This thesis was elaborated and defended at Ghent University within the framework of the European Erasmus Mundus Programme

“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) 

  

                  

Metal‐based engineered nanoparticles in treatment wetlands: interactions with aquatic macrophytes and impact on 

the performance of microbial communities    

Host University: 

 Department  of  Applied  Analytical  and  Physical  Chemistry  

 Yi Xiao 

Promoter:          Co‐promoter:  

Prof. Gijs Du Laing, Ph.D.          Zhuan‐xi Luo, Ph.D.  

Tutors: 

  Frederik Van Koetsem, M.Sc.  Mark Button, Ph.D. 

2011 ‐ 2013

Metal-based engineered nanoparticles in

treatment wetlands: interactions with aquatic

macrophytes and impact on the performance of

microbial communities

Yi Xiao

i

Certification

This is an unpublished M.Sc. thesis and is not prepared for further distribution. The author and the

promoter give the permission to use this thesis for consultation and to copy parts of it for personal

use. Every other use is subject to copyright laws, more specifically the source must be extensively

specified when using results from this dissertation.

Ghent University, August 2013

The Promoter The Author

Prof. Gijs Du Laing, Ph.D.

Yi Xiao

 

 

Dissertation online access release

I hereby authorize the IMETE secretariat to make this dissertation available online on the IMETE

and/or Ghent University website

The Author

Yi Xiao

ii

Acknowledgements

I would like to show my sincere gratitude to my promoter, Prof. Gijs Du Laing for giving me the

chance to work in the Laboratory of Analytical Chemistry and Applied Ecochemistry (Ecochem) as

a master’s student, and also for his valuable comments and suggestions during the whole thesis

work.

I would like to give my sincere thanks to my tutor, Frederik Van Koetsem for his excellent

supervision, friendly assistance and valuable recommendations towards my thesis work. He was

always available for my questions and generously gave his time and vast knowledge to my

questions, leading me into the right direction in successfully completing my thesis work.

I would also like to express my thanks to my co-promoter Zhuan-xi Luo and my co-tutor Mark

Button for their kind help with my thesis work, especially Mark Button. He was so kind to share

many documents with me that were necessary and helpful to my thesis.

I would like to acknowledge the kind assistance of the staff members at the Laboratory of

Analytical Chemistry and Applied Ecochemistry, Joachim, Ria, David, Roseline, Katty and

Hannele Auvinen. Also, I want to take this opportunity to express my thanks to the staff at the

Laboratory of Microbial Ecology and Technology (LabMET) for their kindly guidance and for

providing me with the necessary equipment for my lab work. Also I want to thank Prof. Nico Boon

for providing biogenic Ag NPs for my experiment.

I also wish to express my sincere thanks to the European Union and the board of the IMETE

program for selecting me as a grant student, giving me the chance to study a promising field within

an international atmosphere. I also want to thank the staff members of the IMETE program for their

unreserved help whenever I needed it, during my whole studying period.

Last, but not least, it is my pleasure to thank my parents and my lovely friends for their unreserved

support in my life.

iii

iv

Abstract

Metal-based engineered nanoparticles (ENPs) are already being used on a large scale. This may

lead to the possibility for these nanoparticles (NPs) to enter aquatic systems. However, the fate and

behavior of these ENPs in aquatic environments have not yet been fully studied and the potential

impact these ENPs may have on aquatic plants and microorganisms has been overlooked.

Revealing the interactions between the NPs and aquatic macrophytes, as well as their impact on

microbial communities can help to provide valuable information, which can also be used in the

design of wetlands constructed to treat NPs containing wastewater.

Therefore, batch experiments were set up to study possible interactions of CeO2 NPs and Ag NPs

with a commonly occurring aquatic macrophyte. Dose-response experiments were performed by

spiking different concentrations of CeO2 NPs and Ag NPs, or their corresponding ions as control

into tap water containing Elodea canadensis. Total metals concentration in the water phase, before

and after cultivation, as well as total metal contents in the plants were determined. Additionally,

total nitrogen, total phosphorus and chlorophyll a, b and c contents in the plants were also measured.

In a second experiment, six different extraction reagents were tested for their capacity to desorb

NPs attached to the surface of the plants. In a next experiment, five kinds of water (Milli-Q, tap

water, and the surface waters Mostbeek, Grote Geul and Coupure), differing in physiochemical

characteristics, were used to cultivate the plants for 3 days in presence of either CeO2 NPs, Ag NPs,

or their corresponding ions. This was done to assess whether water composition can affect the

removal efficiency of NPs. In a final set of experiments, mesocosms mimicking a wetland substrate

were set up to examine the potential impact of Ag NPs to microbial communities in wetlands. PVP

coated, citrate coated, and biogenic Ag NPs, as well as Ag+ ions were added to simulated

wastewater used to feed the mesocosms. Community-level physiological profiling (CLPP) was

used to study the impact of the different types of NPs on the microbial community within the

interstitial water and the biofilm growing on the wetland substrate.

v

The study showed that both CeO2 NPs and Ag NPs can stimulate growth of the plants at low

concentrations but inhibit their growth at high concentrations, and generally, NPs are less toxic to

plants than their corresponding ionic forms. Most of the removed NPs were strongly sorbed to the

plants or taken up by the plants, as they cannot anymore be easily extracted using different

extractants after being brought into contact with the plants during one hour. Moreover, water

composition was confirmed to play a role in NPs’ removal processes. PVP coated and citrate

coated Ag NPs have less impact on microbial communities, while biogenic Ag NPs and Ag+ were

more toxic. The results also showed that after previous exposure, microbial communities were

more robust and appeared to have built up some kind of resistance to the disturbance of the

environment with NPs.

Keywords: Engineered nanoparticles (ENPs), aquatic macrophytes, Elodea canadensis, wetland

microbial communities, CeO2 nanoparticles, Ag nanoparticles.

vi

Table of contents

Acknowledgements ................................................................................................................... ii

Abstract .................................................................................................................................... iv

List of figures ........................................................................................................................... ix

List of tables ............................................................................................................................ xii

List of abbreviations ................................................................................................................ xv

Chapter 1: Introduction ............................................................................................................. 1

1.1. Background of the study ............................................................................................ 1

1.2. Potential problems ...................................................................................................... 1

1.3. Objectives ................................................................................................................... 2

Chapter 2: Literature review ...................................................................................................... 3

2.1. Classification of NPs .................................................................................................. 3

2.1.1. Natural NPs ........................................................................................................ 4

2.1.2. Anthropogenic NPs ............................................................................................. 5

2.2. Environmental impact of ENPs .................................................................................. 6

2.2.1. Properties and toxicity of silver NPs .................................................................. 7

2.2.2. Properties and toxicity of CeO2 NPs .................................................................. 8

2.2.3. Metallic NPs and their ionic metal forms ........................................................... 9

2.3. Aquatic macrophytes .................................................................................................. 9

2.3.1. Aquatic macrophytes - Elodea canadensis ....................................................... 10

2.3.2. Characterization of plant performance ............................................................. 11

2.4. Microbial communities and their characterization ................................................... 12

2.4.1. CLPP using BIOLOGTM plates ......................................................................... 13

2.4.2. Denaturing gradient gel electrophoresis (DGGE) ............................................ 15

2.4.3. Fluorescent in situ hybridization (FISH) .......................................................... 16

Chapter 3: Materials and Methods .......................................................................................... 18

3.1. Introduction .............................................................................................................. 18

3.2. Dose Experiment ...................................................................................................... 19

vii

3.2.1. Experimental Setup ........................................................................................... 19

3.2.2. Water analysis................................................................................................... 20

3.2.3. Plants sampling and characterization ................................................................ 21

3.3. Desorption experiment ............................................................................................. 24

3.4. Water composition experiment ................................................................................ 24

3.4.1. Experimental setup ........................................................................................... 24

3.4.2. Water composition analysis .............................................................................. 25

3.5. CLPP experiment ..................................................................................................... 27

Chapter 4: Results ................................................................................................................... 30

4.1. Dose - response experiments .................................................................................... 30

4.1.1. Elodea canadensis plants exposed to CeO2 NPs and Ce3+ ............................... 30

4.1.2. Elodea canadensis plants exposed to Ag NPs and Ag+ .................................... 33

4.2. Desorption experiments ........................................................................................... 36

4.2.1. CeO2 NPs and Ce ions ...................................................................................... 36

4.2.2. Ag NPs and Ag ions ......................................................................................... 37

4.3. Impact of water composition on NPs’ uptake by E. canadensis .............................. 38

4.3.1. Characterization of different (surface) water samples ...................................... 38

4.3.2. Elodea canadensis plants exposed to CeO2 NPs and Ce3+ ............................... 41

4.3.3. Elodea canadensis plants exposed to Ag NPs and Ag ions .............................. 45

4.4. Wetland mesocosm experiment ............................................................................... 48

4.4.1. Selection of time point for further data analysis ............................................... 48

4.4.2. Response of wetland microbial communities on different doses of Ag NPs .... 48

4.4.3. Carbon-Source Utilization Patterns (CSUPs) ................................................... 50

Chapter 5: Discussion .............................................................................................................. 54

5.1. Toxicity of NPs and bulk ions on E. canadensis ...................................................... 54

5.1.1. Toxicity of NPs and bulk ions on E. canadensis .............................................. 54

5.1.2. Uptake and adsorption of plants ....................................................................... 57

5.1.3. Toxicity of NPs and bulk ions on E. canadensis in different water bodies ...... 58

5.2. Mass balance calculations ........................................................................................ 61

viii

5.2.1. Mass balance in tap water ................................................................................. 61

5.2.2. Mass balance in different aquatic environments .............................................. 62

5.3. Effects of Ag NPs and Ag+ on microbial communities ............................................ 63

Chapter 6: Conclusions and recommendations ....................................................................... 66

6.1. Conclusions .............................................................................................................. 66

6.2. Recommendations for further research .................................................................... 67

Chapter 7: Reference ............................................................................................................... 68

ix

List of figures

(Figure) No. (Figure) Title Page

Figure 2- 1 Illustration of Elodea Canadensis (Source:

http://www.evi.com/q/facts_about__elodea_canadensis) ................................................. 10 

Figure 2- 2 Model of BIOLOGTM ECO microplates (Source: http://www.techno-path.co.uk) ........ 13 

Figure 2- 3 Different types of FISH Probes (Source: McNeil and Ried, 2000, Cambridge University

Press) .................................................................................................................................. 17 

Figure 3- 1 Cultivation of Elodea canadensis in water tanks with 10% Hoagland’s solution ……..18 

Figure 3- 2 Setup of the Dose experiment ........................................................................................... 20 

Figure 3- 3 Setup of mesocosms experiment ....................................................................................... 28 

Figure 4- 1 Relationship between theoretical concentration and actual concentration of Ce in

solutions spiked with CeO2 NPs (above) and Ce3+ (below) and containing E. canadensis

plants (0h: before introducing plants to the solutions; 72h: after cultivating plants for 72

hours) (Mean±SD, n=3) ……………………………………………………………...32 

Figure 4- 2 Relationship between theoretical concentration and actual concentration of Ag in

solutions spiked with Ag NPs (above) and Ag+ (below) and containing E. canadensis

plants (0h: before introducing plants to the solutions; 72h: after cultivating plants for 72

hours) (Mean±SD, n=3) .................................................................................................... 35 

Figure 4- 3 Contents of Ce extracted from E. canadensis plant material using different extractants

(Milli-Q water, 0.5M HNO3, 0.02M EDTA, 0.05 mM PVP, 0.05 mM CMC and 0.05

mM Dextran-70); the plants were previously soaked for 1h (Conc. At T1(1h)) in

solutions containing (right) CeO2NPs and (left) Ce3+ (Mean±SD, n=3) ......................... 37 

Figure 4- 4 Contents of Ag extracted from E. canadensis plant material using different extractants

(Milli-Q, 0.5 M HNO3, 0.02 M EDTA, 0.05 mM PVP, 0.05 mM CMC and 0.05 mM

Dextran-70); the plants were previously soaked for 1h (Conc. At T1(1h)) in solutions

containing (right) Ag NPs and (left) Ag+ ions (Mean±SD, n=3) .................................... 38 

Figure 4- 5 Evolution of concentration of Ce in different waters (Tap water, 10% Hoagland’s

solution, Mostbeek water, Grote Geul water, Coupure water) spiked with Ce3+ (above)

x

or CeO2 NPs (below) when exposing E. canadensis plants to these waters (Mean±SD,

n=3) .................................................................................................................................... 44 

Figure 4- 6 Evolution of concentration of Ag in different waters (Tap water, 10% Hoagland’s

solution, Mostbeek water, Grote Geul water, Coupure water) spiked with Ag+ (above)

or Ag NPs (below) when exposing E. canadensis plants to these waters (Mean±SD,

n=3) .................................................................................................................................... 47 

Figure 4- 7 Summary of the microbial development (A=AWCD, B= number of absorbance values

over 2) over a monitoring period of 115 h after inoculation of a Biolog plate of a

wetland mesocosm (from interstitial wastewater) containing 0.1 mg/L Ag supplied as

Biogenic Ag NPs, Citrate Ag NPs, PVP Ag NPs and Ag+ with a blank (no Ag) as

control group. ..................................................................................................................... 48 

Figure 4- 8 Response (AWCD) of microbial communities of interstitial water of a wetland

mesocosm on different concentrations (0.5 mg/L, 1 mg/L, 2 mg/L and 5 mg/L) of (A)

Citrate-Ag NPs, (B) PVP-Ag NPs, (C) Biogenic-Ag NPs and (D) Ag+ after 65 h of

exposure (the control group was not exposed to silver when cultivating the microbial

community in wastewater prior to exposure) (Mean±SD, n=3) ...................................... 50 

Figure 4- 9 PCA using carbon-source utilization data (CSUPs) of a control mesocosm (not exposed

to Ag) and mesocosms exposed to citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs and

Ag+ at 0.1 mg/L; mesocosms from the interstitial water were set up in triplicate (A-C).

Output generated using XLSTAT 2013. .......................................................................... 51 

Figure 4- 10 CLPP of interstitial water from wetland mesocosms exposed to 0.5 mg/L, 1 mg/L, 2

mg/L and 5 mg/L citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs and Ag+; the biofilms

were previously exposed to 0.1 mg/L citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs,

Ag+ and no Ag as control group. Output generated using XLSTAT 2013. .................... 52 

Figure 4- 11 PCA using carbon-source utilization data (CSUPs) of interstitial water from control

mesocosms exposed for 65 h to citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs and

Ag+ at concentrations of 0.5 mg/L, 1 mg/L, 2 mg/L and 5 mg/L Ag . The microbial

communities were not exposed to Ag during preceding growth in the mesocosm.

Output generated using XLSTAT 2013. .......................................................................... 53 

xi

Figure 5- 1 Dose-response of total nitrogen (a: exposed to CeO2 NPs and Ce3+, b: exposed to Ag

NPs and Ag+) and total phosphorus (c: exposed to CeO2 NPs and Ce3+, d: exposed to

Ag NPs and Ag+) in E. canadensis over 72 hours’ cultivation........................................ 55 

Figure 5- 2 Dose-response of chlorophyll a (Chl a) (a: exposed to CeO2 NPs and Ce3+, b: exposed

to Ag NPs and Ag+), chlorophyll b (Chl b) (c: exposed to CeO2 NPs and Ce3+, d:

exposed to Ag NPs and Ag+) and chlorophyll c (Chl c) (e: exposed to CeO2 NPs and

Ce3+, f: exposed to Ag NPs and Ag+) in E. canadensis after 72hours’ exposure (at 0.1

mg/L Ag+, 0.5 mg/L Ag NPs, 0.5 mg/L Ag+, 1 mg/L Ag NPs and 1 mg/L Ag+, Chl c

content was under detection limit of the equipment) ....................................................... 57 

Figure 5- 3 Effect of water composition on the content of total nitrogen, total phosphorus,

chlorophyll a and chlorophyll b in E. canadensis plants upon exposure to 1 mg/L CeO2

NPs, 1 mg/L Ce3+, 0.1 mg/L Ag NPs, 0.1 mg/L Ag+ (Control group: without any

exposure of NPs and ions) ................................................................................................. 59 

xii

List of tables

(Table) No. (Table) Title Page

Table 2- 1 Classification of nanomaterials (Adapted from: Nowack and Bucheli, 2007) ................... 4 

Table 2- 2 Application of metal-based NPs (Adapted from: Bernhardt, Colman et al. 2010) ............ 6 

Table 3- 1 Composition of modified 10% Hoagland's medium…………………………………..19 

Table 3- 2 Composition of Scheel I, II, III solutions used for phosphorus determination ................. 23 

Table 4- 1 Mass change/initial weight (%) of E. canadensis plants after exposure to different

concentrations (0 mg/L, 0.5 mg/L, 1 mg/L, 5 mg/L, 10 mg/L and 50 mg/L) of Ce in

CeO2 NPs and Ce3+ solutions (Mean±SD, n=3)……………………………………….30 

Table 4- 2 Content of total nitrogen (N), total phosphorus (P), Chlorophyll a (Chl a), Chlorophyll b

(Chl b) and Chlorophyll c (Chl c) in E. canadensis plants after being exposed to

different concentrations (0 mg/L, 0.5 mg/L, 1 mg/L, 5 mg/L, 10 mg/L and 50 mg/L) of

Ce in CeO2 NPs and Ce3+ solutions (For P: Mean±SD, n=3) .......................................... 31 

Table 4- 3 Content of Ce in E. canadensis plants (mg Ce/g DM) after cultivation in CeO2 NPs and

Ce3+ solutions containing different concentrations (0 mg/L, 0.5 mg/L, 1 mg/L, 5 mg/L,

10 mg/L and 50 mg/L) of Ce (Mean±SD, n=3) (DLCe=0. 0007mg Ce/g DM) .............. 33 

Table 4- 4 Mass change/initial weight (%) of E. canadensis plants after exposure to different

concentrations (0 mg/L, 0.05 mg/L, 0.1 mg/L, 0.25 mg/L, 0.5 mg/L and 1 mg/L) of Ag

in Ag NPs and Ag+ solution (Mean±SD, n=3) ................................................................. 33 

Table 4- 5 Content of total nitrogen (N), total phosphorus (P), Chlorophyll a (Chl a) , Chlorophyll b

(Chl b) and Chlorophyll c (Chl c) in E. canadensis plants after being exposed to

different concentrations (0 mg/L, 0.05 mg/L, 0.1 mg/L, 0.25 mg/L, 0.5 mg/L and 1

mg/L) of Ag in Ag NPs and Ag+ solutions (For P: Mean±SD, n=3) (DLChl c= 0.003

mg/g FW) ........................................................................................................................... 34 

Table 4- 6 Content of Ag in E. canadensis plants (mg Ag/g DM) after cultivation in Ag NPs and

Ag+ solutions containing different concentrations (0 mg/L, 0.05 mg/L, 0.1 mg/L, 0.25

mg/L, 0.5 mg/L and 1 mg/L) of Ag (Mean±SD, n=3) (DLAg=0.0005 mg Ag/g DM) ... 36 

xiii

Table 4- 7 General properties of different water samples used in the experiments (EC:Electrical

Conductivity; TC: Total Carbon; IC: Inorganic Carbon; TOC: Total Organic Carbon;

DR: Dry Residue; TSS: Total Suspended Solid) (Mean±SD, n=3) (DLTSS=0.33mg/L)39 

Table 4- 8 Concentration of anions in the different water samples used in the experiments

(Mean±SD, n=3) ................................................................................................................ 40 

Table 4- 9 Concentrations of Ca, Mg, K, Na, Ag, and Ce in the different water samples used in the

experiments (Mean±SD, n=3) (DLNa=6.25 mg/L; DLCe=0.04 mg/L; DLAg=0.03 mg/L)

............................................................................................................................................ 41 

Table 4- 10 Mass change/initial weight (%) of E. canadensis plants after cultivation for 72 hours in

different waters (Tap water, 10% Hoagland's solution, Mostbeek water, Grote Geul

water, Coupure water) spiked with either CeO2 NPs or Ce3+ (1 mg/L Ce). Control

group samples were not exposed to NPs nor ions (Mean values ± SD, n = 3) ............... 42 

Table 4- 11 Content of total nitrogen (TN), total phosphorus (TP), Chlorophyll a (Chl a) ,

Chlorophyll b (Chl b) and Chlorophyll c (Chl c) in E. canadensis plants after being

exposed to 1mg/L CeO2 NPs and 1mg/L Ce3+in Tap water, 10% Hoagland’s solution,

Mostbeek, Grote Geul and Coupure (Mean±SD, n=3) (DLChl c= 0.003 mg/g FW) ....... 43 

Table 4- 12 Mass change/initial weight (%) of E. canadensis plants after cultivation for 72 hours in

different waters (Tap water, 10% Hoagland's solution, Mostbeek water, Grote Geul

water, Coupure water) spiked with either Ag NPs or Ag+ (0.1mg/L Ag). Control group

samples were not exposed to NPs nor ions (Mean values ± SD, n = 3).......................... 45 

Table 4- 13 Content of total nitrogen (N), total phosphorus (P), Chlorophyll a (Chl a), Chlorophyll

b (Chl b) and Chlorophyll c (Chl c) in E. canadensis plants after being exposed to Tap

water, 10% Hoagland’s solution, Mostbeek, Grote Geul and Coupure water to which

Ag NPs or Ag+ions were spiked at a concentration of 0.1 mg Ag/L (Mean±SD, n=3)

(DLChl c= 0.003 mg/g FW) ................................................................................................. 46 

Table 5- 1 Mass balance of Ce (M2/M1) in tap water under exposure to different concentrations (0.5

mg/L, 1 mg/l, 5 mg/L, 10 mg/L and 50 mg/L) (M2/M1: actual total Ce/theoretical total

Ce; M3/M2: total Ce in plants/actual total Ce)…………………………………………62 

xiv

Table 5- 2 Mass balance of Ag (M2/M1) in tap water under exposure to different concentrations

(0.05 mg/L, 0.1 mg/l, 0.25 mg/L, 0.5 mg/L and 1 mg/L) (M2/M1: actual total

Ag/theoretical total Ag; M3/M2: total Ag in plants/actual total Ag) ................................ 62 

Table 5- 3 Mass balance (M2/M1) in different aquatic environments (Tap water, 10% Hoagland’s

solution, Mostbeek, Grote Geul and Coupure water) upon exposure to 1 mg/L Ce3+

solution and 1 mg/L CeO2 NPs (M2/M1: actual total mass of Ce/theoretical total mass of

Ce; M3/M2: total Ce in plants/actual total Ce) .................................................................. 63 

Table 5- 4 Mass balance (M2/M1) in different aquatic environments (Tap water, 10% Hoagland’s

solution, Mostbeek, Grote Geul and Coupure water) upon exposure to 0.1mg/L Ag+

solution and 0.1mg/L Ag NPs (M2/M1: actual total mass of Ag/theoretical total mass of

Ag; M3/M2: total Ag in plants/actual total Ag) ................................................................. 63 

xv

List of abbreviations

ENPs

ENMs

NMs

NPs

CeO2

TiO2

SiO2

Fe3O4

Fe2O3

ZnO

CNT

PEG

POD

CAT

SOD

TN

TP

CLPP

DGGE

FISH

AWCD

PCA

CSUPs

ICP-OES

ICP-MS

PVP

CMC

Engineered nanoparticles

Engineered nanomaterials

Nanomaterials

Nanoparticles

Cerium dioxide

Titanium dioxide

Silicon dioxide

Iron(II,III) oxide

Iron(III) oxide

Zinc oxide

Carbon nanotubes

Polyethyleneglycol

Peroxidase

Catalase

Superoxide dismutase

Total nitrogen

Total phosphorus

Community level physiological profiling

Denaturing gradient gel electrophoresis

Fluorescent in situ hybridization

Average well color development

Principal components analysis

Carbon source utilization patterns

Inductively Coupled Plasma Optical Emission Spectrometer

Inductively Coupled Plasma Mass Spectrometer

Polyvinylpyrrolidon

Carboxymethylcellulose

xvi

TOC

TC

IC

TSS

DR

EC

DL

Chl a

Chl b

Chl c

DM

FW

SD

Conc.

Wt

Total organic carbon

Total carbon

Inorganic carbon

Total suspended solids

Dry residue

Electrical conductivity

Detection limit

Chlorophyll a

Chlorophyll b

Chlorophyll c

Dry matter

Fresh weight

Standard deviation

Concentration

Weight

1

Chapter 1: Introduction

1.1. Background of the study

A nanomaterial (NM) is typically defined as a material with at least one characteristic

dimension below 100 nm (Wiesner, Lowry et al. 2006). The ability to design and use

nanoscale materials with their unique properties is now quickly developing in consumer

products, construction business, medical industries, agriculture industries and information

technologies. Maynard (2006) estimated that the production of engineered nanoparticles

(ENPs) will increase to 58,000 tons during 2011-2020, whereas it was only 2000 tons in 2004.

During the last decade, the global market of engineered nanomaterial (ENM) based products

has grown from $7.5 billion to $12.7 billion in 2008 (Fairbrother 2010), and it is expected to

reach $100 billion in 2015 (Podila and Brown 2013).

1.2. Potential problems

The wide use of nanoparticles (NPs) will undoubtedly lead to an increasing release of these

NPs into the environment, especially into the aquatic environment as they can be discharged

into the aquatic system via a number of pathways. Besides the great beneficial applications

that these NPs have, the public started to become skeptical about their subsequent behavior

and toxic impact on the environment and ecological systems. It should be taken into account

whether benefits of nanotechnologies outweigh costs to assure environmental safety and

human health.

However, as the metal-based engineered nanoparticles (ENPs) have different toxicity profiles

compared with large particles and metallic ions, their fate and mobility in the environment are

not predictable. As a result, their potential impact on aquatic systems is still not well known.

In addition, how and to what extent the NPs will be removed from wastewater in water

treatment facilities, like constructed wetlands, is also not well understood yet.

In constructed treatment wetlands, plants can play an essential role in cleaning the

2

environment by accumulating heavy metals and NPs. In addition, they can be used as specific

tools to monitor the degree of toxicity of pollutants to aquatic systems (Johnson, Ostroumov

et al. 2011). Evaluating the toxicity and fate of NPs in aquatic systems may not only help to

ensure safe application of nanotechnologies, but also design functional NPs that have minimal

adverse effects (Wu, Huang et al. 2010).

In addition to aquatic plants, microorganisms also play an important role in the degradation of

contaminants in the environment (Parkinson and Coleman 1991). However, the importance of

microbial ecology in constructed wetlands has often been overlooked (Weber, Gehder et al.

2008), and no information is yet available on the impact of metal-based ENPs on microbial

communities in these constructed wetlands.

1.3. Objectives

This thesis aims at evaluating the potential use of treatment wetlands to remove Ag NPs and

CeO2 NPs. In a first part, specific focus is laid on the role of aquatic macrophytes, revealing

the interactions between the NPs and these macrophytes, and also the factors that affect the

partitioning of NPs between plant tissue and the water phase. Kinetics of removal processes

are also investigated. In a second part, the effect of different types of Ag NPs on wetland

microbial communities was evaluated by the Community Level Physiological Profiling

(CLPP) method.

3

Chapter 2: Literature review

Nanotechnology opens up an opportunity to engineer the properties of different materials and

manufacture nanomaterials (NMs) for multiple uses. NMs can create more economic benefit

than bulk materials because of their unique physical and chemical characteristics, including

small size, high surface area to volume ratio, functional groups, solubility, shape and

aggregation behavior (Schrand, Rahman et al. 2010). With the increasing use of NPs in many

products, they have a great chance to enter the aquatic systems through several pathways

during their manufacture, transport and use. For instance, several researches have proven that

NPs can enter the aquatic system through wastewater treatment plants’ effluents due to the

increasing development of wastewater remediation techniques that are based on the use of

NPs (Vaseashta, Vaclavikova et al. 2007) or through the application of products that contain

NPs on agricultural lands (Gottschalk, Sonderer et al. 2009; Kim, Park et al. 2010; Kaegi,

Voegelin et al. 2011) As a result, in order to use these NPs in a safer way, it is important to

assess their toxicity, fate, mobility and other characteristics in the environment, especially in

aquatic systems.

2.1. Classification of NPs

Generally, NMs can occur in different forms, including one-dimensional fine rods,

two-dimensional ultrathin films and three-dimensional particles (Bernhardt, Colman et al.

2010). Depending on their source and composition, Nowack and Bucheli (2007) have fully

classified the NMs (Table 2-1).

4

Table 2- 1 Classification of nanomaterials (Adapted from: Nowack and Bucheli, 2007)

Formation Examples

Natural

C-containing

Biogenic Organic colloids Humic, fulvic acids

Organisms Viruses

Geogenic Soot Fullerenes

Atmospheri

c Aerosols Organic acids

Pyrogenic Soot

Carbon nanotubes

(CNT)

Fullerenes

Nanoglobules,

onion-shaped

nanospheres

Inorganic

Biogenic Oxides Magnetite

Metals Ag, Au

Geogenic Oxides Fe-oxides

Clays Allophane

Atmospheri

c Aerosols Sea salt

Anthropogenic

(manufactured,

engineered)

C-containing

By-productCombustion

by-products

CNT

Nanoglobules,

onion-shaped

nanospheres

Engineered

Soot

Carbon Black

Fullerenes

Functionalized

CNT, fullerenes

Polymeric NP Polyethyleneglycol

(PEG) NP

Inorganic

By-productCombustion

by-products

Platinum group

metals

Engineered

Oxides TiO2, SiO2

Metals Ag, iron

Salts Metal-phosphates

Aluminosilicate

s

Zeolites, clays,

ceramics

2.1.1. Natural NPs

Natural NPs have a long history which goes far beyond our imagination. Their origin dates

5

back to the formation of the earth. Naturally occurring NPs exist in various forms and have

been widely distributed in the atmosphere, oceans, soil, living organisms and terrestrial water

systems. Furthermore, they are even present in the deep earth, throughout the solar system

and interplanetary space (Wiesner, Lowry et al. 2009). Recent studies show that living

organisms can even synthesize NPs for their own use. Wetland plants Phragmites australis

and Iris pseudoacorus can transform copper into metallic NPs in and near roots with the

assistance of endomycorrhizal fungi when they are grown on contaminated soil. This kind of

process occurs in order to reduce Cu uptake by plants (Manceau, Nagy et al. 2008).

2.1.2. Anthropogenic NPs

Anthropogenic NPs include incidental NPs and engineered nanoparticles (ENPs). The incidental

NPs have risen dramatically since the Industrial Revolution because of the combustion of fossil

fuels, manufacturing emissions, mining, engine exhaust and desertification (Wiesner, Lowry et al.

2009; Bernhardt, Colman et al. 2010). Unlike the natural NPs and incidental NPs which are

heterogeneously formed and diffusely dispersed in the environment, the production of ENPs is

focused on generating pure suspensions or powders of NPs that are as homogeneous in size, shape

and structure as possible (Bernhardt, Colman et al. 2010). The production of different NPs turns out

to be very economically profitable as they can be used in a wide range of fields.

Among different ENPs, the metal-based ENPs, in particular, have received increasing interest for

application in the medical, industrial and military fields (Schrand, Rahman et al. 2010). These

metal-based ENPs usually have all three dimensions between 1 and 100 nm (Ju-Nam and Lead

2008). Schrand, Rahman et al. (2010) have fully reviewed different metal-based ENPs and their

applications (Table 2-2). Metal oxides (e.g., zinc oxide (Zhou, Xu et al. 2006), titanium oxide

(Song, Gao et al. 2012) and cerium oxide) NPs and metal (e.g., gold (Diegoli, Manciulea et al. 2008)

and silver (Gubbins, Batty et al. 2011)) NPs are two main groups in the field of metal-based ENPs.

6

Table 2- 2 Application of metal-based NPs (Adapted from: Bernhardt, Colman et al. 2010)

Nanoparticle Abbreviation Application

Aluminum Al Fuel additive/propellant, explosive, wear resistant

coating additive

Gold Au Cellular imaging, photodynamic therapy

Iron (oxide) Fe, Fe3O4,

Fe2O3 Magnetic imaging, environmental remediation

Silica SiO2

Fabrication of electric and thermal insulators, catalyst

supports, drug carriers, gene delivery, adsorbents,

molecular sieves and filler materials

Silver Ag Antimicrobial, photography, batteries, electrical

Copper Cu

Antimicrobial (i.e., antiviral, antibacterial, antifouling,

antifungal), antibiotic treatment alternatives,

nanocomposite coating, catalyst, lubricants, inks, filler

materials for enhanced conductivity and wear

resistance

Cerium

(oxide) CeO2

Polishing and computer chip manufacturing, fuel

additive to decrease emissions

Manganese

(oxide) Mn Catalyst, batteries

Nickel (oxide) Ni Conduction, magnetic properties, catalyst, battery

manufacturing, printing inks

Titanium

dioxide TiO2

Photocatalyst, antibacterial coating, sterilization, paint,

cosmetics, sunscreens

Zinc (oxide) Zn, ZnO Skin protectant, sunscreen

2.2.Environmental impact of ENPs

Regardless of the positive impacts of ENPs on the economy, the potential impacts of ENPs on

environment and human health have drawn the attention of the public. Recent research shows

that the particles’ behavior is influenced by the particles’ size, shape, surface charge and also

the presence of other materials in the environment (Handy, von der Kammer et al. 2008). To

our knowledge, with the decreasing of the particles’ size, the toxicity of the particles will

change. Johnston, Hutchison et al. (2010) suggested that particle sizes of gold and silver are

influential in observed toxicity response. Smaller particles have greater toxicity than their

larger counterparts. Gaiser, Fernandes et al. (2009) found that the nano-Ag (35 nm) particles

were more toxic than micro-Ag (0.6-1.6 μm) particles in the same aquatic invertebrate and in

7

vitro cell models.

Until now, the potential adverse impact of ENPs on the environment, especially on aquatic

macrophytes, has not been fully studied. Most studies focused on the biological toxicity of

NPs and to date, the probable toxicity of NPs has been proven. The results show that NPs can

be taken up by a wide range of mammalian cell types and cause adverse effects on organs,

tissue, cellular, subcellular and protein levels due to their specific physicochemical properties.

Oxidative damage and causing DNA damage to mammalian cells are the main toxicity

mechanisms of NPs on mammalian species (Bhabra, Sood et al. 2009; Meena and Paulraj

2012). Within mammalian cells, NPs can interfere with the antioxidant defense mechanism

which leads to reactive oxygen species generation, the initiation of an inflammatory response

and perturbation and destruction of the mitochondria causing apoptosis or necrosis (Schrand,

Rahman et al. 2010). NPs also show toxic properties in fish. Asharani, Wu et al. (2008) found

that the toxicity of Ag NPs to aquatic species depends on the concentration of NPs in water,

and NPs can decrease the heart rate, increase the mortality rate and cause hatching delays in

zebrafish embryos. Federici, Shaw et al. (2007) found that after exposure of rainbow trout to

TiO2 NPs, some gill pathologies appeared, including oedema and thickening of the lamellae.

In addition, a trend of decreasing enzyme activity in brain was also observed. Moreover, a

study on terrestrial plants like radish, rape and ryegrass shows that ZnO and Zn NPs can

inhibit root growth when the concentration is 10 mg/L or higher (Lin and Xing 2007).

However, very few researches focused on microorganisms and aquatic macrophytes in

wetlands.

2.2.1. Properties and toxicity of silver NPs

Ag NPs are one of the most valuable metal NPs in the global market. They have been found

in over 250 products in the world (Fabrega, Luoma et al. 2011), including many consumer

products like shampoo, soap, toothpaste, first aid bandages, nutritional supplements, textiles

and so on (Boxall, Chaudhry et al. 2007; Farre, Gajda-Schrantz et al. 2009). This is due to

their antibacterial, antimicrobial, antibiotic, antifungal and partially antiviral properties

8

(Blaser, Scheringer et al. 2008; Farre, Gajda-Schrantz et al. 2009). The wide use of Ag NPs in

the consumer market and remediation technologies have raise the opportunity of the NPs to

enter aquatic systems. According to Blaser, Scheringer et al. (2008), biocidal plastics and

textiles accounted for up to 15% of total silver released to water in the European Union in

2010.

Recent research shows that Ag NPs within the range of 1-10 nm can attach to and inhibit the

HIV-I virus from binding to host cells in vitro (Elechiguerra, Burt et al. 2005). Tian, Wong et

al. (2007) demonstrated that Ag NPs can help wounds heal faster. In their study, Ag NPs

exerted positive effects through their antimicrobial properties, reduction in wound

inflammation and modulation of fibrogenic cytolines. However, some adverse impacts were

also observed, e.g. when using root tip cells of Allium cepa to indicate the toxicity of Ag NPs.

Kumari, Mukherjee et al. (2009) reported that Ag NPs could penetrate plant systems and may

impair stages of cell division causing chromatin briage, stickiness, disturbed metaphase,

multiple chromosomal breaks and cell disintegration.

2.2.2. Properties and toxicity of CeO2 NPs

Meanwhile, CeO2 NPs draw interest from industry because of their high oxygen storage

capacity (Bekyarova, Fornasiero et al. 1998) and their ability to shift easily between the

reduced states (Ce3+) to oxidized states (Ce4+) (Nakagawa, Murata et al. 2007). Nowadays,

CeO2 NPs are used as catalysts and as diesel fuel additive to reduce particulate matter

emissions in the automotive industry (Bekyarova, Fornasiero et al. 1998; Milt, Querini et al.

2003; Gaiser, Fernandes et al. 2012).

A number of toxicity studies suggested that CeO2 NPs have no adverse impact or relatively

low levels of toxicity in vitro, especially on aquatic invertebrates (Gaiser, Fernandes et al.

2009; Gaiser, Biswas et al. 2011; Gaiser, Fernandes et al. 2012). Schubert, Dargusch et al.

(2006) even suggested that CeO2 NPs can be used to modulate oxidative stress in biological

systems as these NPs can protect nerve cells from oxidative stress.

9

2.2.3. Metallic NPs and their ionic metal forms

Recently, studies showed that erosion and runoff are potential pathways for Ag NPs and CeO2

NPs to enter waterways (Limbach, Bereiter et al. 2008; Lowry, Espinasse et al. 2012). Usually,

after NPs enter aquatic systems, they do not just stay in the water in a stable form. Under

some circumstances they will (slowly) dissolve and release metal ions (Kittler, Greulich et al.

2010). In a real environment, ultimate forms of NPs will be influenced by oxidation,

dissolution, sulfidation and so on, and occurrence of these transformations will mainly depend

on local environmental conditions (Lowry, Espinasse et al. 2012).

To our knowledge, silver and cerium are classified as hazardous substances and the toxicity of their

ionic forms has been confirmed through many studies (Mcdonald, Ghio et al. 1995; Ratte 1999). In

fish, the ionic form of silver (Ag+) can cause failure of the organisms to maintain constant Na+ and

Cl- concentrations in blood plasma, which can lead to fish death (Hogstrand and Wood 1998). A

study on primary mouse osteoblasts showed that cerium ions have an adverse impact on the

proliferation, differentiation and mineralization functions after in vitro exposure (Zhang, Liu et al.

2010). All these facts lead to raising concerns on adverse impacts of using NPs and their potential

toxicity to environment and human health.

2.3. Aquatic macrophytes

In aquatic systems, plants are definitely acting as vital parts. They can be primary producers,

provide surface area for bacteria, act as food source for other organisms, provide a habitat for

fish and invertebrates, stabilize the soil, and they are also important in biogeochemical

cycling of nitrogen and carbon (Gubbins, Batty et al. 2011). Using macrophytes for

bioremediation has many advantages. Macrophytes can take up pollutants not only through

their roots but also their leaves. Besides, indeterminate growth of most aquatic macrophytes

also allows for monitoring over time as plants are easy to clone and transplant (Mal, Adorjan

et al. 2002). Previous research on effects of Ag NPs and CeO2 NPs on plants was mainly

focused on algae (Oukarroum, Bras et al. 2012) or floating macrophytes, like duckweed

10

(Lemna Minor L, Landoltia Punctata etc.) (Shi, Abid et al. 2011; Song, Gao et al. 2012). Only

very few studies investigated effects on submerged plants.

2.3.1. Aquatic macrophytes - Elodea canadensis

Elodea canadensis (Figure 2-1) is a classical submergent macrophyte, usually known by

people as an “aquarium plant”. However, it is also a common aquatic macrophyte used in

wetlands to remove organic compounds and heavy metals (Bastviken, Eriksson et al. 2005).

During the past few decades, E. canadensis has been introduced to many places as “aquarium

plant” which made it become one of the key aquatic macrophytes species with a broad

distribution around the world. In addition, Elodea Canadensis has a strong potential for

invasion into new aquatic bodies (Johnson, Ostroumov et al. 2011).

 

Figure 2- 1 Illustration of Elodea Canadensis (Source: http://www.evi.com/q/facts_about__elodea_canadensis)

Previous studies showed that some aquatic plant materials like Azolla filiculoides (Dewet,

Schoonbee et al. 1990; Begum and HariKrishna 2010; Elmachliy, Chefetz et al. 2011) have a

remarkably high capacity to remove heavy metals from wastewater. Generally, there are two

main mechanisms that affect the process. One is a fast metabolism independent of surface

11

reaction, while the other one is a slow metabolism dependent on cellular uptake reactions

(Cho, Lee et al. 1994). In the first process, soluble ions can bind or adsorb to the plants, via

diffusion. The second process is more about mass transfer from the outer to the interior cell

wall (Axtell, Sternberg et al. 2003). Johnson, Ostroumov et al. (2011) suggested that the

removal of metal-based NPs is mainly influenced by the binding process. However, this

process seems to be a species specific process. Zhu, Han et al. (2008) found that magnetite

(Fe3O4) NPs can be adsorbed on pumpkin plants but not adsorbed on lima beans.

2.3.2. Characterization of plant performance

To date, a number of parameters (chlorophyll, peroxidase (POD), catalase (CAT), superoxide

dismutase (SOD), total nitrogen, total phosphorus, etc.) have been developed to evaluate the growth

of plants. Depending on the objective, different parameters can be chosen. In our study, chlorophyll

pigments, total nitrogen and total phosphorus are measured.

Chlorophyll a, b, c

Photosynthesis is the basic function that determines the productivity of plants (Nekrasova,

Ushakova et al. 2011). The process of photosynthesis is mainly affected by photosynthetic

chlorophyll pigments. In green plants, chlorophyll pigments can be present in several forms

(chlorophyll a, b, c etc.) in varying ratios, and the determination of different chlorophyll pigments

can indicate the potential toxicity of the environment to the plants.

Nitrogen and phosphorus

Both nitrogen and phosphorus are essential nutrient elements for plants. Lack of nitrogen and/or

phosphorus is considered to limit the growth of plants (Gerloff and Skoog 1954). Nitrogen

concentration in the leaf can affect the photosynthetic rate because nitrogen is an important

component of photosynthetic enzymes and chlorophyll. Generally, nitrogen and phosphorus have a

great effect on growth, affecting cell number and cell size (Chapin 1980).

12

2.4. Microbial communities and their characterization

Both natural and constructed wetlands have a complex microbial regime. Inside the regime,

micro-organisms play a vital role in ecosystem health, nutrients cycling and the degradation

of contaminants (Aelion and Bradley 1991; Wynn and Liehr 2001). Within the treatment

wetlands, plant tissue, sediments and other objects in the water body can provide surface area

for microbial growth. Hamilton pointed out that bacteria are much more abundant when

grown attached to surfaces rather than just suspended in water (Hamilton 1987).

Usually in constructed wetlands, the performance of purification is based on the combined

action between microbes and plants. To our knowledge, the microbes act actively in

mineralization of organic matter under aerobic and anaerobic conditions (Truu, Juhanson et al.

2009). Therefore, a better understanding of the microbial communities’ behavior upon

exposure to NPs will provide useful information on the performance of constructed treatment

wetlands receiving nanoparticles (Faulwetter, Gagnon et al. 2009).

Currently, there is a variety of methods that can be used to characterize the microbial

communities and each method has its own benefits and limitations. Molecular methods as

well as non-molecular methods are frequently used. Molecular technologies include

polymerase chain reaction followed by denaturing gradient gel electrophoresis (PCR-DGGE)

(Muyzer, Dewaal et al. 1993), fluorescent in situ hybridization (FISH) (Manz, Amann et al.

1992) and so on. The community level physiological profiling (CLPP) method using

BIOLOGTM plates (Weber and Legge 2011) is the most frequently used non-molecular

technique.

The purpose of the experiment is to investigate the overall impact of Ag NPs and Ag+ on

microbial community metabolic functioning in small – scale wetland mesocosms. In this case,

the classical plating and molecular-level techniques seem to be quite time consuming

(Hallberg and Johnson 2005). CLPP is considered as a good choice as it is a relatively rapid

method, does not need specialized expertise and also allows for functional community

characterization at the same time (Weber and Legge 2011).

13

2.4.1. CLPP using BIOLOGTM plates

Microbial communities have great potential for temporal or spatial change in both basic and

applied ecological contexts (Garland 1997). In order to have a better understanding of the

mesocosms, a simple and powerful method needs to be introduced. CLPP offers an easily

applied protocol yielding large amounts of information regarding mixed microbial community

function and functional adaptations over space and time (Weber and Legge 2010). Technically,

CLPP refers to a variety of assays. Currently, the term CLPP is used to describe the collection

of data obtained from BIOLOGTM microplates.

BIOLOGTM microplates (Figure 2-2) consist of 96 wells. In each well, there is a redox dye

indicator (tetrazolium violet) and a different carbon source. During the incubation time, the

color in each well changes due to the production of NADH via cell respiration which can be

detected spectrophotometrically (Weber and Legge 2010).

 

Figure 2- 2 Model of BIOLOGTM ECO microplates (Source: http://www.techno-path.co.uk)

Various types of plates can be used in the Biolog plates’ method. GN2, GP2 and EcoPlates are the

most popular plates. GN2 and GP2 are plates that both contain 95 different carbon sources

(carbohydrates, amino acids, amines, amides, polymers etc.) and with the 96th well as a blank. The

Biolog GN2 plates are designed for gram-negative bacteria while GP2 plates are used to identify

gram-positive bacteria. Both GN2 and GP2 plates are originally designed for species identification.

In contrast to the GN2 and GP2 plates, the ECO plates only have 31 different carbon sources with

14

three replicates for each carbon source. The triplicates are used for better replication and allow for a

better indication of experimental variation (Weber, Gehder et al. 2008). In addition, these ECO

plates are typically designed for ecological study of whole microbial communities instead of

identification of microbial strains (Stefanowicz 2006). The BIOLOGTM ECO plates were first

introduced by Grove, Kautola et al. (2004) to assess the microbial communities in biofilters.

Overall, the Biolog plates’ method is a rapid and convenient method to qualify microbial metabolic

capabilities and hence the functional diversity of microbial communities (Stefanowicz 2006).

However, it still has some drawbacks. First, this method mainly depends on microorganisms that

are active under specific lab conditions, so it may cause bias towards rapidly growing bacteria

(Leckie 2005). Second, as the procedure is conducted under laboratory conditions, it will not be

suitable for all bacteria to grow. Third, there is a potential chance that the carbon sources in

different wells may not always be directly relevant to the communities being studied. In addition,

careful data analysis and interpretation can be challenging in this technique (Insam, Goberna et al.

2004).

There are a number of ways to analyze data collected from Biolog plates, but Weber and

Legge (2010) highlighted the use of average well color development (AWCD), substrate

richness, substrate diversity, and principal components analysis (PCA) using carbon source

utilization patterns (CSUPs) to analyze CLPP data which proved to be very sufficient.

AWCD refers to the average absorbance value (corrected by the blank well) of all 31 wells

which gives an assessment of overall catabolic activity in microbial communities.

131

Where:

AWCD = average well color development

Ai = absorbance reading of well i

A0 = absorbance reading of the blank well (inoculated, but without a carbon source)

15

Sometimes negative standardized absorbance values occur in case there is very little response

in a well. In this case, as data are physically meaningless, they are coded as zeros for further

analysis.

Substrate richness is a measure of the number of different carbon sources utilised by a

microbial population, and is calculated as the number of wells with a corrected absorbance

greater than 0.25 AU. Diversity is expressed here in terms of the Shannon index.

Although absorbance by the wells can be measured regularly in a specific time period, a

single time point can be selected for the evaluation of all plate data according to Weber and

Legge (2010). Selection of this time point should be based on a combination of greatest

variance between well responses and least number of absorbance values above 2 (as these are

not situated within the linear response range).

Weber, Grove et al. (2007) suggest using Taylor power law transform for principle components

analysis. It proved that the transformation can help increase homoscedasticity and normality of the

data. Principle components were extracted and ordinations created from the covariance matrix of

the data using XLSTAT 2013.

2.4.2. Denaturing gradient gel electrophoresis (DGGE)

For a long time, DGGE has been used as a method to investigate the distribution of bacterial

assemblages (Diez, Pedros-Alio et al. 2001). The main purpose of DGGE is to study the

presence and activity of bacterial populations in complex mixtures. In DGGE, DNA

fragments with different sequences but the same length can be separated. The separation is

based on the decreased electrophoretic mobility of a partially melted double-stranded DNA

molecule in polyacrylamide gels containing a linear gradient of DNA denaturants (Muyzer

and Smalla 1998). Among the different DGGE studies, polymerase chain reaction followed by

denaturing gradient gel electrophoresis (PCR-DGGE) is one of the most frequently used

methods. Muyzer, Dewaal et al. (1993) were the first to use 16S rRNA based PCR-DGGE to

profiling the complex microbial populations and they found that this DGGE method can not

16

only be used to determine the genetic diversity among species, but is also useful in diagnosing

the presence and relative abundance of microorganisms. Based on this knowledge, Vilela,

Pereira et al. (2010) used PCR-DGGE to sufficiently evaluate the dynamics of microbial

communities during semi-dry coffee processing. In addition, Raats, Offek et al. (2011) used

16S rRNA based PCR-DGGE to evaluate the microbial communities in raw cow milk.

2.4.3. Fluorescent in situ hybridization (FISH)

FISH is a powerful cytogenetic technique that is usually used to detect and localize specific

DNA sequences on chromosomes, and it turns out to be a highly effective method for

determining the number of specific chromosomes in interphase cells (Feldman, Ebrahim et al.

2000). Usually, a single color FISH is not sufficient enough to estimate the result. As a result,

a multicolor FISH is usually employed (Bischoff, Nguyen et al. 1995). Shin, Ross et al. (1997)

used a multi-color FISH to determine the incidence of somatic chromosomal numeric

alterations in severe/late stage endometriosis.

Generally, there are four types of FISH probes (Figure 2-3)(McNeil and Ried 2000).

Gene-specific probes target specific nucleic acid sequences on chromosomes. Centromeric

probes can bind to repetitive sequences which are specific in this region. Telomeric probes

can be used to visualize all telomeres simultaneously as it has repetitive sequence TTAGGG.

The last one, the chromosome-painting probe consists of pools of chromosome-specific

probes.

17

Figure 2- 3 Different types of FISH Probes (Source: McNeil and Ried, 2000, Cambridge University Press)

18

Chapter 3: Materials and Methods

3.1. Introduction

The whole study was divided into four experiments: a Dose Experiment, a Desorption Experiment,

a Water Composition Experiment and the CLPP Experiment. During the whole experimental

period, Elodea canadensis (Figure 3-1) was cultivated in two water tanks with 10% Hoagland’s

solution (Table 3-1) as cultivation media. The media solution in the tanks was renewed every 1-2

weeks. The Elodea canadensis plants were purchased from Van der Velde Waterplanten B.V.,

Bleiswijk, Holland. The citrate coated Ag NPs (100 ppm) and CeO2 (50000 ppm) NPs were

purchased from PlasmaChem GmbH, Berlin, Germany. The cerium standard solution (Plasma

HIQU, 10000 ± 20 µg Ce3+ mL-1 in 2 – 5 % HNO3) and silver standard solution (Plasma HIQU,

1000 ± 2 µg Ag+ mL-1 in 2 – 5 % HNO3) were obtained from Chem-Lab NV, Belgium. All other

chemical reagents were either obtained from Chem-Lab NV, Belgium or purchased from Merck,

Germany.

Figure 3- 1 Cultivation of Elodea canadensis in water tanks with 10% Hoagland’s solution

19

Table 3- 1 Composition of modified 10% Hoagland's medium

Compound Molecular mass

(g/mol)

Stock solution

(g/500 mL) Use (mL/L)

MgSO47H2O 246.48 61.5 0.2

Ca(NO3)24H2O 236.15 59 0.46

KH2PO4 136.09 34 0.05

KNO3 101.11 25.25 0.25

Micronutrients

H3BO3 61.83 1.43

0.05

MnCl24H2O 197.84 0.91

ZnSO47H2O 287.54 0.11

Na2MoO42H2O 241.95 0.045

CuSO45H2O 249.68 0.045

FeEDTA FeCl36H2O 270.33 0.242

2 EDTA 292.24 0.750

3.2. Dose Experiment

3.2.1. Experimental Setup

In this experiment, pure tap water was used as cultivation media. Before the start of the experiment,

each Elodea canadensis plant was cut to approximately the same size (about 16 cm) and then

incubated in a 100 mL polypropylene recipient (Figure 3-2).

20

Figure 3- 2 Setup of the Dose experiment

The incubation was performed at a fixed illumination (light/dark =12h/12h) with different NPs and

ions concentrations (CeO2 NPs: 0.5 mg/L, 1 mg/L, 5 mg/L, 10 mg/L, 50 mg/L; Ce3+ ions: 0.5 mg/L,

1 mg/L, 5 mg/L, 10 mg/L, 50 mg/L; Ag NPs: 0.05 mg/L, 0.1 mg/L, 0.25 mg/L, 0.5 mg/L, 1 mg/L;

Ag+ ions: 0.05 mg/L, 0.1 mg/L, 0.25 mg/L, 0.5 mg/L, 1 mg/L and control group without any

exposure of NPs/ions) under room temperature for 72 hours. Both Ce3+ ions and Ag+ ions were

spiked from the standard solutions which contained 2 - 5% HNO3. For each concentration, 5

replicates were included (3 for Ce or Ag and phosphorus analysis, 1 for chlorophyll analysis, and 1

for nitrogen analysis). All plants were fully immersed in the solution and the water samples were

taken at T0 (0h) before introducing the plants and T1 (72h) after cultivating plants for 72 hours.

After cultivation, plants were harvested. One replicate was kept fresh for the chlorophyll analysis

and the other four were dried in the oven at 55ºC for the other measurements (total nitrogen, total

phosphorus, and Ce or Ag).

3.2.2. Water analysis

An Inductively Coupled Plasma Optical Emission Spectrometer (ICP-OES) (Varian VISTA-MPX

CCD Simultaneous ICP-OES) and Inductively Coupled Plasma Mass Spectrometer (ICP-MS)

21

(PerkinElmer Elan DRC-e ICP-MS) were used to measure the Ce and Ag concentrations in the

water samples before and after incubation (T0 (0 h) and T1 (72 h)).

Two mL sample and 4 mL 65% HNO3 solution were mixed together in a centrifuge tube and left to

stand in a fume hood overnight. The samples were subsequently digested using a microwave (CEM

MARS-5). In the microwave, first, the samples were ramped to a temperature of 55°C over 5 min

and held at that temperature for 10 min. Second, the temperature was ramped to 75°C over 10 min

and held at that temperature for 10 min. Finally, the samples were ramped to 100°C over 10 min

and held at that temperature for 40 min. This microwave digestion was conducted at a power of 600

W. After the samples were cooled to room temperature, deionized water was added to the digested

solution to reach 20 mL.

3.2.3. Plants sampling and characterization

Fresh weight, dry weight, chlorophyll, total nitrogen and total phosphorus contents were measured

to assess the impact of the different NPs/ions on Elodea canadensis. To assess the removal

efficiency of the plants, the plants were dried to constant weight at a temperature of 55°C. Five mL

65% HNO3 was then added to predigest the dry plant samples overnight. After microwave digestion,

ICP-OES was used to measure the metal concentrations in the plants’ biomass. The digestion and

dilution conditions were the same as for analysis of the water solution (section 3.2.2).

Chlorophyll a, b, c

Approximately 0.5 g of fresh plants was weighed in centrifuge tubes and then mashed using a

pulper together with 15 mL aqueous Acetone-Mg(HCO3)2 solution. After that, the centrifuge tubes

together with the plants and solution were covered with aluminum foil to avoid light exposure. All

samples were allowed to steep at 4°C for a minimum of 2 hours and a maximum of 24 hours.

Subsequently, the samples were centrifuged for 15 min at 3000 rpm. Absorbance of the clarified

extracts was read on the spectrophotometer (JENWAY 6400) at the wavelengths of 750 nm, 664

nm, 647 nm and 630 nm.

22

Ca=11.85(OD664)-1.54(OD647)-0.08(OD630)

Cb=21.03(OD647)-5.43(OD664)-2.66(OD630)

Cc=24.52(OD630)-7.60(OD647)-1.67(OD664)

Where:

Ca, Cb, Cc= concentrations of chlorophyll a, b and c, respectively, in mg/L.

OD647, OD664, OD630= corrected optical densities (with 1 cm light path) at the respective

wavelength.

After determining the concentration of pigment in the extract, the amount of pigment per unit fresh

weight was calculated as follows (take Chlorophyll a as an example):

Chlorophyll a (mg/g FW) =

Total nitrogen

The total nitrogen content was determined through a modified Kjeldahl method (Van Ranst et al.,

1999). First, approximately 0.100 g dry plant material was transferred into the digestion receptacles

and then 7 mL combined reagent (sulfuric/salicylic acid) was added. After reacting for 30 min, 0.5

g sodium thiosulfate was added and after 15 min, 5 mL concentrated sulfuric acid together with 0.2

g catalyst (selenium reagent mixture) and 4 mL H2O2 were added to the solution. Second, all

receptacles were heated at 380°C for at least 1 h until a clear solution was obtained. Third, ±30 mL

of distilled water was added to the clear solution after the solution was cooled down to room

temperature. Subsequently, a base was added to the heated solution to convert ammonium (NH4+)

to ammonia (NH3) and then the obtained solutions were distilled for 8 minutes in a distiller

(Vapodest, Voor ’t Labo, Belgium). Finally, a titrating apparatus (718 STAT Titrino, Metrohm,

Switzerland) was used to titrate the distillate with a N-indicator which contained boric acid. The

amount of total nitrogen was calculated per unit dry matter based on the volume of 0.01 N HCl that

has been used.

14 1000 0.001

23

Where:

DM= weight (dry mass) of the sample (g)

V1= volume of acid added to the distillate during titration (mL)

V0=volume of acid added to the blank during titration (mL)

CHCl=concentration of the acid used for titration (0.01N)

Phosphorus determination

After microwave digestion and the dilution, 1 mL of each solution was transferred into a test tube.

Successively 5 mL milli-Q water, 1 mL Scheel (I) solution, and 1 mL Scheel (II) solution were

added and this mixture was shaken energetically for a perfect homogenization. It was allowed to

react for 15 min. Afterwards, 2 mL Scheel (III) solution was transferred to the tube, after which the

tube was shaken again and allowed to react for another 15 min (Table 3-2).

Table 3- 2 Composition of Scheel I, II, III solutions used for phosphorus determination

Reagent Composition (per liter H2O)

Scheel (I)

1 g Monomethyl-para-aminophenol Sulfate;

5 g Na2SO37 H2O;

150 g NaHSO3 (or 137 g Na2S2O3)

Scheel (II) 50 g (NH4)6Mo7O244H2O;

140 mL H2SO4 (=1.83 g/mL)

Scheel (III) 205 g Sodium Acetate (NaOAc) / 340 g Sodium

Acetate-3-Hydrate (NaOAc4H2O)

After that, the absorbance was read at 700 nm using the spectrophotometer.

mgP ∙1 1 1

1000

Where:

D=dilution factor

ER= extraction ratio (in g dry matter per mL extract)

CP= concentration of P in the extract, calculated from the absorbance.

24

3.3. Desorption experiment

Four target solutions were prepared in advance, including 1 mg/L Ce3+ ions, 1 mg/L CeO2 (0.8

mg/L Ce) NPs, 0.5 mg/L Ag+ ions and 0.5 mg/L Ag NPs solutions. Six different extraction

solutions (Milli-Q water, 0.5 M HNO3, 0.02 M EDTA, 0.05 mM PVP, 0.05 mM CMC and 0.05

mM Dextran 70) were prepared with Milli-Q water.

Eighteen plants (with similar size 16 cm) were directly immersed into 500 mL solutions

containing the NPs/ions for 60 min. After that, the plants were harvested and gently rinsed with

distilled water. Different extraction reagents were distributed into different polypropylene recipients

and the plants that had been exposed to the NPs/ions solutions were directly immersed into the

different extraction solutions for 30 min. Samples of the extraction solutions were taken both before

and after extraction. Sample preparation prior to analysis was similar as in the dose experiment

(Section 3.2). Again, ICP-OES and ICP-MS were used to measure the metal concentrations in the

solutions, indicating the adsorption of NPs/ions to the plants as well as the capacity of the

extractants to remove them again from the plant after adsorption.

3.4. Water composition experiment

3.4.1. Experimental setup

The outline of this experiment was similar to the outline of the dose experiment. However, in this

experiment, 5 different waters, differing in physicochemical characteristics, have been used as

cultivation media instead of just tap water. Next to tap water, 10% Hoagland’s solution and three

surface waters (Mostbeek water, Grote Geul water and Coupure water) were included. The

experiment was divided into 5 batches with each batch containing a certain concentration of

NPs/ions (no NPs/ions, CeO2 NPs (1 mg/L), Ce3+ solution (1 mg/L), Ag NPs (0.1 mg/L) and Ag+

solution (0.1 mg/L)). The exposure period was 3 days. For each combination, 5 replicates were

used to be able to measure different parameters (3 for Ce or Ag and phosphorus contents, 1 for

chlorophyll and 1 for nitrogen).

25

To assess the kinetics of the removal process, 2 mL water samples were taken at 0h, 2h, 6h, 10h,

24h, 48h and 72h. After harvesting the plants that have been immersed in the solution for 3 days,

their fresh and dry weights were recorded. The same methods were used as in the dose experiment

(Section 3.2) for sample preparation and analysis of the content of Chlorophyll a, b, c, total

nitrogen, total phosphorus and the metal concentrations in the plant samples, and metal

concentrations in the water samples.

3.4.2. Water composition analysis

Most methods used to characterize the water samples were based on the Manual for the Soil

Chemistry and Fertility Laboratory (Van Ranst et al., 1999). All measurements were performed in

triplicate.

Electrical conductivity (EC)

Approximately 50 mL of each water sample was transferred into a centrifuge tube, and then a

conductivity meter (LF537, WTW, Weilheim, Germany) was used to directly measure EC.

pH determination

A pH meter (Model 520A, Orion Research Inc., Boston, MA, USA) was used to directly measure

the pH value of different samples. Prior to measurement, a calibration was performed at pH=7 and

pH=4.

Total organic carbon (TOC), total carbon (TC), and inorganic carbon (IC)

All water samples were transferred into specific tubes and then a TOC analyser (TOC-VCPN,

Shimadzu, Kyoto, Japan) was used to directly measure TOC, TC and IC in the different water

samples.

26

Total suspended solids (TSS)

At the beginning, 0.45 μm membrane filters were dried at 105ºC for 30 min in the oven. Then

200-300 mL water samples were filtered over the dried membranes. After filtration, the membrane

filters were dried again in the oven at 105ºC for at least 1 hour. In the end, TSS was calculated

through the mass difference before and after filtration.

TSSmgL

1000 1000

Where:

A = weight of dried membrane (g)

B = after filtration, weight of dried membrane with suspended solids (g)

V = the total volume of water sample which has been filtrated (mL)

Dry residue

Fifty mL water samples were transferred into labeled beakers, and then the beakers were put into

the oven at a temperature of 105ºC until all water was evaporated. The weight of empty beakers,

the weight of the beakers with the water samples and the weight of the beakers and dry residue

were recorded in a worksheet. When the difference between empty beakers and beakers with dry

residue was less than 20 mg, another 50 mL water sample was added, and the procedure was

repeated again.

DRmgL

1000 1000

Where:

A = weight of empty beaker (g)

C = weight of beaker with dry residue (g)

V = total volume of the water sample (mL)

27

K, Na, Ca, Mg, Ag, Ce

ICP-OES was used to analyse the concentrations of K, Na, Ca, Mg, Ag and Ce in the different

water samples.

Anions

Diluted water samples were analysed for anions (F-, Cl-, NO2-, NO3

-, PO43- and SO4

2-) using ion

exchange chromatography (761 Compact IC, Metrohm, Switzerland).

3.5. CLPP experiment

Mesocosms mimicking the gravel bed of a wastewater treatment wetland were constructed as

depicted in Figure 3-3. A perforated tube was placed in the middle of the plastic pot and used to

extract the wastewater with suspended microbial communities. Between the tube and the pot’s wall,

gravels from a local wetland - which could already have biofilms attached on the surface - were

used to fill up the pots to 10-15 cm. These mesocosms were subsequently fed with 0.5 L of

simulated wastewater (1 g/L Molasses, 0.049 g/L Urea and 0.0185 g/L NH4H2PO4), which was

changed every week in order to simulate the 7-day hydraulic retention time in wastewater treatment

wetlands. At the same time, Ag NPs or ions were added to the simulated wastewater to achieve 0.1

mg/L Ag exposure in all groups except the control group. Three different types of Ag NPs were

investigated: biogenic Ag NPs produced with L. fermentum supplied by the Laboratory of

Microbial Ecology and Technology of Ghent University (Prof. Nico Boon), and both citrate- and

PVP-coated Ag NPs from a commercial supplier. Moreover, one positive control (Ag+) and one

negative control (no Ag) were included. All measurements were conducted in triplicate (A-C).

28

Figure 3- 3 Setup of mesocosms experiment

After 30 days’ cultivation, samples were taken and prepared in several steps. Firstly, the interstitial

wastewater containing micro-organisms was extracted from each pot through the central sampling

tube and labeled in the same way as was done for the pots (e.g. control A, B, C; biogenic A, B, C

etc.). Secondly, 18 mL of the sampled liquid was taken and mixed with 2 mL of corresponding

stock solution (Ag+, biogenic Ag NPs, PVP-coated Ag NPs or citrate-coated Ag NPs) in order to

obtain a solution containing 0.5 mg/L, 1 mg/L, 2 mg/L and 5 mg/L Ag. These new samples were

re-labeled with the group name and Ag concentration (e.g. biogenic 0.5, biogenic 1, biogenic 2,

biogenic 5 etc.). Similarly, 18 mL was sampled from the control group (no Ag) and mixed with 2

mL of all stock solutions (Ag+, biogenic Ag NPs, citrate-coated Ag NPs and PVP-coated Ag NPs)

to reach 0.5 mg/L, 1 mg/L, 2 mg/L and 5 mg/L Ag. These samples were labeled as control + group

name + concentration (e.g. control + biogenic 0.5, control + biogenic 1 etc.). In the end, the gravels

from the triplicates of a same treatment were brought together and manually mixed. After that, from

each group 25 g of gravel was weighed and put into a 250 mL phosphate buffer. The mixed

buffering solution was shaken for 3 h at room temperature so that the biofilm could be detached

from the gravels’ surface, generating a suspended microbial community sample of the biofilm. All

these samples were labeled as the group name + biofilm (e.g. biogenic biofilm).

After this sample preparation, each sample was transferred to a BIOLOGTM ECO plate which was

purchased from LABiosytems, The Netherlands. In this assay, 100 μL of mesocosm from

interstitial water or biofilm extract was added to each well of a plate, followed by incubation for

115 h. Absorbance was measured twice per day at 595 nm using a Tecan Infinate M200 plate

29

reader, with 5 seconds of shaking at 2 mm amplitude prior to each reading. After reading the plates,

a time point which gives the most information of the microbial communities was chosen and all

data from interstitial water and biofilms at that time point were used for PCA analysis to indicate

the impact of Ag NPs/ions on wetland microbial communities.

30

Chapter 4: Results

4.1. Dose - response experiments

4.1.1. Elodea canadensis plants exposed to CeO2 NPs and Ce3+

After three days’ cultivation, E. canadensis plants were harvested and characterized. The mass

difference before and after cultivation was determined and presented as percentage of the initial

fresh weight in Table 4-1.

Throughout the whole cultivation period, the fresh weight of the plant did not change too much

compared with the initial fresh weight before introducing plants to the solutions containing various

concentrations of CeO2 NPs and Ce3+.

Table 4- 1 Mass change/initial weight (%) of E. canadensis plants after exposure to different concentrations (0 mg/L,

0.5 mg/L, 1 mg/L, 5 mg/L, 10 mg/L and 50 mg/L) of Ce in CeO2 NPs and Ce3+ solutions (Mean±SD, n=3)

0 mg/L 0.5 mg/L 1 mg/L 5 mg/L 10 mg/L 50 mg/L

CeO2 NPs 0.79±6.22 3.39±3.68 1.37±1.29 1.09±3.38 2.53±0.81 3.28±6.91

Ce3+ 0.79±6.22 1.00±5.53 -2.42±4.75 -2.36±2.07 2.21±1.41 -0.59±0.78

The composition of the plants changes when subjecting them to different concentrations of CeO2

NPs and Ce3+ (Table 4-2). In presence of CeO2 NPs, the total nitrogen and total phosphorus content

did not change too much as function of the exposure. The content of chlorophyll a, b, c varied with

the change in NPs’ concentration, but always Chl a > Chl b > Chl c. The same tendency could also

be found in plants exposed to Ce3+. However, at 50 mg/L Ce3+ solution, the chlorophyll content

decreased sharply, which was especially the case for Chl c, which dropped from 0.037 mg/g FW at

10 mg/L to 0.009 mg/g FW at 50 mg/L. At the same time, the content of phosphorus also dropped

from 8.19±0.55 mg P/g DM at 10 mg/L to 5.51±1.13 mg P/g DM at 50 mg/L. In contrast, the

nitrogen content increased from 20.9 mg N/g DM at 10 mg/L to 35.0 mg N/g DM at 50 mg/L.

31

Table 4- 2 Content of total nitrogen (N), total phosphorus (P), Chlorophyll a (Chl a), Chlorophyll b (Chl b) and

Chlorophyll c (Chl c) in E. canadensis plants after being exposed to different concentrations (0 mg/L, 0.5 mg/L, 1 mg/L, 5

mg/L, 10 mg/L and 50 mg/L) of Ce in CeO2 NPs and Ce3+ solutions (For P: Mean±SD, n=3)

0 mg/L 0.5 mg/L 1 mg/L 5 mg/L 10 mg/L 50 mg/L

CeO2

NPs

N (mg N/g DM) 24.4 29.1 34.3 25.5 26.9 26.5

Chl a (mg/g FW) 0.444 0.393 0.348 0.465 0.458 0.372

Chl b (mg/g FW) 0.394 0.289 0.262 0.401 0.402 0.328

Chl c (mg/g FW) 0.052 0.029 0.031 0.048 0.067 0.057

P (mg P/ g DM) 7.30±3.25 8.31±0.85 7.88±0.88 7.96±0.99 8.89±0.99 8.34±0.65

Ce3+

N (mg N/g DM) 24.4 30.7 29.5 26.4 20.9 35.0

Chl a (mg/g FW) 0.444 0.385 0.403 0.277 0.466 0.455

Chl b (mg/g FW) 0.394 0.265 0.273 0.222 0.346 0.304

Chl c (mg/g FW) 0.052 0.019 0.020 0.033 0.037 0.009

P (mg P/ g DM) 7.30±3.25 8.65±1.43 8.21±1.90 7.94±0.89 8.19±0.55 5.51±1.13

It should be mentioned that the real concentration of Ce in CeO2 NPs and Ce3+ solutions was

always lower than the targeted concentrations (Figure 4-1). For example, in the 50 mg/L treatment,

the measured concentration of Ce in CeO2 NPs solution was only 37.40 ± 0.6 mg/L compared to

47.2 ± 4.0 mg/L for Ce3+. Even though the tests were designed to have the same initial

concentration (Figure 4-1 - X axis), the real Ce concentration in Ce3+ spiked solution (Figure 4-1 -

Y-axis) was always higher than the concentration of Ce in solutions spiked with CeO2 NPs. Figure

4-1 also shows that after introducing plants, the amount of Ce in the water samples decreased. At

50 mg/L, the concentration of Ce in the CeO2 NPs treatment dropped to 24.6 ± 3.2 mg/L while the

concentration of Ce in the Ce3+ treatment decreased to 29.8 ± 8.5 mg/L after the cultivation. In the

control group without the presence of E. canadensis, the Ce concentration slightly changed from

0.062 ± 0.010 mg/L to 0.125 ± 0.006 mg/L (data not shown).

32

Figure 4- 1 Relationship between theoretical concentration and actual concentration of Ce in solutions spiked with

CeO2 NPs (above) and Ce3+ (below) and containing E. canadensis plants (0h: before introducing plants to the solutions;

72h: after cultivating plants for 72 hours) (Mean±SD, n=3)

Part of the CeO2 NPs and Ce3+ were removed from the water phase by the plants (Table 4-3). With

increasing concentration, the amount of Ce in the plants also increased. In the case of CeO2 NPs,

concentrations increased from 0.08±0.02 mg Ce/g DM at 0.5 mg/L to 1.84 ± 0.50 mg Ce/g DM at

50 mg/L. In the presence of Ce3+, the Ce content in plant tissue increased from 0.13 ± 0.01 mg Ce/g

DM at 0.5 mg/L to 18.17 ± 6.38 mg Ce/g DM at 50 mg/L. Plants removed more Ce from Ce3+

spiked solutions than from CeO2 NPs’ spiked solutions. For instance, at 50 mg/L, the amount of Ce

y = 0.498x ‐ 0.5662R² = 0.9946

y = 0.7469x + 0.0805R² = 1

0

5

10

15

20

25

30

35

40

45

0 10 20 30 40 50 60

Actual Ceconcentration (mg/L)

Theoretical Ce concentration (mg/L)

CeO2 NPs

Conc.(72h)

Conc.(0h)

y = 0.9458x ‐ 0.2959R² = 0.9994

y = 0.6089x ‐ 1.2939R² = 0.979

051015

20253035

40455055

0 10 20 30 40 50 60

Actual Ce concentration (mg/L)

Theoretical Ce concentration (mg/L)

Ce3+

Conc.(0h)

Conc.(72h)

33

taken up by plants was almost 10 times higher for plants exposed to Ce3+ ions in comparison to

plants exposed to CeO2 NPs.

Table 4- 3 Content of Ce in E. canadensis plants (mg Ce/g DM) after cultivation in CeO2 NPs and Ce3+ solutions

containing different concentrations (0 mg/L, 0.5 mg/L, 1 mg/L, 5 mg/L, 10 mg/L and 50 mg/L) of Ce (Mean±SD, n=3)

(DLCe=0. 0007mg Ce/g DM)

4.1.2. Elodea canadensis plants exposed to Ag NPs and Ag+

Almost no change in biomass is observed after 72 hours’ exposure of E. canadensis to Ag NPs and

Ag+ solutions (Table 4-4).

Table 4- 4 Mass change/initial weight (%) of E. canadensis plants after exposure to different concentrations (0 mg/L,

0.05 mg/L, 0.1 mg/L, 0.25 mg/L, 0.5 mg/L and 1 mg/L) of Ag in Ag NPs and Ag+ solution (Mean±SD, n=3)

0 mg/L 0.05 mg/L 0.1 mg/L 0.25 mg/L 0.5 mg/L 1 mg/L

Ag NPs 0.79±6.22 1.16±1.89 -1.43±1.34 0.08±2.56 0.07±5.16 0.69±2.59

Ag+ 0.79±6.22 -0.71±0.84 2.23±3.81 -0.48±2.02 2.58±1.93 -2.73±2.15

Total nitrogen, total phosphorus and chlorophyll content were measured to assess the impact of

NPs and ions on performance of the plants (Table 4-5). For both Ag NPs and Ag+ spiked solutions,

the content of nitrogen varies between the different concentrations, and the lowest value was found

at a silver concentration of 0.25 mg/L. Generally, with increasing concentration of Ag NPs and Ag+,

the content of Chl a, b and c decreases, especially for Chl c. According to Table 4-5, at a Ag NPs’

concentration of 0.5 mg/L, the Chl c content dropped below the detection limit (DLChl c=0.003 mg/g

FW), while for Ag+, the Chl c content was below the detection limit at concentrations of 0.1 mg/L,

0.5 mg/L and 1 mg/L. The total phosphorus content in plants that were cultivated in Ag NPs

0 mg/L 0.5 mg/L 1 mg/L 5 mg/L 10 mg/L 50 mg/L

CeO2

NPs < DLCe 0.08±0.02 0.18±0.03 0.67±0.01 1.09±0.12 1.84±0.50

Ce3+ < DLCe 0.13±0.01 0.25±0.02 2.03±0.54 4.80±0.40 18.17±6.38

34

solutions did not differ a lot. The average content was 11.84 mg P/g DM with a standard deviation

of 0.25. On the contrary, the phosphorus content of plants cultivated in Ag+ solutions did vary a lot

between different exposure concentrations. The lowest value (9.22 ± 0.80 mg P/g DM) occurred at

0.05 mg/L, while the highest value (14.00 ± 3.31 mg P/g DM) was obtained at 1 mg/L.

Table 4- 5 Content of total nitrogen (N), total phosphorus (P), Chlorophyll a (Chl a) , Chlorophyll b (Chl b) and

Chlorophyll c (Chl c) in E. canadensis plants after being exposed to different concentrations (0 mg/L, 0.05 mg/L, 0.1 mg/L,

0.25 mg/L, 0.5 mg/L and 1 mg/L) of Ag in Ag NPs and Ag+ solutions (For P: Mean±SD, n=3) (DLChl c= 0.003 mg/g FW)

0 mg/L 0.05 mg/L 0.1 mg/L 0.25 mg/L 0.5 mg/L 1 mg/L

Ag

NPs

N (mg N/g DM) 24.4 39.5 43.0 37.8 47.9 42.4

Chl a (mg/g FW) 0.444 0.337 0.345 0.356 0.195 0.256

Chl b (mg/g FW) 0.394 0.225 0.256 0.231 0.124 0.188

Chl c (mg/g FW) 0.052 0.015 0.028 0.009 < DLChl c < DLChl c

P (mg P/ g DM) 7.30±3.25 11.80±0.17 12.20±1.19 11.54±0.14 11.72±0.20 11.93±1.40

Ag+

N (mg N/g DM) 24.4 30.8 40.8 22.3 38.3 40.4

Chl a (mg/g FW) 0.444 0.435 0.204 0.300 0.220 0.136

Chl b (mg/g FW) 0.394 0.317 0.127 0.209 0.132 0.082

Chl c (mg/g FW) 0.052 0.031 < DLChl c 0.017 < DLChl c < DLChl c

P (mg P/ g DM) 7.30±3.25 9.22±0.80 11.85±1.72 9.85±0.62 12.13±0.82 14.0±3.31

Like with CeO2 NPs and Ce3+ spiked solutions, the actual measured concentrations of Ag in the Ag

NPs and Ag+ treatments (Figure 4-2 - Y-axis) were different from the targeted concentrations

(Figure 4-2 - X-axis). After introducing plants, the concentration of Ag in water phase decreased,

both for solutions with Ag NPs and Ag+. For example, at the targeted concentration of 1 mg/L, the

real initial concentration was 1.12 ± 0.05 mg/L in the case of Ag NPs. After 72 hours, the

concentration dropped to 0.28 ± 0.05 mg/L. While for Ag+, the real initial concentration was 0.80 ±

0.02 mg/L, and then dropped to 0.21 ± 0.03 mg/L after 72 hours.

35

Figure 4- 2 Relationship between theoretical concentration and actual concentration of Ag in solutions spiked with

Ag NPs (above) and Ag+ (below) and containing E. canadensis plants (0h: before introducing plants to the solutions; 72h:

after cultivating plants for 72 hours) (Mean±SD, n=3)

Part of the silver added in both the Ag NPs and Ag+ solutions was removed by the plants (Table

4-6). The amount of silver in plant tissue increased with increasing spike concentrations. When

comparing the same dose, plants exposed to Ag+ contained more silver than those exposed to Ag

NPs. At 1 mg/L, the amount of Ag in plants was 0.712±0.266 mg Ag/g DM when Ag NPs were

added, which is almost 7 times higher than when Ag+ spiked solution was used (0.111±0.008 mg

Ag/g DM).

y = 1.1213x ‐ 0.0046R² = 0.9998

y = 0.244x + 0.0309R² = 0.8865

0.0

0.2

0.4

0.6

0.8

1.0

1.2

0 0.2 0.4 0.6 0.8 1 1.2

Actual Ag concentration (mg/L)

Theoretical Ag concentration (mg/L)

Ag NPs

Conc.(0h)

Conc.(72h)

y = 0.7957x + 0.0072R² = 0.9974

y = 0.2015x + 0.0156R² = 0.9814

0.0

0.2

0.4

0.6

0.8

1.0

0 0.2 0.4 0.6 0.8 1 1.2

Actual Agconcentration (mg/L)

Theoretical Ag concentration (mg/L)

Ag+

Conc.(0h)

Conc.(72h)

36

Table 4- 6 Content of Ag in E. canadensis plants (mg Ag/g DM) after cultivation in Ag NPs and Ag+ solutions

containing different concentrations (0 mg/L, 0.05 mg/L, 0.1 mg/L, 0.25 mg/L, 0.5 mg/L and 1 mg/L) of Ag (Mean±SD, n=3)

(DLAg=0.0005 mg Ag/g DM)

4.2. Desorption experiments

4.2.1. CeO2 NPs and Ce ions

During the desorption experiments, different extraction reagents showed a different extraction

capacity (Figure 4-3). The results were presented as the concentration of Ce in fresh weight of

plants. For Ce3+, 0.5 M HNO3 and 0.02 M EDTA solutions showed a very strong capacity to extract

Ce3+ ions from the plants. After soaking pre-exposed plants for 1h, 0.5 M HNO3 extracts contained

44.4 μg Ce/g FW and 0.02 M EDTA extracts contained 40.3 μg Ce/g FW. Compared to these two

extractants, the others did not have a great capacity to extract Ce3+ ions, with none of them resulting

in a final Ce concentration above 3 μg Ce/g FW.

For CeO2 NPs, a 0.05 mM CMC solution extracted the highest amount (1.16 μg Ce/g FW) of Ce

after soaking the plant for 1h in extractant. The other solutions (0.05 mM PVP solution, Milli-Q

water, 0.05 mM Dextran-70 solution, 0.5 M HNO3 and 0.02 M EDTA) did not show a strong

ability to extract CeO2 NPs from plants. As the results for these extractants varied around the

detection limit, they should be considered as not very reliable.

0 mg/L 0.05 mg/L 0.1 mg/L 0.25 mg/L 0.5 mg/L 1 mg/L

Ag NPs < DLAg 0.023±0.003 0.039±0.003 0.074±0.010 0.070±0.011 0.111±0.008

Ag+ < DLAg 0.026±0.004 0.078±0.015 0.090±0.032 0.440±0.126 0.712±0.266

37

Figure 4- 3 Contents of Ce extracted from E. canadensis plant material using different extractants (Milli-Q water,

0.5M HNO3, 0.02M EDTA, 0.05 mM PVP, 0.05 mM CMC and 0.05 mM Dextran-70); the plants were previously soaked

for 1h (Conc. At T1(1h)) in solutions containing (right) CeO2NPs and (left) Ce3+ (Mean±SD, n=3)

4.2.2. Ag NPs and Ag ions

The extraction capacity of six extractants for Ag NPs and Ag+ ions was presented in Figure 4-4.

Like in previous experiment, the results were presented as the concentration of Ag in plants’ fresh

weight. The nitric acid solution (0.5 M HNO3) extracted significantly higher Ag+ amounts from the

plants (1.00 μg Ag /g FW) than the other reagents (Milli-Q: 0.38 μg Ag /g FW, 0.02 M EDTA:

0.49 μg Ag /g FW, 0.05 mM PVP: 0.43 μg Ag /g FW, 0.05 mM CMC: 0.52 μg Ag /g FW, 0.05

mM Dextran-70: 0.45 μg Ag /g FW).

For Ag NPs, generally 0.05 mM PVP solution and 0.5 M HNO3 solution have a higher extraction

capacity with the extracted amount being 0.23 μg Ag/g FW and 0.19 μg Ag/g FW, respectively.

The remaining three solutions have a similar extraction capacity for Ag NPs with a concentration of

around 0.10 μg Ag/g FW.

38

Figure 4- 4 Contents of Ag extracted from E. canadensis plant material using different extractants (Milli-Q, 0.5 M

HNO3, 0.02 M EDTA, 0.05 mM PVP, 0.05 mM CMC and 0.05 mM Dextran-70); the plants were previously soaked for 1h

(Conc. At T1(1h)) in solutions containing (right) Ag NPs and (left) Ag+ ions (Mean±SD, n=3)

4.3. Impact of water composition on NPs’ uptake by E. canadensis

4.3.1. Characterization of different (surface) water samples

Different waters were employed in this part of the study. These waters were characterized as shown

in Table 4-7. The 10% Hoagland’s solution had the lowest EC value which was 108 μs/cm, and

water from Grote Geul had the highest EC value of 2240 μs/cm. The EC values for tap water, water

sampled from Mostbeek and Coupure Canal were 614 μs/cm, 711 μs/cm and 871 μs/cm,

respectively. The 10% Hoagland’s solution turned out to be slightly acid with a pH of 5.43. On the

contrary, all other waters had a pH value higher than 7, of which Coupure water had the highest

(pH 8.92).

Grote Geul had the highest value of TC (72.8 mg/L) and IC (67.15 mg/L), followed by Coupure

water (TC: 56.0 mg/L, IC: 54.09 mg/L). The 10% Hoagland’s solution had the lowest TC (0.7

mg/L) and IC (0.19 mg/L) value. The highest TOC occurred in the water sample from Mostbeek

(10.71 mg/L), followed by water from Grote Geul (5.65 mg/L), while tap water had the lowest

TOC value (0.25 mg/L).

Dry residue in the three surface waters was quite high (Mostbeek: 497 mg/L, Grote Geul: 1274

mg/L, Coupure: 548 mg/L), but it was only 298 mg/L for tap water and 64 mg/L for a 10%

Hoagland’s solution. Total suspended solids (TSS) showed the same tendency with high values for

39

the sampled surface waters (Mostbeek: 13.83 mg/L, Grote Geul: 6.00 mg/L, Coupure: 16.33 mg/L)

and low values for tap water and 10% Hoagland’s solution (both below detection limit).

Table 4- 7 General properties of different water samples used in the experiments (EC:Electrical Conductivity; TC:

Total Carbon; IC: Inorganic Carbon; TOC: Total Organic Carbon; DR: Dry Residue; TSS: Total Suspended Solid)

(Mean±SD, n=3) (DLTSS=0.33mg/L)

Tap water 10%

Hoagland’s Mostbeek Grote Geul Coupure

EC (μs/cm) 614 ±1.00 108±0.15 711±1.00 2240±0.01 871±1.00

pH 7.72±0.09 5.43±0.20 8.05±0.19 8.62±0.10 8.92±0.09

TC (mg/L) 35.2±0.1 0.7±0.1 30.4±0.1 72.8±1.0 56.0±1.2

IC (mg/L) 34.99±0.14 0.19±0.01 19.64±0.05 67.15±0.06 54.09±0.02

TOC (mg/L) 0.25±0.17 0.54±0.09 10.71±0.17 5.65±0.88 1.91±1.22

DR (mg/L) 298±10 64±10 497±8 1274±17 548±26

TSS (mg/L) <DLTSS <DLTSS 13.83±0.58 6.00±2.00 16.33±0.29

The concentrations of anions present in the water samples are presented in Table 4-8. Grote Geul

water had the highest concentrations of F- (0.40 mg/L), Cl- (489.5 mg/L), NO2- (4.19 mg/L) and

NO3- (36.4 mg/L) among the 5 waters. The amounts of F- and NO2

- of the other four water samples

were below the detection limit. The 10% Hoagland’s solution had the least amount of Cl-, and

Mostbeek had the least amount of NO3-. The amount of PO4

3- could only be detected in the 10%

Hoagland’s solution, where it was 2.26 mg/L. Mostbeek water samples had the highest value of

SO42- which was 192.3 mg/L, followed by Grote Geul water samples (97.9 mg/L), Coupure water

(76.6 mg/L) and tap water (68.1 mg/L). The 10% Hoagland’s solution had the lowest value of SO42-

which was 9.8 mg/L.

40

Table 4- 8 Concentration of anions in the different water samples used in the experiments (Mean±SD, n=3)

Tap water

10%

Hoagland’s Mostbeek Grote Geul Coupure

F- (mg/L) < DL (0.25) < DL (0.25) < DL (0.25) 0.40±0.01 < DL (0.25)

Cl- (mg/L) 34.2±2.9 2.3±1.0 46.2±0.2 489.5±3.4 78.6±1.2

NO2- (mg/L) < DL (0.125) < DL (0.125) < DL (0.125) 4.19±0.28 < DL (0.125)

NO3- (mg/L) 21.9±1.1 36.4±0.2 13.6±0.9 36.4±0.8 26.7±0.6

PO43- (mg/L) < DL (1) 2.26±0.06 < DL (1) < DL (1) < DL (1)

SO42- (mg/L) 68.1±0.6 9.8±0.1 192.3±1.2 97.9±0.6 76.6±0.3

In general, water samples from Mostbeek, Grote Geul and Coupure had higher Ca, Mg, K and Na

concentrations than tap water and 10% Hoagland’s solution (Table 4-9). The highest values of Ca,

Mg, K and Na were found in water from Mostbeek (113.6 mg Ca/L), Coupure (10.69 mg Mg/L),

Grote Geul (16.20 mg K/L) and Grote Geul (204.7 mg Na/L), respectively. The lowest values of

Ca, Mg and Na were found in 10% Hoagland’s solution (11.1 mg Ca/L, 2.78 mg Mg/L, <DL (6.25

mg Na/L), respectively). Tap water had the lowest value of K, which was 2.73 mg K/L. Both Ag

and Ce concentrations in the different water samples were below the detection limit of ICP-OES

(DLCe=0.04 mg/L; DLAg=0.03 mg/L).

41

Table 4- 9 Concentrations of Ca, Mg, K, Na, Ag, and Ce in the different water samples used in the experiments

(Mean±SD, n=3) (DLNa=6.25 mg/L; DLCe=0.04 mg/L; DLAg=0.03 mg/L)

Tap water 10%

Hoagland’s Mostbeek Grote Geul Coupure

Ca (mg/L) 81.3±0.4 11.1±0.0 113.6±0.2 103.0±0.1 105.6±0.4

Mg (mg/L) 7.97±0.03 2.78±0.02 9.32±0.10 45.11±0.36 10.69±0.04

K (mg/L) 2.73±0.01 5.90±0.13 8.90±0.24 16.20±0.21 9.55±0.03

Na (mg/L) 22.8±0.6 <DLNa 22.5±0.4 204.7±3.5 43.8±0.7

Ag (mg/L) <DLAg <DLAg <DLAg <DLAg <DLAg

Ce (mg/L) <DLCe <DLCe <DLCe <DLCe <DLCe

4.3.2. Elodea canadensis plants exposed to CeO2 NPs and Ce3+

After cultivating the plants for 72 hours, the mass difference/initial weight (fresh weight) was

measured (Table 4-10). Generally, without any exposure to CeO2 NPs, Ag NPs, Ce3+ or Ag+, plants

appear to have grown faster in surface water (Mostbeek, Grote Geul and Coupure) than in tap water

and nutrient solution. The same situation was seen even after the introduction of CeO2 NPs and

Ce3+ to the water samples. In Coupure water, plants have grown most significantly after 72 hours

(Control group: 5.42%, CeO2 NPs: 6.15%, Ce3+: 8.85%). Generally, plants grown in presence of

CeO2 NPs showed a higher mass difference than those grown in Ce3+ solution. For example, in 10%

Hoagland’s solution, plants in CeO2 NPs on average increased 2.10% while in Ce3+ solution, on

average it was -2.45%.

42

Table 4- 10 Mass change/initial weight (%) of E. canadensis plants after cultivation for 72 hours in different waters

(Tap water, 10% Hoagland's solution, Mostbeek water, Grote Geul water, Coupure water) spiked with either CeO2 NPs

or Ce3+ (1 mg/L Ce). Control group samples were not exposed to NPs nor ions (Mean values ± SD, n = 3)

Tap water 10%

Hoagland’s Mostbeek Grote Geul Coupure

Control group -0.89±0.39 1.06±2.57 3.11±1.34 3.20±0.09 5.42±0.66

CeO2 NPs 1.26±3.01 2.10±1.88 3.10±2.68 5.48±3.34 6.15±3.23

Ce3+ 0.48±1.38 -2.45±0.30 -0.14±1.41 3.67±2.54 8.85±5.17

The properties of plants also differed under the different exposure conditions (Table 4-11). When

there is no exposure to CeO2 NPs, Ag NPs, Ce3+ and Ag+, plants that were grown in Grote Geul

water had the lowest total nitrogen (33.3 mg N/g DM) and total phosphorus (8.9 mg P/g DM)

content, but the highest chlorophyll content (Chl a: 0.269 mg/g FW, Chl b: 0.177 mg/g FW, Chl c:

< DLChl c). In general, the plants that were cultivated in surface water had a higher chlorophyll

content but lower nitrogen and phosphorus content than those grown in tap water or nutrient

solution.

When the plants were immersed in 1 mg/L CeO2 NPs and 1 mg/L Ce3+ solutions, total nitrogen and

total phosphorus content did not fluctuate much, and were generally higher in tap water and

nutrient solution. In presence of CeO2 NPs and Ce3+, plants grown in Coupure water had the

highest chlorophyll content, and overall, the chlorophyll content was higher in surface water than in

tap water or nutrient solution.

43

Table 4- 11 Content of total nitrogen (TN), total phosphorus (TP), Chlorophyll a (Chl a) , Chlorophyll b (Chl b) and

Chlorophyll c (Chl c) in E. canadensis plants after being exposed to 1mg/L CeO2 NPs and 1mg/L Ce3+in Tap water, 10%

Hoagland’s solution, Mostbeek, Grote Geul and Coupure (Mean±SD, n=3) (DLChl c= 0.003 mg/g FW)

Tap water 10%

Hoagland’sMostbeek Grote Geul Coupure

Control

Group

TN (mg N/g DM) 46.0 46.2 50.4 33.3 38.4

Chl a (mg/g FW) 0.164 0.157 0.195 0.269 0.191

Chl b (mg/g FW) 0.088 0.095 0.113 0.177 0.113

Chl c (mg/g FW) < DLChl c < DLChl c < DLChl c < DLChl c < DLChl c

TP (mg P/ g DM) 12.2±0.162 13.9±0.422 13.2±0.644 8.9±0.071 10.9±0.617

CeO2

NPs

TN (mg N/g DM) 36.7 49.3 41.7 36.9 44.2

Chl a (mg/g FW) 0.144 0.171 0.259 0.172 0.334

Chl b (mg/g FW) 0.094 0.113 0.187 0.111 0.257

Chl c (mg/g FW) < DLChl c < DLChl c < DLChl c < DLChl c < DLChl c

TP (mg P/ g DM) 13.1±0.705 14.7±0.602 13.5±1.212 10.2±0.788 10.4±0.364

Ce3+

TN (mg N/g DM) 41.8 44.6 41.1 33.3 38.6

Chl a (mg/g FW) 0.195 0.241 0.222 0.265 0.288

Chl b (mg/g FW) 0.104 0.138 0.119 0.148 0.160

Chl c (mg/g FW) < DLChl c < DLChl c < DLChl c < DLChl c < DLChl c

TP (mg P/ g DM) 11.3±0.921 12.8±1.741 11.8±1.161 9.6±0.410 9.8±0.167

To reveal the kinetics of Ce removal by E. canadensis in the different waters, the amount of Ce was

measured at certain time intervals (0h, 2h, 6h, 10h, 24h, 48h and 72h) for each kind of water

(Figure 4-5). After spiking 10% Hoagland’s solution with 1 mg/L Ce3+, the Ce concentration

decreased sharply from 0.867 mg/L to 0.326 mg/L within the first 2 hours, after which it slowly

decreased to 0.058 mg/L within the next 22 hours. In the end, only 0.031 mg Ce/L was left in the

water phase. For the other four water samples, the concentration of Ce decreased slowly during a

lag phase of about 10 hours, but after 10 hours it started to decrease quickly until 72 hours.

44

In water samples spiked with 1 mg/L CeO2 NPs, the evolution of the Ce concentration in the water

phase was very similar for all five types of water. During the first 10 hours, the concentration

decreased slowly, but after 10 hours the concentration started to drop quickly, with a final

concentration around 0.05 mg/L after 72 hours.

Figure 4- 5 Evolution of concentration of Ce in different waters (Tap water, 10% Hoagland’s solution, Mostbeek

water, Grote Geul water, Coupure water) spiked with Ce3+ (above) or CeO2 NPs (below) when exposing E. canadensis

plants to these waters (Mean±SD, n=3)

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

1

0 2 6 10 24 48 72

Ce concentration (mg/L)

Time (h)

Ce3+

Tap water

10% Hoagland's

Mostbeek

Grote geul

Coupure

‐0.1

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0 2 6 10 24 48 72

Ce concentration (mg/L)

Time (h)

CeO2 NPs

tap water

10% Hoagland's

Mostbeek

Grote geul

Coupure

45

4.3.3. Elodea canadensis plants exposed to Ag NPs and Ag ions

Compared to the mass changes/initial weight obtained from exposure to CeO2 NPs and Ce3+, the

same situation can be found in the exposure to Ag NPs and Ag+ solution (Table 4-12), which

means plants that were grown in surface waters tend to show a higher mass change than those

grown in tap water or nutrient solution. In addition, plants exposed to 0.1 mg/L Ag NPs have a

higher mass difference than those exposed to Ag+ at the same concentration. For example, in Grote

Geul water, the mass of plants exposed to 0.1 mg/L Ag NPs increased by 8.79%; however, in Ag+

solution, on average the plants’ mass decreased by 1.68%.

Table 4- 12 Mass change/initial weight (%) of E. canadensis plants after cultivation for 72 hours in different waters

(Tap water, 10% Hoagland's solution, Mostbeek water, Grote Geul water, Coupure water) spiked with either Ag NPs or

Ag+ (0.1mg/L Ag). Control group samples were not exposed to NPs nor ions (Mean values ± SD, n = 3)

Tap water 10%

Hoagland’s Mostbeek Grote Geul Coupure

Control group -0.89±0.39 1.06±2.57 3.11±1.34 3.20±0.09 5.42±0.66

0.1 mg/L Ag NPs 3.93±1.82 -0.33±3.77 -0.75±3.05 8.79±6.15 3.61±6.15

0.1 mg/L Ag+ ions -0.86±3.06 -1.76±1.33 0.46±3.69 -1.68±1.00 0.09±3.75

Nitrogen, chlorophyll and phosphorus contents in the plants are presented in Table 4-13. For plants

exposed to 0.1 mg/L Ag NPs, the content of nitrogen and phosphorus did not deviate too much

from the average of 40.4±2.4 mg N/g DM and 11.4±0.5 mg P/g DM. Plants that were immersed in

Mostbeek water have the highest chlorophyll content (Chl a: 0.337 mg/g FW, Chl b: 0.261 mg/g

FW, Chl c: <DLChl c). However, chlorophyll contents in plants from Grote Geul water and Coupure

water were very low, even lower than those from plants grown in tap water and nutrient solution.

Plants grown in surface water containing 0.1 mg/L Ag+, contain less nitrogen and phosphorus than

those grown in tap water and 10% Hoagland’s solution with the same Ag+ concentration. The

chlorophyll content of plants grown in surface water were generally higher than those of plants

grown in tap water and nutrient solution, but chlorophyll contents were still quite low for plants

grown in water from Grote Geul.

46

Table 4- 13 Content of total nitrogen (N), total phosphorus (P), Chlorophyll a (Chl a), Chlorophyll b (Chl b) and

Chlorophyll c (Chl c) in E. canadensis plants after being exposed to Tap water, 10% Hoagland’s solution, Mostbeek,

Grote Geul and Coupure water to which Ag NPs or Ag+ions were spiked at a concentration of 0.1 mg Ag/L (Mean±SD,

n=3) (DLChl c= 0.003 mg/g FW)

Tap water 10%

Hoagland’s Mostbeek Grote Geul Coupure

Control

group

N (mg N/g DM) 46.0 46.2 50.4 33.3 38.4

Chl a (mg/g FW) 0.164 0.157 0.195 0.269 0.191

Chl b (mg/g FW) 0.088 0.095 0.113 0.177 0.113

Chl c (mg/g FW) < DLChl c < DLChl c < DLChl c < DLChl c < DLChl c

P (mg P/ g DM) 12.2±0.1 13.9±0.4 13.2±0.6 8.9±0.1 10.9±0.6

Ag

NPs

N (mg N/g DM) 39.5 40.1 42.0 37.1 43.4

Chl a (mg/g FW) 0.180 0.044 0.337 0.034 0.037

Chl b (mg/g FW) 0.118 0.042 0.261 0.029 0.035

Chl c (mg/g FW) < DLChl c < DLChl c < DLChl c < DLChl c < DLChl c

P (mg P/ g DM) 11.1±0.3 11.5±1.3 12.1±0.9 11.2±1.1 11.0±1.0

Ag+

ions

N (mg N/g DM) 50.2 43.6 32.0 39.4 37.8

Chl a (mg/g FW) 0.048 0.282 0.231 0.066 0.201

Chl b (mg/g FW) 0.035 0.177 0.146 0.056 0.125

Chl c (mg/g FW) < DLChl c < DLChl c < DLChl c < DLChl c < DLChl c

P (mg P/ g DM) 15.7±0.4 16.3±0.8 14.2±2.0 14.2±0.3 12.3±0.6

To study the kinetics of Ag removal, the concentration of Ag in the water samples was also

measured at 0h, 2h, 6h, 10h, 24h, 48h and 72h (Figure 4-6). After exposing the plants to water

containing Ag+, the Ag concentration in 10% Hoagland’s solution dropped dramatically in the first

2 hours, from 0.105 mg/L to 0.031 mg/L after 2h. After this rapidly decrease, the concentration

decreased slightly to 0.010 mg/L during the next 70 hours. The other four kinds of water followed a

similar trend, although the initial drop in concentration was less pronounced. A rapid decrease to

half of the initial concentration can be observed in the first 6 hours, and a slower decrease

afterwards until the end of the experiment (72 hours). When Ag NPs were used, the concentration

47

of Ag did not change too much within the first 10 hours, but it started to drop quickly in the next 62

hours.

Figure 4- 6 Evolution of concentration of Ag in different waters (Tap water, 10% Hoagland’s solution, Mostbeek

water, Grote Geul water, Coupure water) spiked with Ag+ (above) or Ag NPs (below) when exposing E. canadensis plants

to these waters (Mean±SD, n=3)

0

0.02

0.04

0.06

0.08

0.1

0.12

0 2 6 10 24 48 72

Ag concentration  (mg/L)

Time (h)

Ag+

Tap water

10%Hoagland's

Mostbeek

Grote geul

Coupure

0

0.02

0.04

0.06

0.08

0.1

0.12

0 2 6 10 24 48 72

Ag concentration (mg/L)

Time (h)

Ag NPs

Tap water

10%Hoagland'sMostbeek

Grote geul

Coupure

48

4.4. Wetland mesocosm experiment

4.4.1. Selection of time point for further data analysis

A summary of the rate of microbial development following BIOLOG plate inoculation is shown in

Figure 4-7. For all samples, the average well color development (AWCD) and the number of

absorbance values over 2 both increased over the monitoring period following plate inoculation.

During the first 48 h of incubation on the plate, AWCD values seem to be low and stable, and

almost no well has an absorbance value over 2. However, after 48 hours, AWCD started to increase,

reaching the highest values at the end of the monitoring period (115 h). With increasing AWCD,

the number of absorbance values over 2 also increased. It also reaches its highest value after 115 h.

Based on the principle that a time point that can be used for analysis should be a time point at

which the AWCD is relatively high while the number of absorbance values over 2 is low, 65 h was

chosen as the time point for data analysis in the further experiments with BIOLOG plates.

Figure 4- 7 Summary of the microbial development (A=AWCD, B= number of absorbance values over 2) over a

monitoring period of 115 h after inoculation of a Biolog plate of a wetland mesocosm (from interstitial wastewater)

containing 0.1 mg/L Ag supplied as Biogenic Ag NPs, Citrate Ag NPs, PVP Ag NPs and Ag+ with a blank (no Ag) as

control group.

4.4.2. Response of wetland microbial communities on different doses of Ag NPs

A series of concentrations (0.5 mg/L, 1 mg/L, 2 mg/L and 5 mg/L) of Citrate and PVP coated Ag

NPs, Biogenic Ag NPs and Ag+ were used to examine their potential impact on microbial

communities commonly present in wetlands. These communities were sampled from a wetland

microcosm setup established in the laboratory, in which the communities were exposed to the

49

different NPs and ions in the different concentrations. A control group which was not exposed to

any form of silver during the microcosm experiment was also included to assess whether resistance

of the community against the silver was built up in the exposed communities during the mesocosm

experiment (Figure 4-8).

In the dose-response curve of the microbial communities from interstitial water that was previously

exposed to 0.1 mg/L citrate coated Ag NPs (Figure 4-8, A-Citrate Ag NPs), the average well color

development (AWCD) decreased rapidly from 1.09 (at 0 mg/L) to 0.35 (at 5 mg/L) with increasing

citrate Ag NPs’ concentration, while in the control group (Figure 4-8, A-Control + Citrate Ag NPs),

it decreased from 0.97 to 0.17.

For PVP coated Ag NPs, the AWCD value remained stable in both the control group (Figure 4-8,

B-Control + PVP Ag NPs) and the previously exposed group (Figure 4-8, B-PVP Ag NPs).

However, a slight decrease was observed for the control group at the highest dose (5 mg/L).

In the case of biogenic Ag NPs, AWCD dropped quickly to 0.01 (at 1 mg/L) from 1.01 at 0 mg/L,

but remained constant and low with further increase in silver concentration after prior exposure

(Figure 4-8, C-Biogenic Ag NPs). The control group (Figure 4-8, C-Control + Biogenic Ag NPs)

showed a similar tendency, with a sharp decrease from 0.97 at 0 mg/L to 0.02 at 1 mg/L. With

doses increasing above 1 mg/L, the AWCD value kept stable around 0.

Finally, after spiking with Ag+, the AWCD in both the control group (Figure 4-8, D-Control + Ag+)

and the sample group (Figure 4-8, D- Ag+) showed a downward trend with increasing exposure

concentrations. When the concentration in the solution that was previously exposed to Ag+

increased from 0 mg/L to 1 mg/L, a sharp drop from 1.11 at 0 mg/L to 0.01 at 1 mg/L can be

observed. At higher doses, the AWCD value varied around 0 with increasing concentration of Ag+.

The control group experienced slightly lower values of AWCD at the lowest doses. The AWCD

value in the control group quickly dropped from 0.97 (at 0 mg/L) to 0.01 (at 1 mg/L), and remained

low at higher doses.

50

Generally, at the same Ag NPs or Ag+ concentration, AWCD values of the control groups were

always lower than those mesocosms that were previously exposed for 4 weeks to low

concentrations (0.1 mg/L) of Ag+ or Ag NPs.

Figure 4- 8 Response (AWCD) of microbial communities of interstitial water of a wetland mesocosm on different

concentrations (0.5 mg/L, 1 mg/L, 2 mg/L and 5 mg/L) of (A) Citrate-Ag NPs, (B) PVP-Ag NPs, (C) Biogenic-Ag NPs and

(D) Ag+ after 65 h of exposure (the control group was not exposed to silver when cultivating the microbial community in

wastewater prior to exposure) (Mean±SD, n=3)

4.4.3. Carbon-Source Utilization Patterns (CSUPs)

A principal component analysis (PCA) was performed to compare the CSUPs for each treatment.

Data were assessed for normality and homoscedasticity, and normalized using a Taylor

transformation where necessary.

In Figure 4-9, data from the interstitial water at the time point of 65 h are presented. These

microbial communities were only exposed to 0.1 mg/L Ag during the previous cultivation for 4

weeks, and not anymore exposed during the incubation on the Biolog plates. The PCA ordination

shows that no clear groups can be defined among the different mesocosms exposed to Ag NPs

having different coatings and Ag+ at 0.1 mg/L.

51

Figure 4- 9 PCA using carbon-source utilization data (CSUPs) of a control mesocosm (not exposed to Ag) and

mesocosms exposed to citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs and Ag+ at 0.1 mg/L; mesocosms from the

interstitial water were set up in triplicate (A-C). Output generated using XLSTAT 2013.

When stock solutions of citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs and Ag+ were added to the

mesocosms of interstitial water which have been exposed to 0.1 mg/L Ag during previous

incubation in order to reach the concentrations of 0.5 mg/L, 1 mg/L, 2 mg/L and 5 mg/L Ag, the

PCA ordination of both the sample group and control group changed (Figure 4-10 and Figure 4-11).

In Figure 4-10, mesocosms exposed to citrate-coated Ag NPs and PVP-coated Ag NPs group well

with each other and with the biofilm group exposed to 0.1 mg/L citrate-coated Ag NPs, biogenic

Ag NPs, PVP-coated Ag NPs and Ag+ solution. However, mesocosms exposed to biogenic Ag NPs

and Ag+ solutions did not group well with the other mesocosms. Even those exposed to the same

compound at different concentrations differed a lot.

Control A

Control B

Control C

Biogenic A

Biogenic B

Biogenic C

Citrate A

Citrate B

Citrate C

Ag A

Ag B

Ag C

PVP A

PVP B

PVP C

‐1

‐0.5

0

0.5

1

1.5

2

‐1.5 ‐1 ‐0.5 0 0.5 1 1.5 2 2.5 3

F2 (18.01 %)

F1 (30.95 %)

Observations (axes F1 and F2: 48.96 %)

52

Figure 4- 10 CLPP of interstitial water from wetland mesocosms exposed to 0.5 mg/L, 1 mg/L, 2 mg/L and 5 mg/L

citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs and Ag+; the biofilms were previously exposed to 0.1 mg/L citrate Ag NPs,

biogenic Ag NPs, PVP Ag NPs, Ag+ and no Ag as control group. Output generated using XLSTAT 2013.

When samples of the control mesocosms that were not exposed to Ag during growth in the

mesocosm were exposed to citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs and Ag+ at

concentrations of 0.5 mg/L, 1 mg/L 2 mg/L and 5 mg/L, the PCA ordination changed again (Figure

4-11). Again, those exposed to citrate Ag NPs and PVP Ag NPs aggregated in the right part of the

plot close to the center, except one exposed to 5 mg/L citrate Ag NPs (Control + Citrate 5), and the

CSUPs from the communities exposed to biogenic Ag NPs and Ag+ shifted to the left part of the

chart. However, in contrast to what was observed for the exposed communities, all controls

exposed to biogenic Ag NPs and Ag+ are now grouped well within the plot.

control biofilm

pvp biofilmbiogenic biofilmAg+ biofilmcitrate biofilm

citrate 0.5

citrate 1

citrate 2

citrate 5

biogenic 0.5

biogenic1

biogenic 2

biogenic 5

PVP 0.5

PVP 1PVP2

PVP 5

Ag 0.5Ag 1

Ag 2

Ag 5

‐1.5

‐1

‐0.5

0

0.5

1

1.5

2

‐2.5 ‐2 ‐1.5 ‐1 ‐0.5 0 0.5 1 1.5 2 2.5 3

F2 (15.19 %)

F1 (48.33 %)

Observations (axes F1 and F2: 63.52 %)

53

Figure 4- 11 PCA using carbon-source utilization data (CSUPs) of interstitial water from control mesocosms

exposed for 65 h to citrate Ag NPs, biogenic Ag NPs, PVP Ag NPs and Ag+ at concentrations of 0.5 mg/L, 1 mg/L, 2 mg/L

and 5 mg/L Ag . The microbial communities were not exposed to Ag during preceding growth in the mesocosm. Output

generated using XLSTAT 2013.

control +Ag 0.5control +Ag 1

control +Ag 2control +Ag 5

control+bio 0.5

control+bio 1

control+bio 2control+bio 5

control+pvp 0.5control+pvp 1control+pvp 2

control+pvp 5

Control+Citrate 0.5

Control+Citrate 1

Control+Citrate 2

Control+Citrate 5‐3

‐2

‐1

0

1

2

‐4 ‐3 ‐2 ‐1 0 1 2 3 4

F2 (17.11 %)

F1 (49.01 %)

Observations (axes F1 and F2: 66.12 %)

54

Chapter 5: Discussion

5.1.Toxicity of NPs and bulk ions on E. canadensis

According to Table 4-1 and Table 4-4, all CeO2 NPs, Ce3+, Ag NPs and Ag+ had an effect on the

growth of E. Canadensis. In general, at low spike concentrations, plant growth was stimulated

while at higher spike concentrations, the growth of the plants was inhibited.

5.1.1. Toxicity of NPs and bulk ions on E. canadensis

Both nitrogen and phosphorus are essential elements in plants’ growth, and may relate to cell

division and the synthesis of chemicals that are important to plants (Jiang, Li et al. 2012). The

content of nitrogen and phosphorus in plants is considered as one of the most likely factors that can

limit the plant growth, especially in the case of aquatic macrophytes (Gerloff and Skoog 1954). As

a result, nitrogen and phosphorus content are used as an indicator of the degree of toxicity.

In Figure 5-1, total nitrogen (TN) seemed to be more stable in plants exposed to NPs compared to

those exposed to ions. In the presence of Ce3+, total nitrogen continued decreasing until the Ce3+

concentration reached 10 mg/L, with a significant increase at 50 mg/L Ce3+. In the case of Ag+,

total nitrogen in plants started to increase after 0.25 mg/L Ag+. However, as there was only one

sample plant used for total nitrogen determination, a standard deviation is not available here. In this

case, the total nitrogen results could be biased.

No significant difference was found in the total phosphorus content in plants with increasing

concentration of CeO2 NPs and Ag NPs, when compared to the control group (total phosphorus:

7.3 mg P/g Dry wt) (Table 4-2; Table 4-5). Even for Ce3+ solution, total phosphorus content

remained quite stable when the concentration of Ce3+ increased from 0.5 mg/L to 10 mg/L.

However, a significant reduction was observed at 50 mg/L Ce3+ with only 75% of the phosphorus

remaining compared to the control group. Total phosphorus content in plants exposed to Ag+

showed a similar trend as the observed change in total nitrogen content, i.e., the content kept

55

increasing from 0.25 mg/L Ag+. But again, results could be biased as standard deviations are not

available.

Figure 5- 1 Dose-response of total nitrogen (a: exposed to CeO2 NPs and Ce3+, b: exposed to Ag NPs and Ag+) and

total phosphorus (c: exposed to CeO2 NPs and Ce3+, d: exposed to Ag NPs and Ag+) in E. canadensis over 72 hours’

cultivation

Photosynthesis is the basic progress that determines the productivity of green plants. Furthermore,

the photosynthetic rate of plants can be affected by the chlorophyll content (Nekrasova, Ushakova

et al. 2011).

As shown in Figure 5-2, when plants were exposed to CeO2 NPs and Ce3+, generally the content of

Chl a, b and c kept increasing from low (0.5 mg/L) to relatively high (10 mg/L). Overall, Chl a

increased up to 105% of the control level, Chl b increased up to 88-102% of the control level and

Chl c increased to 70-130% of the control level. At 50 mg/L of CeO2 NPs, Chl a, b and c dropped

to 84%, 83% and 110% of the control level, respectively. While for Ce3+, Chl a, b and c dropped to

102%, 77% and 17% of the control level.

56

In contrast, when plants were exposed to Ag NPs and Ag+, the content of Chl a, b and c in plants

kept decreasing with the increasing silver concentration. In Ag+ solution, Chl a, b and c was 98%,

80% and 60% of the control level at lowest concentration (0.05 mg/L) but dropped to 31%, 21%

and 0% at the highest concentration (1 mg/L). As presented in Figure 5-2, there was almost no Chl

c in plants when Ag+ reached 0.1 mg/L and Ag NPs reached 0.5 mg/L. In the presence of Ag NPs,

Chl a, b and c was 80%, 65% and 54% of the control level at lower concentration (0.25 mg/L) and

then reduced to 44%, 31% and 0 at higher concentration (0.5 mg/L). On the whole, content of Chl a,

b and c was more stable in plants exposed to NPs. This is because NPs’ toxicity to E. canadensis

mainly depends on the released ions. Zhao and Wang et al. (2011) also suggested Ag NPs toxicity

to D. magna is influenced by the released Ag ions. In addition, Chl c is more sensitive to the change

of Ce3+ and Ag+ compared to Chl a and b. The concentrations of Chl c here and also in later

chapters were very low and statically have negative values; this may be attributed to the fact that

the amount of Chl c is under the detection limit of the equipment.

The increasing values in chlorophyll pigments in the presence of CeO2 NPs and Ce3+ indicated that

low concentrations of CeO2 NPs and Ce3+ could stimulate the growth of E. canadensis. While, at

high concentrations (50 mg/L) the plants’ growth was inhibited, especially for Ce3+. If increase the

CeO2 NPs’ concentration, further inhibition could be expected. In general, Chl a, b and c in plants

that have been in contact with Ag NPs and Ag+, kept decreasing, which means that both Ag NPs

and Ag+ are toxic to plants even at very low concentrations (0.05 mg/L). From this point of view, it

is clear that Ag NPs and Ag+ are more toxic than CeO2 NPs and Ce3+ for E. canadensis as plants

exposed to Ag NPs and Ag+ had lower chlorophyll contents. Gaiser, Fernandes et al. (2009) also

concluded that Ag NPs are more toxic than CeO2 NPs for the aquatic invertebrate D. magna, and

are more cytotoxic to both trout primary hepatocytes and human hepatocyte cell line in vitro.

Overall, the accumulation of NPs and ions in E. canadensis can cause a decrease in chlorophyll

content, and also a slight decrease of total nitrogen and total phosphorus content. Moreover, the

decrease in nitrogen and phosphorus could further inhibit the plants growth (Gerloff and Skoog

1954) as they are essential elements in chlorophyll pigments synthesis. As a result, the

accumulation of NPs and ions in plants turns out to inhibit the photosynthetic rate and decrease the

biochemical reactions, and in the end, may cause the death of the plants.

57

Figure 5- 2 Dose-response of chlorophyll a (Chl a) (a: exposed to CeO2 NPs and Ce3+, b: exposed to Ag NPs and

Ag+), chlorophyll b (Chl b) (c: exposed to CeO2 NPs and Ce3+, d: exposed to Ag NPs and Ag+) and chlorophyll c (Chl c) (e:

exposed to CeO2 NPs and Ce3+, f: exposed to Ag NPs and Ag+) in E. canadensis after 72hours’ exposure (at 0.1 mg/L Ag+,

0.5 mg/L Ag NPs, 0.5 mg/L Ag+, 1 mg/L Ag NPs and 1 mg/L Ag+, Chl c content was under detection limit of the equipment)

5.1.2. Uptake and adsorption of plants

Six different solutions (Milli-Q, 0.5 M HNO3, 0.02 M EDTA, 0.05 mM PVP, 0.05 mM CMC and

0.05 mM Dextran-70) were employed to examine the amount of NPs and ions that have been

adsorbed on plants’ surface. Results in Figure 4-3 showed that only a small amount of NPs and ions

58

that were brought into contact with the plants during one hour could be removed from the plants

again by means of the applied extractants, which means that most of the removed NPs and ions

were either taken up by the plants and accumulated in tissues, or were irreversibly sorbed onto the

plants surface. The extracted NPs’ amount was lower than the extracted ions’ amount. Due to their

more neutral charge, the particles may enter the plants more easily or sorb stronger to the surface of

the plants.

5.1.3. Toxicity of NPs and bulk ions on E. canadensis in different water bodies

Tap water, 10% Hoagland’s nutrient solution and three kinds of surface waters (Mostbeek, Grote

Geul, and Coupure canal) were used as basic cultivation media, to examine whether water

composition can affect the removal efficiency of NPs and ions from the water phase. According to

Figure 5-3, water composition has an effect on the content of total nitrogen, total phosphorus and

chlorophyll pigments.

59

Figure 5- 3 Effect of water composition on the content of total nitrogen, total phosphorus, chlorophyll a and

chlorophyll b in E. canadensis plants upon exposure to 1 mg/L CeO2 NPs, 1 mg/L Ce3+, 0.1 mg/L Ag NPs, 0.1 mg/L Ag+

(Control group: without any exposure of NPs and ions)

In Figure 5-3, no difference in nitrogen and phosphorus can be distinguished and according to

Table 4-10 and Table 4-12, the mass change before and after the cultivation did not increase or

decrease too much; in this case, the toxicity of NPs and ions is mainly determined by changes in the

chlorophyll content. Based on the content of Chl a and b (Chl c was under detection limit (0.003

mg/g FW)), it can be concluded that the 0.1 mg/L Ag+ solution was the most toxic compound in the

tap water group with only 29.5% Chl a and 39.6% Chl b of the control level remaining, and an

average fresh weight loss of -0.86% (Table 4-12). This result supports the conclusion that Ag+ is

more toxic than CeO2 NPs and Ce3+ which was already concluded from the dose-response

experiment.

In the 10% Hoagland’s solution, 0.1 mg/L Ag NPs turned out to be the most toxic for E.

canadensis. With exposure to 0.1 mg/L Ag NPs, the content of Chl a and b dropped dramatically,

with only 27.8% and 43.9% of the value of the control group remaining. This is because the small

60

particle size (10 nm) of Ag NPs allows the particles to pass through cell membranes and

accumulate, which in turn can lead to cell malfunction (Choi, Deng et al. 2008) and can further

inhibit plants’ growth. The reduction of growth can partially also be explained by the extremely

low IC values, which meant that plants did not have enough carbon sources for their growth. Also,

the 10% Hoagland’s solution provided an acid environment which could not be suitable for the

growth of plants and further inhibit their growth.

In Mostbeek water, all added NPs and ions turned out to stimulate the growth of E. canadensis,

with 0.1 mg/L Ag NPs having the most obvious effects. The Chl a and b contents were 172.3% and

229.9% of the control level, respectively. At 0.1 mg/L Ag+, E. canadensis’s Chl a content was

118.2% while Chl b was 129.0% of the control level. In the presence of 1 mg/L CeO2 NPs, Chl a

and b contents increased to 132.6% and 164.5% of the control level, and at 1 mg/L Ce3+, Chl a and

b levels increased to 113.8% and 104.7%.

On the contrary, 1 mg/L Ce3+, 1 mg/L CeO2 NPs, 0.1 mg/L Ag+ and 0.1 mg/L Ag NPs all had a

negative effect on the growth of E. canadensis when considering water sampled from Grote Geul.

Ag NPs seemed to be the most toxic in this scenario, with 12.7% Chl a and 16.4% Chl b of the

control value. The toxicity varied from highest to lowest in Grote Geul water as follows: Ag NPs >

Ag+ > CeO2 NPs > Ce3+.

In Coupure water, 1 mg/L CeO2 NPs showed a positive effect on the growth of E. canadensis. In

this culture medium, Chl a increased to 174.9% of the control value while Chl b increased to 227.0%

of the control value. Ag NPs at a concentration of 0.1 mg/L showed a strong inhibition effect on the

plants’ growth through the reduction of Chl a and b contents, which were 19.2% and 30.9% of the

control level, respectively.

Overall, E. canadensis turned out to be more sensitive to NPs and ions present in surface water, as

they induced more significance changes in content of chlorophyll pigments in these media. This

might be due to the surface water have higher values of IC, anions etc. than tap water and 10%

Hoagland’s solution providing more carbon and nutrients for plants’ growth. Moreover, it is also

clear that the toxicity of NPs and their ionic forms varies widely with water composition. It is

61

possible that the different kinds of ions (Cl-, SO42- etc.) occurring in water phase can affect the

actual concentration of NPs and their ions, available for the plants. For example, in presence of high

concentrations of Cl-, part of the Ag+ could react with Cl- and precipitate so that the potential

toxicity of Ag+ is decreased. Moreover, the toxicity does not only depend on the plants’ species

(Oukarroum, Polchtchikov et al. 2012) but also on the type of element. For example, Ag NPs at a

lower concentration (0.1 mg/L) showed a higher toxicity than 1 mg/L CeO2 NPs.

5.2. Mass balance calculations

5.2.1. Mass balance in tap water

Mass loss always occurred during the experiments (Table 5-1 and 5-2).The actual mass (calculated

from the measured concentration through ICP-OES/ICP-MS) in the water phase varied from 28.7%

to 100% of the theoretical mass (calculated from the theoretical concentration) content. This loss of

mass could be caused by loss during gentle rinsing of the plants with distilled water after 72 h

cultivation, sorption to recipients and/or deviations from the theoretical mass due to spiking errors.

Generally, after 72 h cultivation, plants grown in solutions spiked with ions contained higher

amounts of Ag and Ce compared to plants grown in solutions containing NPs. This is possibly

because ions usually have smaller diameters (<2 nm) than their corresponding NPs (CeO2 NPs: 4

nm, Ag NPs: 10 nm). As a result, ions can pass more easily through cell walls and membranes, and

have a higher chance to be accumulated in plant tissue. However, when the environment becomes

too toxic for plants to survive or the plants accumulate too much toxic compounds in their tissues,

plants will start to degrade the heavily polluted parts. As a result, the accumulated NPs and ions

will be released back into water phase.

62

Table 5- 1 Mass balance of Ce (M2/M1) in tap water under exposure to different concentrations (0.5 mg/L, 1 mg/l, 5

mg/L, 10 mg/L and 50 mg/L) (M2/M1: actual total Ce/theoretical total Ce; M3/M2: total Ce in plants/actual total Ce)

0.5 mg/L 1 mg/L 5 mg/L 10 mg/L 50 mg/L

Ce3+

M2/M1 (%) 76.0 62.2 93.7 100 96.9

M3/M2 (%) 58.4 67.6 76.3 85.0 48.0

CeO2

NPs

M2/M1 (%) 77.7 52.0 70.6 58.5 76.4

M3/M2 (%) 38.7 61.9 28.4 28.5 8.5

Table 5- 2 Mass balance of Ag (M2/M1) in tap water under exposure to different concentrations (0.05 mg/L, 0.1 mg/l,

0.25 mg/L, 0.5 mg/L and 1 mg/L) (M2/M1: actual total Ag/theoretical total Ag; M3/M2: total Ag in plants/actual total Ag)

0.05 mg/L 0.1 mg/L 0.25 mg/L 0.5 mg/L 1 mg/L

Ag+

M2/M1 (%) 100 87.4 81.6 94.0 76.3

M3/M2 (%) 60.2 64.2 17.4 66.9 65.5

Ag

NPs

M2/M1 (%) 91.3 77.6 78.9 28.7 31.3

M3/M2 (%) 34.3 38.1 35.7 29.0 19.0

5.2.2. Mass balance in different aquatic environments

A similar mass loss could also be observed in different aquatic environments (Table 5-3 and 5-4).

Besides explanations given above, the relatively higher mass loss in the different aquatic

environments could also be caused by the formation of precipitates through reactions with the ions

present in the surface waters. For example, Ag+ could react with Cl- to form AgCl precipitates,

hereby reducing the actual Ag content in the water phase as AgCl cannot be easily dissolved even

with acidification by means of HNO3 solution. In addition, the sampled surface waters had high

TSS and DR values, which provides an opportunity for NPs and ions to adsorb onto particulates or

large clusters so that their concentration in water phase could be reduced. In all aquatic

63

environments under study, plants seemed to accumulate more ions than their corresponding NPs,

which support also the conclusions drawn from the previous experiment.

Table 5- 3 Mass balance (M2/M1) in different aquatic environments (Tap water, 10% Hoagland’s solution,

Mostbeek, Grote Geul and Coupure water) upon exposure to 1 mg/L Ce3+ solution and 1 mg/L CeO2 NPs (M2/M1: actual

total mass of Ce/theoretical total mass of Ce; M3/M2: total Ce in plants/actual total Ce)

Tap water 10%

Hoagland’s Mostbeek Grote Geul Coupure

Ce3+

M2/M1 (%) 59.3 79.6 59.9 53.1 52.6

M3/M2 (%) 82.8 95.1 82.1 78.4 76.0

CeO2

NPs

M2/M1 (%) 63.6 66.0 62.5 62.6 59.6

M3/M2 (%) 75.6 74.2 70.2 79.4 70.7

Table 5- 4 Mass balance (M2/M1) in different aquatic environments (Tap water, 10% Hoagland’s solution,

Mostbeek, Grote Geul and Coupure water) upon exposure to 0.1mg/L Ag+ solution and 0.1mg/L Ag NPs (M2/M1: actual

total mass of Ag/theoretical total mass of Ag; M3/M2: total Ag in plants/actual total Ag)

Tap water 10%

Hoagland’s Mostbeek Grote Geul Coupure

Ag+

M2/M1 (%) 71.9 68.5 69.2 64.5 72.1

M3/M2 (%) 70.8 82.2 69.0 59.9 72.7

Ag

NPs

M2/M1 (%) 62.1 54.7 63.6 56.4 57.4

M3/M2 (%) 48.6 52.9 34.6 43.2 49.1

5.3. Effects of Ag NPs and Ag+ on microbial communities

The overall value of AWCD from highest to lowest follows the sequence PVP Ag NPs > Biogenic

Ag NPs > Citrate Ag NPs > Ag+ (Figure 4-7A), which indicates that the Ag NPs coated with PVP

only have a moderate effect, while the most toxic compound to the wetland’s microbial

communities is Ag+. Bacterial uptake of Ag+ can result in disruption of ATP production and DNA

64

replication, and consequently cause damage to cell walls (Mirzajani, Ghassempour et al. 2011;

Mirzajani, Askari et al. 2013). A similar conclusion can be drawn when looking at the

dose-response curves (Figure 4-8). Even though all Ag NPs and Ag+ seemed to have a negative

effect on the microbial communities, reflected in decreasing values of AWCD with increasing

concentrations, the degree of the toxicity is different between the different compounds. Differences

between the different NPs can be explained by their difference in size. With larger size, their

toxicity seems to be lower. This is because small-sized particles can easily pass through cell walls

or membranes, and finally reach the interface of cells where they can inhibit cell division etc. and

furthermore inhibit the activity of the microbial communities. According to PCS results, the PVP

Ag NPs have an average size of 148.7 nm. Probably that is why they do not show significant

toxicity to microbial communities when compared with much smaller particles, like the biogenic

Ag NPs which have an average size of 11.2 nm. Previous study also suggested that in suspensions

of biogenic Ag NPs, some Ag+ remained in the solution as these Ag NPs were synthesized from

Ag+ and the ions can be preserved for some time (De Gusseme, Sintubin et al. 2010). In addition,

the different coatings could also affect the rate of aggregation, the size of the aggregates and the

way they interact with the microorganisms, which could in turn affect the observed toxicity. Kowk,

Auffan et al. (2012) found that citrate-coated Ag NPs are most likely to stay in the water column

and increase the risk to pelagic organisms while PVP-coated Ag NPs tend to aggregate quickly and

settle onto the sediment. As a result, PVP-coated Ag NPs are considered to be more dangerous to

bottom-feeding species in aquatic systems.

In the dose-response curves (Figure 4-8), the control group (no exposure in microcosm cultivation)

and the previously exposed group (exposed to 0.1 mg/L Ag for 4 weeks) behaved differently.

Generally, the previously exposed groups have higher AWCD values than the control groups,

which indicate that some kind of resistance has been built up by prior exposure to Ag. This is

reasonable as mesocosms of the control group did not have enough time to adjust or adapt to the

new environment. In communities that have previously been exposed to 0.1 mg/L Ag, part of the

organisms that cannot adapt to the 0.1 mg/L Ag may have already been eliminated. According to

Kowk, Auffan et al. (2012) PVP-coated Ag NPs can easily aggregate and settle onto sediments

under particular conditions. This may result in concentrations of individual particles in solution

65

remaining low (or even decreasing) at increasing concentrations, which may explain slight

increases of AWCD at intermediate concentrations, as presented in Figure 4-8B. However, the

citrate-coated Ag NPs (Figure 4-8, A) may stay as individual particles in suspension during the

whole test period, maximizing the total surface area exposed to the solution and leading to the

highest possible effect per unit of silver (Shahrokh and Emtiazi 2009).

In the PCA analysis, no defined groups could be identified when evaluating CSUPs from

mesocosm communities exposed to citrate-coated Ag NPs, biogenic Ag NPs, PVP-coated Ag NPs

and Ag+ solution at 0.1 mg/L Ag (Figure 4-9). This indicates that at low concentrations (0.1 mg/L),

similar communities composed of similar species were developed (Weber and Legge 2011) and it

also suggests that citrate-coated Ag NPs, biogenic Ag NPs, PVP-coated Ag NPs and Ag+ solution

did not have a significant difference in toxicity towards wetlands microbial communities at low

concentration. However, when the concentration was increased (Figure 4-10 and Figure 4-11),

mesocosms exposed to Ag NPs and Ag+ solution were grouped on the left side of the plot. These

findings suggest that microbial communities are more capable of handling disturbances induced by

citrate-coated Ag NPs and PVP-coated Ag NPs. In another words, biogenic Ag NPs and Ag+

solution are more toxic to the microbial communities, compared with citrate-coated Ag NPs and

PVP-coated Ag NPs. The results also suggest that Ag+ and biogenic Ag NPs have some common

characteristics. These results are probably caused partly by presence of ionic Ag in the biogenic

stock solution, as suggested by De Gusseme, Sintubin et al (2010). Moreover, probably the

biogenic Ag NPs are easier to release Ag+ to the water phase while other coated Ag NPs are not. As

a result, microbial communities respond similarly to biogenic and ionic Ag on the BIOLOGTM

ECO plates.

Further work could be done to examine whether CeO2 NPs and Ce3+ generate a similar response of

the microbial communities and also plants can be introduced to see whether they could help to

increase the resistance of microbial communities to NPs.

66

Chapter 6: Conclusions and recommendations

6.1. Conclusions

In dose-response experiments, the effects of NPs (CeO2 NPs and Ag NPs) and their corresponding

ions (Ce3+ and Ag+) on the aquatic macrophyte E. canadensis were studied. This response seemed

to vary with the dose. Generally, high concentrations of NPs and their corresponding ions resulted

in growth inhibition, while low concentrations could even stimulate plant growth. In addition, when

considering the growth of E. canadensis, ions seemed to be more toxic to plants compared to NPs.

This is mainly caused by the size effect, as small sized ions can penetrate cell walls more easily

than the bigger sized NPs. Besides that, NPs can also adsorb on the plants’ cell walls, which in turn

can interfere with their own penetration activity. As a result, more ions are accumulated in plants

and subsequently inhibit plant growth upon exposure to NPs. Nanoparticles brought into contact

with the plants for a short time (1 h) are either directly accumulated in plant tissues or strongly

sorbed on the plants’ surface, as they cannot be extracted anymore using different extractants

afterwards. These findings indicate that the degradation of plants previously exposed to NPs or ions

could cause a “secondary pollution” to the environment, as they can release the accumulated NPs

and ions.

The toxicity of NPs and ions also depends on water composition. E. canadensis plants are more

sensitive to NPs and ions in surface waters that have higher TOC, IC, TC, pH etc. With varying

concentrations of ions (Cl-, SO42- etc.), the actual concentration of NPs and their corresponding ions

remaining in the water phase may also vary, as for instance precipitates can be formed. As a result,

in constructed wetlands, the presence of large suspended solids and some anions can help to reduce

the concentration of NPs and their corresponding ions even before plant remediation processes take

place.

Not only aquatic macrophytes, but also microbial communities in wetlands can be affected by NPs.

The CLPP method was employed to evaluate the potential toxicity of Ag NPs with different

coatings as well as Ag+. The results suggest that all types of silver have a negative effect on

67

microbial communities, but overall, the toxicity from highest to lowest is: Ag+ > biogenic Ag NPs >

citrate Ag NPs > PVP Ag NPs. This result is mainly due to the size effect, as PVP Ag NPs have the

largest particle size while Ag+ is much smaller. Moreover, also the type of coating can influence the

rate and size of NPs’ aggregation, and subsequent uptake and toxicity to microbial communities. In

addition, with a prior exposure to NPs during several weeks, microbial communities become more

resistance and robust to disturbance by NPs.

6.2. Recommendations for further research

Based on the results from the current study, some recommendations are made for future work:

The NPs species tested should be extended to a wider range so that more precise

conclusions can be drawn regarding the potential toxicity of NPs and their corresponding

ions to aquatic plants and the environment.  

More extensive concentration sets should be included, e.g., to evaluate the LC 50 of the

aquatic plants. As a result, the maximum concentration of NPs that can be treated in

constructed wetlands could be assessed. NPs occuring at higher concentrations will most

likely cause the death of the plants, and will thus strongly inhibit the removal efficiency

of NPs from wetlands.  

A longer exposure period should be employed in order to evaluate the chronic toxicity of

NPs to aquatic macrophytes.  

More frequently occurring wetland macrophytes other than E. canadensis should also be

studied, as more than one plant species will occur in wetlands in real cases.  

A constructed wetland at pilot-scale or full scale is suggested to be employed to evaluate

the long-term performance and response of plants and microbial communities after being

exposed to NPs and ions.  

68

Chapter 7: Reference

Aelion, C. M. and P. M. Bradley (1991). "Aerobic Biodegradation Potential of Subsurface

Microorganisms from a Jet Fuel-Contaminated Aquifer." Applied and Environmental

Microbiology 57(1): 57-63.

Asharani, P. V., Y. L. Wu, et al. (2008). "Toxicity of silver nanoparticles in zebrafish models."

Nanotechnology 19(25).

Axtell, N. R., S. P. K. Sternberg, et al. (2003). "Lead and nickel removal using Microspora and

Lemna minor." Bioresource Technology 89(1): 41-48.

Bastviken, S. K., P. G. Eriksson, et al. (2005). "Potential denitrification in wetland sediments with

different plant species detritus." Ecological Engineering 25(2): 183-190.

Begum, A. and S. HariKrishna (2010). "Bioaccumulation of trace metals by aquatic plants."

Internat J of Chem Tech Research 2(1): 250-254.

Bekyarova, E., P. Fornasiero, et al. (1998). "CO oxidation on Pd/CeO2–ZrO2 catalysts." Catalysis

Today 45(1–4): 179-183.

Bernhardt, E. S., B. P. Colman, et al. (2010). "An Ecological Perspective on Nanomaterial Impacts

in the Environment." Journal of Environmental Quality 39(6): 1954-1965.

Bhabra, G., A. Sood, et al. (2009). "Nanoparticles can cause DNA damage across a cellular

barrier." Nature nanotechnology 4(12): 876-883.

Bischoff, F. Z., D. D. Nguyen, et al. (1995). "Estimates of Aneuploidy Using Multicolor

Fluorescence in-Situ Hybridization on Human Sperm (Vol 66, Pg 237, 1994)."

Cytogenetics and Cell Genetics 69(3-4): 189-189.

Blaser, S. A., M. Scheringer, et al. (2008). "Estimation of cumulative aquatic exposure and risk due

to silver: Contribution of nano-functionalized plastics and textiles." Science of the Total

Environment 390(2-3): 396-409.

Boxall, A., Q. Chaudhry, et al. (2007). Current and future predicted environmental exposure to

engineered nanoparticles. York, UK, Central Science Laboratory.

Chapin, F. S. (1980). "The Mineral-Nutrition of Wild Plants." Annual Review of Ecology and

Systematics 11: 233-260.

69

Cho, D. Y., S. T. Lee, et al. (1994). "Studies on the Biosorption of Heavy-Metals onto

Chlorella-Vulgaris." Journal of Environmental Science and Health Part a-Environmental

Science and Engineering & Toxic and Hazardous Substance Control 29(2): 389-409.

Choi, O., K. K. Deng, et al. (2008). "The inhibitory effects of silver nanoparticles, silver ions, and

silver chloride colloids on microbial growth." Water Research 42(12): 3066-3074.

De Gusseme, B., L. Sintubin, et al. (2010). "Biogenic Silver for Disinfection of Water

Contaminated with Viruses." Applied and Environmental Microbiology 76(4): 1082-1087.

Dewet, L. P. D., H. J. Schoonbee, et al. (1990). "Bioaccumulation of Selected Heavy-Metals by the

Water Fern, Azolla-Filiculoides Lam in a Wetland Ecosystem Affected by Sewage, Mine

and Industrial-Pollution." Water Sa 16(4): 281-286.

Diegoli, S., A. L. Manciulea, et al. (2008). "Interaction between manufactured gold nanoparticles

and naturally occurring organic macromolecules." Science of the Total Environment

402(1): 51-61.

Diez, B., C. Pedros-Alio, et al. (2001). "Application of denaturing gradient gel electrophoresis

(DGGE) to study the diversity of marine picoeukaryotic assemblages and comparison of

DGGE with other molecular techniques." Applied and Environmental Microbiology 67(7):

2942-2951.

Elechiguerra, J., J. Burt, et al. (2005). "Interaction of silver nanoparticles with HIV-1." Journal of

Nanobiotechnology 3(1): 1-10.

Elmachliy, S., B. Chefetz, et al. (2011). "Removal of Silver and Lead Ions from Water Wastes

Using Azolla filiculoides, an Aquatic Plant, Which Adsorbs and Reduces the Ions into the

Corresponding Metallic Nanoparticles Under Microwave Radiation in 5 min." Water Air

and Soil Pollution 218(1-4): 365-370.

Fabrega, J., S. N. Luoma, et al. (2011). "Silver nanoparticles: Behaviour and effects in the aquatic

environment." Environment International 37(2): 517-531.

Fairbrother, H. (2010). Environmental, health, and safety effects of engineered nanomaterials:

challenges and research needs.

Farre, M., K. Gajda-Schrantz, et al. (2009). "Ecotoxicity and analysis of nanomaterials in the

aquatic environment." Analytical and Bioanalytical Chemistry 393(1): 81-95.

70

Faulwetter, J. L., V. Gagnon, et al. (2009). "Microbial processes influencing performance of

treatment wetlands: A review." Ecological Engineering 35(6): 987-1004.

Federici, G., B. J. Shaw, et al. (2007). "Toxicity of titanium dioxide nanoparticles to rainbow trout

(Oncorhynchus mykiss): Gill injury, oxidative stress, and other physiological effects."

Aquatic toxicology 84(4): 415-430.

Feldman, B., S. A. D. Ebrahim, et al. (2000). "Routine prenatal diagnosis of aneuploidy by FISH

studies in high-risk pregnancies." American Journal of Medical Genetics 90(3): 233-238.

Gaiser, B., T. Fernandes, et al. (2009). "Assessing exposure, uptake and toxicity of silver and

cerium dioxide nanoparticles from contaminated environments." Environmental Health

8(Suppl 1): 1-4.

Gaiser, B. K., A. Biswas, et al. (2011). "Effects of silver and cerium dioxide micro- and nano-sized

particles on Daphnia magna." Journal of Environmental Monitoring 13(5): 1227-1235.

Gaiser, B. K., T. F. Fernandes, et al. (2012). "Interspecies comparisons on the uptake and toxicity

of silver and cerium dioxide nanoparticles." Environmental Toxicology and Chemistry

31(1): 144-154.

Garland, J. L. (1997). "Analysis and interpretation of community-level physiological profiles in

microbial ecology." Fems Microbiology Ecology 24(4): 289-300.

Gerloff, G. C. and F. Skoog (1954). "Cell Contents of Nitrogen and Phosphorous as a Measure of

Their Availability for Growth of Microcystis Aeruginosa." Ecology 35(3): 348-353.

Gottschalk, F., T. Sonderer, et al. (2009). "Modeled Environmental Concentrations of Engineered

Nanomaterials (TiO2, ZnO, Ag, CNT, Fullerenes) for Different Regions." Environmental

Science & Technology 43(24): 9216-9222.

Grove, J. A., H. Kautola, et al. (2004). "Assessment of changes in the microorganism community in

a biofilter." Biochemical Engineering Journal 18(2): 111-114.

Gubbins, E. J., L. C. Batty, et al. (2011). "Phytotoxicity of silver nanoparticles to Lemna minor L."

Environmental Pollution 159(6): 1551-1559.

Hallberg, K. B. and D. B. Johnson (2005). "Microbiology of a wetland ecosystem constructed to

remediate mine drainage from a heavy metal mine." Science of the Total Environment

338(1-2): 53-66.

71

Hamilton, W., Ed. (1987). Biofilms: microbial interactions and metabolic activities. Ecology of

microbial communities. Cambridge, Cambridge University Press.

Handy, R. D., F. von der Kammer, et al. (2008). "The ecotoxicology and chemistry of

manufactured nanoparticles." Ecotoxicology 17(4): 287-314.

Hogstrand, C. and C. M. Wood (1998). "Toward a better understanding of the bioavailability,

physiology and toxicity of silver in fish: Implications for water quality criteria."

Environmental Toxicology and Chemistry 17(4): 547-561.

Insam, H., M. Goberna, et al. (2004). "Use of Biolog® for the community level physiological

profiling (CLPP) of environmental samples." Molecular microbial ecology manual.

Volumes 1 and 2(Ed. 2): 853-860.

Jiang, H. S., M. Li, et al. (2012). "Physiological analysis of silver nanoparticles and AgNO3

toxicity to Spirodela polyrhiza." Environmental Toxicology and Chemistry 31(8):

1880-1886.

Johnson, M. E., S. A. Ostroumov, et al. (2011). "Study of the interactions between Elodea

canadensis and CuO nanoparticles." Russian Journal of General Chemistry 81(13):

2688-2693.

Johnston, H. J., G. Hutchison, et al. (2010). "A review of the in vivo and in vitro toxicity of silver

and gold particulates: Particle attributes and biological mechanisms responsible for the

observed toxicity." Critical reviews in toxicology 40(4): 328-346.

Ju-Nam, Y. and J. R. Lead (2008). "Manufactured nanoparticles: An overview of their chemistry,

interactions and potential environmental implications." Science of the Total Environment

400(1-3): 396-414.

Kaegi, R., A. Voegelin, et al. (2011). "Behavior of Metallic Silver Nanoparticles in a Pilot

Wastewater Treatment Plant." Environmental Science & Technology 45(9): 3902-3908.

Kim, B., C.-S. Park, et al. (2010). "Discovery and Characterization of Silver Sulfide Nanoparticles

in Final Sewage Sludge Products." Environmental Science & Technology 44(19):

7509-7514.

72

Kittler, S., C. Greulich, et al. (2010). "Toxicity of Silver Nanoparticles Increases during Storage

Because of Slow Dissolution under Release of Silver Ions." Chemistry of Materials 22(16):

4548-4554.

Kumari, M., A. Mukherjee, et al. (2009). "Genotoxicity of silver nanoparticles in Allium cepa."

Science of the Total Environment 407(19): 5243-5246.

Kwok, K. W. H., M. Auffan, et al. (2012). "Uptake of silver nanoparticles and toxicity to early life

stages of Japanese medaka (Oryzias latipes): Effect of coating materials." Aquatic

toxicology 120: 59-66.

Leckie, S. E. (2005). "Methods of microbial community profiling and their application to forest

soils." Forest Ecology and Management 220(1-3): 88-106.

Limbach, L. K., R. Bereiter, et al. (2008). "Removal of Oxide Nanoparticles in a Model

Wastewater Treatment Plant: Influence of Agglomeration and Surfactants on Clearing

Efficiency." Environmental Science & Technology 42(15): 5828-5833.

Lin, D. H. and B. S. Xing (2007). "Phytotoxicity of nanoparticles: Inhibition of seed germination

and root growth." Environmental Pollution 150(2): 243-250.

Lowry, G. V., B. P. Espinasse, et al. (2012). "Long-Term Transformation and Fate of

Manufactured Ag Nanoparticles in a Simulated Large Scale Freshwater Emergent

Wetland." Environmental Science & Technology 46(13): 7027-7036.

Mal, T. K., P. Adorjan, et al. (2002). "Effect of copper on growth of an aquatic macrophyte, Elodea

canadensis." Environmental Pollution 120(2): 307-311.

Manceau, A., K. L. Nagy, et al. (2008). "Formation of metallic copper nanoparticles at the soil-root

interface." Environmental Science & Technology 42(5): 1766-1772.

Manz, W., R. Amann, et al. (1992). "Phylogenetic Oligodeoxynucleotide Probes for the Major

Subclasses of Proteobacteria - Problems and Solutions." Systematic and Applied

Microbiology 15(4): 593-600.

Maynard, A. D. (2006). "A research strategy for addressing risk." Nanotechnology, Woodrow

Wilson International Center for Scholars.

73

Mcdonald, J. W., A. J. Ghio, et al. (1995). "Rare-Earth (Cerium Oxide) Pneumoconiosis -

Analytical Scanning Electron-Microscopy and Literature-Review." Modern Pathology

8(8): 859-865.

McNeil, N. and T. Ried, Eds. (2000). Examples of different types of fluorescence in situ

hybridisation (FISH) probes. Expert Reviews in Molecular Medicine. Cambridge,

Cambridge University Press.

Meena, R. and R. Paulraj (2012). "Oxidative stress mediated cytotoxicity of TiO2 nano anatase in

liver and kidney of Wistar rat." Toxicological and Environmental Chemistry 94(1):

146-163.

Milt, V. G., C. A. Querini, et al. (2003). "Abatement of diesel exhaust pollutants: NOx adsorption

on Co,Ba,K/CeO2 catalysts." Journal of Catalysis 220(2): 424-432.

Mirzajani, F., H. Askari, et al. (2013). "Effect of silver nanoparticles on Oryza sativa L. and its

rhizosphere bacteria." Ecotoxicology and environmental safety 88: 48-54.

Mirzajani, F., A. Ghassempour, et al. (2011). "Antibacterial effect of silver nanoparticles on

Staphylococcus aureus." Research in Microbiology 162(5): 542-549.

Muyzer, G., E. C. Dewaal, et al. (1993). "Profiling of Complex Microbial-Populations by

Denaturing Gradient Gel-Electrophoresis Analysis of Polymerase Chain

Reaction-Amplified Genes-Coding for 16s Ribosomal-Rna." Applied and Environmental

Microbiology 59(3): 695-700.

Muyzer, G. and K. Smalla (1998). "Application of denaturing gradient gel electrophoresis (DGGE)

and temperature gradient gel electrophoresis (TGGE) in microbial ecology." Antonie Van

Leeuwenhoek International Journal of General and Molecular Microbiology 73(1):

127-141.

Nakagawa, K., Y. Murata, et al. (2007). "Formation and reaction activity of CeO2 nanoparticles of

cubic structure and various shaped CeO2–TiO2 composite nanostructures." Materials

Chemistry and Physics 104(1): 30-39.

Nekrasova, G. F., O. S. Ushakova, et al. (2011). "Effects of Copper(II) Ions and Copper Oxide

Nanoparticles on Elodea densa Planch." Russian Journal of Ecology 42(6): 458-463.

74

Nowack, B. and T. D. Bucheli (2007). "Occurrence, behavior and effects of nanoparticles in the

environment." Environmental Pollution 150(1): 5-22.

Oukarroum, A., S. Bras, et al. (2012). "Inhibitory effects of silver nanoparticles in two green algae,

Chlorella vulgaris and Dunaliella tertiolecta." Ecotoxicology and environmental safety 78:

80-85.

Oukarroum, A., S. Polchtchikov, et al. (2012). "Temperature influence on silver nanoparticles

inhibitory effect on photosystem II photochemistry in two green algae, Chlorella vulgaris

and Dunaliella tertiolecta." Environmental Science and Pollution Research 19(5):

1755-1762.

Parkinson, D. and D. C. Coleman (1991). "Microbial communities, activity and biomass."

Agriculture, Ecosystems & Environment 34(1–4): 3-33.

Podila, R. and J. M. Brown (2013). "Toxicity of engineered nanomaterials: a physicochemical

perspective." Journal of biochemical and molecular toxicology 27(1): 50-55.

Raats, D., M. Offek, et al. (2011). "Molecular analysis of bacterial communities in raw cow milk

and the impact of refrigeration on its structure and dynamics." Food Microbiology 28(3):

465-471.

Ratte, H. T. (1999). "Bioaccumulation and toxicity of silver compounds: A review." Environmental

Toxicology and Chemistry 18(1): 89-108.

Schrand, A. M., M. F. Rahman, et al. (2010). "Metal-based nanoparticles and their toxicity

assessment." Wiley Interdisciplinary Reviews: Nanomedicine and Nanobiotechnology

2(5): 544-568.

Schubert, D., R. Dargusch, et al. (2006). "Cerium and yttrium oxide nanoparticles are

neuroprotective." Biochemical and biophysical research communications 342(1): 86-91.

Shahrokh, S. and G. Emtiazi (2009). "Toxicity and unusual biological behavior of Nanosilver on

gram positive and negative bacteria assayed by microtiter-plate." Eur J Biol Sci 1(3):

28-31.

Shi, J. Y., A. D. Abid, et al. (2011). "To duckweeds (Landoltia punctata), nanoparticulate copper

oxide is more inhibitory than the soluble copper in the bulk solution." Environmental

Pollution 159(5): 1277-1282.

75

Shin, J. C., H. L. Ross, et al. (1997). "Detection of chromosomal aneuploidy in endometriosis by

multi-color fluorescence in situ hybridization (FISH)." Human genetics 100(3-4): 401-406.

Song, G., Y. Gao, et al. (2012). "Physiological effect of anatase TiO2 nanoparticles on Lemna

minor." Environmental Toxicology and Chemistry 31(9): 2147-2152.

Stefanowicz, A. (2006). "The biolog plates technique as a tool in ecological studies of microbial

communities." Polish Journal of Environmental Studies 15(5): 669-676.

Tian, J., K. K. Y. Wong, et al. (2007). "Topical Delivery of Silver Nanoparticles Promotes Wound

Healing." ChemMedChem 2(1): 129-136.

Truu, M., J. Juhanson, et al. (2009). "Microbial biomass, activity and community composition in

constructed wetlands." Science of the Total Environment 407(13): 3958-3971.

Vaseashta, A., M. Vaclavikova, et al. (2007). "Nanostructures in environmental pollution detection,

monitoring, and remediation." Science and Technology of Advanced Materials 8(1–2):

47-59.

Vilela, D. M., G. V. D. Pereira, et al. (2010). "Molecular ecology and polyphasic characterization

of the microbiota associated with semi-dry processed coffee (Coffea arabica L.)." Food

Microbiology 27(8): 1128-1135.

Weber, K. and R. Legge (2010). Community-Level Physiological Profiling. Bioremediation. S. P.

Cummings, Humana Press. 599: 263-281.

Weber, K. P., M. Gehder, et al. (2008). "Assessment of changes in the microbial community of

constructed wetland mesocosms in response to acid mine drainage exposure." Water

Research 42(1–2): 180-188.

Weber, K. P., J. A. Grove, et al. (2007). "Data transformations in the analysis of community-level

substrate utilization data from microplates." Journal of Microbiological Methods 69(3):

461-469.

Weber, K. P. and R. L. Legge (2011). "Dynamics in the bacterial community-level physiological

profiles and hydrological characteristics of constructed wetland mesocosms during

start-up." Ecological Engineering 37(5): 666-677.

Wiesner, M. R., G. V. Lowry, et al. (2006). "Assessing the risks of manufactured nanomaterials."

Environmental Science & Technology 40(14): 4336-4345.

76

Wiesner, M. R., G. V. Lowry, et al. (2009). "Decreasing Uncertainties in Assessing Environmental

Exposure, Risk, and Ecological Implications of Nanomaterials†‡." Environmental Science

& Technology 43(17): 6458-6462.

Wu, B., R. Huang, et al. (2010). "Bacterial responses to Cu-doped TiO2 nanoparticles." Science of

the Total Environment 408(7): 1755-1758.

Wynn, T. M. and S. K. Liehr (2001). "Development of a constructed subsurface-flow wetland

simulation model." Ecological Engineering 16(4): 519-536.

Zhang, J. C., C. L. Liu, et al. (2010). "Effect of cerium ion on the proliferation, differentiation and

mineralization function of primary mouse osteoblasts in vitro." Journal of Rare Earths

28(1): 138-142.

Zhao, C. M. and W. X. Wang (2011). "Comparison of Acute and Chronic Toxicity of Silver

Nanoparticles and Silver Nitrate to Daphnia Magna." Environmental Toxicology and

Chemistry 30(4): 885-892.

Zhou, J., N. S. Xu, et al. (2006). "Dissolving Behavior and Stability of ZnO Wires in Biofluids: A

Study on Biodegradability and Biocompatibility of ZnO Nanostructures." Advanced

materials 18(18): 2432-2435.

Zhu, H., J. Han, et al. (2008). "Uptake, translocation, and accumulation of manufactured iron oxide

nanoparticles by pumpkin plants." Journal of Environmental Monitoring 10(6): 713-717.