.2.3.1 Comparison of Commercial DNA Isolation Kits .............................................................. 21

3.2.3.2 Liquid Nitrogen (LN) Enhanced Cell Lysis ...................................................................... 22

3.2.4 DNA Yield and Purity ............................................................................................................. 23

3.2.5 PCR Amplification ................................................................................................................... 24

3.2.6 DGGE ...................................................................................................................................... 26

3.3 Results ............................................................................................................................................. 28

Table of Contents

IX

3.3.1 DNA Isolation and Purity ........................................................................................................ 28

3.3.2 PCR Amplification ................................................................................................................... 32

3.3.3 DGGE ...................................................................................................................................... 35

3.4 Discussion ....................................................................................................................................... 37

3.4.1 Waste Generation, Processing Time, and Cost .................................................................... 37

3.4.2 DNA Yield ........................................................................................................................... 38

3.4.3 Purity .................................................................................................................................... 39

3.4.4 Phylogenetic Comparison of Extracts .................................................................................. 41

3.5 Conclusions ..................................................................................................................................... 43

Chapter 4 In Situ Detection of Immobilized Bacteria in Laboratory Scale Partial Nitrification

SBRs

4.1 Objectives ....................................................................................................................................... 44

4.2 Materials and Methods .................................................................................................................... 45

4.2.1 Partial Nitrification Reactor Setup and Operation ................................................................... 45

4.2.2 Chemical Analyses ................................................................................................................... 48

4.2.2.1 Ammonia Nitrogen (NH4+) ............................................................................................... 48

4.2.2.2 Nitrite Nitrogen (NO2-) ..................................................................................................... 48

4.2.2.3 Nitrate Nitrogen (NO3-) ..................................................................................................... 49

4.2.2.4 Hydroxylamine (NH2OH) ................................................................................................. 49

4.2.3 FISH ......................................................................................................................................... 49

4.2.3.1 Reagents and Probes ......................................................................................................... 50

4.2.3.2 Fixation with Paraformaldehyde (based on Amann 1995 for Gram-negative bacteria) ... 51

4.2.3.3 Hybridization .................................................................................................................... 51

4.2.3.4 Imaging ............................................................................................................................. 52

4.2.4 Live/Dead Staining .................................................................................................................. 52

4.2.5 DNA Isolation, PCR, DGGE, and Sequencing ........................................................................ 52

4.3 Results ............................................................................................................................................. 53

4.3.1 FISH Images, Inorganic Nitrogen Speciation, and Live/Dead Staining .............................. 53

4.3.2 DGGE and Sequencing ........................................................................................................ 62

4.4 Discussion ....................................................................................................................................... 63

4.4.1 Inhibition of Nitratation ....................................................................................................... 63

4.4.2 In situ detection and characterization of NOB community .................................................. 65

4.5 Conclusions ..................................................................................................................................... 68

Chapter 5 Summary of Conclusions...................................................................................................69

References...............................................................................................................................70

Appendix 1…………………………………………………………………………………...79

Appendix 2…………………………………………………………………………………...80

List of Figures

X


Figure 1. Lentikat Biocatalyst structure from Bouskova et al. (2011) .................................................... 9


Alcohol Biocarriers

Figure 2. 2% Agarose gel profiles of PCR products ............................................................................. 33

Figure 3. DGGE Profile of 16S rDNA amplicons from all samples. .................................................... 35

Figure 4. UPGMA Dendrogram and BSI for Sample N1. .................................................................... 35

Figure 5. UPGMA Dendrogram and BSI for Sample N2. .................................................................... 36

Figure 6. UPGMA Dendrogram and BSI for Sample D1. .................................................................... 36

Figure 7. UPGMA Dendrogram and BSI for Sample D2. .................................................................... 36

Figure 8. UPGMA Dendrogram and BSI for Sample L1...................................................................... 37

Figure 9. UPGMA Dendrogram and BSI for Sample P1 ...................................................................... 37


SBRs

Figure 10. Image of nitrifying SBRs in laboratory in ICT Prague. ....................................................... 45

Figure 11. Hydroxylamine dosing regimes for treatment reactors........................................................ 47

Figure 12. FISH images of newly manufactured nitrification biocatalyst ............................................ 53

Figure 13. Nitrogen speciation in control reactor (Reactor C) effluent throughout operational life. ... 54

Figure 14. FISH images from control reactor (Reactor C) biocatalysts sampled on Day 30. ............... 55

Figure 15. Inorganic nitrogen speciation and hydroxylamine dosing in Reactor A. ............................ 57

Figure 16. FISH images of biocatalysts sampled from treatment Reactor A. ....................................... 58

Figure 17. Inorganic nitrogen speciation and hydroxylamine dosing in Reactor B .............................. 59

Figure 18. FISH images of biocatalysts sampled from treatment Reactor B. ....................................... 60

Figure 19. Live/Dead Images of biocarriers sampled from Reactor B. ................................................ 62

Figure 20. DGGE Profile of amplified 16S rDNA extracted from Reactor B biocarriers on Day 274. 62

Appendix 2. Omission of Q-D2-LN Justification

Figure 21. DGGE Profiles for Sample D2 and Sample D1…………………………………………...80

Figure 22. UPGMA Dendrogram of Sample D2 Including Q-D2-LN………………………………..80

List of Tables

XI


Table 1. Typical nitrogen values for various wastewaters adapted from Zanetti et al. (2012) ............... 1

Table 2. Stoichiometric comparison of BNR pathways taken from Zanetti et al. (2012) ....................... 8


Alcohol Biocarriers

Table 3 Touchdown PCR Master Mix Formula .................................................................................... 25

Table 4. Preparation of ureumformamide (UF)/6% acrylamide solutions employed in denaturing

gradient gels. ................................................................................................................................. 27

Table 5. DGGE Gel Casting Reagents .................................................................................................. 27

Table 6. Plzeň and Lochovice Activated Sludge Characteristics .......................................................... 28

Table 7. Average cost, processing time, and mass of waste generated during a 9 sample DNA

Isolation......................................................................................................................................... 28

Table 8. DNA yields (µg/g), absorbance 260 nm/280nm, and absorbance 260nm/230nm.

...................................................................................................................................................... 29

Table 9. PCR amplification of 16S rDNA isolated from Lentikat Biocatalysts® and activated sludge

samples .......................................................................................................................................... 32

Table 10. Mean BSI of all DGGE extracts excluding MB-LN-D2 ....................................................... 41


SBRs

Table 11. Summary of treatments administered to Reactors A and B throughout their operations. ..... 47

Table 12. Hybridization probes employed. Table adapted from Nielsen et al. 2009. ........................... 51

Table 13. Residual hydroxylamine concentrations measured in each reactor in µM ........................... 55

Abbreviations

XII

Amo Ammonia Monooxygenase

Anammox Anaerobic Ammonia Oxidation

AOA Ammonia Oxidizing Archaea

AOB Ammonia Oxidizing Bacteria

ARDRA Amplified Ribosomal DNA Restriction Analysis

ATAD Autothermal Aerobic Sludge Digester

BABE Bioaugmentation Batch Enhanced

BNR Biological Nitrogen Removal

BSA Bovine Serum Albumin

BSI Band Sharing Index (Phylogenetic richness compared to synthetic lane)

CANON Completely Autotrophic Nitrogen Removal over Nitrite

CARD Catalyzed Reporter Deposition

CLSM Confocal Laser Scanning Microscopy

COD Chemical Oxygen Demand

DEMON De-ammonification

DNA Deoxyribonucleic Acid

dsDNA Double stranded DNA

DO Dissolved Oxygen

DGGE Denaturing Gradient Gel Electrophoresis

FISH Fluorescent in situ Hybridization

Hao Hydroxylamine Oxidoreductase

Hh Hydrazine Hydrolase

Hzo Hydrazine Oxidoreductase

IQR Inner Quartile Range

LN Liquid Nitrogen

MAR Microautoradiography

MSW Municipal Solid Waste

MSc Master of Science

Nar Nitrate Reductase

Nir Nitrite Reductase

NirK Nitrite Reductase variation K

NirS Nitrite Reductase variation S

NOB Nitrite Oxidizing Bacteria

Nor Nitric Oxide Reductase

Nos Nitrous Oxide Reductase

Nxr Nitrite Oxidoreductase

OLAND Oxygen Limited Autotrophic Nitrification-Denitrification

OTU Operational Taxonomic Unit

PCR Polymerase Chain Reaction

PFA Paraformaldehyde

PNA Peptide Nucleic Acid

PVA Poly Vinyl Alcohol

qPCR Quantitative Real Time Polymerase Chain Reaction

Abbreviations

XIII

RBC Rotating Biological Contactor

RNA Ribonucleic Acid

rRNA Ribosomal Ribonucleic Acid

SBR Sequencing Batch Reactor

scBNR Short Cut Biological Nitrogen Removal

SEM Scanning Electron Microscopy

SHARON Single Reactor System for High Ammonium Removal over Nitrite

SNAP Single Stage Nitrogen Removal Using Anammox and Partial Nitrification

TKN Total Kjeldahl Nitrogen

UF Ureum-formamide

t-RFLP Terminal-Restriction Fragment Length Polymorphism

WWTP Waste Water Treatment Plant

WW Waste Water

Ch 1. Introduction

1

1.1 Introduction

In the early days of activated sludge wastewater treatment, the primary objectives were

the removal of carbonaceous organic material and the transformation of ammonia to nitrate

(Seviour and Nielsen, 2010). Nowadays, a greater understanding of the environmental

impacts of nitrogen and phosphorus compounds as well as the identification of ecologically

sensitive areas with economic and recreational importance has led to an emphasis on their

removal in wastewater treatment plants (WWTPs). Despite the importance of phosphorus

removal, the subsequent review and research presented will focus on the applied

microbiology of nitrogen removal.

As our understanding of microbiologically mediated nitrogen removal (BNR) processes

advances, techniques continue to emerge to facilitate the proliferation of beneficial organisms

to optimize WWTP performance. While some technologies require the construction of new

facilities, many also focus on retrofitting existing activated sludge WWTPs by altering

process flows, bioaugmenting with biocarriers in existing reactors, and/or by adding new side

stream reactors. Regardless of the approach taken, the ability to characterize and monitor the

community of nitrogen removing microorganisms is essential for maintaining and optimizing

the environmental and economic performance of WWTPs as well as diagnosing the cause of

problems.

The research presented in this MSc. Thesis is meant to 1) develop an internal protocol for

total genomic DNA extraction from nitrifying and denitrifying microorganisms immobilized

in porous polyvinyl alcohol pellets (Lentikat Biocatalysts®) and activated sludge by

comparing commercial DNA isolation kits and 2) to assess the development of nitrifying

bacteria communities immobilized in Lentikat Biocatalysts® and exposed to a known nitrite

oxidizing bacteria (NOB) inhibitor, hydroxylamine, in lab scale sequencing batch reactors

(SBRs) using Fluorescence in situ Hybridization (FISH) and chemical analyses. The

following review is meant to establish the current state of knowledge regarding: biological

nitrogen removal (BNR) pathways, the organisms involved in them, the utility of biocarriers

namely Lentikat Biocatalysts®, and the molecular tools at our disposal for characterizing

microbial communities in environmental and WWTP derived samples.

Ch 2. Literature Review

1

2.1 The Importance of Nitrogen Removal in Wastewater Treatment

The importance of Nitrogen removal from wastewater prior to discharge has become

increasingly clear in light of the negative impacts caused by high concentrations of inorganic

nitrogen species on aquatic environments. Although nonpoint sources such as agricultural

runoff and atmospheric deposition are considered far more problematic, WWTP effluents are

recognized as a significant point source contributing to nitrogen enrichment of both surface

and ground waters (Smith et al., 1999). Some typical values for nitrogen content expressed in

terms of Total Kjeldahl Nitrogen (TKN) and ammonium nitrogen (NH4-N) in various waste

streams requiring treatment are given in Table 1. The three primary environmental impacts

resulting from inorganic nitrogen enrichment are: acidification of freshwater ecosystems;

eutrophication of fresh, brackish and marine ecosystems; and direct toxicity to aquatic

organisms and even humans (Smith et al., 1999; Kloep et al. 2000; Camargo and Alonso,

2006).

Table 1. Typical nitrogen values for various wastewaters adapted from Zanetti et al. (2012)

TKN (mg/L) NH4-N (mg/L)

Raw Domestic Wastewater 60-110 50-100

Swine Wastewater 996-1,520 534-1154

Anaerobic Digester Reject

Water

3,000-5,000 2,000-4,000

While nitrogen induced acidification is most often attributed to atmospheric deposition of

nitrous acid (HNO3) into freshwater ecosystems, the discharge of ionized ammonia (aka

ammonium; NH4+) into systems with low buffering capacities can also be a cause. This in

particular, is due to the release of hydrogen ions (H+) during the naturally occurring oxidation

of ammonium (Camargo and Alonso, 2006). Though worth mentioning, acidification

resulting from WWTP effluent is not as great of a concern as eutrophication or direct toxicity

to aquatic organisms.

High concentrations of ammonia, nitrite, and/or nitrate all exert direct toxic effects on

aquatic organisms (Kloep et al., 2000). Unionized ammonia, NH3, can be acutely or

chronically toxic particularly to fish and is present in significant quantities at elevated pH

(>8) and temperatures (>20oC) (Randall and Tsui, 2002). Notable effects on fish include:

immune system repression, inhibition of ATP production, reduction in blood oxygen carrying

capacity, and suppression of gill tissue function leading to asphyxiation (Camargo and

Alonso, 2006). Likewise, nitrite (NO2-) toxicity has been attributed with causing severe

electrolyte imbalances, disruption of membrane potentials, neurotransmission, muscle


2

contractions, and repression of immune function in fish and crustaceans (Camargo and

Alonso, 2006). However the most important toxic effect of nitrite and nitrate (NO3-) in fish

(and humans) is the oxidation of iron atoms in hemoglobin thus converting it to

methemoglobin and rendering it useless for oxygen distribution. Because toxicity varies by

species, the following water quality criteria for long term and short term maximum exposure

concentrations for aquatic organisms to ammonia, nitrite and nitrate are recommended by

Camargo and Alonso (2006): 0.02-0.35 mg NH3-N/L, 0.35- 3 mg NO2-N/L, and 2-5 mg

NO3-N/L.

The last and perhaps most significant impact of nitrogen enrichment in aquatic

ecosystems is eutrophication. This is defined as a state of being “well nourished” in regards

to the concentrations of growth-limiting nutrients, most often nitrogen and to an even greater

extent phosphorus (Smith et al., 1999). The result of such nutrient enrichment is the excessive

proliferation of algae and macrophytes followed by oxygen depletion as dead biomass

accumulates and decomposes (Camargo and Alonso, 2006). Additional problems induced by

eutrophication include: shifts in macroinvertebrate, vascular plant, and algae composition,

toxic algal blooms, fish kills, reduced water clarity, elevated pH in the water column, loss of

coral reef communities (marine), disruption of drinking water treatment processes, recreation

restriction, and many more (Smith et al., 1999; Camargo and Alonso, 2006). As reported by

Smith et al. (1999) the following values for total nitrogen (mg/m3) may constitute a

“eutrophic” state in lakes, streams and coastal marine systems respectively:

650-1,200; >1,500; and >400.

2.2 Biological Nitrogen Removal (BNR) Pathways

2.2.1 Nitrogen Speciation

The most common forms of nitrogen entering municipal and industrial wastewater

treatment plants are ammonium (NH4+) and ammonia (NH3), in pH dependent equilibrium

skewed towards NH4+

below pH 9, and organic nitrogen compounds including urea, proteins

and amino acids (Seviour and Nielsen, 2010). As biologically catalyzed reactions proceed, a

portion of ammonium and organic nitrogen are directly incorporated into heterotrophic

biomass however a majority is mineralized into NH3/NH4+ in a process referred to as

ammonification (Zeng et al., 2012). Further microbial processes, namely nitrification, give

rise to aqueous oxidized inorganic nitrogen species including nitrite (NO2-), and nitrate (NO3

-

) while denitrification and anammox processes further transform these into gaseous species,


3

primarily dinitrogen (N2) and to a lesser extent nitric oxide (NO) and nitrous oxide (N2O)

which are off-gassed to the atmosphere particularly in oxygen limiting conditions (Seviour

and Nielsen, 2010). It’s worth noting that in conditions of high aeration, typical of many open

air biological reactors, some ammonia (NH3) volatilization to the atmosphere will also occur.

2.2.2 Nitrification

Nitrification is an aerobic two-stage biologically catalyzed process through which

ammonia is fully oxidized to nitrate by way of nitrite as an intermediate.

NH3/NH4+ → NO2

- → NO3

-

Each stage of nitrification is primarily catalyzed by separate functional groups of

chemolithoautotrophic bacteria that obtain energy from oxidizing inorganic compounds and

carbon from CO2 fixation and in some cases the degradation of simple organic compounds.

Unfortunately, the oxidation of inorganic nitrogen species yields relatively low energy, thus

nitrifying bacteria are notoriously slow growers in comparison with most heterotrophic,

organic compound degrading, microbes present in WWTPs (Seviour and Nielsen, 2010;

Whang et al., 2009). As a result, special design and operational parameters must be taken into

consideration in order to facilitate their presence and activity. Furthermore, the stability of

nitrification in wastewater treatment, particularly in activated sludge plants, is highly

sensitive to alterations in environmental and operational parameters. Sudden or even seasonal

changes in temperature, pH, dissolved oxygen (DO), wastewater composition and

concentrations can cause disruptions in nitrification performance that may require many

weeks to recover from (Whang et al., 2009; Zeng et al., 2012). Nevertheless nitrification,

particularly ammonia oxidation, remains a critical component of our wastewater treatment

strategy in terms of nitrogen removal, as no known viable alternatives exist at this time

(Seviour and Nielsen, 2010).

2.2.2.1 Ammonia Oxidation (Nitritation)

Ammonia oxidation to nitrite is the first, and rate limiting, stage of nitrification. It is

mostly attributed to the activity of a functional group of microbes referred to as Ammonia

Oxidizing Bacteria (AOB) that employ the following metabolic reaction pathway (Seviour

and Nielsen, 2010):

NH4+ + O2 + 2H

+ + 2e

- → NH2OH

NH2OH + H2O → HNO2 + 4H

+ + 4e

-


4

The first step of this reaction, the oxidation of ammonia to hydroxylamine, is catalyzed by the

enzyme ammonia monooxygenase (Amo) while the second step, the oxidation of

hydroxylamine to nitrite, is catalyzed by hydroxylamine oxidoreductase (Hao) (Simon and

Klotz, 2013; Seviour and Nielsen, 2010). Based on extensive research, AOB are credited with

carrying out a majority of ammonia oxidation in WWTPs, however it is worth noting that

recently discovered Ammonia Oxidizing Archaea (AOA) and Anaerobic Ammonia Oxidizers

(Anammox) are also thought to play a role in some nitrifying reactors and marine biofiltration

systems (Sakami et al., 2012; Kawagoshi et al., 2012). The significance of the role played by

AOA however, remains poorly characterized therefore continued discussion will focus on

AOB. Anammox will be discussed further in section 2.2.4.

Most AOB are members of the phylum Proteobacteria and more specifically the classes

Betaproteobacteria and Gammaproteobacteria and are widespread throughout terrestrial,

freshwater, and marine ecosystems (Seviour and Nielsen, 2010; Whang et al. 2009). Two

species belonging to the genus Nitrosococcus are the only known AOB from the class

Gammaproteobacteria, and are obligate halophiles widely found in brackish and marine

environments and occasionally brackish biofilters (Seviour and Nielsen, 2010; Kumar et al.,

2013). On the other hand, according to Seviour and Nielsen (2010), Betaproteobacterial

AOB belonging to the genera Nitrosomonas and Nitrosospira are far more diverse and widely

distributed across terrestrial and aquatic ecosystems. Among these, Nitrosospira are believed

to be the dominant AOB in soil ecosystems while Nitrosomonas seem to include a broader

array of physiologically diverse/tolerant species with greater presence in a various aquatic

environments, especially WWTPs. This being said, Whang et al. (2009) and Sakami et al.

(2012) report that Nitrosospira-like AOB do occasionally exist and play a role in nitrifying

WWTP reactors and marine aquaculture biofilters.

2.2.2.2 Nitrite Oxidation (Nitratation)

Nitrite oxidation to nitrate is the second and final stage in the aerobic process of

nitrification and is mostly carried out by a functional group referred to as Nitrite Oxidizing

Bacteria (NOB). Unlike AOB, NOB are not all strict chemolithoautotrophs in that some are

also known to oxidize simple organic compounds like acetate and pyruvate (Seviour and

Nielsen, 2010). Furthermore, they possess greater diversity of enzymatic machinery for the

oxidation of nitrite than AOB do for ammonia. The only well characterized nitrite oxidation

enzyme is nitrite oxidoreductase (Nxr) belonging to the genus Nitrobacter which catalyzes


5

the following metabolic reaction (Seviour and Nielsen, 2010; Vanparys et al., 2007; Simon

and Klotz, 2013):

NO2- + H2O

↔ NO3

- + 2H

+ + 2e

-

Phylogenetically speaking, NOB, include at least one member from the phylum Chloroflexi

as well as Proteobacteria genera from the classes Alphaproteobacteria (Nitrobacter),

Deltaproteobacteria (Nitrospina), and Gammaproteobacteria (Nitrococcus) as well as

Nitrospira (name of class and genus) from the phylum Nitrospirae (Vanparys et al., 2007;

Seviour and Nielsen, 2010; Sorokin et al., 2012).

Of these genera, Nitrococcus and Nitrospina, have only been isolated from marine

ecosystems while Nitrobacter and Nitrospira have been found to tolerate a wide swath of

aquatic and terrestrial environments and are the most commonly detected NOB in WWTPs

(Seviour and Nielsen, 2010; Han et al., 2012).

2.2.3 Denitrification

Denitrification is a four step anaerobic/anoxic process whereby nitrate and nitrite are

transformed to gaseous species through the following metabolic reaction pathway (Seviour

and Nielsen, 2010):

NO3- → NO2

- → NO

→ N2O

→ N2

While the end product of N2 is most desirable because of its inert nature in the atmosphere,

releases of the intermediates NO and N2O (both greenhouse gasses) can be significant

depending on the denitrifying community present and environmental conditions (Seviour and

Nielsen, 2010; Kong et al., 2013).

Bacteria carrying out denitrification are far more diverse and heterogeneous a group

than AOB or NOB. Typically, denitrifiers are facultative anaerobes that utilize nitrate and

nitrite as alternative terminal electron acceptors in their respiration when ample oxygen is not

available. The ability to do this has been found across the spectrum of prokaryotes from

organotrophs, lithotrophs, and diazotrophs, to halophiles and thermophiles (Seviour and

Nielsen, 2010). This heterogeneity makes denitrifiers a more difficult functional group to

define, however common genera reported in literature include: Bacillus, Pseudomonas,

Paracoccus, Hyphomicrobium, Azoarcus, Marinobacter, Halomonas, Methylophaga, and

many more (Yoshie et al., 2004; Song et al., 2012; Osaka et al., 2008).

The metalloenzymes essential for denitrification are Nitrate reductase (Nar), Nitrite

reductase (Nir), Nitric oxide reductase (Nor), and Nitrous oxide reductase (Nos). As is the


6

case for Amo in AOB, there are different forms of these enzymes coded for by different gene

sequences, for example the nitrite reductase variations NirS and NirK. Due to the diversity of

denitrifying organisms, molecular tools discussed in section 2.5, enable the quantification of

these gene sequences and have become the preferred method for assessing the abundance of

denitrifying bacteria in environmental samples and WWTPs (Warneke et al., 2011).

2.2.4 Anaerobic Ammonia Oxidation (Anammox)

Anaerobic ammonia oxidation is a nitritation-denitritation alternative to the classical

nitrification-denitrification BNR pathway discussed previously. As the name suggests,

anammox, is an anaerobic microbially catalyzed process whereby ammonia serves as the

electron donor, nitrite as the electron acceptor, and dinitrogen gas is the final product (Feng et

al., 2007). The process proceeds via the following reaction pathway (Feng et al., 2007; Jetten

et al., 2009; Seviour and Nielsen, 2010):

Overall Reaction: NH4+ + NO2

- N2 + H2O

Metabolic Pathway: NO2- → NO, NO + NH4

+ → N2H4

→ N2 + H2O

Anammox Stoichiometry: NH4

+ + 1.32NO2

- + 0.066HCO3

- + 0.13H

+ 1.02N2 + 0.26NO3- + 0.066CH2O0.5N0.15 + 2.03H2O

First, nitrite is reduced to nitric oxide by nitrite reductase (NirS); next hydrazine hydrolase

(Hh) reduces nitric oxide while simultaneously oxidizing ammonium to form hydrazine.

Lastly, hydrazine is oxidized by hydrazine oxidoreductase (Hzo) to produce dinitrogen gas,

water, and a very small amount of byproduct nitrate (Feng et al., 2007; Jetten et al., 2009).

Anammox bacteria are slow growing chemolithoautotrophs belonging to phylum

Planctomycetes. They are obligate anaerobes with high sensitivity to oxygen (>2 µM O2 is

fatal), they rely on CO2 fixation and bicarbonate as their carbon sources, have doubling times

between 10-20 days (lower in situ), and have relatively low biomass yields (Jetten et al.,

2009; Seviour and Nielsen, 2010). Owing to their low growth rate, they tend to thrive in

natural environments that have low substrate concentrations. On the other hand they do not

all adhere strictly to chemolithoautrophic metabolism as some are known to employ ferrous

iron and/or specific organic compounds as alternative electron donors and ferric iron,

manganese oxides, and/or nitrate as alternative electron acceptors (Jetten et al., 2009). To

date, 5 candidatus genera have been described including Kuenenia, Brocadia, Jettenia,

Anammoxoglobus, and Scalindua. The first four were all enriched from activated sludge


7

while Scalindua was found in marine sediments. Anammox is estimated to be responsible for

33-50% of global nitrogen removal from marine ecosystems (Dalsgaard et al., 2005).

2.3 Applications of Biological Nitrogen Removal (BNR) Pathways in

Wastewater Treatment and the Case for Partial Nitrification

The earliest activated sludge WWTPs focused on the removal of organic carbon

compounds and the transformation of ammonia to less toxic species like nitrite and nitrate

(Seviour and Nielsen, 2010). As discussed in section 2.1, the negative environmental impacts

of these species have since been well documented thus necessitating the implementation of

more comprehensive nitrogen removal systems. To accommodate this requirement, there has

been a massive proliferation of reactor types, multistage configurations, process control

strategies, and bioaugmentation techniques used at WWTPs. From an engineering standpoint,

there are far too many nitrogen removal approaches to go into any detail here, however from

the standpoint of BNR pathways the options are far more limited.

Nitrification-denitrification via nitrate has long been the dominant BNR pathway

employed in WWTPs around the world (Kuenan and Robertson, 1994; Zanetti et al., 2012;

Seviour and Nielsen, 2010). Its rise to dominance is due to its simplicity in regards to

providing more or less stable performance, while relying on a simple staged aerobic/anoxic

configuration (though modern systems can be far more complex), the occasionally necessary

dosing of simple organic substrates (COD) to facilitate adequate denitrification, and

prescribed (fixed) operational controls (Seviour and Nielsen, 2010; Zanetti et al., 2012).

Although effective, this strategy is quite expensive due to the high cost of aeration and the

dosing of COD when needed. This has fueled great interest in the development of potentially

cheaper alternatives aimed at employing so called “short cut” biological nitrogen removal

(scBNR) pathways particularly for use on waste waters low in COD.

There are two known scBNR pathways that both rely on partial nitrification, namely

nitritation facilitated by AOB, as their initial step. The first option is nitritation followed by

heterotrophic denitritation which cuts out the conversion of nitrite to nitrate thereby reducing

the amounts of oxygen and COD consumed (Aslan and Dahab, 2008; Zanetti et al., 2012).

The second option employs only partial nitritation followed by anammox which not only

further reduces oxygen consumption but also eliminates COD consumption (though

bicarbonate and/or CO2 are needed) (Lan et al., 2011; Zanetti et al., 2012). The O2 and COD

requirements of the three BNR pathways are summarized in Table 2.


8

Table 2. Stoichiometric comparison of BNR pathways taken from Zanetti et al. (2012)

gO2 / gN gCOD/ gN

Nitrification-Denitrification 4.57 4

Nitritation-Denitritation 3.43 2.4

Partial Nitritation-Anammox 1.72 NA

The key to achieving partial nitrification is selecting for AOB over NOB. Currently

established strategies for accomplishing this include: maintaining reactor temperatures over

25oC, maintaining free ammonia (NH3) levels above 1 mg/L, maintaining a free nitrous acid

(H-NO2-N) concentration between 0.011-0.10 mg/L, limiting oxygen concentrations to

between 0.5-1.5 mg/L (though difficult to control), and chemical inhibition with

hydroxylamine (Zanetti et al., 2012; Park and Bae, 2009; Xu et al., 2012). Several reactors

have been developed that employ one or more of these strategies including the high

temperature nitritation Single Reactor System for High Ammonium Removal over Nitrite

(SHARON), the oxygen limited AOB/anammox single reactor Completely Autotrophic

Nitrogen Removal over Nitrite (CANON) and Oxygen Limited Autotrophic Nitrification-

Denitrification (OLAND) systems, as well as the De-ammonification (DEMON), Single

Stage Nitrogen Removal Using Anammox and Partial Nitrification (SNAP), and

Bioaugmentation Batch Enhanced (BABE) systems (Kumar and Lin, 2010).

Chemical inhibition of NOB via the dosing of hydroxylamine (NH2OH) is a less

studied approach that has yet to be applied outside of laboratory settings. Kindaichi et al.

(2004) followed up on previous studies that established NH2OH dosing as a method for

stimulating AOB growth but noted inhibitory effects on NOB by examining this inhibition.

They found that the addition of 250 µM NH2OH completely inhibited the growth on NOB in

rotating biological contactor (RBC) biofilms. Xu et al. (2012) then investigated the use of

NH2OH inhibition as a strategy for establishing a partial nitrification reactor using aerobic

granules that are not compatible with the free ammonia or oxygen limitation strategies.

Though they were unsuccessful at achieving the appropriate ratios of NH4+ and NO2

- for

subsequent anammox treatment, they reported great compatibility of inhibiting NOB with

NH2OH in aerobic granules. Extrapolating upon this conclusion, it seems as though

controlled NH2OH dosing has great potential as a strategy to facilitate the start-up and

performance of partial nitrification reactors thus further research is warranted.


9

2.4 Lentikat Biocatalysts

Biocatalyst, also called biocarrier, technology is based on the encapsulation of

functional microorganisms within porous hydrogel matrices for use in biologically mediated

processes. This concept has been successfully applied for the production of bioethanol,

pharmaceutical enzymes, artificial seeds, and artificial cells in addition to medical treatments

and bioaugmentation of BNR at WWTPs (Park and Chang, 2000).

Lentikat’s Inc. is a market leading producer of biocatalysts based in Prague, Czech

Republic. Their patented biocatalysts (Lentikat Biotechnology: German Patent

# DE 198 27 552) are made from nontoxic, non-biodegradable, non-abrasive, and highly

elastic polyvinyl alcohol (PVA) (Bouskova et al., 2011; Park and Chang, 2000; Vackova et

al., 2012). Each lens shaped PVA pellet is approximately 3-4 mm in diameter and 200-

400 µm in width as shown in Figure 1. They are produced at room temperature by blowing a

mixed solution, containing the cell suspension and dissolved polymer, through a jet nozzle

into a rotating wire wheel composed of 1 µm wires. The solution is cut into appropriately

sized beads which land on a moving film where they harden with the help of a warm air

blower (Park and Chang, 2000; Bouskova et al., 2011). Following immobilization, pellets

undergo a 6 week cultivation period in the laboratory (Vackova et al., 2011).

Figure 1. Lentikat Biocatalyst structure, the porous PVA hydrogel is depicted in blue while the orange

spots represent encapsulated microorganisms (Bouskova et al., 2011)

Lentikat’s wastewater treatment biocatalysts are specially designed to facilitate

enhanced nitrogen removal. They currently produce three separate pellets, one for

nitrification that contains encapsulated Nitrosomonas europaea and Nitrobacter winogradskyi

and the other two for denitrification containing either encapsulated Paracoccus denitrificans

or Pseudomonas fluorescens (Bouskova et al., 2011). Because the production technique is

carried out at room temperature, nitrifying organisms in Lentikat’s Biocatalysts retain far

greater levels of activity compared to PVA biocatalysts produced at higher temperatures,

which most nitrifiers do not tolerate (Park and Chang, 2000).


10

Biocatalyst technologies have demonstrated a number of advantages when compared

with conventional suspended culture bioaugmentation technologies. They are easy to handle

and recover from solutions and provide enhanced volumetric nitrogen removal by facilitating

a high density of cells without the threat of washout and with reduced susceptibility of

nitrifiers to predation (Vackova et al., 2011; Ravnjak et al., 2013). Bouskova et al. (2011)

reported nitrogen removal efficiencies of 98% in well maintained systems (provided with

ample oxygen and organic substrate) treating concentrations of N-NH4+ and N-NO3

- as high

as 2500 mg/L and 4000 mg/L respectively. They also demonstrated that Lentikat’s

Biocatalysts retained high nitrogen removal efficiency when applied to separate wastewaters

containing high concentrations of inorganic salts (20 g/L NaCl and 2 g/L Na2SO4) and toxic

compounds (Aniline) at concentrations far greater than previously reported to cause

inhibition. These results correlate well those of Barber and Stuckey (1999) who found that

Nitrobacter immobilized in porous PVA beads demonstrated decreased susceptibility to

unionized Ammonia (NH3) concentrations previous reported to be inhibitory.

These data support the claims that encapsulation of biomass enhances adaptability to

harsh conditions and increases robustness towards fluctuating environmental parameters such

as chemical shocks, pH changes, and temperature shifts (Bouskova et al., 2011; Trogl et al.,

2011; Ranjak et al., 2013; Barber and Stuckey, 1999; Park and Chang, 2000). Consequently,

nitrifying bacteria immobilized within PVA seem to possess great potential to enhance the

performance of new and existing WWTPs as legislative restrictions on nitrogen emissions get

tighter.

2.5 Molecular Methods for Assessing Microbial Communities in WWTP

Molecular biological techniques for assessing microbiological communities in

environmental samples and WWTPs have proven effective at overcoming the biases and

shortfalls of the culture based approaches that previously dominated such analyses (Gilbride

et al., 2006). Some of the powerful advantages of molecular biological approaches include:

the ability to characterize the structure of complex mixed communities in situ as well as

identify representative populations of difficult to culture anaerobes, currently unculturable

species, and functional groups, such as nitrifiers and denitrifiers, based on unique DNA

sequences and/or genes coding for functional enzymatic machinery. These tools can therefore

provide engineers with a greater ability to optimize WWTP performance by facilitating


11

beneficial organisms while combating problematic ones on a continual basis (Gilbride et al.,

2006; Sanz and Kochling, 2007).

2.5.1 DNA Isolation

The isolation, or extraction, of pure genomic DNA from environmental or WWTP

samples is a critical initial step in all molecular analyses of microbial populations that require

subsequent polymerase chain reaction (PCR) DNA amplification (Mahmoudi et al., 2011).

The process of DNA isolation includes cell lysis and homogenization which typically

involves heating, detergents, and/or mechanical force followed by the stepwise removal of all

non DNA constituents and the eventual elution of DNA in a suitable storage buffer

(Whitehouse and Hottel, 2006; Mahmoudi et al., 2011). Isolating acceptable quantities of

high purity DNA from soil and activated sludge samples can be particularly challenging due

to presence of compounds that may inhibit downstream PCR such as humic acids and other

recalcitrant organic compounds and/or pollutants (Whitehouse and Hottel, 2006; Guobin et

al., 2008; Mahmoudi et al., 2011).

Nowadays, numerous commercial DNA extraction kits are available that enable the

processing of high volumes of samples with relatively lower cost and time consumption in

comparison with previous established methods (Whitehouse and Hottel, 2006; Dauphin et al,

2009). An additional advantage of these commercial kits is the incorporation and continuing

improvement of PCR inhibitor removal techniques (Whitehouse and Hottel, 2006). On the

other hand, previous investigations report differential suitability of kits in their applications to

environmental and WWTP derived samples and some degree of “extraction bias” when

different kits are applied to the same samples. This implies that microbial community

assessments obtained from subsequent analyses may not fully represent reality and therefore

adequate investigation is needed to determine the most suitable DNA extraction protocol for

each particular sample type (e.g. activated sludge or Lentikat Biocatalysts)(Gilbride et al.,

2006; Mahmoudi et al., 2011).

Despite the apparent limitations, DNA Isolation remains essential for the

characterization of microbial communities in soils and WWTPs using molecular techniques

including: Amplified Ribosomal DNA Restriction Analysis (ARDRA), Denaturing Gradient

Gel Electrophoresis (DGGE), Terminal-restriction Fragment Length Polymorphism (t-

RFLP), Multiplex PCR, Real Time PCR (qPCR), and Nucleic Acid Microarrays (Gilbride et

al., 2006; Sanz and Kochling, 2007).


12

2.5.2 Polymerase Chain Reaction (PCR)

PCR is a nucleic acid amplification technique developed in the 1980’s that has

become a cornerstone of many molecular analyses employed across nearly the entire

spectrum of biological sciences. PCR proceeds via the cyclical denaturation of DNA

molecules by heating, followed by cooling which triggers hybridization with target sequenced

primers (most often targeting genes coding for 16S rRNA in Prokaryotes and 18S rRNA in

Eukaryotes) that are assembled into complimentary strands, or copies, of the original

fragment by DNA polymerase (Mullis et al., 1986). During each subsequent cycle the DNA

fragments generated in all previous cycles also serve as templates for further amplification

thus resulting in the exponential increase in the concentration of target DNA molecules

(Mullis et al., 1986). While the input concentrations of nucleic acid may be extremely small,

the sensitivity of successful PCR enables the output of sufficient DNA concentrations for

comprehensive analyses including the assessment of subdominant species (Gilbride et al.,

2006; Postollec et al., 2011).

Since its inception numerous PCR techniques and PCR based analyses have been

developed and successfully employed to assess microbial communities in WWTP’s. Among

the most commonly used for nitrifying bacteria and archaea are quantitative real time PCR

(qPCR) and conventional PCR followed by denaturing gradient gel electrophoresis (PCR-

DGGE) (Sanz and Kochling, 2007; Gilbride et al., 2006). qPCR involves the generation of an

amplification curve by quantifying the fluorescence given off by amplicons at every cycle

throughout the amplification process. This provides an estimate of the initial concentration of

the target DNA molecule and thus its source organism’s abundance in the initial sample

(Postellec et al., 2011). This technique is frequently applied to assess the populations of total

bacteria, AOB, NOB, anammox, and bulking organisms in WWTPs (van den Akker et al,.

2010; Davery et al., 2013; Wang et al., 2010; Wang et al., 2012 ). A major advantage of

qPCR is that it does not require post PCR manipulation and therefore minimizes additional

contamination risk (Postellec et al., 2011). On the other hand, while its inherently high

sensitivity enables the identification of subdominant species with high accuracy, it also

dictates that extra controls over experimental design must be applied to eliminate errors

(Postellec et al., 2011). The present study undertaken in this MSc. thesis employs PCR-

DGGE which is described in more detail in the following section (2.5.3).

As mentioned in section 2.5.1, PCR is vulnerable to inhibition by a variety of

materials and chemical constituents associated with WWTP and environmentally derived


13

samples. Besides insufficient initial DNA concentration, inhibition is the greatest cause of

PCR failure (Alaeddini, 2012). Substances reported to cause PCR inhibition include: humic

compounds, polysaccharides in feces, heme in blood samples, proteinases and phenols used

in DNA extraction, heavy metals, certain divalent ions, nanoparticles, excessive DNA, urea,

certain microfluidic chips, and many others (Alaeddini, 2012; Kodzius et al., 2011). While

the precise mechanisms of PCR inhibition are not yet well understood, several methods for

overcoming inhibition are routinely used starting with selecting the most appropriate DNA

extraction protocol. As noted by Whitehouse and Hottel (2006), many commercial DNA

extraction kits are designed to work with particular sample types (e.g. soil or stool) by

incorporating washing buffers that target commonly associated inhibitors for removal.

Additional, techniques used to overcome PCR inhibition include: DNA dilution, addition of

amplification facilitators like bovine serum albumen (BSA), the addition of extra polymerase

enzymes, or DNA purification (Alaeddini, 2012).

2.5.3 Denaturing Gradient Gel Electrophoresis (DGGE)

DGGE is a genetic finger printing technique for determining the dominant members

of microbial communities in environmental samples and WWTP sludges with high precision

(Sanz and Kochling, 2007; Hesham et al., 2011). The principle of this technique is that PCR

amplified DNA fragments of the same length are separated in a polyacrylamide gel

containing a gradient of DNA denaturants. The basis of this denaturation is the differing

sequence of base pairs in each fragment giving rise to unique melting domains. This refers to

a section of the DNA fragment with identical melting temperature. Once one of these sections

reaches its melting temperature the helical DNA structure breaks down creating drag that

severely restricts continued migration through the gel (Muyzer and Smalla, 1998). The end

result is a series of bands in the gel, each representing an accumulation of DNA fragments

with an identical sequence of base pairs thus representing an individual “species”. DGGE

profiles can be used to make simultaneous comparisons of samples (e.g. in time series

investigation) and/or bands can be cut from the gel and the DNA therein can be sequenced to

determine the species represented (Muyzer and Smalla, 1998).

The applications of PCR-DGGE in the analyses of WWTP communities have been

rapidly increasing over the past decade. Successful applications include: characterizing the

communities carrying out essential functions such as anaerobic sludge digestion (Kim et al.,

2009) and autothermal aerobic sludge digestion (Hayes et al., 2011); analyzing the effects of


14

changing operational parameters such as temperature (Niu et al,. 2012) and aeration regimes

(Tocchi et al., 2012.) in aerobic activated sludge reactors; comparing raw sewage and aerobic

sludges from different WWTPs to assess performance differences (Liu et al., 2006); and

comparing compartments and treatment trains within a single WWTP to assess bulking

problems (Hesham et al., 2011).

Particularly relevant to this study, DGGE has also long been applied to detect

differences in DNA extraction protocols (e.g. differential cell lysis techniques) and PCR

biases (Muyzer and Smalla, 1998; Sanz and Kochling, 2007). Quigley et al. (2012) and

Mahmoudi et al. (2011) used DGGE profiles to compare DNA isolated using different

commercial DNA extraction kits on raw milk and soil samples respectively in terms of yield,

purity, and the presence of PCR inhibitors. Both studies found that significant differences

occurred between kits in terms of purity, yield (total and variability), and species composition

in the case of soil. The aim of this current study is to use DGGE as a tool to help establish an

internal DNA extraction protocol for future characterization and long term evaluation of

communities encapsulated within and fixed upon Lentikat Biocatalysts® and activated sludge

flocs.

2.5.4 Fluorescence in Situ Hybridization (FISH)

FISH is a cultivation independent technique employed to assess the phylogenetic and

spatial composition of microbial communities derived from environmental samples and

WWTP sludges. The principle behind this method is that samples are immediately fixed, then

the cells are permeabilized, nucleic acids are hybridized with fluorescently labeled

oligonucleotide probes, thoroughly washed and examined using flow cytometry or

epifluorescent or confocal laser scanning microscopy (CLSM) (Amman et al., 1995; Gilbride

et al., 2006). FISH is extremely useful in WWTP studies in that it enables the characterization

of the structure and quantity of morphologically in-tact microorganisms present in complex

communities down to the single cell level (Amman et al., 1995; Nielsen et al., 2009). Some

additional highly publicized advantages of FISH over other molecular techniques are: the

speed with which it can be carried out; the simplicity of microscopic analyses; the specificity

of RNA, DNA, and more recently PNA (peptide nucleic acid) probes to target whole domains

(e.g. Bacteria, Archaea, or Eukarya) on down to single species and sub-species; and its

relative immunity to inhibition or extraction and amplification biases that can affect PCR

based analyses (Amman et al., 1995; Nielsen et al., 2009; Okten et al., 2012; Machado et al.,


15

2013). Problems commonly encountered while using FISH include: autofluorescence of cells

and surrounding materials, nonspecific binding of fluorescent labeled oligonucleotides, and

low signal intensity (Amman et al., 1995; Okten et al., 2012).

While a majority of the primers used in PCR target DNA sequences coding for 16S

ribosomal RNA (rRNA) in prokaryotes and 18S rRNA for eukaryotes, the oligonucleotide

probes used in FISH target the rRNA itself (Gilg et al., 2010; Machado et al., 2013). The

highly conserved nature of these sequences makes them well suited for defining operational

taxonomic units (OTUs), or the microbiological equivalent of “species”, and is the primary

reason for their targeting by FISH probes. Another reason for targeting rRNA is that it is

present in cells in far greater quantities than DNA and thus gives off a more pronounced

signal (Nielsen et al., 2009). The establishment of extensive and periodically scrutinized

databases, such as the University of Vienna’s “probeBase” (Loy et al., 2003), has provided a

foundation for the development of a wide variety of probes now used in FISH (Lucker et al.,

2007; Nielsen et al., 2009). Refinement of probes is also continually occurring as researchers

identify regions of rRNA with highly limited access to oligonucleotide diffusion (Okten et

al., 2012) and PNA probes demonstrate greater specificity and thermal stability than some

RNA and DNA alternatives (Machado et al., 2013).

Fluorescent microscopy employed in FISH is based on the principle of exposing

hybridized specimens to short wavelength visible light to excite the fluorescent dyes,

fluorophores, which in turn leads to them emitting longer wave light which can be detected

(Seviour and Nielsen, 2010). The most common fluorophores used in FISH include:

fluorescein (FITC), tetramethylrhodamine, and indocarbocyanines (CY3, CY5, and Cy7)

(Amman et al., 1995). Recent advances in FISH image analysis are due in large part to the

use of CLSM and associated computer software that combine the ability to take three

dimensional images with automated and/or semi-automated quantification of hybridized cells

(Seviour and Nielsen, 2010).

The application of FISH in investigations of WWTP microbial communities is

extremely widespread. A majority of these studies focus on characterizing populations of

functional groups such as nitrifiers, denitrifiers, polyphosphate accumulators, glycogen

accumulators, as well problematic filamentous organisms (Nielsen et al., 2009). While

traditional FISH is still most frequently used as a standalone molecular method for analysis,

newer variations that include catalyzed reporter deposition (CARD-FISH) and


16

Microautoradiography (MAR-FISH) are becoming increasing popular (Seviour and Nielsen,

2010).

The use of FISH to assess nitrifying bacteria in WWTPs has been an extremely

common practice since the late 1990’s (Gilbride et al., 2006). More recently, FISH has

emerged the method of choice for assessing AOB vs. NOB present in partial nitrification

reactors as demonstrated by Cho et al. (2010), Zhang et al. (2012), Kong et al. (2013), Okabe

et al. (2011), and Gu et al. (2012) just to name a few.

Perhaps the most interesting trend, evident in a number of the partial nitrification

investigations mentioned above, is the evolving tendency to employ FISH in combination

with other molecular tools. FISH is increasingly being combined with scanning electron

microscopy (SEM), qPCR, and PCR-DGGE in order to provide comprehensive analyses of

microbial communities that combine the advantages of each, while attempting to mitigate and

identify their shortcomings. This trend is not confined to partial nitrification but is becoming

prevalent throughout microbial ecology with examples including: Yasin et al. (2012), who

used a combination of FISH and PCR-DGGE to characterize the hydrogen producing

communities in food waste fermenters; Cardinali-Rezende et al (2012), who combined FISH,

CARD-FISH, qPCR, and DGGE to track the evolution of the microbial community in a full

scale municipal solid waste (MSW) anaerobic digester from start up to steady state;

Fernandes et al. (2013) who used FISH to track phosphorus accumulators, nitrifiers and

sulphate reducers while using DGGE to track overall community structure in a full scale

sequencing batch reactor treating domestic wastewater for 180 days; Hayes et al. (2011) who

combined FISH, FISH-MAR, and PCR-DGGE to assess the microbial ecology of an

autothermal thermophilic aerobic sludge digester (ATAD); and Ferrero et al. (2010) who

combined PCR-DGGE and FISH to characterize the microbial populations in high altitude

soils.

Interestingly, Cardinali-Rezende et al. (2012) noted that for a majority of their

overlapping analyses, qPCR yielded lower cell enumerations than FISH by 1-4 orders of

magnitude. While this trend was not universal, they attributed the difference to potential

losses of DNA during extraction and purification. In a more complimentary combination,

Kong et al. (2013) used FISH to determine that Nitrosomonas was the dominant genus of

AOB in his lab scale partial nitrification reactor and PCR-DGGE on genes coding for

variations of Amo to determine the evolution of dominant Nitrosomonas species over time. If

nothing else, the complimentary and sometimes conflicting results presented in these articles


17

indicate that combining FISH with additional molecular tools for microbial ecology

investigations is becoming mainstream.

Ch 3. Genomic DNA Isolation and Amplification from Bacteria Immobilized in Poly Vinyl Alcohol Biocarriers

18

3.1 Objectives

The objectives of this investigation are to compare the effectiveness of four commercial DNA

isolation kits at isolating bacterial DNA from PVA biocarriers (Lentikat’s Biocatalysts) for

use in downstream PCR based applications. The primary criteria for comparison are DNA

yield, purity, waste generation, processing time, cost per sample, successful PCR

amplification, and phylogenetic richness in extracts.


19

3.2 Materials and Methods

3.2.1 Poly Vinyl Alcohol Pellets and Activated Sludge Samples

A total of 6 unique samples were analyzed in this project, 4 of Lentikat’s Biocatalysts

and 2 of aerobic activated sludge. Paracoccus cultures for immobilization in PVA

biocatalysts were obtained from the collection of Deutsche Sammlung von Microorganismen

and Zellkulturen GmbH while nitrifiers were enriched from activated sludge and were

cultivated at the Slovak University of Technology in Bratislava. The specific PVA employed

for immobilization was Mowiol® 28-99 (Kuraray America, Inc., Houston, USA) with a 99%

degree of saponification (hydrolysis) and relative molecular mass of 145,000 g/mol. The

preparation of lens-shaped biocatalysts (diameter, 3 - 4 mm; thickness 200–300 μm) was

described in section 2.4 and was carried out by LentiKat´s Incorporated (Prague, Czech

Republic).

Two of the biocatalyst samples, N1 and N2, were nitrification biocarriers containing

immobilized Nitrosomanas europaea and Nitrobacter winogradskyi. Sample N1 was

obtained from LentiKat’s Inc. immediately after immobilization and prior to the subsequent 6

week in lab cultivation period that all commercially distributed biocatalysts undergo. Sample

N2 was obtained following 4 months of implementation treating effluent from a municipal

root zone WWTP in Kotenčice, Czech Republic. Both samples, N1 and N2 were collected

and transported to ICT Prague on February 15, 2013

The other 2 biocatalyst samples, D1 and D2, were both denitrification biocarriers

containing immobilized Parococcus denitrificans (strain DSM 1403). Sample D1 was

obtained following 38 months of implementation performing post denitrification on effluents

from mixed sewage and industrial waste water at a private pharmaceutical production plant.

Sample D2 was obtained following 9 months of implementation performing post

denitrification of chemically treated underground water from uranium mining (DIAMO

Corp). Organic substrate augmentation at both of these of these facilities was carried out with

the controlled dosing of Brenntaplus vp1 (Brenntag N.V., Deerlijk, Belgium) carbon rich

nutrient blend. Both samples, D1 and D2, were collected and transported to ICT Prague on

February 15, 2013.


20

The two activated sludge samples, L1 and P1, were collected from aeration basins in

conventional suspended growth activated sludge WWTPs. Sample L1 was obtained on March

20, 2013 from the Lochovice, CZ WWTP operated by Envi-Pure Inc. (Prague, Czech

Republic). This plant treats primarily Industrial WW. Sample P1 was obtained on March 20,

2013 from the Plzeň, CZ municipal WWTP operated by Veolia Water (Paris, France). This

plant treats primarily MSW and industrial WW primarily from Beer Breweries.

Once in the laboratory, all biocatalyst samples were transferred to 50 mL Falcon

tubes, residual water was removed by pipetting, and tubes were stored at -20oC until DNA

extraction. Activated sludge samples were stored briefly in 3L aerated carboys. Prior to DNA

extraction samples were thoroughly homogenized, small amounts were then transferred to

sterile 2 mL microcentrifuge tubes, centrifuged for 1 min after which residual water was

removed by pipetting and the thickened product was used for isolation.

3.2.2 Activated Sludge Characterization

The TSS and VSS of activated sludge samples were determined following protocols

adapted from 2540 D in Standard Methods for the Examination of Water and Wastewater.

22nd

Edition (2012):

TSS

1. Filter 10 mL of thoroughly homogenized sludge under vacuum onto a pre-

weighed Pragapor #6 membrane filter (50 mm diameter, 0.4 µm thick)

(Pragochema spol. s.r.o, Prague, CZ)

2. Place filter in over to dry at 105oC for 2 hours and weigh

3. Calculate TSS with the following formula:

TSS =

TSS total suspended solids [g/L];

SF weight of filter and sludge after drying [mg];

F weight of filter [mg];

V homogenized sample volume used for analysis [mL].


21

VSS

1. Following determination of TSS place the filter in a pre-weighed porcelain dish

2. Add 2-3 drops of glycerin and place in an electric oven at 550oC for 2 hours, dry

in the desiccator and weigh.

3. Calculate VSS with the following formula:

VSS =

VSS … volatile suspended solids [g/L];

SFD weight of dish, filter, and residue after drying at 105 °C but prior to burning

[mg]

SBD weight of dish, filter, and residue after burning [mg];

F weight of the filter [mg];

V homogenized sample volume used for analysis [mL].

3.2.3 DNA Isolation

3.2.3.1 Comparison of Commercial DNA Isolation Kits

Bacterial DNA was isolated from all 6 samples in triplicate, using four different

commercial DNA extraction kits selected to include several nucleic acid extraction

methodologies. The manufacturer´s protocols for gram-negative bacteria were followed for

each kit and can found online (as of 3/6/2013) at the links provided in Appendix 1. Slight

modifications to some protocols were made based upon manufacturer recommended

troubleshooting following a test run to familiarize ourselves with each procedure. The

average processing time of a 9-sample run was determined for each kit beginning with the

addition of the first reagent and ending with the final elution of DNA and calculated based

upon two runs. In addition, the cost per sample was determined based upon the ratio of kit

price/preps and the mass of waste generated in each 9 sample run was recorded and averaged.

The cost of pipette tips, eppendorf tubes, standard laboratory equipment, and inexpensive

(< €200) kit specific equipment (Chemagic: Magnetic stand; PowerSoil®: Vortex adapter

tube holder) were excluded from these cost analyses. All extracted DNA was eluted in the

buffers provided by the manufacturers and subsequently stored at -20 °C until downstream

use. The kits evaluated and methodologies employed include:


22

The QIAamp® DNA Stool kit (50 preps) (QIAGEN Inc., Valencia, CA, USA)

Principle: Lyse bacterial cells with enzymes and heating (70oC); bind Impurities

to inhibit-ex tablets (a fluidized adsorption media); bind DNA to membrane and

wash; elute DNA in buffer.

Protocol Modifications: Reduced elution buffer from 200 μL to 100 μL

Justification: Preliminary results yielded very low concentrations of DNA in

eluate and thus unreliable Nanodrop™ results.

The PowerSoil® DNA Isolation kit (50 preps) (MoBio Laboratories Inc., Carlsbad, CA,

USA) Bead Beating and Membrane.

Principle: Lyse bacterial cells with combined mechanical and enzymatic force;

precipitate initial impurities; bind DNA to membrane and wash; elute DNA in

buffer.

Protocol Modifications: A) Increased step 5 (mechanical lysis) vortexing time

from 10 to 15 minutes, B) Reduced elution buffer from 100 μL to 50 μL

Justification: A) Optional manufacturer recommendation, B) Preliminary results

yielded very low concentrations of DNA in eluate and thus unreliable Nanodrop™

results.

The Chemagic DNA Bacteria Kit (100 preps) (PerkinElmer chemagen Technologe

GmbH, Baesweiler, Germany)

Principle: Lyse bacterial cells with enzymes and mild heating (37oC); bind DNA

to magnetic beads; washout proteins, RNA, Lipids, etc.; elute DNA in buffer.

Protocol Modifications: Used 4 μL of RNAse A (5 mg/ml) in lysis (Step 1)

Justification: Optional Manufacturer Recommendation

The MasterPure™ DNA Purification kit (200 preps) (Epicentre Biotechnologies,

Madison, WI, USA)

Principle: Lyse cells with enzymes and heating (65oC); cool and precipitate

impurities; precipitate DNA with alcohol; rinse with alcohol and elute DNA in

buffer.

Protocol Modifications: None

3.2.3.2 Liquid Nitrogen (LN) Enhanced Cell Lysis

Prior to this study, attempts to extract bacterial DNA from Lentikat’s biocatalysts

using the UltraClean® Microbial DNA Isolation Kit (MoBio laboratories, Carlsbad, CA

USA) consistently yielded very low concentrations of DNA (>1µg DNA/g sample). For this

reason, during preliminary tests to familiarize ourselves with the different DNA Isolation

protocols we performed liquid nitrogen (LN) enhanced cell lysis to determine if it would


23

increase DNA yields. As these tests were considered preliminary they were only carried out

in duplicate and only using sample D2. At the time, the resulting difference in yields between

LN treated biocatalysts and untreated biocatalysts were not considered significant and the

treatment was abandoned. Upon revisiting the data months later it became apparent that LN

treatment seemed to deliver higher yields and comparable purity across the board. Although

there is not enough data to draw definitive conclusions we have chosen to present it. Also,

LN treated samples extracted with the Powersoil® and QIAmp® kits were subjected to PCR-

DGGE for further phylogenetic comparison.

We developed the following protocol for LN treatment of polyvinyl alcohol

biocarriers:

Wear thick thermally protective gloves and full face masks!

1. Transfer approximately 10 g of fresh biocatalyst pellets to a 50 mL falcon tube and

remove as much water as possible using a 1 mL pipette

2. Transfer 2-3 g of dewatered pellets to a sterile smooth ceramic mortar

3. Pour enough liquid nitrogen into the mortar to cover to pellets, once it has evaporated

completely, repeat this process. After multiple repetitions (4-6) the rate of evaporation

will decrease significantly.

4. Once all liquid nitrogen has evaporated thoroughly grind the pellets with the pestle.

Note: They should be extremely brittle at this point, if they remain rubbery and resist

grinding into a wet powder repeat step 3.

5. Transfer ground up pellets to a clean 50 mL falcon tube and process immediately or

store at -20oC

3.2.4 DNA Yield and Purity

Isolated DNA was evaluated using a NanoDrop™ 1000 Spectrophotometer with the

software package ND-1000 Version 3.8 (Thermo Fisher Scientific Inc., Waltham, MA,

USA). The operating protocol was as follows:

1. Thaw frozen samples (if necessary), vortex for 3-5 seconds, and spin briefly to

remove drops from the tube walls and lid.

*Note: To calibrate the instrument follow steps 2-5 using only fresh elution buffer

corresponding to that in each sample and by selecting “Blank” instead of “Measure”.

Recalibration was performed after every 15-20 measurements.

2. Load 1.5-2 µL of sample onto the measurement pedestal


24

3. Close the sampling arm and initiate the measurement by selecting “Measure” on the

PC operating software

4. Upon completion of the measurement open the sample arm and clean the upper and

lower sampling pedestals with a clean lab wipe

5. If errors occur clean the upper and lower pedestals with a lab-wipe lightly moistened

with distilled water and repeat measurement.

This instrument measures isolated DNA concentration based upon the measured

absorbance at 260 nm (A260) using Beer’s Law, and has lower and upper detection limits for

dsDNA of 2 ng/μL and 3,700 ng/μL respectively. To estimate the purity of extracted nucleic

acid, the absorbencies at 280 nm (A280) and 230 nm (A230) were measured and the ratios of

averages between the A260 nm and A280 nm (A260/A280) and the A260 nm and A230 nm

(A260/A230) were calculated for each measurement. Samples with A260/A280 ratios

between 1.8 and 2.0 were presumed to be free of contamination (Dauphin et al., 2009).

Samples with mean A260/280 ratios below 1.8 were presumed to contain protein or other

contaminants, whereas samples with ratios above 2.0 were presumed to be contain RNA

contamination (Thermo Fisher Scientific, 2010; Mahmoudi et al., 2011; UMTK 322).

A260/A230 ratios are a supplementary measure of purity where values between 2-2.2 are

considered pure. Lower A260/A230 values often attributed to the presence of humic

compounds and/or residual phenol or other extraction reagents while higher values are often

attributed to errors in blanking the instrument (Mahmoudi et al., 2011; Thermo Fisher

Scientific, 2010).

Each sample of isolated DNA (6 samples>4 kits>Triplicate of each) was measured a

minimum of 5 times and the values for total DNA concentration, A260/A280, and

A260/A230 were statistically analyzed in order to characterize the products of each isolation.

3.2.5 PCR Amplification

The DNA extracts were used as a templates for Touchdown PCR amplification of the

variable V3 region of 16S rDNA (product 566 bp long + 40 bp GC clamp). The nucleotide

sequences of eubacterial-specific universal primers were as follows: 341F (5´-

CTACGGGAGGCAGCAG-3´) and 907R (5´-CCGTCAATTCMTTTGAGTTT-3´), (Schäfer

and Muyzer, 2001) to facilitate subsequent DGGE, we employed a forward primer, which


25

included a 40 bp GC clamp on its 5´ end; 341F-GC (5´-

CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCACGGGGGGGCCTACGGGAGGC

AGCG-3´), (Muyzer et al. 1993). Primers were purchased from Generi Biotech s.r.o. (Hradec

Králové, Czech Republic). PCR Grade water (Ref 03315932001) was purchased from

Hoffman-La Roche Ltd. (Basel, Switzerland). PCR reactions were carried out according to

the FastStart Taq DNA Polymerase, dNTPack (Hoffmann-La Roche Ltd., Basel, Switzerland)

manufacturer recommendations in 25 μL total volumes containing:

Table 3 Touchdown PCR Master Mix Formula

Reagent Quantity Concentration

Template DNA 1 μL Variable

PCR Grade Water 18.7 μL NA

PCR Buffer 2.5 μL 50 mM Tris/HCl, 10 mM KCl, 5 mM

(NH4)2, pH 8.3/25 °C

MgCl2 1.5 μL 1.5 mM

dNTP mix 0.5 μL 0.2 mM

Forward Primer 0.25 μL 0.5 µM

Reverse Primer 0.25 μL 0.5 µM

Fast Start Taq DNA Polymerase 0.3 μL 1U

The Following reagents were included in the FastStart Taq DNA Polymerase, dNTPack (Ref

04738357001) (Hoffmann-La Roche Ltd., Basel, Switzerland):

10x PCR buffer [500 mM Tris/HCl, 100 mM KCl, 50 mM (NH4)2, pH 8.3/25 °C]

25 mM MgCl2 solution

10 mM of the dNTP mix

5U/µL FastStart Taq DNA Polymerase

PCR cycling was carried out using a T-Personal Thermocycler (Biometra GmbH, Goettingen,

Germany) with the following conditions:

For initial denaturing heat to 95oC for 4 minutes

1. 95oC for 30 seconds (denaturing)

2. 65oC for 30 seconds (annealing)

3. 72oC for 2 minutes (elongation)

Annealing temperature was reduced by 1oC on each subsequent cycle until reaching

58 oC. After this point annealing was carried out at 58

oC for 21 cycles.

o **Template DNA removed from acrylamide gel following DGGE was subject

to 15 instead of 21 cycles**

During the final elongation step 72oC was maintained for 10 minutes.


26

PCR products were run on 2% agarose gel (m/v) stained with of Nancy-520 DNA Gel Stain

(Sigma-Aldrich Corporation, St. Louis, MO, USA) to evaluate successful amplification.

Results were visualized with the Major Science documentation system (Major Science,

Saratoga, CA, USA) provided with Canon G11 camera (Canon Inc., Tokyo, Japan). Samples

that failed to amplify were diluted 20x and 50x with PCR grade water and re-amplified.

3.2.6 DGGE

Bacterial PCR amplicons were run through a 6% polyacrylamide gel with a

denaturing gradient of 20% - 70% ureumformamide (80% denaturant solution contained

5.6 M urea and 32% (v/v) formamide). Visual quantification of PCR products was carried out

by comparing bands in the 2% agarose gel with the GeneRuler™ 100 bp Plus DNA Ladder

(Thermo Fisher Scientific Inc., Waltham, MA, USA). Between 10-13 μL of each sample

were mixed with 2 μL of 6x loading dye (Thermo Fisher Scientific Inc., Waltham, MA, USA)

and loaded into wells in the stacking gel. Electrophoresis was performed in a 17 L bath of 1x

Tris-acetate-EDTA buffer at 200 V and 60°C for 5 h using an INGENYphorU-2x2 DGGE

apparatus (Ingeny, Goes, The Netherlands).

The step by step manufacturer’s protocol for the INGENYphorU-2x2 DGGE

apparatus can be found online (as of 3/6/2013) at the link provided in Appendix 1. This

protocol was followed closely with only the following modifications:

Step 3.2: We applied a small amount of High-Vacuum Grease (Dow Corning®,

Midland, MI USA) to the U-shaped spacer in the glass plate sandwich to create a

better seal and prevent the gradient gel from leaking prior to polymerization.

Step 5.9: Electrophoresis was run for 5 hrs. at 200 V

Step 7.2: Stock solutions were prepared in 50 mL batches and were adapted from 9%

acrylamide in the manufacturer’s protocol to 6% acrylamide for our purposes.

The ureumformamide/acrylamide stock solutions and reagent mixtures employed in gel

casting (section 4 of the manufacturer’s protocol) are listed in Tables 3 and 4 below.


27

Table 4. Preparation of ureumformamide (UF)/6% acrylamide solutions employed in denaturing gradient

gels.

Reagent [Sigma Aldrich Serial #] 0% UF Acrylamide solution 80% UF Acrylamide solution

Urea [U6504] ---- 16.8 g

1% TAE, pH 8.3 [E5134] 1 mL 1 mL

40 % Bis-Acrylamide [A76168] 7.5 mL 7.5 mL

Formamide [F9037] ---- 16 mL

Molecular H2O Dilute to 50 mL Dilute to 50 mL

Table 5. DGGE Gel Casting Reagents

Reagent [Sigma Aldrich Serial#] Denaturant Concentration

0% (Stacking Gel) 20% 70%

0% UF solution 10 mL 18 mL 3 mL

80% UF solution ---- 6 mL 21 mL

99% Tetramethylethylenediamine

(TEMED) [T9281]

5 µL 8 µL 8 µL

20% Ammonium Persulfate (APS)

[A3678]

25 µL 50 µL 50 µL

The gel was subsequently stained with 10 mL of Sybr® Green I Nucleic Acid Gel Stain (Life

Technologies, Carlsbad, CA, USA) 1:10,000 solution and visualized with the Major Science

documentation system (Major Science, Saratoga, CA, USA) provided with Canon G11

camera (Canon Inc., Tokyo, Japan). Digital image cropping was carried out using GNU

Image Manipulation Program (GIMP) version 2.8 (GNU Development Team, Berkeley, CA

USA) and image analysis was carried out using Phoretix 1D Gel Analysis Software

(TotalLab Ltd., Newcastle, United Kingdom). DGGE profiles were analyzed separately for

each sample. The procedure for image analysis included:

1. Inversion

2. Lane Creation

3. Background Subtraction

4. Band Detection

5. Profile Deconvolution

6. Reference Line Calibration

7. Band Matching to Synthetic Lane

8. UPGMA Dendrogram and Similarity Index Generation

As stated above band matching was achieved through the generation of a synthetic lane

composed of all unique bands present across all lanes in each sample’s profile. The UPGMA

dendrograms generated compare the similarity between each lane in the profile while the

band sharing index (BSI), also called Dice similarity, represents each individual lane’s direct

similarity to the synthetic lane containing all bands in the profile.


28

3.3 Results

3.3.1 DNA Isolation and Purity

Table 6. Plzeň and Lochovice Activated Sludge Characteristics

Sample TSS VSS VSS% (VSS/TSS)

P1 (Plzeň) 11.9 g/L 9.4 g/L 85.3%

L1 (Lochovice) 5.7 g/L 4.2 g/L 73.8%

Upon visual inspection the Plzeň sludge was medium to dark brown in color and when

left undisturbed would rapidly and completely settle to the bottom of the vessel. In contrast

the Lochovice sludge was dark brown to black in color and when left undisturbed would

separate into a floating fraction and a settleable fraction. Table 5 shows that the Plzeň sludge

had more than double the TSS and VSS of the Lochovice sludge however both were

comparable in terms of VSS/TSS ratio at 85.3% and 73.8% respectively.

Table 7. Average cost, processing time, and mass of waste generated during a 9 sample DNA Isolation.

Isolation Kit Processing Time Waste (g) Recovery

Volume (µL)

Cost

(€/Sample) PowerSoil® 111 min 17 sec ± 12 sec. 123.75 ± 0.95 50* 3.2

QIAamp® 113 min 47 sec ± 9 min 44 sec 136.85 ±0.35 100* 5.1

Chemagic 111 min 28 sec ± 4 min 11 sec 97.44 ± 9.04 100 2.4

MasterPure™ 152 min 59 sec ± 5 min 29 sec 41.97 ± 6.77 35 1.2

*Half of manufacturer recommended volume.

Processing time and waste generation were calculated based upon the average of 2

separate 9 sample runs. Table 6 shows that processing times for all kits were very similar

except for MasterPure™, which took significantly longer. The fastest overall run was

achieved with the QIAmp® stool kit at 104 minutes and 3 seconds. The PowerSoil® kit was

the most consistent with only 12 seconds difference between the two separate run times.

Waste generation differed greatly between each kit. The greatest mass of wastes were

generated by the two spin filter kits, QIAmp® and PowerSoil®. Waste generation with the

Chemagic kit varied the greatest in between runs due in most part to employing techniques to

reduce the number of pipette tips used without compromising purity used during the second

run. Waste generated by the MasterPure™ kit was significantly lower than for any other kit.


29

Table 8. DNA yields (µg/g), absorbance 260 nm/280nm, and absorbance 260nm/230nm. [

]

PowerSoil® QIAamp®

Yield

µg/g

A260/A280 A260/A230 Yield

µg/g

A260/A280 A260/A230

N1 1.79

1.60

1.86

1.95

1.84

2.00

1.51

0.84

1.58

1.29

1.23

1.61

1.73

1.65

2.12

0.93

0.83

1.06

N2 6.23

3.79

6.56

1.91

1.89

1.95

1.86

1.53

2.04

9.95

7.84

10.57

1.99

1.95

2.05

1.89

1.14

1.95

D1 3.93

2.70

4.59

1.89

1.82

1.96

1.47

1.37

1.69

3.97

3.39

4.45

1.73

1.72

1.81

1.38

1.10

1.44

D2 7.37

6.52

8.53

1.93

1.90

1.96

1.82

1.78

1.87

6.87

6.56

7.26

1.91

1.84

1.95

1.59

1.50

1.73

D2 LN*

10.47

9.87

11.20

1.98

1.96

2.02

1.65

1.59

1.70

11.84

11.19

12.02

2.03

1.98

2.04

1.30

1.22

1.30

L1 24.17

19.54

26.22

1.93

1.92

1.95

1.99

1.97

2.04

15.23

14.05

15.72

1.91

1.85

1.95

1.25

0.80

1.30

P1 17.04

13.32

17.37

1.91

1.90

1.92

1.97

1.76

1.99

30.66

30.35

38.96

2.14

2.13

2.16

2.08

1.76

2.12

Chemagic MasterPure™ Yield

µg/g

A260/A280 A260/A230 Yield

µg/g

A260/A280 A260/A230

N1 67.69

58.17

173.53

1.68

1.39

1.72

1.11

0.90

1.34

35.45

28.82

95.83

1.85

1.81

2.20

1.23

1.16

1.92

N2 160.88

156.02

165.89

1.69

1.67

1.71

1.11

0.95

1.34

298.83

138.05

311.46

2.02

1.96

2.05

1.95

1.90

1.97

D1 299.53

283.46

353.88

1.80

1.77

1.82

1.08

1.03

1.10

236.98

203.72

288.90

1.93

1.92

1.95

1.64

1.60

1.71

D2 173.40

113.86

182.97

1.42

1.42

1.44

0.74

0.73

0.80

334.51

308.32

351.17

1.83

1.81

1.84

1.44

1.40

1.47

D2

LN* 276.73

269.27

290.58

1.59

1.58

1.59

0.75

0.73

0.76

385.02

279.55

436.48

1.79

1.76

1.85

1.36

1.33

1.45

L1 179.52

154.24

563.28

1.64

1.53

1.66

1.26

1.24

1.27

512.95

404.09

527.56

1.76

1.70

1.76

1.41

1.28

1.45

P1 221.70

170.19

230.56

1.64

1.63

1.67

1.21

1.18

1.23

557.83

493.85

623.77

1.85

1.82

1.86

1.57

1.53

1.61

* Values for liquid nitrogen treated samples represent duplicate rather than triplicate samples.


30

DNA yields varied significantly between sample origin, extraction kits, and even in

some case triplicate isolations. Due to variability in yield and purity ratios obtained between

triplicate isolations the assumption of normal distribution could not always be made and thus

median and interquartile range were determined to best describe the data in Table 7. A

common trend was that 2/3 triplicate isolations would return very similar results while the

third would differ greatly and cause either positive of negative skew. The extent and direction

of such skews can be observed by examining the Q1 and Q3 values provided in Table 7.

DNA yield varied significantly between both isolation kits and sample origin however

many trends in the data were clearly evident and are summarized below. Note that LN

treatment treated biocarriers are considered separately from all other analyses:

Liquid Nitrogen Treatment (D2-LN vs. D2)

o Sample D2-LN Yielded higher median amounts of DNA for all isolation kits

and Q1 values greater than the Q3 values of D2 for all kits except

MasterPure™

Sample N1 yielded the lowest amount of DNA for each kit across the board

Sample D1 yielded the next lowest amount of DNA for all kits except Chemagic for

which it was the highest.

Sample L1 yielded the highest amounts of DNA for the PowerSoil® kit and the

second highest amounts for both the QIAmp® stool kit and the MasterPure™ kit

Sample P1 yielded the highest amount of DNA for the QIAmp® stool kit and the

MasterPure™ kit and the second highest for the PowerSoil® kit

Samples N2 and D2 were in the middle of the pack for median DNA yield although

sample N2 yielded lower amounts of DNA from each kit except the QIAmp® stool

kit

The QIAmp® stool kit yielded median DNA concentrations of 15.23 – 30.66 µg/g for

activated sludge samples and below 9.95 µg/g for all biocarrier samples

The PowerSoil® kit yielded median DNA concentrations of 17.04 - 24.17 µg/g for

activated sludge samples and below 7.37 µg/g for all biocarrier samples


31

The Chemagic kit yielded median DNA concentrations of 179.52-221.70 µg/g for

activated sludge samples and 67.69 - 299.53 µg/g for all biocarrier samples with only

1 median yield below 160.88 µg/g

The MasterPure™ kit yielded median DNA concentrations of 512.95 - 557.83 µg/g

for activated sludge samples and 35.45 - 334.51 µg/g for all biocarrier samples with

only 1 median yield below 236.98 µg/g

DNA purity was assessed based upon two ratios, the absorbance at 260nm to the

absorbance at 280 nm (A260/A280) and the absorbance at 260 nm to the absorbance at

230 nm (A260/A230). A260/A280 values between 1.8–2.0 (considered pure) were achieved

by all kits for at least 1 sample. Median A260/A230 values of 2.0-2.2 (considered pure) were

achieved only in one instance thus this measure of purity was assessed based upon proximity

to this range. Data presented in Table 7 imply that purity is more highly correlated with

particular isolation kits rather than with sample origin. Purity results are summarized below:

Liquid Nitrogen Treatment D2-LN vs. D2

o The median values of A260/A280 for sample D2-LN were 0.03-0.12 higher for

each kit except MasterPure™ which was 0.04 lower. Only the PowerSoil® kit

achieved a pure A260/A280 ratio of 1.98 while others were 0.01 (MasterPure™),

0.03 (QIAmp®), and 0.21 (Chemagic) out of the range. Interestingly, median

A260/A230 values of LN treated biocarriers were lower by 0.17 (PowerSoil®),

0.2 (MasterPure™), 0.29 (QIAmp®) and 0.01 (Chemagic). It’s worth noting that

the despite the miniscule drop in A260/A230 for the Chemagic D2-LN, the

untreated D2 extract was already well below 1.0 and represented the worst purity

measurement obtained for any other extract.

The PowerSoil® kit achieved median pure A260/A280 ratios across the board and ranked

first with the lowest average inner quartile range (IQR) of 0.092. It also and had the

highest A230/A260 ratios for samples N1, D2, and L1 and second highest for D2 and P2.

Overall, all median A260/A230 ratios exceeded 1.46, four exceeded 1.8, and two exceed

1.95, however the average IQR of 0.33 ranked third out of the four kits.

The MasterPure™ kit achieved median pure A260/A280 ratios for 4/6 samples, was

within 0.04 of the desired range for the remaining two samples (N2 and L1) and ranked


32

third with an average IQR of 0.11. It also achieved the highest median A260/A230 ratios

for samples N2 and D1 and the second highest values for samples N1, D2 and L1.

Overall, all A260/230 ratios exceeded 1.22 with 3 exceeding 1.5, one equaling 1.95, and

ranked second with an average IQR of 0.21.

The Chemagic Kit only achieved a median pure A260/A280 ratio for sample D2 with the

remaining values between 0.11 – 0.38 below the desired range. Despite the comparatively

poor A260/A280 purity it did rank second with an average IQR of 0.098. In terms of

A230/A260 ratios it also yielded the lowest values across the board with no value

exceeding 1.26 and two values below 1.0 despite the top ranked (smallest) IQR of 0.18.

The QIAmp® stool kit achieved median pure A260/A280 ratios for 3/6 samples with the

remainder falling 0.07 – 0.14 above or below the desired range. The average A260/A280

IQR was larger than for any of the other kits at 0.15. In terms of A260/A230, this was the

only kit to yield a median ratio in the desired 2.0-2.2 range which was 2.08 for sample P1.

Overall however, it yielded median A260/A230 values ranking third lowest out of the

four kits for 4/6 samples, ranked second once, and even had one value below 1.0. The

average A260/A230 IQR was also worse than any other kit at 0.41.

3.3.2 PCR Amplification

Table 9. PCR amplification of 16S rDNA isolated from Lentikat Biocatalysts® and activated sludge

samples

Sample PowerSoil® QIAamp® Chemagic MasterPure™

N1 + + + +

N2 + + + -/+

D1 + + + +

D2 + + -/-/+ -/-/+

D2-LN +/- +/- NA NA

L1 + + + -/+

P1 + + + -/+ Note: + Indicates successful amplification of undiluted template DNA; -/+ Indicates failed

amplification of undiluted template DNA but successful amplification at 1:20 dilution; -/-/+ Indicates

failed amplification of undiluted and 1:20 dilution but successful amplification at 1:50 dilution.


33

Figure 2. 2% Agarose gel profiles of PCR products

A. PCR Products optimized for downstream DGGE. B. PCR Products optimized for downstream DGGE

C. PCR products of PowerSoil® and QIAmp® with full buffer D. PCR products of QIAmp® with full buffer

K= Concentrated (½ manufacturer recommended elution buffer); Q= QIAamp®; MB= (MoBIO)

PowerSoil®; CH= Chemagic; MP= MasterPure™; L= Ladder; NC= Negative control Hy= Sample

from partial nitrification SBR (Section 4.2.5). LN = indicates treatment with liquid nitrogen prior to

DNA extraction. Samples presented /20 or /50 (e.g. D2/50) indicate dilution ratios where applied. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 A L NC K-Q

N1 K-MB-N1

CH N1

MP N1

K-Q N2

K-MB N2

CH N2

MP N2/20

K-Q D1

K-MB D1

CH D1

MP D1

B L K-Q-

D2

K-MB

D2

CH

D2/50

MP

D2/50

K-Q

L1

K-MB

L1 CH

L1

MP

L1/20

K-Q

P1

K-MB

P1 CH

P1

MP

P1/20

K-MB

Hy

C L NC MB

N1

MB

N2

MB

D1

MB

D2

MB

L1

MB

P1

MB

D2-LN

MB-D2 LN/20

Q

N1

Q

N2

Q

D1

Q

D2

D L Q

L1

Q

P1

Q-D2

LN

Q-D2

LN/20

L

PCR Amplification was performed on one of the three DNA extracts from each

sample chosen based upon NanoDrop® data to best represent the median values for yield and

purity. In the case of incongruence between yield and purity values, the extract was selected

that best represented median purity.

Table 8 and Figure 2 Images A and B indicate that all kits isolated DNA that

successfully amplified via Touchdown PCR for all samples, but that dilution was necessary in

2 3 4 5 6 7 8 9 10 11 12 13 14 2 3 4 5

2 3 4 5 6 7 8 9 10 11 12 13 14 2 3 4 5 6 7 8 9 10 11 12 13 14


34

some cases. Table 8 and Figure 2 Images C and D indicate that DNA extracted from all

samples using the PowerSoil® and QIAmp® stool kits successfully amplified when the

manufacturers recommended volume of elution buffer was used. Unfortunately due to an

error in programming the thermocycler, images C and D depict an amplification that was cut

short by 6 cycles. However, these results are still interesting in that these images indicate that

the concentration of PCR products obtained from the QIAmp® stool kit extracts were lower

than those extracted with the PowerSoil® kit possibly indicating some PCR interfering

contaminants.

Table 8 and Figure 2 Images C and D also show that while undiluted extracts from

sample D2 treated with LN readily amplified, 1:20 dilutions with PCR grade water did not.

Note that both were tested as insurance to avoid running PCR again in the event that the

undiluted extracts would not amplify.

As indicated above and show in Table 8, the undiluted MasterPure™ extracts from

samples N2, D2, L1, and P1 as well as the Chemagic extract for sample D2 failed to amplify

despite successful positive control and other sample amplification (not shown). Another

round of PCR was then performed on these samples using a 1:20 dilution with PCR grade

water which resulted in successful amplification for all but two samples which amplified after

a final round using 1:50 dilution ratio.


35

3.3.3 DGGE

Figure 3. DGGE Profile of amplified V3 region of 16S rDNA from all samples. From left to right:

Samples L1, N1, D2, Reactor B (Ch. 4), D1, P1, N2 (Order: QIAmp®, PowerSoil®, Chemagic,

Masterpure™). *Note the image of Sample L1 was taken from a separate gel.*

Figure 3 shows the DGGE profiles from all samples and kits examined. For each

sample the order of extracts in each lane from left to right is QIAmp®, Powersoil®,

Chemagic, and Masterpure™. The only deviation from this ordering is for sample D2, where

LN treated biocarrier extracts from QIAmp® and Powersoil® were added adjacent to their

untreated analogues. Note that Q-D2-LN is marked with an X. Upon close investigation it

was determined that Q-D2-LN was in fact a mistakenly amplified extract of Sample D1.

Evidence of this mistake and justification of its omission are presented in Appendix 2.

Figure 4. UPGMA Dendrogram and BSI for Sample N1.

Figure 4 shows that for sample N1, all extracts shared 90% similarity with eachother. A total

of 15 distict bands were identified across all N1 extracts. Both Masterpure™ and Chemagen

represented 93% of total phylogenetic richness (BSI) in the synthetic lane while QIAmp®

represented the least at 85%.

BSI (%)

93

89

85

93

N1 N2 D1 D2 L1 P1 Reactor B

-Ch.4

X MB

LN


36

Figure 5. UPGMA Dendrogram and BSI for Sample N2.

Figure 5 shows that all profiles for sample N2 shared 87% similarity with eachother overall.

A total of 22 distinct bands were identified across all extracts. The Chemagic kit extract

represented the greatest amount of phylogenetic richness at 95% while the Masterpure™

extract represented the lowest at 90%.

Figure 6. UPGMA Dendrogram and BSI for Sample D1.

Figure 6 shows that all profiles for sample D1 shared 87% similarity with eachother overall.

A total of 17 distinct bands were identified across all D1 extracts. The QIAmp® stool kit

represented the greatest percentage of phylogenetic richness at 97% while the Powersoil® kit


Figure 7. UPGMA Dendrogram and BSI for Sample D2.

Figure 7 shows that that all extracts from sample D2 shared only 78% similarity with

eachother overall. A total of 20 distinct bands were identified across all D2 extracts. The

Chemagic and Masterpure™ kits represented the least amount of phylogentic richness at 71%

and 75% respectively, while the LN treated Powersoil® extract represented the most at 95%.

Interestingly the LN treated Powersoil® extract also shared only 91% similarity with its non-

LN treated analogue.

BSI (%)

90

93

95

93

BSI (%)

90

90

97

87

BSI (%)

75

71

95

86

80


37

Figure 8. UPGMA Dendrogram and BSI for Sample L1

Figure 8 shows that all extracts from sample L1 shared 84% similarity overall. A total of 25

distinct bands were identified across all L1 extracts. The QIAmp® and Masterpure™ extracts

represented 98% and 96% of total phylogentic richness respectively while the Powersoil®

extract represented the least at 81%.

Figure 9. UPGMA Dendrogram and BSI for Sample P1

Figure 9 shows that all extracts from sample P1 shared 93% similarity overall. A total of 21

distinct bands were identified across all extracts. The Masterpure™ kit extract represented the

greatest percentage of total phylogenetic richness at 98% while the Powersoil® kit


3.4 Discussion

3.4.1 Waste Generation, Processing Time, and Cost

The differences in waste generation between the kits can be attributed primarily to the

difference in techniques employed for DNA isolation. The Powersoil® and QIAmp® kits

were the most similar. The greatest contributing factor to QIAmp® producing the most waste

overall was the use of a combined total of 8x 1.5-2 mL microcentrifuge tubes and 2 mL spin

collection tubes as well as all the pipette tips necessary to make all transfers. Furthermore the

“InhibitEx” tablets add extra weight as well. In contrast the Powersoil® kit uses only 4x 2mL

microcentrifuge tubes plus 1x bead beating tube, however the dense plastic construction of

the bead beating tubes plus the beads themselves add significant mass to the waste

generation. While the Chemagic kit required only 2x microcentrifuge tubes its relatively high

BSI (%)

96

98

86

81

BSI (%)

98

95

89

95


38

mass of waste generated was due to weight of the magnetic beads and washing reagents

required but more importantly the very high number of pipette tips required. Lastly, the very

best kit in terms of waste generation was Masterpure™. This kit was quite efficient in terms

of pipette tip usage, volume of reagents used, and required only 2x microcentrifuge tubes.

The processing times for the Powersoil®, QIAmp®, and Chemagic kits were all very

similar. On the other hand, the Masterpure™ kit took significantly longer for a 9 sample run

due to 50 minutes of incubation time throughout the protocol with periodic pauses for

vortexing and 20 additional minutes of refrigerated centrifugation.

In terms of cost, the QIAmp® stool kit was the most expensive at €5.1 per sample

followed by the Powersoil® kit at €3.2 per sample. It is worth noting that the Powersoil® kit

was the only kit tested that included all necessary bead beating, microcentrifuge, and

collection tubes in this price. On the other hand, the Chemagic kit cost €2.4 per sample

excluding microcentrifuge tubes and RNAse. The Masterpure™ kit was by far the least

expensive at a cost of only €1.2 per sample however it cannot be purchased in quantities

fewer than 200 preps unlike Powersoil® (50 prep minimum), QIAmp® (50 preps), and

Chemagic (100 preps). Furthermore, in considering costs it’s important to take into account

the greater processing time required by the Masterpure™ kit. While the fixed cost per sample

is the lowest the additional costs in terms of salary of the technician performing the

extractions may significantly reduce this benefit especially when only running a small

number of samples.

3.4.2 DNA Yield

Our results indicate that for touchdown PCR and DGGE applications all kits yielded

sufficient quantities of DNA from all samples tested when we followed the manufacturer

protocols. Before discussing and comparing the extraction kits it’s worth noting a few trends

evident between the individual samples. As expected sample N1, yielded the lowest amount

of DNA across the board, yet all extracts amplified successfully. Also as expected, both

activated sludge samples, which contain biomass to total mass ratios higher than biocarriers,

yielded the greatest amounts of DNA for each kit except Chemagen. Lastly, the data implies

that LN treatment of biocarriers prior to DNA isolation was effective at increasing the

quantity of DNA yielded. This is a logical finding as the LN treatment thoroughly reduced


39

the biocarriers to a homogenous mass with characteristics similar to those activated sludge.

This serves to effectively eliminate any barriers that may have impeded lysis enzymes from

contacting the biomass most deeply embedding in the biocarrier material. Furthermore, it is

consistent with the findings of Alain et al. (2011) who obtained greater DNA yields from

deep sea sediments by integrating a similar cryogenic grinding step into their lysis procedure.

In regards to the extraction kits, we did reduce the amount of elution buffer for the

QIAmp® and Powersoil® extracts in order to obtain more reliable NanoDrop® readings,

however samples diluted to the manufacturer recommended volumes amplified in touchdown

PCR without impediment. In fact, the only real trouble we encountered was that a number of

Masterpure™ extracts had too high DNA concentrations for successful amplification without

dilution. Based upon the purity ratios obtained for nearly all of these samples it seems likely

that these failed amplifications were due to excessive DNA concentrations, however the

failure of the 1:20 dilution from MP-D2 most likely indicated the presence of PCR inhibitors.

Similarly, the undiluted Chemagen extract from sample D2 also failed to amplify and

although it may have been related to excessive DNA concentration, poor purity or a

combination of both factors seems very likely. Fortunately, in both of these cases the DNA

concentration was sufficient to withstand 1:50 dilution and still amplify, while the inhibitor’s

influence was overcome.

The Powersoil® and QIAmp® stool kits both yielded similarly low quantities of DNA

from all biocarrier samples and slightly higher amounts for activated sludge samples. Despite

DNA yields below 2 µg/g of sample N1, both kits produced extracts that amplified well.

While this was an encouraging finding it raised the question of whether or not such low

yields of DNA contain enough nucleic acid diversity to be representative of the total

phylogenetic diversity present in this sample. To answer this question we moved forward

with DGGE analysis which is discussed in a later section.

Based upon these results, the Masterpure™ kit appears to be the best suited to isolate

DNA for downstream applications that require very high concentrations.

3.4.3 Purity

Overall the Powersoil® kit performed the best in terms of DNA extract purity for both

A260/A280 and A260/A230 ratios. Not only did Powersoil® extracts meet the desired


40

A260/A280 purity standards for all samples but they also demonstrated the lowest variability

compared to all other kits. In terms of A260/A230 purity standards, Powersoil® extracts were

again among the best nearly across the board however they show greater variability in IQR

than Chemagic or Masterpure™ extracts in this measure.

The Masterpure™ kit performed nearly as well as the Powersoil® kit both in terms of

A260/A280 and A260/A230 ratios. While the IQR of A260/A280 demonstrated greater

variability than Powersoil® the opposite was true for A260/A230 IQR. Despite purity ratios

greater that many samples that did amplify, the Masterpure™ sample D2 extract failed to

amplify at 1:20 dilution, which would almost certainly rule out excessive template DNA as a

cause. Given that sample D2 was implemented in the remediation of groundwater

contaminated with sulfuric acid from uranium mining operations, it seems most likely that

multivalent cations such as heavy metals may be the culprit. Sample D2 was also extremely

dark in color and “dirty” in appearance so humic compounds shouldn’t be ruled out entirely

either. In any event, this case should be considered as a cautionary reminder that while the

purity ratios yielded on Nanodrop® may be positive indicators, they don’t guarantee that

concentrations of all impurities are below inhibitory levels.

The QIAmp® stool kit performed fairly well despite ranking 3rd

in terms of both

A260/A280 and A260/A230 ratios. What was troubling, however was that the QIAmp®

extracts demonstrated the greatest degree of variability of all kits tested. IQR values for

A260/A230 demonstrated a particularly high degree of inconsistency, which may or may not

be the reason for the difference in the reduced cycle PCR product concentrations compared to

the Powersoil® extracts show in Figure 2 images C and D.

The Chemagic kit performed the worst by far in terms of both measures of purity.

While the A260/A280 ratios indicated the presence of impurities for all but one sample they

were not so low as to cause major concern that the DNA would not be usable for touchdown

PCR amplification. On the other hand, the A260/A230 purity ratios were dismal across the

board but particularly for biocarrier samples. The extract purity values for sample D2,

discussed earlier, were the worst of any sample measured in this project.

Overall, each of the kits tested were able to successfully extract DNA of acceptable

yield and purity for downstream applications, in this case PCR-DGGE. The most consistently


41

Table 10. Mean BSI of all DGGE extracts

excluding MB-LN-D2

QIAmp® Stool Kit 91.3% σ = 7%

Powersoil® 87.5% σ = 4%

Chemagic 88.3% σ = 9%

Masterpure™ 90.3% σ = 8%

troubling purity results were obtained with the Chemagic kit, however a simple and obvious

modification to the protocol may logically improve these results. The manufacturer’s protocol

for this kit is designed for the extraction of DNA from laboratory bacterial cultures and thus

is performed entirely in a single microcentrifuge tube until elution. Logically, the assumption

is that any culture medium would easily be washed out and removed with other impurities

once the cells are lysed and DNA is bound to the magnetic beads, thus there is no pressing

necessity to use extra microcentrifuge tubes. In our case, when applying the Chemagic kit to

extract DNA from biocarriers, soils (data not included in analyses), and to a lesser extent

activated sludge, the sample residual remains in the microcentrifuge tube until the very last

step. It seems logical that briefly centrifuging the sample after the initial enzymatic lysis step

and transferring the lysate supernatant into a clean microcentrifuge tube could drastically

reduce the potential for contamination by humic acids, other complex organic compounds,

extraction reagents and/or washing buffers. It seems likely that the particularly poor purity of

Chemagic extracts may be due to impurities adsorbing to or infiltrating the pores of

biocarriers and being unintentionally carried through to subsequent steps in the isolation

process. This may explain the low A260/A230 ratios which are often attributed to phenol or

other extraction reagents remaining in the eluted DNA. This scenario is also supported by

fact that A260/A230 ratios were better for activated sludge samples, which would more easily

be removed along with reagents in washing steps than biocarriers that remain until the final

stage.

Based upon these results, it seems that all kits tested can be applied to Lentikat’s

Biocatalysts in order to extract DNA of sufficient purity for use in downstream Touchdown

PCR applications. Furthermore, these data indicate that the Powersoil® kit is the best suited

for isolating DNA for use in applications that require very high purity.

3.4.4 Phylogenetic Comparison of Extracts

The results from our DGGE analyses imply that

all kits performed comparably well in terms of

phylogenetic richness and similarity. Despite some

variability between the BSI values of different samples

Table 9 shows that overall the mean BSI values for all


42

kits were not significantly different. Furthermore, the similarity in banding patterns between

kits ranged from 78-93% with a mean of 86.5% which is highly encouraging considering the

results of Mahmoudi et al. (2011) who reported similarity of only 2-10% in a similar study

examining soil samples. Even the result of 78% overall similarity yielded for sample D2 was

deflated by the superior performance of the LN treated biocarrier extract. Overall, these

results demonstrate a far lower degree of extraction bias than had been expected.

When considered in the context of our results for the LN treatment, it seems that

suggesting superiority of one DNA isolation kit over another in terms of phylogenetic

comparison would be misguided. While the QIAmp® stool kit did demonstrate the greatest

consistency of high BSI values, the results for MB-LN-D2 suggest that obtaining the most

comprehensive phylogenetic richness from each sample may best be achieved through

optimizing cell lysis rather than kit selection. While DGGE does not provide comprehensive

phylogenetic richness as much as it is known for characterizing dominant community

members, the differences in the intensity of identical bands between extracts apparent in

Figure 3 suggests that dominant community members may be absent or less pronounced in

some extracts. This raises the question of whether or not any DNA extraction provides a true

representation of phylogenetic dominance within a community or just susceptibility to the

cell lysis techniques employed in DNA isolation. While this is an extremely bold conclusion

to draw from such highly limited data it is supported by the findings of Alain et al. (2011)

mentioned previously and Jiang et al. (2011) who found that integrating lysis techniques

drastically reduced DNA extraction biases from mangrove sediments. Interestingly, they also

noted that more intense lysis lead to a greater release of humic compounds and thus

decreased extract purity. A similar trend was apparent in both purity ratios for LN treatments

with all kits except Chemagic, which had already yielded the lowest purity of any sample D2

extract. The mistake that prevented Q-LN-D2 from being included in these analyses is even

more disappointing in light of the data yielded by MB-LN-D2.

Based upon the results of DGGE phylogenetic comparisons, no DNA isolation kit

performed significantly better or worse than any other. On the other hand, some extraction

biases were discovered in these analyses and though limited, the data for LN treatment and


43

available literature suggest that these biases may be minimized through continued evaluations

of enhanced lysis techniques.

3.5 Conclusions

All commercial DNA Isolation kits tested were compatible with Lentikat’s Biocatalysts

and yielded DNA of sufficient quantities and purities for downstream PCR-DGGE analyses.

The processing times for each kit were highly comparable except for the Masterpure™ DNA

purification kit which took significantly longer. On the other hand, the Masterpure™ DNA

purification kit produced less than half the mass of waste and cost only half as much per

sample than its next closest competitor which was the Chemagic DNA Bacteria kit in both

instances.

The Masterpure™ DNA purification kit and Chemagic DNA Bacteria kit both yielded the

highest median quantities of DNA in µg/g for all samples across the board. In terms of

isolated DNA purity, the Powersoil® DNA Isolation kit outperformed all others across the

board with the Masterpure™ DNA Purification kit ranking second.

All kits demonstrated a high degree of similarity and thus low degree of extraction bias in

DGGE phylogenetic comparisons. The QIAmp® DNA Stool kit performed the best in terms

of phylogenetic richness, however no other kit was significantly worse. Though the data is

limited, LN enhanced lysis of PVA biocarriers showed promise in mitigating extraction

biases and increasing the phylogenetic richness of isolated DNA. Given the high

phylogenetic similarity between all DNA isolation kit extracts, the potential of enhanced lysis

techniques for optimizing community characterization, DNA purity, and to a lesser extent

yield should be considered most heavily in establishing laboratory protocol for working with

Lentikat’s Biocatalysts. Therefore, the Powersoil® DNA isolation kit is recommended as the

best option from those examined for applications that require the highest degree of purity as

well as further investigations of enhanced cell lysis techniques. This holds true especially

when high DNA yield is not essential and sample mass is abundant. For applications where

higher DNA yield is required or sample mass is limited, the Masterpure™ DNA Purification

kit is recommended.

Ch 4. In Situ Detection of Immobilized Bacteria in Laboratory Scale Partial Nitrification SBRs

44

4.1 Objectives

The objectives of this investigation are to observe and characterize the effects of

hydroxylamine on NOB immobilized in Lentikat’s Nitrifying Biocatalysts in laboratory scale

partial nitrification SBRs. This is accomplished through a combination of water chemistry

analyses and fluorescence in situ hybridization and supplemented through PCR-DGGE and

16S rDNA sequencing.

Please note that the operation of these reactors and chemical analyses were performed

by Petr Kelbich, MSc. and Iva Johanidesová, MSc. and were the subject Mrs. Johanidesová’s

MSc. thesis titled "Evaluation of the possibility of maintaining partial nitrification using

immobilized microbial cultures” (Johanidesová, 2013).


45

4.2 Materials and Methods

4.2.1 Partial Nitrification Reactor Setup and Operation

Three SBRs, Labeled A, B and C, were set

up in 1.5L internal volume glass bioreactors

with external water circulation chambers in

the laboratory at ICT Prague on July 24,

2012 (Day 1). All three reactors were

situated on top of magnetic stir plates with

constant stirring, continuously aerated, and

linked in series to a Julabo F250 water bath

(Julabo GmbH., Seelbach, Germany) that

maintained constant temperature at 30oC

through continuous circulation. Each reactor

was loaded with 100 g (66.7 g/L) of wet

LentiKat’s nitrifying biocatalysts obtained following manufacture and subsequent cultivation

or “Grow out” on July 12, 2012.

During the startup phase (day 1-14), each reactor was drained and re-filled with 1L of

synthetic WW containing 50 mg N-NH4/L, 2.3 g/L KH2PO4, 2.9 g/L K2HPO4, and 0.5 g/L

NaHCO3 five days per week. Once steady state nitrification was achieved (day 15), the

concentration of N-NH4 was increased to 300 mg/L (600 mg/L prior to weekends) and

operation was again continued until steady state nitrification was reached (day 43). At this

point, Reactors A and B were subject to separate treatment regimes by dosing hydroxylamine

hydrochloride (NH2OH·HCl) obtained from Sigma-Aldrich Spol. s.r.o (Prague, Czech

Republic), while the operation of reactor C was maintained as a control until it suffered a

catastrophic mechanical failure on day 112. The treatments for Reactors A and B are detailed

in the following paragraphs and summarized in Table 10.

Beginning on day 43, Reactor A was dosed to a concentration of 0.5 mg NH2OH/L

once daily. Dosing was automated with a peristaltic pump that delivered the appropriate

quantity of hydroxylamine over a 15 minute time period. On day 73 the dose of

hydroxylamine was increased to 8 mg/L per day which was delivered in 1 mg aliquots over a

30 minute time period every 3 hours. This dosing was automated using a storage solution

Figure 10. Nitrifying SBRs in laboratory in ICT Prague.

Note: This image is of a subsequent experiment that

employed the same physical apparatus minus 1 reactor.

Photo Credit: Petr Kelbich


46

with a concentration of 200 mg NH2OH/L, the peristaltic pump, and a Kanlux Cyber TM-6

digital timer. On day 87, the thermostat unit critically failed and could not be immediately

replaced so the remainder of the investigation was carried out with the reactors operating at

room temperature (20-25oC). On day 112 the dose of hydroxylamine was increased to 16 mg

NH2OH/L per day delivered in 2 mg doses, every three hours, over a 15 minute time period.

On day 121, the dose of hydroxylamine was reduced to 4 mg/L per day delivered in 2 mg

doses, every 12 hours, over a 15 time period. The experiment was concluded on day 169 and

Reactor was shut down.

Beginning on day 43, Reactor B was dosed to a concentration of 5 mg NH2OH/L once

daily. Dosing was automated with the same apparatus and delivery time used for Reactor A.

On day 73, the dose of hydroxylamine was increased to 40 mg NH2OH/L per day which was

delivered in 5 mg aliquots over a 30 minute time period every 3 hours. This dosing was

automated using a storage solution with a concentration of 40g NH2OH/L, the peristaltic

pump, and a Kanlux Cyber TM-6 digital timer. On day 87, the thermostat unit critically failed

and could not be immediately replaced so the remainder of the investigation was carried out

with the reactor operating at room temperature (20-25oC). The dose of hydroxylamine was

also reduced on this date to 10 mg/L delivered in 1.25 mg aliquots every 3 hours over a 30

minute time period. On day 91, hydroxylamine dosing was ceased and ammonia nitrogen

concentration in the synthetic WW influent was reduced to 50 mg/L. The concentration of

ammonia in the reactor influent was increased to 75 mg/L on day 97, to 150 mg/L on day 99,

and back to 300 mg/L on day 107. While this experiment was concluded on day 169, Reactor

B remains operational as of June 3, 2013 as part of a separate experiment.

Beginning on day 174, the synthetic WW used over the previous 6 months was

replaced with the liquid fraction of post anaerobically digested sludge from Prague Municipal

WWTP (Prague, Czech Republic). This digestate was diluted in order to achieve an influent

N-NH4+

concentration of approximately 300 mg/L (600 mg/L prior to weekends). Operation

was again held steady until it was decided to reduce the weekend dose of N-NH4+ to 300 g/L

on day 284.


47

Table 11. Summary of treatments administered to Reactors A and B throughout their operations.

Day # Reactor A Reactor B

1 Startup 30oC.

1L/d [50 mg N-NH4/L, 2.3 g/L

KH2PO4, 2.9 g/L K2HPO4, and 0.5

g/L NaHCO3]

Startup 30oC.

1L/d [50 mg N-NH4/L, 2.3 g/L

KH2PO4, 2.9 g/L K2HPO4, and 0.5

g/L NaHCO3]

14 N-NH4 increased to 300 mg/L N-NH4 increased to 300 mg/L

43 NH2OH dosing 0.5 mg*d

1 dose delivered over 15 min period

NH2OH dosing 5 mg*d

1 dose delivered over 15 min period

73 NH2OH dosing 8 mg*d

1 dose every 3 hrs.; 30 min delivery

NH2OH dosing 40 mg*d


87 Temp from 30oC to Room Temp Temp from 30

oC to Room Temp

NH2OH dosing 10 mg*d


91 NH2OH dosing Terminated

N-NH4 decreased to 50 mg/L

97 N-NH4 increased to 75 mg/L






1 dose every 12 hrs.; 15 min

delivery

169 Experiment concluded and reactor

disassembled

Experiment concluded however

reactor operation maintained

174 Influent switched from Synthetic

WW to liquid digester effluent

diluted to ~300 mg/L N-NH4+

285 Weekend dose of N-NH4+ reduced

from 600 mg/L to 300 mg/L

Figure 11. Hydroxylamine dosing regimes for treatment reactors.


48

4.2.2 Chemical Analyses

The concentrations of inorganic nitrogen species in reactor effluents were analyzed on a

regular basis. Parameters assessed included NH4+, NO2

-, NO3

-, and NH2OH and were

determined using the following protocols:

4.2.2.1 Ammonia Nitrogen (NH4+)

This protocol was adapted from Standard Methods in the Examination of Water and

Wastewater 4500-NH3 (APHA, 1992).

Nessler Reagent Seignett Salt Solution

25 g HgI2

17.5 g KI

40 g NaOH (in 125 mL dH2O)

dH2O Fill to 250 mL

2.5 g KNaC4H4O6

50 mL dH2O

1. Load 5 mL of sample into a test tube

2. Add 100 µL of Seignett salt solution and 100 µL

3. Seal the test tube and mix thoroughly.

4. After 10 minutes measure the absorbance at 425 nm with a photoLab® 6100 VIS

spectrophotometer (WTW GmbH, Weilheim, DE)

5. Calculate the concentration of Ammonia using a calibration curve.

4.2.2.2 Nitrite Nitrogen (NO2-)

This protocol was adapted from Standard Methods in the Examination of Water and

Wastewater 4500-NO2- (APHA, 2012).

SANED Reagent

10 g Sulfanilamide

0.5 g N-(1-napthyl)-1,2-ethyendiamine-dichloride

25 mL H3PO4 [CONC]

dH2O to 250 mL

1. Load 5 mL of sample into a test tube

2. Add 125 µL of SANED reagent and 1.1 mL of dH2O

3. Seal the tube and mix thoroughly



5. Calculate the concentration of Ammonia using a calibration curve.


49

4.2.2.3 Nitrate Nitrogen (NO3-)

This protocol was derived from ISO 7890-1:1986 (ISO, 1986).

Amide-Sulfuric Acid Acid Mixture DMP

0.8 g amide-sulfuric acid

dH2O to 100 mL 2 mL concentrated H2SO4

2 mL concentrated H3PO4

0.24 g 2,6-dimethylphenol

200 mL glacial acetic acid

1. Load 0.5 mL of sample into a test tube

2. Add 50 µL of amide-sulfuric acid solution, 3.5 mL acid mixture, and 500 µL DMP

solution

3. Seal the tube and mix thoroughly



4.2.2.4 Hydroxylamine (NH2OH)

This Protocol was adapted from Frear and Burrell (1955).

8-Quinolinol Solution

o 1 g 8-Quinolinol

o 100 mL 99% Ethanol

10-4

M NADH Dehydrogenase

0.025 mM NH2OH

1M Sodium Carbonate 0.001 M Manganese Chloride

12% Trichloroacetic Acid 0.05 M PBS pH 6.8

1. Add 1 mL of Sample to a test tube

2. Add 1 mL of PBS, 800 µL of dH2O, and 200 µL of trichloroacetic acid

3. Add 1 mL of 8-quinolinol and mix gently

4. Add 1 mL of 1 M sodium carbonate solution and shake vigorously

5. Seal the tube and place in boiling water bath for 1 minute.

6. After cooling for 15 minutes measure the absorbance at 705 nm photoLab® 6100

VIS spectrophotometer (WTW GmbH, Weilheim, DE)

4.2.3 FISH

This internal protocol has been adapted from Amman (1995) in order to hybridize

nitrifying microorganisms immobilized within Lenticat biocatalysts. Hybridization probes

were purchased from Generi Biotech (Hradec Králové, Czech Republic) and were pre labeled

with fluorophores listed in Table 11. The stock probes arrived in varying concentrations and

were diluted with sterile distilled water to 50mM working solutions prior to hybridization.

To establish a baseline for comparison, biocatalysts were sampled from all reactors and

subsequently fixed for hybridization on day 30 of the experiment. At this stage all three

reactors were exhibiting similar performance in regards to inorganic nitrogen speciation.

Treatments were commenced on day 43 and Reactors A and B were then sampled again on

day 130. Reactor B was sampled again on day 203. Unfortunately Reactor C suffered a

critical vessel failure on day 111 before a second round of biocatalysts could be sampled and


50

fixed. All biocatalysts were fixed with paraformaldehyde (see section 4.2.3.2), stored at -

20oC, and hybridized in February 2013. For supplemental comparison, nitrifying biocatalyst

samples N1 and N2 (See Section 3.2.1) we also hybridized and imaged.

4.2.3.1 Reagents and Probes

1 x Phosphate-Buffer-Saline

(PBS)

3 x Phosphate-Buffer-Saline

(PBS)

4% Paraformaldehyde (PFA)-

PBS Solution

8 g NaCL

0.2 g KCl

1.44 g Na2HPO4

0.2 g NaH2PO4

1000 mL dH2O

pH 7

24 g NaCL

0.6 g KCl

g Na2HPO4

0.6 g NaH2PO4

pH 7

66 mL ddH2O ( 60oC)

4 g PFA

2-3 drops Conc. NaOH

34 mL 3 x PBS

pH 7-7.4 (HCl)

Tris-HCl Buffer (1 M) NaCl Stock (5 M) SDS-solution (10%)

15.8 g Tris/HCl

100 mL dH2O

pH 8

29.2 g NaCl

100 mL dH2O

10 g SDS

100 mL H2Obidest

TE Buffer EDTA (0.5 M) Ethanol

1.6 g (10 mM) Tris/HCl

0.37 g (1 mM) EDTA

(Na2)-EDTAxH2O

1000 mL dH2O

pH 7.2/ pH 8

18.6 g EDTA

100 mL dH2O

50%

80%

96%

100%

Hybridization Buffer (35% Formamide) Hybridization Buffer (40% Formamide)

360 mL NaCl (5M)

40 µL Tris/HCl buffer

800 µL Formamide

800 µL dH2O

2 µL SDS (10%)

360 mL NaCl (5M)


900 µL Formamide

700 µL dH2O

2 µL SDS (10%)

Washing Buffer (35% Formamide) Washing Buffer (40% Formamide)


460 µL NaCl (5 M)

500 µL EDTA (0.5 M)

50 µL SDS (10%)

Dilute to 50 mL with dH2O


700 µL NaCl (5 M)

500 µL EDTA (0.5 M)

50 µL SDS (10%)

Dilute to 50 mL with dH2O


51

Table 12. Hybridization probes employed. Table adapted from Nielsen et al. 2009.

Probes Target Formamide % Florophore Reference

NSO 1225 Betaproteobacterial

AOB

35 FITC Mobarry et al. 1996

NSO 190 Betaproteobacterial

AOB

35 FITC Mobarry et al. 1996

NTSPA 712 Phylum

Nitrospirae

35 Cy3 Daims et al. 2001

cNTSPA 712

(Competitor)

NTSPA 712 non

target organisms

35 None Daims et al. 2001

NTSPA 662 Genus Nitrospira 35 Cy3 Daims et al. 2001

cNTSPA 662

(Competitor)

NTSPA 662 non

target organisms

35 None Daims et al. 2001

NIT 3 Genus Nitrobacter 40 Cy3 Wagner et al. 1996

cNIT 3

(Competitor)

NIT 3 non target

organisms

40 none Wagner et al. 1996

4.2.3.2 Fixation with Paraformaldehyde

1. Collect samples of 2-10 pellets from each reactor and place in separate 2 mL eppendorf

tubes

2. Add 1.5 mL of 4% PFA-PBS solution, then add tap water until final volume reaches 2 mL

3. Mix by shaking, then refrigerate at 4oC for 4 hrs.

4. Remove and discard supernatant in PFA waste container.

5. Add 2 mL of 1xPBS washing buffer and invert several times to mix.

6. Repeat washing step 2 additional times.

7. Add 500 µL of 1xPBS and 500 µL of 100% Ethanol (not denatured)

8. Vortex to mix and store in the freezer at -20oC

4.2.3.3 Hybridization

1. Transfer a minimum of 2 pellets in a sterile 2 mL eppendorf tube

2. Dehydrate samples through successive ethanol rinses. First add enough 50% Ethanol to

cover the pellets (300 µL) and allow them to rest for 3 minutes before removing and

discarding the supernatant.

3. Repeat step 2 using 80% Ethanol

4. Repeat step 3 using 96-99% Ethanol

5. Remove the supernatant and incubate the sample tubes at 46oC for 20 minutes (or until

dry)

6. Add 50 µL of Hybridization buffer (Probe specific formula: See FISH Reagent list above)

7. Add Fluorescent probe and equal volume of complementary competitor probe (if needed)

and tap gently to mix.

a. 6.25 µL for Cy 3 labeled Probes

b. 8 µL for FITC labeled Probes

8. Incubate at 46oC for 3 hours.

9. Remove from incubator and discard the supernatant into PFA waste.

10. Add 2 mL of Washing Buffer and place in hot water bath at 48oC for 20 minutes


52

11. Remove from hot water bath and discard the supernatant.

12. Add 2 mL of cold Distilled H2O, shake gently and then discard the supernatant

13. Add 200-300 µL of TE Buffer (enough to cover the pellets in liquid)

14. Store refrigerated at 4oC for no more than 2 weeks.

4.2.3.4 Imaging

Fluorescence detection was carried out with an Olympus BX51-RFAA microscope,

images were captured with an Olympus MX10 CCD Camera using the fluorescence imaging

software CELLF all products of Olympus Corporation (Tokyo, Japan). Image manipulations

including: automatic white balance, manual brightness and contrast adjustment, and auto

color sharpening were carried out using GNU Image Manipulation Program (GIMP)

version 2.8 (GNU Development Team, Berkeley, CA USA).

4.2.4 Live/Dead Staining

Live/dead staining was carried out on biocatalysts sampled from Reactor B on day

224 with the LIVE/DEAD® BacLight™ Bacterial Viability kit, for microscopy and

quantitative assays (L7012, Lot 950626) (Life Technologies Corporation, Carlsbad, CA)

using the following protocol:

1. Lay Fresh PVA pellet on a sterile glass slide.

2. Apply 3 µL of a 1:1 mixture of 3.34 mM SYTO 9 dye and 20 mM Propium Iodide

(included in kit) to the pellet.

3. Incubate in the dark at room temperature for 15 minutes

4. Observe using fluorescent microscope (see section 4.2.3.4 for microscope

specifications)

4.2.5 DNA Isolation, PCR, DGGE, and Sequencing

DNA Isolation was carried out on biocatalysts sampled from Reactor B on day 274

with the PowerSoil® DNA Isolation kit (MoBio Laboratories Inc., Carlsbad, CA, USA) using

the manufacturer recommended protocol modified as detailed in Section 3.2.2 of this report.

Touchdown PCR was carried out using the protocol detailed in Section 3.2.4 prior to DGGE.

DGGE was carried out with the protocol detailed in Section 3.2.5. Bands were cut from the

acrylamide gel, soaked in PCR grade water (Hoffmann-La Roche Ltd., Basel, Swiss) for 36

hours, and subsequently re-amplified using Touchdown PCR. During this final round of PCR

amplification, the protocol detailed in section 3.2.4 was shortened by 6 cycles and an

analogue forward primer without GC-Clamp was employed. These samples were then sent to


53

the Institute for Inherited Metabolic Disorders in the First Faculty of Medicine and General

Hospital at Charles University in Prague, Czech Republic for purification and sequencing.

4.3 Results

4.3.1 FISH Images, Inorganic Nitrogen Speciation, and Live/Dead Staining

Figure 12. Newly manufactured nitrification biocatalyst (see sample N1 in Chapter 3) and a used

nitrification biocatalyst (see sample N2 in Chapter 3). AOB are hybridized with FITC (green) labeled

NSO 190 & NSO 1225 while NOB are hybridized with Cy3 (red) labele

A. New Nitrification Biocatalyst (N1) B. New Nitrification Biocatalyst (N1)

C. New Nitrification Biocatalyst (N1) D. Used Nitrification Biocatalyst (N2)


54

E. Used Nitrification Biocatalyst (N2) F. Used Nitrification Biocatalyst (N2): Cy3 only

Figure 12 Images A-F depict both new and used nitrification biocarriers hybridized

with AOB and NOB targeted probes. These images clearly show vibrant coloration consistent

with bacteria containing high concentrations of rRNA thus implying a high level of activity.

Nearly all of these images demonstrate that the distribution of AOB and NOB in Lentikats

Biocatalysts is not uniform but highly variable between individual pellets. Regardless, they

also demonstrate that each biocarrier contains substantial populations of both functional

groups. Images E and F both depict the same area of the same pellet. Image E, shows both

AOB and NOB while Image F shows only NOB. These images demonstrate the high

percentage of the biocarriers hydrogel matrix that is saturated with bacterial biomass while

also validating our hybridization technique. A close examination of both images reveals that

the bright green clusters of AOB in image E are merely blank spaces in image F, however

this is not always clear because of the 3 dimensional nature of the biocarriers.

Figure 13. Nitrogen speciation in control reactor effluent throughout operational life.

Throughout its operational life, Reactor C demonstrated complete nitrification of

ammonium to nitrate thus indicating the activity of both AOB and NOB. These results are


55

corroborated by Figure 14 Images A and B which clearly show that members of both

functional groups were present in biocarriers on day 30 of the experiment. In fact, the spike in

nitrite concentration to 45.07 mg/L on day 15 represents the only data point for this reactor

where the nitrite concentration exceeded 0.13 mg/L. Unfortunately, the next data point was

taken on day 49, so it is unclear exactly when the concentration of nitrite returned below

0.1 mg/L although it is likely to have occurred prior to sampling on day 30.

Table 12 below shows the residual concentrations of NH2OH in each reactor. Because

the protocol for measuring NH2OH was not discovered until this experiment was under way

there are only four data points from which to establish a baseline molar concentration of

hydroxylamine under control conditions in Reactor C. While this is too few data points to

draw definitive conclusions from, it is worth noting that the concentration never exceeded

6.05 µM NH2OH.

Figure 14. FISH images from control reactor (Reactor C) biocatalysts sampled on Day 30 of the

experiment. AOB are hybridized with FITC (green) labeled NSO 190 & NSO 1225 while NOB are

hybridized with Cy3 (red) labeled NIT3 and NTSPA 662 & NTSPA 712.

A. Day 30 B. Day 30

Table 13. Residual hydroxylamine concentrations measured in each reactor in µM

Day # Reactor A Reactor B Reactor C

87 18.16 2,664.24 6.05

91 15.13 605.51 6.05

93 51.47 3.03 3.94

107 4.54 3.33 4.54

112 13.02 3.02 NA

114 58.74 3.63 NA

122 363.31 6.05 NA

128 86.29 5.75 NA

132 4.48 5.44 NA


56

Figure 15 (below) shows the inorganic nitrogen speciation and NH2OH dosing in

Reactor A throughout its operational life. Note a similar spike in nitrite concentration

corresponding with the initial increase of influent ammonia to 300 mg/L on day 14.

Unfortunately, it is again unclear when this concentration returned to baseline levels of below

0.13 mg/L but it is presumed to be prior to sampling of biocarriers for hybridization on

day 30. It is also evident that the thermostat failure on day 87 corresponded with a massive

spike in ammonia to 624 mg/L and crash in nitrate production down to 10.8 mg/L. We

believe that the complete inhibition of both AOB and NOB evident at this time was due

mostly to temperature shock, but could also be a result of the combination of NH2OH dosing

and the high weekend dose (600 mg/L) of influent ammonia as well. The residual

concentration of NH2OH of 18.16 µM is above the expected baseline concentration however

it is still well below the 250 µM concentration reported by Kindaichi et al (2004) to inhibit

NOB in suspended cultures. Regardless, nitrification did recover significantly over the

following 12 days before the dose of NH2OH was increased to 16 mg/L*d on day 99, after

which nitrate concentrations fell steadily reaching their minimum levels of 5.3 mg/L on

day 128. Interestingly, this comes following the day 122 measurement in which the highest

concentration of residual NH2OH of 363.06 µM was measured in Reactor A. Perhaps most

interesting is that the dose of NH2OH was reduced on day 120, just prior to nitrate levels

bottoming out. As residual NH2OH then decreased back towards “background levels” of

below 10 µM despite continued dosing, nitrate production began a steady increase doubling

approximately every 20 days through the end the reactors operation, reaching 22.8 mg/L on

day 169.


57

Figure 15. Inorganic nitrogen speciation and hydroxylamine dosing in Reactor A throughout its

operational life. Note: All concentrations given in mg/L, Hydroxylamine (NH2OH) dose corresponds with

the secondary Y-axis (right), while all other parameters correspond

Figure 16 images A and B demostrate that biocatalyst in Reactor A did contain

significant populations of both AOB and NOB at the outset of this experiment which is

corroboarted by the nitrogen speciation in Figure 15 discussed previously. Images D and F in

particular show that significant populations of NOB still existed in Reacor A biocarriers

approximately 2 weeks after the peak inhibition of nitratation. The corresponding nitrogen

speciation implies that nitratation was in recovery and was likely to be between

7.5-10.5 mg/L*d at this point in time. One drawback of FISH is that dead cells containing

rRNA may still hybridize and thus we are unable to determine what percentage of the NOB

present were actually alive and viable.


58

Figure 16. FISH images of biocatalysts sampled from treatment Reactor A. AOB are hybridized with

FITC (green) labeled NSO 190 & NSO 1225 while NOB are hybridized with Cy3 (red) labeled NIT3 and

NTSPA 662 & NTSPA 712.

A. Day 30 B. Day 30

C. Day 141 D. Day 141: Cy3 Only

E. Day 141 F. Day 141


59

Figure 17. Inorganic nitrogen speciation and hydroxylamine dosing in Reactor B throughout its

operational life. Note: All concentrations given in mg/L, Hydroxylamine (NH2OH) dose corresponds with

the secondary Y-axis (right), while all other parameters correspond

Figure 17 shows that Reactor B exhibited nearly identical nitrogen speciation to

Reactors A and C prior to NH2OH dosing. The presence of NOB in Reactor B biocarriers is

also confirmed by Figure 18 Images A and B. Similar to Reactor A, Reactor B showed

inhibition of both AOB and NOB corresponding with the thermostat failure on day 87,

however in Reactor B this inhibition was more comprehensive and furthermore it coincided

with the highest dosing levels and highest residual concentrations of NH2OH. The amount of

residual NH2OH measured in Reactor B on day 87 was 88 mg/L or 2,664.24 µM which is just

over 10 times the 250 µM reported by Kindaichi et al. (2004) to inhibit NOB and 1.3 times

the 2000 µM they reported to inhibit AOB. Hydroxylamine dosing was then reduced by ¾

and subsequently completely ceased 4 days later. Interestingly, day 87 represented the lowest

measured level of nitrate in the reactor of 4.01 mg/L and these levels did not exceed 8.2 mg/L

until day 164, after which they hovered between 1-28 mg/L until commencing a steady

increase on day 274. Overall the data shows that nitrite levels exceeded nitrate levels in

Reactor B for as many as 182 days.

Figure 18 Images C-F show that despite a high degree of inhibition over the previous

54 days, NOB remained present in fairly large numbers in Reactor B biocarriers. These

images also show that the conditions inside of the reactor caused a high degree of mechanical

stress and breakdown of the PVA biocarriers, which is supported to a greater extent by


60

images G and H from day 203 and the finding on day 169 that the weight of biocarriers in

Reactor B had been reduced from 100g to 50 g over the first period of the experiment. Images

G-K from day 203, go further than images C-F by showing a seemingly large population of

NOB present in Reactor B despite over 116 days of very high nitratation inhibition. The

rebound of nitrate production in Reactor B, albeit slow, lends credibility to the accuracy of

these images however Live/Dead staining was performed to investigate whether or not the

NOB were in fact still alive or merely inactive in regards to nitrite oxidation.

Figure 18. FISH images of biocatalysts sampled from treatment Reactor B. AOB are hybridized with

FITC (green) labeled NSO 190 & NSO 1225 while NOB are hybridized with Cy3 (red) labeled NIT3 and

NTSPA 662 & NTSPA 712.

A. Day 30 B. Day 30

C. Day 141 D. Day 141


61

E. Day 141 F. Day 141: Cy3 Only

G. Day 203 H. Day 203: Cy3 Only

J. Day 203 K. Day 203


62

Figure 19. Live/Dead Images of biocarriers sampled from Reactor B on Day 224, approximately 137 days

after inhibition of nitritation. Live bacteria appear green in color while dead bacteria appear red/orange.

A. Day 224 B. Day 224

Figure 19 Images A and B from day 224 indicate that it is indeed

possible that a significant portion of the bacteria within the biocarriers

were dead. Unfortunately with neither the mechanism of hydroxylamine

inhibition of nitratation or the ability to target only dead NOB it is not

possible to identify the population of dead bacteria. Still the fact remains

that nitratation in Reactor B had been severely restricted for over

137 days prior to Live/Dead staining and thus NOB would somehow

have to sustain their maintenance energy in order to survive this long a

period. Thus, it seems plausible that a majority of the dead bacteria in

these images are NOB although it cannot be stated with certainty.

4.3.2 DGGE and Sequencing

Figure 20 shows the DGGE profile of amplified 16S rDNA

extracted from Reactor B on day 274. The numbers in the image

correspond with the labels given to each band that was successfully

removed from the gel, reamplified, and sent for sequiencing. The results

from sequencing were obtained on May 29, 2013 and were inconclusive

due to very high background signals most likely caused by excessive

template DNA concentration due to noteable evaporation during PCR.

Alternative possibilites that should not be ruled out include

contamination, non-specific binding of primers, and UV damage.

Samples will be purified and resent for sequencing (if possible) and we

hope to present the results at the defense of this thesis on June 20, 2013.

Figure 20. DGGE

Profile of amplified

16S rDNA extracted

from Reactor B

biocarriers on Day

274.


63

4.4 Discussion

4.4.1 Inhibition of Nitratation

The mechanism through which hydroxylamine inhibits nitratation has not been well

characterized using modern techniques. Xu et al. (2012) cite a study by Deturk et al. (1958)

stating that hydroxylamine prevents the induction of the enzyme Nxr and affects protein

synthesis, however there do not seem to be any more recent follow up studies that definitively

prove or disprove this theory. Regardless, Xu et al. (2012) cites Hao et al. (1994) and Van der

Star et al. (2008) in stating that the inhibition of NOB by hydroxylamine is irreversible. This

is a critical assumption in assessing the performance of the partial nitrification reactors in this

experiment.

Kindaichi et al (2004) established that maintaining a hydroxylamine concentration of

250 µM in an RBC reactor fed with synthetic WW was sufficient to completely inhibit

nitratation. Xu et al. (2012) employed a slightly higher concentration of 300 µM NH2OH

dosed every 2 days to completely inhibit nitratation in an aerobic granule SBR. During their

experiment, Xu et al. (2012) ceased hydroxylamine dosing after complete nitritation had been

sustained and after 16 days nitratation began to exceed nitritation. This was attributed to the

presence and subsequent proliferation of NOB deeply embedded within aerobic granules

(confirmed by FISH) and thus shielded from exposure to and/or the effects of hydroxylamine

as well as washout. FISH images indicate that this population was extremely small which

would explain why the amount of nitrate produced throughout hydroxylamine dosing was

insignificant. The existence of such microenvironments free from sufficient hydroxylamine

concentrations to inhibit NOB is feasible considering that AOB consume hydroxylamine,

thus a reduced concentration deep within aerobic granules or suspended flocs is logical and

supported by the findings of Han et al. (2012).

Excluding the reactor crash on day 87, the inhibition of NOB in Reactor A peaked

after 37 days of hydroxylamine dosing at 242 µM followed by 10 days at 484 µM. After this

period the daily doses were dropped to 121 µM which is reported Hao et al. (1994) to be

sufficient to sustain established nitritation over nitratation in submerged aerobic filters.

Interestingly, Xu et al. (2012) performed batch tests which indicated that 150 µM was not

sufficient to suppress nitratation in an aerobic granule SBR. While nitrate production in

Reactor A did steadily increase (doubling every 20 days) through the end of the experiment,

nitrite production followed a seemingly proportional increase while ammonium decreased


64

over that same period. This may simply be a result of the reactor’s biota recovering from the

nearly continuous shocks evident in Figure 16 that occurred between days 60-125. These data

may therefore indicate a scenario similar to that described by Xu et al. (2012) whereby NOB

more deeply embedded in biocarriers were shielded from the peak hydroxylamine doses. As

conditions stabilized in the reactor, the may have been able to acclimate and continue

nitratation activity. It could also imply that the subsequent decrease in the daily dose to

121 µM was insufficient to prevent unaffected NOB from starting to grow outwards hence

the steady increase in nitratation.

Unfortunately, it seems unlikely that Reactor A had reached stable state equilibrium

with regards to nitritation performance at the conclusion of the experiment on day 169, thus it

is not appropriate to conclude whether or not sustained nitritation/nitratation was achieved.

Regardless, on the final day of operation, nitrate production was 23% as high as nitrite

production while Xu et al. (2012) reported < 1% in their reactors. If this did represent a stable

state without proliferation of NOB it could mean that NOB immobilized in Lentikat’s

Biocatalysts were in fact more resilient towards hydroxylamine inhibition than those in

aerobic granules or RBC biofilms. FISH images presented in Figure 16 show that NOB were

present in comparable quantities before and 14 days after peak hydroxylamine induced

inhibition.

The peak of nitratation inhibition in Reactor B occurred following 23 days of

hydroxylamine dosing at 1,211 µM. On the day of the thermostat failure and subsequent

reactor crash, the residual hydroxylamine concentration in Reactor B was high enough to

inhibit even AOB activity. Interestingly, nitritation recovered and reached more or less stable

performance 34 days later, which it sustained for an additional 48 days until the conclusion of

the synthetic WW portion of experiment on day 169. Despite the fact that hydroxylamine

dosing was ceased on day 91, nitratation remained severely inhibited over the final 78 days

preceding the switch from synthetic WW to digestate.. Following steady sate nitritation,

nitratation jumped as high as 19% of nitritation for one measurement but spent most of the

time from day 121-169 around 5% and ended on day 169 at 4.7%. Even after Reactor B was

switched to diluted digester effluent, nitratation did not exceed nitritation until a total of 182

days had passed since peak inhibition began.

The diffusion of hydroxylamine into the biocarriers may logically be a critical factor

in the difference between nitratation inhibition observed both Reactors A and B. The higher


65

concentration of hydroxylamine reached in Reactor B should have been accompanied with

higher concentrations diffusing deeper within biocarriers. It is possible that the only NOB

spared from exposure to inhibitory concentrations of hydroxylamine were so deeply

embedded in the biocarrier’s rigid hydrogel matrix and crowded by other organisms that

despite having ample substrate, the lack of space to reproduce and expand prevented them

from proliferating soon after dosing was ceased. With inhibitory concentrations of

hydroxylamine not diffusing as far into biocarriers in Reactor A, less deeply embedded NOB

as well as a greater proportion of the population overall may have avoided the full effects of

inhibition, which might explain the more steady recovery of nitratation observed.

The results do not clearly demonstrate threshold hydroxylamine concentrations

needed to achieve long term stable nitritation over nitratation using Lentikat’s Nitrifying

Biocatalysts. For this reason a cost analysis of large scale hydroxylamine dosed SBR’s for

partial nitrification would not be appropriate. The results do however show that the short term

high doses in Reactor B were far more effective at achieving a greater degree of long term

NOB inhibition than the lower peak dose combined with chronic low doses in Reactor A.

4.4.2 In situ detection and characterization of NOB community

The results of fluorescence in situ hybridization from both Reactors A and B indicate

that despite successful inhibition of nitratation, NOB washout was not significant. This is

most strongly demonstrated by Figure 18 Images C-K which show that significant

populations of NOB were present in Reactor B biocarriers both 54 and 116 days after peak

inhibition. Given the extremely low level of nitratation over this period, these finding imply

that one or more of the following scenarios was likely occurring: 1) the signal represented

non-specific binding of oligonucleotide probes, 2) the signal represented dead NOB cells

containing rRNA and thus remained susceptible to hybridization, 3) NOB were in fact present

and active and had switched to alternative metabolic pathways for survival 4) the signal

represented NOB that were alive and continued to oxidize nitrite to some degree. In the

absence of results from rDNA sequencing, it is impossible to determine exactly which species

of NOB were present following hydroxylamine treatments. Furthermore the DGGE results in

Figure 3 indicate that even freshly manufactured biocarriers contain more than the mere two

nitrifying organisms advertised, thus this discussion will consider the possibility that a

broader array of NOB genera were potentially present.


66

The non-specific binding of the oligonucleotide hybridization probes seems unlikely.

According to Nielsen et al (2009), the probes employed in this study are among the most

thoroughly tested and reliable available for use in in situ identification of NOB. The

hybridization of dead NOB cells on the other hand is likely occurring to at least some extent

particularly in Figure 18 Images C-F. At this stage in the reactors operation, it was still being

fed with synthetic wastewater, thus there should not have been any seeding of heterotrophic

organisms that might degrade dead cell material. Following the switch from synthetic WW to

anaerobic digester effluent, it is more likely that such organisms would be present in the

reactor to remove dead biomass, however the structure of the biocarriers may still have

impeded heteretrophic organisms from accessing and clearing out all dead NOB. While the

Live/Dead stains shown in Figure 19 were performed following the switch in reactor feed,

they indicate that a potentially significant portion of biomass present was dead. While it’s

unclear if this dead biomass is on the surface of the biocarrier, within the biocarrier, or even

how deep within the biocarrier it may be, it is possible, although not confirmable, that a

significant proportion are NOB. Given the fact that nitratation was not 100% inhibited

however, this potential scenario alone cannot account for all NOB present.

The results of chemical analyses clearly indicate that some NOB remained actively

performing nitratation in both reactors following peak inhibition. Based upon the extremely

low production of nitrate in Reactor B during this period however, it’s likely that only very

deeply embedded NOB continued nitratation, thus it’s possible that only a fraction if any of

the NOB shown in FISH images fit this description. An interesting alternative is that some

proportion of NOB inhibited by hydroxylamine were able to shift to an alternative substrate

for survival. The arguments in favor of this scenario include the well documented metabolic

flexibility of Nitrobacter which are known to metabolize simple organic compounds like

pyruvate or reduce nitrite to nitric oxide under anaerobic conditions (Deni et al., 2004; Ahlers

et al., 1990). Given that the reactor was maintained under constant aeration it is unlikely that

anaerobic nitrite reduction was occurring. On the other hand, the ability of Nitrobacter to

metabolize simple organic compounds may have been significant to their survival and may

explain spikes in activity just prior to, and especially in the months following the switch from

synthetic WW to digester effluent.

Deni et al. (2004) note that Nitrobacter populations were more active in soils

containing organic metabolites from diesel fuel degraded by other organisms compared to


67

populations in uncontaminated soils, implying that they survive better under mixotrophic

conditions. While Xu et al. (2012) found that most NOB were washed out of aerobic sludge

granules following hydroxylamine inhibition, it is possible that the rigid matrix of biocarriers

and lack of heterotrophic competition allowed them to maintain a significant presence despite

conditions in which they would normally be out competed due to retarded growth. A major

problem with this hypothesis is that the COD of synthetic WW was insignificant and thus the

only source of organic substrate should have come from dead biomass within the reactor.

Given the numerous shocks experience by Reactor B it is likely that dead biomass was

available, however the theoretical lack of heterotrophs needed to metabolize complex

organics into the simple compounds usable by Nitrobacter casts doubt on this theory.

Another detractor from this theory is that if the NOB present were alive but starving due to

substrate limitation a decrease in rRNA content would be expected and would result in a drop

in FISH signal strength while cells that were suddenly killed by shock could retain higher

levels of rRNA if not lysed (Hawkins et al., 2011). Unfortunately, it is not clear from our

FISH analyses whether or not such a drop in signal strength occurred.

The plausibility of this alternative substrate scenario increases substantially following

the switch from synthetic WW to anaerobic digestate. The first reason for this is that the

digestate theoretically contained a higher COD concentration and a diverse array of

microorganisms not present in the synthetic WW, although there is no data to show this. In

addition to the boost in simple organic compounds directly useable by Nitrobacter, the

addition of heterotrophic organisms to the reactor likely increased the degradation rate of

dead biomass already present in/on biocarriers. This could not only free up space for NOB

expansion but could also carve channels that link deeply embedded NOB performing

nitratation to such newly cleared spaces opening the door for proliferation. The 5 independent

and unsustained spikes in nitrate concentration in Reactor B between days 150-250 could be

explained by spikes in the availability of simple COD compounds to the limited population of

Nitrobacter performing nitratation and in turn those NOB surviving solely on COD. The

eventual sustained increase in nitratation following day 254 could be indicative of either

seeding of fresh NOB from digestate or the proliferation of endemic NOB freed from the

depths of biocarriers by biologically created channels, the mechanical breakdown of the

biocarriers themselves, or a combination of both. It’s difficult to conclude which is more

likely considering that the presence of NOB in digestate is rare but occasionally happens


68

(Regueiro et al., 2012) and mechanical breakdown of biocarriers should have manifested

itself sooner. If neither is the case, the most likely alternative would be that the effects of

hydroxylamine are in fact reversible over time, possibly following reproduction which most

NOB subjected to hydroxylamine inhibition likely don’t get the opportunity to undergo prior

to washout. In essence, this may be the first study in which a long term investigation of the

effects of hydroxylamine on affected NOB has been possible because they do not appear to

washout from biocarriers unlike RBC biofilms and aerobic granules.

4.5 Chapter 4 Conclusions

These results demonstrate that up to 180 days of significant nitritation inhibition is

achievable after treating Lentikat’s Nitrifying Biocatalysts with only short term highly

concentrated doses of hydroxylamine. On the other hand, even when subjected to lower doses

comparable to those used in studies by Kindaichi et al (2004) and Xu et al (2012), nitratation

recovery by immobilized NOB proceeded more slowly than in RBC biofilms or aerobic

granules respectively.

These results also indicate that despite a high degree of hydroxylamine induced

nitratation inhibition, immobilized NOB were not significantly washed out of Lentikat’s

Nitrifying Biocatalysts. This finding may present a unique opportunity to investigate the long

term effects of hydroxylamine on NOB as well as characterizing the mechanisms through

which inhibition and potential nitratation recovery occur.

NOB detected by FISH within biocatalysts following inhibition are believed to

include: a small fraction performing limited nitratation, a potentially significant proportion of

dead biomass, an unknown fraction that switched to alternative organic substrates for survival,

and in one reactor an unknown fraction that may have been seeded from anaerobic digester

effluent.

Finally, the eventual recovery of nitratation observed in Reactor A was likely due to

biota acclimating to system shocks and less likely, but potentially, the proliferation of

endemic NOB unaffected by peak hydroxylamine doses. The recovery of nitratation in

Reactor B is most likely due to seeding of NOB from digester effluent, the proliferation of

endemic NOB unaffected by peak hydroxylamine inhibition, and/or the reversal of

hydroxylamine effects over time.

Ch 5. Summary of Conclusions

69

5.1 Genomic DNA Isolation and Amplification from Bacteria Immobilized in

Poly Vinyl Alcohol Biocarriers

All commercial DNA Isolation kits tested were compatible with Lentikat’s Biocatalysts

and yielded DNA of sufficient quantities and purities for downstream PCR-DGGE analyses.

The processing times for each kit were highly comparable except for the Masterpure™ DNA

purification kit which took significantly longer. On the other hand, the Masterpure™ DNA

purification kit produced less than half the mass of waste and cost only half as much per

sample as its next closest competitor which was the Chemagic DNA Bacteria kit in both

instances.

The Masterpure™ DNA purification kit and Chemagic DNA Bacteria kit both yielded the

highest median quantities of DNA in µg/g for all samples across the board. In terms of

isolated DNA purity, the Powersoil® DNA Isolation kit outperformed all others across the

board with the Masterpure™ DNA Purification kit ranking second.

All kits demonstrated a high degree of similarity and thus low degree of extraction bias in

DGGE phylogenetic comparisons. The QIAmp® DNA Stool kit performed the best in terms

of phylogenetic richness, however no other kit was significantly worse. Though the data is

limited, LN enhanced lysis of PVA biocarriers showed promise in mitigating extraction

biases and increasing the phylogenetic richness of isolated DNA. Given the high

phylogenetic similarity between all DNA isolation kit extracts, the potential of enhanced lysis

techniques for optimizing community characterization, DNA purity, and to a lesser extent

yield should be considered most heavily in establishing laboratory protocol for working with

Lentikat’s Biocatalysts. Therefore, the Powersoil® DNA isolation kit is recommended as the

best option from those examined for applications that require the highest degree of purity as

well as further investigations of enhanced cell lysis techniques. This holds true especially

when high DNA yield is not essential and sample mass is abundant. For applications where

higher DNA yield is required or sample mass is limited, the Masterpure™ DNA Purification

kit is recommended.

5.2 In Situ Detection of Immobilized Bacteria in Laboratory Scale Partial

Nitrification SBRs

The investigation of laboratory partial nitrification SBRs demonstrated that up to 180

days of significant nitritation inhibition is achievable after treating Lentikat’s Nitrifying

Biocatalysts with only short term highly concentrated doses of hydroxylamine. On the other

hand, even when subjected to lower chronic doses comparable to those used in studies by

Ch 5. Summary of Conclusions

70

Kindaichi et al (2004) and Xu et al (2012), nitratation recovery by immobilized NOB

proceeded more slowly than in RBC biofilms or aerobic granules respectively.

Results also indicate that despite a high degree of hydroxylamine induced nitratation

inhibition, immobilized NOB were not significantly washed out of Lentikat’s Nitrifying

Biocatalysts. This finding may present a unique opportunity to investigate the long term

effects of hydroxylamine on NOB as well as characterizing the mechanisms through which

inhibition and potential nitratation recovery occur.

NOB detected by FISH within biocatalysts following inhibition are believed to include: a

small fraction performing limited nitratation, a potentially significant proportion of dead

biomass, an unknown fraction that switched to alternative organic substrates for survival, and

in one reactor an unknown fraction that may have been seeded from anaerobic digester

effluent.

Finally, the eventual recovery of nitratation observed in Reactor A was likely due to biota

acclimating to system shocks and less likely, but potentially, the proliferation of endemic

NOB unaffected by peak hydroxylamine doses. The recovery of nitratation in Reactor B is

most likely due to seeding of NOB from digester effluent, the proliferation of endemic NOB

unaffected by peak hydroxylamine inhibition, and/or the reversal of hydroxylamine effects

over time.

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Appendix 1. Links to Manufacturer Protocols Online

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Manufacturers protocols used in this report can be found at the following links as of June 3, 2013:

1. QIAmp® DNA Stool Kit

(http://www.qiagen.com/Products/Catalog/Sample-Technologies/DNA-Sample-

Technologies/Genomic-DNA/QIAamp-DNA-Stool-Mini-Kit#resources)

2. PowerSoil® DNA Isolation Kit

(http://www.mobio.com/images/custom/file/protocol/12888.pdf)

3. Chemagic DNA Bacteria Kit

(http://www.chemagen.com/fileadmin/downloads/chemagic_DNA_Bacteria_Kit.pdf)

4. MasterPure™ DNA Purification Kit

(http://www.epibio.com/docs/default-source/protocols/masterpure-dna-purification-

kit.pdf?sfvrsn=4)

5. INGENYphorU-2x2 DGGE Apparatus

(http://www.ingeny.com/Manuals_files/INGENYphorU%20manual.pdf)

http://www.qiagen.com/Products/Catalog/Sample-Technologies/DNA-Sample-Technologies/Genomic-DNA/QIAamp-DNA-Stool-Mini-Kit#resources

http://www.qiagen.com/Products/Catalog/Sample-Technologies/DNA-Sample-Technologies/Genomic-DNA/QIAamp-DNA-Stool-Mini-Kit#resources

http://www.mobio.com/images/custom/file/protocol/12888.pdf

http://www.chemagen.com/fileadmin/downloads/chemagic_DNA_Bacteria_Kit.pdf

http://www.epibio.com/docs/default-source/protocols/masterpure-dna-purification-kit.pdf?sfvrsn=4

http://www.epibio.com/docs/default-source/protocols/masterpure-dna-purification-kit.pdf?sfvrsn=4

http://www.ingeny.com/Manuals_files/INGENYphorU%20manual.pdf

Appendix 2. Omission of Q-D2-LN Justification

80

Figure 21. DGGE Profiles for Sample D2 and Sample D1. Extract Q-D2-LN is outlined in Red.

The first indication that Q-D2-LN was mislabeled was a visual inspection of the DGGE profiles

shown in Figure 21 above. The banding pattern clearly appears more closely related to the profiles for

Sample D1 than for Sample D2. There is some similarity between the Q-D2-LN profile and the other

Sample D2 profiles, however this can be explained by the fact that both are denitrification biocarriers

that theoretically contained the same mixture of immobilized organisms at the time of manufacture.

Figure 22. UPGMA Dendrogram of Sample D2 Including Q-D2-LN

Figure 22 shows that Q-D2-LN shared only 30% similarity with all other Sample D2 extracts which

shared at least 70% similarity with each other. Furthermore, Q-D2-LN shared only 50% similarity

represented only 50% phylogenetic richness with the synthetic lane comprised of all bands detected

across Sample D2. The combination of these factors gave me a high degree of certainty that Sample

Q-D2-LN had been mislabeled and that omitting it from the analyses presented in Chapter 3 was

justified.

BSI (%) 60 50 76 80 76 50

D2 D1

genomic dna extraction and detection of bacteria ... · immobilized in polyvinyl alcohol host...

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