Molecular Identification of Rhizosphere Bacteria from Phragmites Sp

Home , Rhizosphere

Molecular identification of rhizosphere bacteria from Phragmites sp. growing in constructed wetlands treating benzene derivative compounds

Melissa Pena Matos

Thesis to obtain the Master of Science Degree in Microbiology

Supervisors: Professor Dr. Susete Maria Martins Dias Professor Dr. Jorge Humberto Gomes Leitão

Examination Committee Chairperson: Prof. Dr. Nuno Gonçalo Pereira Mira Supervisor: Prof. Dr. Susete Maria Martins Dias Members of the Committee: Prof. Dr. Ana Cristina Anjinho Madeira Viegas

October 2016

Acknowledgments

First of all, I would like to acknowledge Professor Susete Martins Dias for giving me the opportunity to join her Laboratory, for her guidance, patience, support, availability, determination, advice and encouragement.

To Professor Jorge Leitão for also giving me the opportunity to join his laboratory, patience guidance, advice and support.

I thanks to Renata and Rui for all the support, laughs, help, availability, shared knowledge and for putting up with me. You made the work in the lab more fun and better.

To all the people in the labs (Laura, Rita, Nídia, Isabel, Sofia, João S., João B., Joana, Silvia, Soraia and Tiago) for all the help you gave in the lab and all the conversations that we shared.

A special thanks to Sara and Alicia for always being there, your support and friendship and concern over me.

I would like to specially thanks Tânia for always being there, all of your support, patience and most special all the encouragement words for when the motivation was not the strong enough.

And finally, but not least would like to express my gratitude to my family, specially my parent who were always present for me when I needed the most, and supporting me unconditionally. This thesis is for you.

Abstract

Constructed wetlands (CWs) are an eco-friendly approach in treatment of industrial effluents and have versatile operation conditions. In this context, the present work aimed to contribute to a better understanding of the microbial diversity present in the rhizosphere of Phragmites sp. rooted on light expanded clay aggregates (LECA®) or soil from two different pilot CWs treating industrial wastewaters containing benzene derivative compounds.

Non-selective solid media was used to isolate viable microorganisms from Phragmites sp. rhizosphere samples and a total of 27 bacterial and 3 fungal isolates were obtained. Sequencing of amplimers obtained with primers for the bacterial 16S ribosomal RNA gene or fungal Internal Transcribed Spacer (ITS) allowed the identification of the isolates as belonging to the phyla Firmicutes, Proteobacteria, Bacteroidetes, Actinobacteria, and Ascomycota.

From the 27 bacterial isolates obtained, those belonging to the genera Acinetobacter (isolated from LECA® and soil CWs), Lysinibacillus (LECA®), Cohnella (soil), Microbacterium (soil), Dermacoccus (soil), Staphylococcus (soil) and Micrococcus (soil) could grow in minimal medium (MM) containing 1000 ppm of nitrobenzene as the sole carbon and nitrogen sources. Interestingly, growth of mixed cultures under these conditions was observed for the bacterial strains unable to grow as single isolates, highlighting the importance of microbial consortia metabolism in aromatic compounds biodegradation.

Catechol dioxygenase activity was detected in crude extracts prepared from bacterial cells grown on solid MM supplemented with 1000 ppm nitrobenzene. Since no nitrobenzene reductase activity was detected in these extracts, bacterial nitrobenzene removal in the studied constructed wetland may occur by the oxidative pathway.

Keywords: constructed wetlands, catechol dioxygenase, microbial consortia, nitrobenzene, nitrobenzene reductase, rhizosphere.

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Resumo

As zonas húmidas artificiais (ZHA) são uma abordagem amiga do ambiente para tratamento de efluentes industriais. Neste contexto, o presente trabalho pretendeu contribuir para uma melhor compreensão da diversidade microbiana presente na rizosfera de Phragmites sp. enraizada em agregados leves de argila expandida (LECA®) ou em solo presente em duas ZHA piloto distintas a tratar efluentes que contêm compostos derivados do benzeno.

De amostras da rizosfera de Phragmites sp. foram isolados microrganismos viáveis em meio não- seletivo, resultando um total de 27 bactérias e 3 fungos isolados. A sequenciação dos amplicões obtidos com oligonucleótidos iniciadores para o gene ribossomal 16S bacteriano e o espaçador interno transcrito (ITS) de fungos, permitiram a identificação dos isolados como pertencentes aos filos Firmicutes, Proteobacteria, Bacteroidetes, Actinobacteria e Ascomycota.

Dos 27 isolados bacterianos, os que pertencem aos géneros Acinetobacter (isolado a partir das ZHA com LECA® e com solo), Lysinibacillus (LECA®), Cohnella (solo), Microbacterium (solo), Dermacoccus (solo), Staphylococcus (solo) e Micrococcus (solo) cresceram em meio mínimo contendo 1000ppm de nitrobenzeno como única fonte de carbono e azoto. Curiosamente, observou-se o crescimento nestas condições de culturas contendo a mistura de estirpes bacterianas anteriormente incapazes de crescer individualmente, destacando assim a importância dos consórcios microbianos na biodegradação de compostos aromáticos.

Nos extratos proteicos preparados a partir de células bacterianas crescidas em meio mínimo com 1000ppm de nitrobenzeno foi detetada atividade da enzima catecol dioxigenase. A não deteção de atividade de nitrobenzeno redutase nestes mesmos extratos sugere que a remoção do nitrobenzeno nas ZHAs estudadas ocorra pela via oxidativa.

Palavras-chave: catechol dioxigenase, consórcios microbianos, nitrobenzeno, nitrobenzeno redutase, rizosfera, zonas húmidas artificiais

Table of Contents

Acknowledgments ...... i

Abstract...... iii

Resumo ...... v

Table of Contents ...... vii

Table List ...... ix

Figure List ...... x

Abbreviations ...... xii

1. Introduction ...... 1

1.1. Wetlands and Constructed wetlands ...... 1

1.1.1. Types of Constructed Wetlands ...... 1

1.1.2. Wetland ecology ...... 3

1.1.2.1. Soil ...... 3

1.1.2.2. Plants used in Constructed Wetlands...... 4

1.1.2.2.1. Phragmites sp...... 5

1.1.2.3. Constructed wetland microorganisms ...... 5

1.1.2.3.1. Microbial metabolism ...... 6

1.2. Nitroaromatic compounds ...... 6

1.2.1. Nitrobenzene ...... 7

1.3. Remediation of nitroaromatic compounds ...... 7

1.3.1. Physical methods...... 7

1.3.2. Bioremediation ...... 8

1.3.2.1. Aerobic pathways mediated by monooxygenases and dioxygenases ...... 9

1.3.2.2. Aerobic pathways mediated by nitroreductases ...... 10

1.3.3. Mechanisms involved in nitrobenzene biodegradation...... 13

1.3. Aims of this thesis ...... 15

2. Materials and methods ...... 17

vii

2.1. Sampling ...... 17

2.2. Isolation of culturable microorganisms ...... 17

2.3. DNA extraction ...... 17

2.4. Polymerase chain reaction (PCR) ...... 18

2.5. Nitrobenzene as the sole carbon and nitrogen source for bacteria ...... 19

2.6. Presence of plasmids in bacterial isolates ...... 19

2.7. Total protein extracts ...... 19

2.9. Nitroreductase assay ...... 20

2.10. Catechol 2,3-dioxygenase assay ...... 20

3. Results ...... 22

3.1. Microorganism isolation and identification ...... 22

3.5. Nitrobenzene as the sole carbon and nitrogen source for bacteria ...... 27

3.6. Presence of plasmids in bacterial isolates ...... 27

3.7. Nitroreductase and catechol 2,3-dioxygenase assays ...... 28

4. Discussion ...... 30

5. Conclusion and Future work ...... 36

7. References ...... 38

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Table List

Table 1 - Number of isolates purified in each bed, depth and corresponding dilution ...... 22

Table 2 - Summary of the information gathered on the isolates for bed L5: Fungi Bacteria...... 24

Table 3 - Summary of the information gathered on the isolates for bed L3...... 25

Figure List

Figure 1 – Types of Constructed Wetlands: A) Free-Water Surface; B) Horizontal Subsurface Flow; C) Vertical Subsurface Flow. Images adapted from Tilley et al (2014)...... 2

Figure 2 - Examples of reactions catalyzed by (A) styrene monooxygenase and (B) 2- nitropropanedioxygenase. Image adapted from Torres Pazmiño et al. (2010) ...... 10

Figure 3 – Mechanism of action of type I and II nitroreductases. Type I nitroreductases transfer two electrons from NAD(P)H to a nitroaromatic compound (1) forming nitroso (2) and hydroxylamino (3) intermediates and lastly an amino group (4)(a). Type II nitroreductases transfer a single electron to the nitro group, forming a nitro anion radical (5), which in the presence of oxygen will generate the superoxide anion in a futile redox cycle, regenerating the nitro group (b). Image adapted from Oliveira et al. (2010)...... 11

Figure 4 – Oxidative (red) and reductive (purple) nitrobenzene degradative pathways. Adapted from Arora and Bae (2014)...... 14

Figure 5 - Example of one of the agarose gels (0.8% agarose) obtained with the results from the plasmid extractions from the microorganisms isolated. M, marker; Lane number corresponds to the isolated bacteria as presented in tables 3 and 4: 2, Brevibacillus Brevis; 4, Acinetobacter radioresistens; 10, Lysinibacillus sp.; 12, Lysinibacillus sphaericus; 16, Acinetobacter sp.; 15, Castellaniella sp.; 19, Lysobacter sp.; 21, Bacillus sp.; 23, Dermacoccus sp.; 25, Micrococcus sp.; 26, Micrococcus sp.; 29, Bacillus megaterium...... 28

Figure 6 - Catechol 2,3-dioxigenase activity assay. The reaction mixture contains6.021 ×10-5 U/mg of a mixture of the protein extracts obtained from the isolates that grew in MM with 1000 ppm of nitrobenzene, potassium phosphate buffer (50 mM, pH 8), catechol 0.33 mM in a total volume of 1 mL...... 29

Abbreviations

atm Atmosphere

Blast Basic local alignment search tool

Bp Base pair

CW Constructed Wetland

DNA Deoxyribonucleic acid dNTP Deoxynucleotide triphosphates

EDTA Ethylenediamine tetraacetic acid

FAD Flavin adenine dinucleotide

FMN Flavin mononucleotide

FWS Free water surface

HF Horizontal flow

HLR Hydraulic loading rate

HRT Hydraulic retention time

ITS Internal transcribed spacer kDa kilo-Dalton

L3 Constructed wetland bed 3

L5 Constructed wetland bed 5

LECA® Light expanded clay aggregate

MgSO4 Magnesium sulphate

MM Minimal medium

NaCl Sodium chloride

NADH Nicotinamide adenine dinucleotide

NADPH Nicotinamide adenine dinucleotide phosphate

P. austraulis Phragmites austrailis

PDA Potato dextrose agar

PCR Polymerase chain reaction

xii

PBS Phosphate buffered solution ppm parts per million

TAE Tris acetate EDTA

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1. Introduction

1.1. Wetlands and Constructed wetlands

Wetlands are transitional environments that lie between dry land and open water, ecologically intermediate the terrestrial and aquatic ecosystems. Since wetlands often form part of a continuum of a type community, it is difficult to set boundaries (Vymazal and Kröpfelová, 2008).

Wetlands are unique among the major ecosystem groups due to their specific proprieties: abundant water, which is important for most forms of biological productivity, and plants that are adapted to take advantage of this water supply while overcoming the periodic shortage of other essential elements, such as oxygen. As a result, wetlands have a higher rate of biological activity than most ecosystems, which makes possible to transform many of the common pollutants that occur in conventional wastewater into harmless by-products or essential nutrients that can be used for additional biological productivity. Wetlands are one of the least expensive treatments systems to operate and maintain (Kadlec and Wallace, 2009).

Even though natural wetlands are able to provide high levels of wastewater treatment the effect of toxic material and pathogens in water and on the long-term degradation of wetlands due to additional nutrient and hydraulic loadings from wastewater are still of concern. To overcome these, constructed wetlands (CWs) have been built for wastewater treatment (Kivaisi, 2001). CWs are therefore engineered systems designed and constructed to mimic natural wetland systems for treating wastewater, using natural processes involving wetland vegetation, soil and the associated microbiota to aid in treating wastewater (Vymazal and Kröpfelová, 2008; H. Wu et al., 2015).

CWs technology depends on natural occurring energies making it a cheaper technology with less operation and maintenance requirements than conventional treatment systems. CWs are therefore applicable for wastewater treatment in small communities and areas that do not have public sewage systems or economically underdeveloped and public effluents (Kivaisi, 2001; Haberl et al., 2003; Saeed and Sun, 2012; H. Wu et al., 2015; S. Wu et al., 2015)).

1.1.1. Types of Constructed Wetlands

CWs can be divided in two basic types according their hydrology: free water surface (FWS) CWs and subsurface flow systems (SSF) CWs. In FWS systems, the wastewater flow is shallow and the substrate is over saturated, identical to natural wetlands (Figure 1 A)). In SSF systems, the wastewater flow through the substrate can be either horizontal (Figure 1 B)) or vertical (Figure 1 C)), according to the flow direction, which leads to different maximum hydraulic retention times

1 and dissolved oxygen availability. Thus, SSF CWs are further divided in vertical flow (VF) and horizontal flow (HF) CWs. Various wetland systems can also be combined, named as hybrid CWs, which consist generally in two stages of several parallel CWs in series, for example: VF-HF CWs, HF-VF CW, HF-FWS and FWS-HF CWs (H. Wu et al., 2015).

Figure 1 – Types of Constructed Wetlands: A) Free-Water Surface; B) Horizontal Subsurface Flow; C) Vertical Subsurface Flow. Images adapted from Tilley et al (2014).

The demand for a better understanding of the mechanisms responsible for wastewater treatment led to this variety of designs and configurations. Since each industrial effluent has a specific composition and characteristics, when planning a CW, both the design and configuration must be carefully considered. Design and operation of CWs should take into account the following items: 1) the site selection, 2) rooting substrate selection, 3) plant selection, 4) wastewater type, 5) hydraulic loading rate (HLR), 6) hydraulic retention time (HRT), 7) water depth, 8) operation mode and 9) maintenance procedures. All these items affect removal mechanisms such as

2 sedimentation, filtration, precipitation, volatilization, adsorption, plant uptake and various microbial processes important in contaminants removal (Wu et al., 2014; H. Wu et al., 2015).

1.1.2. Wetland ecology

Due to the presence of ample water, wetlands are home to a variety of microbial and plant species. A wide range of life forms, from the smallest viruses to the largest trees is a result from the diversity of physical and chemical niches present in wetlands. Interspecific relations are created due to this biological diversity, which results in more complete utilization of energy inflows, and lastly on the treatment proprieties of the wetland ecosystem. Both genetic diversity and functional adaptation of grant living organisms to use constituents of wastewaters for their growth and reproduction mediate/induce physical, chemical and biological transformations in pollutants, thereby modifying water quality (Kadlec and Wallace, 2009).

One of the most important zones in CWs is the root zone (or rhizosphere), since it is the place where physicochemical and biological processes occur and are induced by the interaction of plants, microorganism, soil and pollutants (Stottmeister et al., 2003). The rhizosphere is the zone around a root where many complex biological interactions exist, many of them involving microorganism (Shaw et al., 2006). In order to treat wastewater as efficiently as possible, detailed knowledge on the effectiveness of various plant species, the colonization characteristics of certain groups of microorganisms, and how specific contaminants interact with the filter bed material is of critical importance when designing CWs (Stottmeister et al., 2003).

Scientists and engineers have dealt with technological design issues, with the reactive zone of the rhizosphere being treated as a “black box” where the inlet and outlet loads are the only issues considered. The complexity of the microbiology of treatment wetland accounts for the lack of suitable testing systems and study methods. However the recent application of novel molecular biology techniques of metagenomics and the development of small scale process modelling experiments has opened a new era in the research of wetland wastewater treatment (Stottmeister et al., 2003; Faulwetter et al., 2009).

1.1.2.1. Soil

A suitable substrate in CWs is important, because it can provide an adequate growing medium for plant and microbial films. The soil matrix is also important since it has a decisive influence on the hydraulic process (Stottmeister et al., 2003; S. Wu et al., 2015).

Chemical soil composition and physical parameters such as grain-size distributions, interstitial pores space, effective grain sizes, degrees of irregularity and the coefficient of permeability are also important factors that influence the bio-treatment system (Stottmeister et al., 2003).

The most used substrates include natural material, artificial media and industrial by-products such as gravel, sand, clay, vermiculite, calcite, marble, slag, fly ash, bentonite, dolomite, limestone, shell, zeolite, wollastonite, activated carbon, light weight aggregates (S. Wu et al., 2015).

1.1.2.2. Plants used in Constructed Wetlands

Numerous studies of wastewater treatment with and without plants have been carried out. These studies concluded that the performance is higher when plants are present. Therefore, macrophytic plants are considered an essential component of CW treatments design. Indeed macrophytic plants act as intermediates for purification reactions, contributing to erosion control, providing a surface area and oxygen for the growth of microorganisms and, insulate the bed surface during winter (Vymazal and Kröpfelová, 2008; Kadlec and Wallace, 2009; Saeed and Sun, 2012; S. Wu et al., 2015).

Macrophytes can be divided in 4 groups based on their morphology and physiology (Kadlec and Wallace, 2009; Saeed and Sun, 2012):

1. Emergent: members of these group can grow on water saturated or submersed soil, as well as in water depth of 0.5 metres or more above the substrate surface. A few examples of these are Acorus calamus, Carex rostrata, Phragmites australis, Scirpus Iacustris, and Typha latifólia. 2. Floating-leaved: this group comprises plants that are rooted in submersed sediments in water depths of approximately 0.5 to 3 meters and have either floating or slightly aerial leaves. A few examples of these plants are Nymphaea odorata and Nuphar lutea. 3. Submerged: members of this group have the photosynthetic tissue submersed in water, grow well in oxygenated water and are mainly used for polishing secondary treated wastewater. Vascular angiosperms (e.g. Myriophyllum spicatum, Ceratophyllum demersum) occur in water only to about 10 meters of water depth (1 atmosphere (atm) hydrostatic pressure) and non-vascular macro-algae can occur to the lower limit of the photic zone (up to 200 meters, e.g. Rhodophyceae). 4. Free floating: members of this group are not rooted to the substratum, i.e., they float freely on surface water, and are usually restricted to non-turbulent protected areas. These plants are capable of removing nitrogen and phosphorus through denitrification, and incorporate these two components into plant biomass. In addition, these plants also remove suspended solids. Examples include Lemna minor, Spirodela polyrhiza and Eichhornia crassipes).

The diverse physical components of wetland macrophytes (aerial tissues, plant tissue in water, and roots) contribute to the wetland performance optimisation to some extent. For example, aerial tissues store nutrients, provide insulation to the system during winter and add aesthetic values. The plant tissue in water acts as a filter medium, excretes oxygen and reduces water velocity and

4 thus enhancing sedimentation. The roots perform a supplementary role by fostering biodegradation routes inside wetland systems (Saeed and Sun, 2012).

Several adaptations of plants allow their growth in water or wetlands. These adaptations include reproductive strategies and behavioural responses, physiological and morphological responses. One of the most important adaptations was the development of aerenchymous plant tissues that transport gases to and from the roots through the vascular tissues of the plant above water and in contact with the atmosphere. This adaptation provides an aerated root zone (Kadlec and Wallace, 2009). Node-like segmentation and diaphragms, which are gas permeable structures that provide a secure barrier by preventing liquids from penetrating the plant tissues and protect the gas chamber in the rhizome area (Stottmeister et al., 2003).

1.1.2.2.1. Phragmites sp.

Among different macrophytes Phragmites sp. promotes higher, above ground biomass and rigours root penetration inside the media (Saeed and Sun, 2012).

Phragmites, also known as common reed is a perennial flood tolerant grass with an extensive rhizome system. This species is a dominant component almost all over the world in freshwater, brackish, and often in marine littoral communities being, widely distributed throughout Europe, Africa, Asia, Australia and North America. However, Phragmites is considered as an invasive introduced pest species in North America, New Zealand and some parts of Australia. Although it is normally associated to lowlands, Phragmites has also been described at 3000 meters in Tibet. (Vymazal and Kröpfelová, 2008; Engloner, 2009; Vymazal, 2013).

P. austraulis habitat description range from heavily polluted to oligotrophic lakes. In Europe this species is regarded as an ecologically beneficial plant that provides a habitat for endangered wildlife, buffers zones for nutrient retention and stabilizes shores banks. It also has an economical importance since it may be harvested for roof thatch, fence material, fuel etc. (Vymazal and Kröpfelová, 2008).

1.1.2.3. Constructed wetland microorganisms

The CWs provide a suitable environment that supports growth and reproduction of microscopic organisms capable of breaking down pollutants. Bacteria and fungi are two important groups of microorganisms, because of their role in assimilation, transformation and recycling of chemical constituents present in various wastewaters. Generally, microbes have the first access to the dissolved components present in wastewater. Accomplishing sorption or transformation of these components directly or live symbiotically with other plants and animals by capturing the dissolved elements thus making them accessible to their symbiont or host (Kadlec and Wallace, 2009).

In CWs, the most stable microbiota is found in the biofilm that is associated with the plant roots and/or attached to the surface of the filter bed material. This microbiota is a complex microbial community that is created through the interaction with wastewater, being mainly responsible for the degradation performance of the system (Adrados et al., 2014). Thus the diversity of microorganisms may be critical for the proper functioning and maintenance of the system (Ibekwe, Grieve and Lyon, 2003).

CWs microbiota consists of autochthonous (indigenous) and allochthonous (foreign) microorganisms. The first ones exhibit adaptive features, they are able to possess metabolic activity, survive and grow in wetland systems participating in purification processes. While the second ones (including pathogens entering with wastewater) usually do not survive or may not have any functional importance in the wetland environment (Truu, Juhanson and Truu, 2009).

The characterization of these communities is of the utmost interest, not only because it provides a better understanding of the microbial diversity in these habitats, but it also helps to determine their biological activity (Bouali et al., 2013). This knowledge is also expected to allow the improvement of the design and performance of CWs (Truu, Juhanson and Truu, 2009).

1.1.2.3.1. Microbial metabolism

The majority of the important chemical transformations that are carried out by microbes are performed by enzymes, proteins that catalyse chemical reactions. Wetland microbial communities use enzymes to break down complex organic compounds into simpler compounds, releasing energy (catabolism). Besides the presence of appropriated enzymes environmental conditions such as temperature, dissolved oxygen (DO), and hydrogen ion concentration (pH) also affect the microbial communities biodegradative activity. The concentration of the chemical substrate undergoing the transformation is crucial to determinate reaction rates (Kadlec and Wallace, 2009).

1.2. Nitroaromatic compounds

Toxic compounds can be divided into natural products which are developed as a consequence of the metabolic activities of living organisms, and xenobiotics that are produced by industrial processes or other human activities. Due to the presence in their structure of chemical groups that are not normally present in natural compounds, xenobiotics persist and are recalcitrant and their chemical properties and quantities determine their toxicity and persistence in the environment. The releasing of xenobiotics by humans, is very recent and their interaction with targeted and non-targeted organisms causes extensive damage in the ecosystems due to the imbalance in the food chain.

The net result has been bioaccumulation and biomagnification of xenobiotics in aquatic and terrestrial organisms. The ecological stress imposed by massive amounts of xenobiotics is partially surmounted through microbial attenuation (Kulkarni and Chaudhari, 2007; Roldán et al., 2008).

One of the largest and most important group of chemicals in use today is constituted by nitroaromatic compounds. These compounds are organic molecules characterized by the presence of at least one nitro group attached to an aromatic ring. Most nitroaromatic compounds are xenobiotic chemicals. Due to the toxic, mutagenic and carcinogenic natures of these compounds and their incomplete degradation products, the United States Environmental Protection Agency (EPA) has listed some of them as priority pollutants. Nevertheless, these compounds have been used in multiple applications as pharmaceuticals, antimicrobial agents, food additives, pesticides, explosives dyes and raw materials in several industrial processes. One consequence of the widespread use of these compounds is environmental contamination of soil and water. This contamination results from intentional application to the environment (pesticides), improper handling and/or storage practices by both producers and users (Ju and Parales, 2010; Oliveira et al., 2010; Singh, Kaur and Singh, 2012).

1.2.1. Nitrobenzene

Nitrobenzene is one of the aromatic compounds having a lot of applications including its use for pharmaceuticals and aniline production (a precursor for the synthesis of polyurethane) among others (Arora and Bae, 2014).

The reported half-lives for nitrobenzene in environments such as surface water, ground water, soil and activated sludge range from two to more than 625 days, showing the persistence of this chemical. A few aerobic bacteria capable of degrading nitrobenzene have been isolated, despite the recalcitrance of this compound to biodegradation (Zhao and Ward, 1999). Some of these bacteria belong to the genera: Streptomyces, Corynebacterium, Stapylococcus, Streptococcus, Alcaligenes, Acinetobacter, and Flavocaterium (Zheng et al., 2009).

1.3. Remediation of nitroaromatic compounds

1.3.1. Physical methods

The development of cost-effective technologies for the remediation of nitroaromatic compounds was encouraged by the increase of public awareness about the hazards of toxicity and risks associated with these compounds. Several conventional (physical) clean-up methods like, hydrolysis, photo-oxidation, incineration and absorption are used for the removal of nitroaromatic

7 compounds from contaminated soil. However these methods are inefficient, expensive and non- sustainable (Kulkarni and Chaudhari, 2007).

1.3.2. Bioremediation

Numerous aerobic bacterial strains, isolated from contaminated soils and groundwaters, are capable of utilizing specific nitroaromatic compounds as their sole carbon, nitrogen and energy sources (Parales et al., 2005).

Techniques like bioremediation, which uses biological systems to catalyse degradation or transformation of these recalcitrant molecules to less toxic or non-toxic compounds, are captivating attention worldwide as simple tools to decontaminate nitroaromatic-polluted environments since they are eco-friendly and sustainable (Kulkarni and Chaudhari, 2007).

Bioremediation is the use of organisms, such as plants or microbes, to degrade or detoxify hazardous materials on contaminated sites. Many factors that can affect the biodegradation process depends on the nature of molecules to be degraded (e.g., molecule size, charge number and position of functional groups, solubility and toxicity) as well as the environmental conditions.

The use of biological processes to degrade hazardous materials became a viable and acceptable possibility due to the discovery that many soil microorganisms are capable of metabolizing xenobiotic compounds. The oldest inhabitants of Earth, microbes, are versatile and adaptive to changing environments and are cost-effective in xenobiotic removal. Due to their diverse metabolic enzymes microbes, have been employed for safe removal of environment contaminants, either through direct destruction or indirectly through transformation of the contaminant into a safer intermediate. Microbial transformation of nitroaromatics in nature occurs at a relatively slow pace. A consortium of microorganisms is often required for a complete biodegradation, since a single species generally is not capable of metabolizing all the products resulting from the initial biodegradation reactions (Kulkarni and Chaudhari, 2007; Singh, Kaur and Singh, 2012).

When exposed to nitroaromatic compounds, microbial species may either utilize the compound directly in their catabolism (being the nitroaromatic compound used as a sole source of carbon, nitrogen and energy), or by co-metabolism, a non-specific transformation by enzymes specific for other substrates (where the nitroaromatic compound can be transformed only in the presence of another substrate and may or may not serve as a carbon and nitrogen source). The reactions in co-metabolism may proceed under aerobic or anaerobic conditions, and can be catalysed by a whole range of organisms either by oxidation or by reduction in the presence of an exogenous carbon. These transformations produce organic derivatives, which can be as toxic as the parent molecules. Thus, these reactions are not preferred over mineralization for final bioremediation. Biological processes for degradation of nitroaromatics can therefore be divided into anaerobic

8 and aerobic processes as described by Kulkarni and Chaudhari (2007) and Singh, Mishra and Ramanthan (2015):

 Anaerobic: anaerobic transformation of nitroaromatics is receiving increased attention. It involves reduction of nitro groups to aromatic amines through a six-electron transfer mechanism. The reduction of nitro groups to nitroso derivatives, hydroxyl amine or amines is catalysed by nitroreductases. The degradation of most of the poly-nitroaromatics compounds occurs only under anaerobic conditions. The complete mineralization of nitroaromatics by an individual species of anaerobe is rare and the synergistic participation of a microbial consortium is required for a partial or complete degradation of several compounds;

 Aerobic: degradation of nitroaromatic compounds by aerobic bacteria involves mainly mono- and di-nitroaromatics as a source of carbon and/or nitrogen and energy by complete mineralization. In the past few decades, several reports came up with the isolation of microbes mineralizing different nitroaromatic compounds and their degradation pathway. However only few of them have been extensively studied and characterized. There are different strategies in the aerobic degradation of nitroaromatics: o Monooxygenase catalysed reactions: monooxygenases add a single oxygen atom and causes elimination of the nitro groups from mono-nitrophenols; o Dioxygenase catalysed reactions: dioxygenases introduce two hydroxyl groups into the aromatic ring with the removal of a nitro group as nitrite from the aromatic ring; o Meisenheimer complex formation: the addition of a hydride ion to the aromatic ring of the nitroaromatic compound leads to the formation of a Meisenheimer complex. The complex re-aromatizes after the release of the nitrite anion; o Partial reduction of aromatic ring: the nitro group is partially reduced to the corresponding hydroxylamine, which upon hydrolysis yields ammonia.

1.3.2.1. Aerobic pathways mediated by monooxygenases and dioxygenases

Oxygenases are enzymes that are ubiquitous in nature and play an important role in the metabolism of a broad range of compounds.

Monooxygenases are highly specialized enzymes that have evolved to catalyse the incorporation of one atom of molecular oxygen into an organic compound. These enzymes generally use NADH or NADPH as cofactors to provide reducing power for the supply of electrons to the substrate and can be ion metal-, haem- or Flavin-dependent (Nolan and O’Connor, 2008; Torres Pazmiño et al., 2010).

Dioxygenases are multicomponent enzyme systems. In dioxygenase reactions both atoms from a single molecule of molecular oxygen are incorporated in the organic substrate. These enzymes include two major classes: haem-dependent iron sulphur dioxygenases and Rieske iron-sulphur

9 non-haem dioxygenases the majority of which are NADH dependent (Gibson and Parales, 2000; Nolan and O’Connor, 2008) .

The initial step in the biodegradation of numerous aromatic compounds by aerobic bacteria often involves the incorporation of molecular oxygen into the aromatic ring, in a reaction mediated either by monooxygenase or dioxygenase (Nolan and O’Connor, 2008; Iwai et al., 2010).

Figure 2 - Examples of reactions catalyzed by (A) styrene monooxygenase and (B) 2- nitropropanedioxygenase. Image adapted from Torres Pazmiño et al. (2010)

1.3.2.2. Aerobic pathways mediated by nitroreductases

Nitroreductases are members of the NAD(P)H/FMN oxidoreductase family. They catalyse the reduction of nitro compounds to hydroxylamino or amino derivatives. Nitroreductases constitute a family of proteins with conserved sequences and have been grouped together based on their sequence similarity (Roldán et al., 2008; Oliveira et al., 2010; Yang, Lin and Wei, 2016). Nitroreductases were firstly described in eubacteria, but they have been discovered in all kingdoms. These enzymes are capable of catalysing the reduction of nitro-substituted compounds using flavin mononucleotide (FMN) or flavin adenine dinucleotide (FAD) as prosthetic groups and nicotinamide adenine dinucleotide (NADH) or nicotinamide adenine dinucleotide phosphate (NADPH) as electron donors. Recently these proteins have raised interest among environmental engineering scientists, due to their role in mediating nitroaromatic toxicity and their potential use in bioremediation and biocatalysts. The medical community has also paid attention to these enzymes due to their potential interest as agents used to activate prodrugs in direct anticancer therapies (Oliveira et al., 2010; Roldán et al., 2008; Yang et al., 2016).

According to their ability to reduce nitro groups in the presence of oxygen by one or two electron transfers (Figure 1) nitroreductases can be grouped in two categories:

 Type I (oxygen-insensitive) nitroreductases catalyse the sequential transfer of two electrons from NAD(P)H to the nitro groups of nitro-substituted compounds, in the presence or absence of oxygen, resulting in nitroso and hydroxylamine intermediates and finally primary amines. However, the hydroxylamine derivative may be the end product of the reaction and usually, the nitroso intermediate is not detected because the second two-electron reaction has a much faster rate than the first two-electron transfer (Figure 2 a).  Type II (oxygen-sensitive) nitroreductases catalyse one-electron reduction of the nitro group in the presence of oxygen, producing a nitro anion radical that subsequently reacts with molecular oxygen, forming a superoxide radical and regenerating the original nitroaromatic compound (Figure 2 b). This “futile redox cycle” can cause oxidative stress due to the production of large amounts of superoxides. This type of nitroreductases can mediate the reduction of nitroaromatics by two-electron transfers only under anaerobic conditions.

NAD(P)H NAD(P) NAD(P)H NAD(P) NAD(P)H NAD(P)

11 oxygen will generate the superoxide anion in a futile redox cycle, regenerating the nitro group (b). Image adapted from Oliveira et al. (2010).

Most bacteria contain several types of nitroreductases. Almost all nitroreductases share similar biochemical properties. They usually occur as homodimers (24-30 kDa subunits), have broad substrate specificity, contain FMN as cofactor and catalyse the reduction of the different nitrocompounds using a ping-pong bi-bi kinetic mechanism. Nitroreductases are strongly inhibited by dicoumarol, a diaphorase activity inhibitor, and by p-hydroxymercuribenzoate, a sulphydryl group reagent. Other specific inhibitors are p-iodosobenzoic acid, sodium azide and Cu2+ ions (Oliveira et al., 2010; Roldán et al., 2008; Yang et al., 2016).

Bacteria may contain both types of nitroreductases, although the most described are the type I nitroreductase since a great variety of them has been purified and/or their genes cloned and characterized. The phylogenetic analysis of putative bacterial nitroreductases genes suggests oxygen-insensitive nitroreductases can be classified in two main groups or families that are represented by the Escherichia coli nitroreductases NfsA (group A) and NfsB (group B). Group A nitroreductases are usually NADPH-dependent, while group B nitroreductases may use both NADH or NADPH as electron donors (Oliveira et al., 2010; Roldán et al., 2008; Yang et al., 2016).

Bacterial nitroreductases have received increasing attention in the last few decades because they can be used in the production of intermediates for the synthesis of commercial materials. Bacterial nitroreductases have also attracted considerable interest as nitro prodrug activators used in cancer therapy in the techniques known as antibody-directed enzyme prodrug therapy (ADEPT), gene-directed enzyme prodrug therapy (GDEPT) and virus-directed enzyme prodrug therapy (VDEPT) (Oliveira et al., 2010; Roldán et al., 2008; Yang et al., 2016).

The genes required for degradation of xenobiotics by a particular organism may be acquired by various horizontal transfer mechanisms, so that lateral gene transfer could be involved in the broad distribution of nitroreductases in prokaryotic organisms. Transmissible, plasmid-borne nitroreductase genes have also been reported in different bacteria (Oliveira et al., 2010; Roldán et al., 2008; Yang et al., 2016).

It is believed that nitroaromatic compounds are not the physiological substrate of nitroreductases, since only recently these compounds have been released into the environment. However, under selective pressure of environmental pollution, the capacity to degrade these compounds may have evolved in microorganisms, giving them a selective advantage to survive in polluted environments (Oliveira et al., 2010; Roldán et al., 2008; Yang et al., 2016).

Until now only a few microbial nitroreductases have been explored an identified by genomics and metagenomics approaches for developing the biotechnological applications (Oliveira et al., 2010; Roldán et al., 2008; Yang et al., 2016).

1.3.3. Mechanisms involved in nitrobenzene biodegradation

Bacteria with the ability of using nitrobenzene as the sole source of carbon and energy, degrade this compound aerobically, by one of the two major pathways, a partial reductive pathway and a dioxygenase catalysed pathway (Figure 3).

In the oxidative pathway, the degradation of nitrobenzene starts with the action of nitrobenzene- 1,2-dioxygenase, which converts nitrobenzene into catechol, that is further cleaved by the action of catechol 2,3-dioxygenase and metabolized by the meta-cleavage pathway. This pathway is documented in Comamonas JS 765, Acidovovax sp. JS42, and Micrococcus sp. Strain SMN1 (Arora and Bae, 2014; Singh, Mishra and Ramanthan, 2015).

The reductive pathway has been studied in mineralizing strains like Pseudomonas pseudoalcaligenes JS54 and Pseudomonas putida HS12. The initial step of this pathway involves nitrobenzene reduction to hydroxylaminobezene by a nitrobenzene nitroreductase. A mutase then isomerases hydroxylaminobenzene to 2-amoniphenol by intramolecular transfer of hydroxyl groups. The 2-aminophenol is further metabolized by a meta-cleavage pathway. Similarly to catechol 2,3-dioxygenase, 2-aminophenol 1,6-dioxygenase opens the aromatic ring of 2- aminophenol to produce 2-aminomuconic semialdehyde. This product is then oxidized to 2- aminomuconate in a NADH-dependent reaction, and then deaminated to form 4-oxalocrotonate. Metabolization proceeds through decarboxylation, followed by hydrolysis and cleavage by an aldolase, to eventually yield pyruvate and acetaldehyde. Finally, acetaldehyde dehydrogenase scavenges the acetaldehyde by oxidation into acetate, which feeds the tricarboxylic acid (TCA) cycle.

Several other bacteria have been cultured that are also able to grow on nitrobenzene, using similar pathways and enzymes.

The reduction of the nitro group is a highly favourable reaction, and therefore the reductive pathway is thought to be prevalent in most strains that have been isolated by growth on nitrobenzene (Ju and Parales, 2010; Arora and Bae, 2014).

Some bacteria cannot use nitrobenzene as sole source of carbon and energy, requiring the presence of an additional carbon source in order to transform nitrobenzene (Arora and Bae, 2014).

Figure 4 – Oxidative (red) and reductive (purple) nitrobenzene degradative pathways. Adapted from Arora and Bae (2014).

1.3. Aims of this thesis

CWs are an eco-friendly approach for the treatment of industrial effluents with a unique ecosystem. Therefore, the main goal of this study is to gain insights into the diversity and function of the microbial community present in the rhizosphere of a Phragmites sp. from two SSHF CWs treating benzene derivative compounds, one of the two pilot beds uses as substrate soil and the other LECA®. Another goal is to understand how these microorganisms perform nitrobenzene removal and whether they are able to use the nitrobenzene as sole source of carbon and nitrogen.

In order to accomplish these goals, cultivable microorganisms that were present in the rhizosphere of a Phragmites sp. from the CW treating benzene derivative compounds were isolated in solid medium. The microbes were then identified based in their 16S rDNA region in the case of bacteria, or the ITS region in the case of fungi. Their ability to grow with nitrobenzene as sole source of carbon was tested by growing the isolated bacteria in MM with nitrobenzene as sole source of carbon and nitrogen. Since the genes required for xenobiotics have been reported to be present in plasmids, the presence of plasmids in the isolated bacteria was also checked. Nitroreductase and catechol 2,3-dioxygenase assays were also performed in order to understand how nitrobenzene is degraded.

2. Materials and methods

2.1. Sampling

Samples from Phragmites sp. rhizosphere were taken from two different pilot CW beds (20m2/each) treating benzene derivative compounds. One of the beds had LECA® and was called L5 and one sample was taken from this bed. The other bed had soil and was called L3 and two samples were withdrawn, one from the surface and the other 10 cm below ground. All samples were collected near by the inlet of the horizontal beds.

2.2. Isolation of culturable microorganisms

The isolation of culturable microorganism was based on the method described by Weyens et al. (2009), using about 1 gram of rhizosphere that was resuspended in 10 mL of 10mM magnesium

-7 sulphate (MgSO4). Serial dilutions up to 10 were prepared using MgSO4. One hundred microliters of the 10-3, 10-4, 10-5, 10-6, 10-7 dilutions were spread onto the surface of triplicate Petri dishes containing 1/10 diluted 869 media (Eevers et al., 2015). Petri dishes were incubated at 30°C for 7 days.

Morphologically different colonies were purified in 1/10 diluted 869 medium in the same conditions as above, until the cultures were considered pure. Colonies were stored at -80°C in 40% glycerol, in duplicates.

2.3. DNA extraction

Bacterial individual colonies were suspended in liquid 1/10 diluted 869 medium and incubated overnight. Total genomic DNA was extracted using the High Pure PCR Template Preparation Kit (Roche Diagnostics GmbH, Mannheim, Germany) following the protocol for Isolation of Nucleic Acids from Bacteria or Yeast, with the following alterations: 15 µL of lysozyme (10 mg/mL) were used followed by incubation for more than 30 minutes, 50 µL of pre-warmed Elution Buffer were added to the upper reservoir of the filter tube, the tubes assembly were centrifuged for 1 minute at 8000 × g, these two last steps were repeated twice.

Fungi cultures were grown on potato dextrose agar (PDA) for 15 days at 30ºC. The mycelium was scraped from agar cultures, introduced in the tubes from the UltraClean Soil DNA Isolation Kit (MoBio Laboratories Inc., Carlsbad, USA) and placed in liquid nitrogen for 5 minutes. Total fungal genomic DNA was extracted, using the kit, according to the manufacturer’s instructions.

The DNA concentration and purity were assessed by measuring the absorbance at 260 nm in a spectrophotometer (NanoDrop®, ND-1000 Technologies).

The extracted genomic DNA integrity was checked visually after agarose electrophoresis (0.8%) in 1×Tris-acetate-EDTA (TAE) (TAE 50x: 242 g/L Tris Base, 57,1 mL/L acetic acid, 100 mL/L of 0.5 EDTA, pH 8) buffer at 100 volts for approximately 30 minutes and GelRed staining (11.69 grams of NaCl, 2 L of distilled water, 200 µL of GelRed) for 15 minutes. Gels were visualized and photographed using a transilluminator UV Gel DOC XR (BioRAD, Universal hood II) coupled to an image capture system.

2.4. Polymerase chain reaction (PCR)

For bacteria the amplified region was the 16S rDNA, and for fungi was the ITS region (ITS1, 5.8S, ITS2). The amplification was done using the extracted DNA. The used primers were: a) bacteria E334F (5-CCAGACTCCTACGGGAGGCAGC-3) and E939R (5- CTTGTGCGGGCCCCCGTCAATTC-3) (Rudi et al., 1997); b) fungi ITS5 (5- GGAAGTAAAAGTCGTAACAAGG-3) (White et al., 1990) and LR6 (5- CGCCAGTTCTGCTTACC-3) (Vilgalys and Hester, 1990).

The PCR reactions were conducted in a total volume of 20 µL volume. Each reaction contained:

1.5 mM MgCl2, 0.1 mM dNTP, primers 0.02 mM for bacteria and in 0.01 mM for fungi, 1 U Taqmed DNA polymerase and 1× of the corresponding buffer, provided by the polymerase manufacture (Citomed), 100 ng of DNA template and water to fulfil the volume.

The amplification reactions were conducted using the following conditions: a) bacteria :5 minutes at 95°C, followed by 30 cycles of 45 seconds at 95°C, 1 minute at 58.2 °C and 45 seconds at 72°C, and finally one cycle of 7 minutes at 72°C; b) fungi: 5 minutes at 95°C, 30 cycles of 30 seconds at 95°C, 30 seconds at 50°C and 72°C for 2 minutes and a final cycle of 72°C for 7 minutes.

In order to evaluate if the PCR reaction was successful, the PCR products were electrophoresed on a 1.5% agarose gel in 1×TAE buffer at 100 volts for about 45 minutes, and stained with GelRed solution (11.69 grams of sodium chlorine (NaCl), 2 L of distilled water, 200 µL of GelRed). Gels were visualized and photographed using a transilluminator UV Gel DOC XR (BioRAD, Universal hood II) coupled to an image capture system. The bands of interest were cut from the gel and purified using an NZYGelpure Kit (NZYTech genes & enzymes, Lisbon, Portugal), according to the manufacturer’s instructions with minor changes: Binding Buffer was added until the fragment of gel was covered and, instead of using the Elution Buffer, 30 µL of distilled water were used. The purified PCR products were sequenced by a commercial company (Operon MWG Eurofins, Germany), using the primers: a) bacteria E334F; b) fungi ITS5.

The sequences with more than 200 base pairs (bp) were compared to other sequences deposited in GenBank, using the tool Blast, and the highest hits were selected for identification. Whenever the sequences obtained were shorter than 200 bp in length, the PCR reaction was repeated and sent again for sequencing.

The identity considered for identification of the microorganisms was above 99%.

In cases, where more than one microorganism presented the same identification, one was selected and considered as representative of those with the same identity.

2.5. Nitrobenzene as the sole carbon and nitrogen source for bacteria

In order to evaluate the ability of bacterial isolates to grow using nitrobenzene as the sole carbon and nitrogen source bacterial isolates were spread individually in minimal solid medium (1 mM

MgSO4, 1 mM CaCl2 and M9 minimal salts containing Na2HPO4 (6.8 g/L), KH2PO4 (3.0 g/L), NaCl

(0.5 g/L), NH4Cl (1.0 g/L) with 1000 or 200 ppm of nitrobenzene and incubated at 30°C for 3 days in nitrobenzene saturated atmosphere.

2.6. Presence of plasmids in bacterial isolates

From 72-hour liquid culture of the bacterial isolates grown in MM with 200 ppm nitrobenzene, 10 mL were taken and centrifuged at the maximum velocity during 5 minutes. The supernatant was removed and 250 µL of 1× phosphate buffered Saline (PBS) with lysozyme (2.5 mg/mL) and RNAse (0.5 µg/mL) were added and incubated for 1 hour at 37ºC. The kit NZYMiniprep (NZYTech genes & enzymes, Lisbon, Portugal) was used to lyse the cells, according to the manufacture instructions. To visualize if plasmids were present, an electrophoresis of extracted samples was performed using a 0.8% agarose gel 1×TAE buffer at 100 volts for about 45 minutes, and stained with GelRed solution (11.69 grams of NaCl, 2 L of distilled water, 200 µL of GelRed). Gels were visualized and photographed using a transilluminator UV Gel DOC XR (BioRAD, Universal hood II) coupled to an image capture system.

2.7. Total protein extracts

Ten Petri dishes of each bacterial isolate were grown in minimal solid medium with 1000 ppm of nitro-benzene at 30ºC for 3 days in nitrobenzene saturated atmosphere.

The microorganisms’ colonies were scraped from the medium using about 3 mL of a NaCl 0.9% solution and a L shape glass spreader. The resulting suspension was collected into a centrifuge

19 tube and placed on ice. Samples were centrifuge for 5 minutes at 4°C at 14000 × g. The pellets were resuspended together in the same tube, in potassium phosphate buffer (200 mM, pH7), and the suspension with all of the isolates was then placed in a sonication tube. The tube was placed on ice and was immersed the sonicator probe in the sample. 5 cycles of 30 seconds ON/OFF, 50% duty cycles and an output control of 6 was carried out. The sonicated samples were centrifuged for 45 minutes at 4°C and 20000 × g. The supernatants were collected to a new tube and kept in ice until being used.

2.8. Protein quantification

Protein concentration was estimated using the Bradford (1976) method. Briefly in a glass tube 50 µL of the protein extract and 2.5 mL of Coomassie Blue were mixed and vortexed. A blank was prepared using 50 µL of water instead of protein extract. After 10 minutes, the absorbance was registered at a wavelength of 595 nm. Results presented are the mean of triplicate experiments. A calibration curve was used, previously prepared using bovine serum albumin solution of known concentration

2.9. Nitroreductase assay

The protein crude use for the nitroreductase assay was prepared according to the previously described protocol for total protein extraction. Nitroreductase activity was performed according to Somerville et al (1995) where the activity was determined spectrophotometrically by measuring the decrease of absorbance at a wavelength of 340 nm due to the consumption of NADPH. The reaction mixture contained protein extract, nitrobenzene (0.1 mM), NADPH (0.5 mM), phosphate buffer (20 mM, pH 8) and water in a total volume of 1 mL. Single protein crude extracts and the mixture of protein extracts obtained for several microorganisms were tested.

2.10. Catechol 2,3-dioxygenase assay

The protein crude use for the catechol 2,3-dioxygenase assay was prepared according to the previously described protocol for total protein extraction. Crude extract was prepared by mixing the protein obtained for each of the bacterial isolates that grew in MM with 1000 ppm of nitrobenzene. Catechol 2,3- dioxygenase activity was determined as described as Nozaki (1970) where the catechol 2,3- dioxygenase activity was determined spectrophotometrically by measuring the formation of 2-hydroxymuconic semialdehyde at a wavelength of 375 nm. Solutions of catecol 3.3 mM, potassium phosphate buffer 200 mM (ph 8) and protein extract

(656.38 µg/mL) were prepared. The reaction mixture contained in a total volume of 1 mL, protein extract, potassium phosphate buffer (50 mM, pH 8), catechol 0.33 mM and water. The extinction coefficient of hydroxymuconic semialdehyde is 4.4 × 104 M-1 cm-1. One unit (U) of enzyme activity was defined as the amount of protein that converts 1 micromole of catechol per minute into 2- hydroxymuconic semialdehyde under the experimental conditions used. Specific activity is expressed as U per milligram of protein (U/mg).

3. Results

3.1. Microorganism isolation and identification

Microorganisms were isolated from the rhizosphere of a Phragmites sp. present in two pilot CWs treating benzene derivative compounds. These microorganisms were obtained from two pilot different CWs, one had LECA® (L5) and the other had soil (L3). From the CW bed L5, only one sample was taken, while for bed L3, two different samples were taken one from the surface and the other from 10 cm below the ground.

Afterwards the samples were processed and plated in solid medium in order to obtain the viable, cultivable microorganisms present in each of the beds. The solid medium used was 1/10 diluted 869 medium, since according to Eevers et al. (2015) this medium enables the recovery of highest numbers of cultivable microorganisms as well as ensures the highest diversity. Multiple microorganism grew in all the plates of both beds. A total of 30 microorganisms that seem to be morphological different were isolated from the 2 beds (Table 1). The isolation of the microorganisms was done in the same medium as the one used above. The pure colonies were then grown in the same medium with the exception of fungi that were grown in PDA, and further stored in 40% glycerol at -80°C.

Table 1 - Number of isolates purified in each bed, depth and corresponding dilution

Bed Dilution Number of isolates obtained

-3 10 5 10-4 1

L5 10-5 5 10-6 2

10-7 1 10-3 1

L3 10-4 3

(0 cm) 10-5 2 10-7 4

-3 10 3 L3 10-5 2 (10 cm) 10-6 1

Microorganisms were isolated in bigger number in L5 (14 isolates) than in L3 surface (10 isolates) and L3 10 cm in depth (6 isolates).

The next step was to extract total genomic DNA from each of the isolates obtained previously. For this purpose, commercial kits were used following manufacturer’s instructions, with a few alterations. The extracted genomic DNA concentration of each isolate was measured, and to verify its integrity, an agarose gel was performed. Then the extracted DNA was submitted to PCR amplification, to amplify the 16S rDNA in bacteria or the ITS region in fungi. The obtained DNA fragments from the PCR reaction were separated in an agarose gel and later extracted from the gel. The bands obtained were purified, and sent for sequencing in order to study the diversity of the microbial population present in each isolate.

Each of the obtained sequences were compared with sequences deposited in the GenBank using the Blast tool. Only the sequencing results over 200 bp were considered as sufficient for identification of the isolated microorganisms.

Each organism was identified based on the sequence with the highest identity (above 99%) in GenBank, as shown in Tables 2 and 3. In case of a tie between two sequences in which one was a species and the other was only a genus, the genus was the one that was considered. When the identity of one sequence was the same and the same species was obtained but different strains were observed the organism was identified only at the species level.

Table 2 - Summary of the information gathered on the isolates for bed L5: Fungi Bacteria

Isolate Sequence Genera/ Species Phylum name length (identity ≥ 99%)

1 410 Albifimbria viridis Ascomycota

2 576 Brevibacillus brevis Firmicutes

3 534 Brevibacillus brevis Firmicutes

Proteobacteria; 4 309 Acinetobacter radioresistens Gammaproteobacteria

5 479 Aspergillus niger Ascomycota

Proteobacteria; 6 532 Xanthobacter sp. Alphaproteobacteria

7 560 Pedobacter sp. Bacteroidetes

8 545 Rhodococcus sp. Actinobacteria

9 548 Microbacterium sp. Actinobacteria

10 466 Lysinibacillus sp. Firmicutes

11 563 Lysinibacillus sp. Firmicutes

12 219 Lysinibacillus sphaericus Firmicutes

13 235 Lysinibacillus sphaericus Firmicutes

14 288 Microsphaeropsis arundinis Ascomycota

Table 3 - Summary of the information gathered on the isolates for bed L3.

Depth Isolate Sequence Genera/ Species Phylum (cm) name length (identity ≥ 99%)

Proteobacteria; 15 213 Castellaniella sp. Betaproteobacteria

Proteobacteria; 16 515 Acinetobacter sp. Gammaproteobacteria

17 570 Staphylococcus sp. Firmicutes

18 521 Microbacterium sp. Actinobacteria

Proteobacteria; 0 19 562 Lysobacter sp. Gammaproteobacteria

20 347 Bacillus sp. Firmicutes

21 293 Bacillus sp. Firmicutes

22 320 Cohnella sp. Firmicutes

23 296 Dermacoccus sp. Actinobacteria

24 251 Dermacoccus sp. Actinobacteria

25 298 Micrococcus sp. Actinobacteria

26 518 Micrococcus sp. Actinobacteria

27 226 Dermacoccus sp. Actinobacteria 10 28 452 Staphylococcus sp. Firmicutes

29 490 Bacillus megaterium Firmicutes

30 223 Micrococcus sp. Actinobacteria

The microorganisms isolated from L5 belong to the phyla Ascomycota, Firmicutes, Proteobacteria, Bacteroidetes and Actinobacteria. Fungi and the phyla Bacteroidetes were only found in this CW bed. At the surface of L3 the microorganisms present belong to the phyla Firmicutes, Proteobacteria and Actinobacteria. While in L3 at a depth of 10 cm the microorganisms present belong to the phyla Actinobacteria and Firmicutes.

In some cases, the microorganisms presented the same identification (Table 3 and 4). In these cases, only one was further used. The situations where identical identification was achieved were: the isolates 2 and 3, and the one chosen to be the representative was the isolate 2; the isolates

10 and 11, and the one chosen to be the representative was the isolate 10; the isolates 12 and 13, and the one chosen to be the representative was the isolate 12; the isolates 23 and 24, and the one chosen to be the representative was the isolate 23.

In Figure 4 it is possible to observe which microorganisms belongs to each CW bed and the microorganisms common between then. Thus, L5 is the one with most isolated microorganisms (3 fungus and 11 bacteria), followed by L3 at the surface (10 bacteria) and lastly L3 at 10 cm depth (6 bacteria). It is also possible to observe that L5 and L3 at the surface have the genus Microbacterium in common, while L5 and L3 at a depth of 10 cm do not have any microorganism in common. The bed L3 at a surface and L3 at a depth of 10 cm have the genera Dermacoccus and Staphylococcus. The 3 do not have any microorganisms in common.

Figure 4 – Analysis of the microbial diversity extracted from CWs L3 (soil) and L5 (LECA®). In yellow is represented L5 isolated microorganisms, highlighting the fungus.

3.5. Nitrobenzene as the sole carbon and nitrogen source for bacteria

In order to evaluate if the isolated bacteria were able to use nitrobenzene as sole source of carbon and nitrogen, they were grown in minimal solid medium with 1000 ppm of nitrobenzene.

From all of the isolated bacteria, a total of 14 out of the 27 isolates were able to grow in the previously described media. The isolated microorganisms that were able to grow individually in this medium were the isolates:  From Phragmites sp. rooted in LECA®: 4, 10 and 12;  From Phragmites sp. rooted in soil o Surface: 16, 17, 18, 20, 21, 22, and 23; o 10 cm depth from surface: 26 ,27, 28 and 30.

The bacteria that were not able to grow in this medium grew in solid MM with 200 ppm of nitrobenzene, which were the isolates 2 ,6, 7, 8, 9, 15, 19, 25 and 29. Afterwards, these bacteria that did not grew individually in the solid MM with 1000 ppm, were cultivated together in the same medium, and growth was observed.

3.6. Presence of plasmids in bacterial isolates

In order to understand if nitrobenzene degradation capabilities were inherent of the bacteria isolated a search for plasmids was undertaken, since the genes required for degradation of xenobiotics can be recruited by various horizontal transfer mechanism, and transmissible plasmid-borne nitroreductase genes have been reported in different bacteria (Roldán et al., 2008). In order to carry this test, a sample from 72-hour liquid cultures for each the isolates grown in MM with 200 ppm of nitrobenzene was taken and, with the aid of a kit, plasmids were extracted. Afterwards, to visualize if plasmids were present in the isolates, an agarose gel was performed.

As shown in Figure 5, no plasmid bands were observed, for each isolate.

3.7. Nitroreductase and catechol 2,3-dioxygenase assays

Since nitrobenzene can be degraded by microorganisms through two pathways, a partial reductive pathway were nitroreductases are involved, and a dioxygenase catalysed pathway, assays to access how nitrobenzene is degraded by the consortium of microorganism isolated previously were carried out.

An extract containing total proteins obtained from cells harvested from 10 Petri dishes of each isolate grown individually in MM supplemented with 1000 ppm nitrobenzene was used. The crude extract had a protein concentration of 656.38 µg/mL.

In the nitroreductase assay, nitroreductase activity was not detected in the crude extract. However, catechol 2,3-dioxygenase assay activity was detected in the crude extract as shown in Figure 6. The specific activity was estimated as 6.021 ×10-5 U/mg.

Figure 6 - Catechol 2,3-dioxigenase activity assay. The reaction mixture contains 6.021 ×10-5 U/mg of a mixture of the protein extracts obtained from the isolates that grew in MM with 1000 ppm of nitrobenzene, potassium phosphate buffer (50 mM, pH 8) and catechol 0.33 mM in a total volume of 1 mL.

4. Discussion

CWs are engineered systems for wastewater system with a unique ecosystem (Vymazal and Kröpfelová, 2008; Wu et al., 2014). These CWs are a cheap technology, with low operation and maintenance requirement and are eco-friendly (Saeed and Sun, 2012). One of the most important zones in CWs is the rhizosphere, since it is the place where physicochemical and biological processes occur (Stottmeister et al., 2003). Bacteria and fungi are two important groups of microorganisms, generally they are the first one to have access to the dissolved component present in wastewater (Kadlec and Wallace, 2009). The microbiota present in plant roots is a complex community that may be critical for the proper functioning and maintenance of the system. Therefore characterization of these communities is of interest, since it can provide a better understanding of the microbial diversity and can help determine their biological activity (Ibekwe, Grieve and Lyon, 2003; Bouali et al., 2013).

In order to comprehend better these systems an analysis of the microbial community composition and its diversity in the rhizosphere of a Phragmites sp. of two CWs treating benzene derivative compounds was conducted.

According to Faulwetter et al. (2009) one of the most important factors that influences the removal of pollutants in treatment wetlands it is their microbiology. The application of molecular techniques in the study of treatment wetlands has opened a new era in this field of research. Results published until now confirm the existence of appropriate microbial functional groups (Faulwetter et al., 2009).

In this study, the amplification of the 16S rDNA gene in bacteria and the ITS region in fungi was used to study the microbial population diversity present in the rhizosphere of Phragmites sp. present in two CWs treating benzene derivative compounds.

Two microbial kingdoms were found: bacteria and fungi. Bacteria were dominant in both beds L5 and L3, suggesting that these microorganisms may be responsible for most of the pollutants removal.

Microbial population isolated from Phragmites sp. rhizosphere rooted in LECA® indicate the presence of the phyla Firmicutes, Proteobacteria, Bacteroidetes, Actinobacteria and Ascomycota.

The Firmicutes genus organisms isolated in this work belong to the genera Brevibacillus and Lysinibacillus. In the genus Brevibacilus, Brevibacillus brevis, is one of the bacterial species detected. This species has been described as arsenic tolerant bacteria (Banerjee et al., 2013), capable of degrading pyrene intracellularly (Liao et al., 2015). In the genus Lysinibacillus, isolates identified as Lysinibacillus sp. and Lysinibacillus sphaericus were found. A particular strain of L. sphaericus was described as possessing an azoreductase capable of catalysing the reduction of azo dyes and nitrocompounds to their respective amines (Misal et al., 2014), and in this study both isolated species were able to grow in nitrobenzene1000 ppm.

The Proteobacteria isolated in this work were Gammaproteobacteria and Alphaproteobacteria. From the Gammaproteobacteria the species Acinetobacter radioresistens was isolated. This species was reported as able to grow on phenol or benzoate as sole carbon and energy source (Caposio et al., 2002) and in this study the isolated species could grow in nitrobenzene1000 ppm. From the Alphaproteobacteria the genus Xanthobacter was found. This genus has stains that fix nitrogen (Garrity et al., 2005) and oxidize benzene, toluene and phenol (Zhou et al., 1999). Nevertheless, the species isolated was not able to grow in 1000 ppm nitrobenzene.

The isolated Bacteroidetes is represented by the genus Pedobacter which has strains that degrade a wide range of organic compounds such as the volatile organic compounds ethyl benzene and m-xylene (Krieg et al., 2011). Nonetheless the species isolated was not able to grow in 1000 ppm nitrobenzene.

Members of the genera Rhodococcus and Microbacterium, belonging to the phylum Actinobacteria were identified during this work. The genus Rhodococcus is composed of organisms with remarkable metabolic diversity, which makes them ideal candidates for enhancing the bioremediation of contaminated sites, and as biocatalysts for a wide range of biotransformations, and also show an exceptional ability to degrade xenobiotics including, polycyclic aromatic hydrocarbons and nitroaromatic compounds (Goodfellow et al., 2012). The genus Microbacterium has a strain that possess textile effluent-degrading and plant growth- promoting activities (S. Wu et al., 2015).

From the Ascomycota phylum Albifimbria viridis was one of species, and since it is a new species only little information is known about it (Lombard et al., 2016). One of the other was Aspergillus niger that may have a free radical pathway of nitroaromatic and possibly certain chlorinated hydrocarbon degradation under anaerobic conditions (Metosh-Dickey, Mason and Winston, 1998).

Microorganisms from the phyla Firmicutes, Proteobacteria and Actinobacteria were isolated from the L3 at a depth of 0 cm.

The isolated Firmicutes belong to the genera Staphylococcus, Bacilus and Cohnella. From the genus Staphylococcus there are strains capable of degrade pyrene (Valsala et al., 2014) and that have nitroreductases (Oliveira et al., 2010) and in this study the isolated species could grow in nitrobenzene1000 ppm. A Bacillus species metabolise a wide range of aromatic compounds (Willetts, 1974) one of them being nitrobenzene (Li et al., 2014) and in this study the isolated species could grow in nitrobenzene1000 ppm. There is a Cohnella strain that can hydrolyse p- nitrophenyl (De Vos et al., 2009).

The Proteobacteria isolated were Betaproteobacteria and Gammaproteobacteria. From the Betaproteobacteria the genus Castellaniella, has denitrifying and hydrolyses p-nitroanilide species (Kämpfer et al., 2006). In the Gammaproteobacteria there is the genera Acinetobacter and Lysobacter. The Acinetobacter genus has strains capable of multiple degradation of hydrocarbons dibenzothiophene, flourene, dibenzofuran, benzyl sulphide and sodium benzoate

(Thangaraj, Kapley and Purohit, 2008) and in this study the isolated species could grow in nitrobenzene 1000 ppm. In the Lysobacter genus there is a strain that has the ability to degrade naphthalene, phenanthrene and carbazole (Maeda et al., 2009).

From the Actinobacteria phylum the genera Microbacterium and Dermacoccus. In concern to the genus Microbacterium, above it was already mention some characteristics of this genus. The Dermacoccus genus has a strain with azoreductase activity (Lang et al., 2014) and in this study the isolated species could grow in nitrobenzene 1000 ppm.

From the L3 bed, samples taken from 10 cm depth were found to contain microorganisms of the phyla Firmicutes and Actinobacteria.

The phylum Firmicutes is represented by the genera Staphylococcus and Bacillus. The genus Staphylococcus was already characterized above. The Bacillus genus is represented by the species Bacillus megaterium, which has strains with the ability of degrade an azo dye (Shah, 2014) and can potentially be used in bioremediation of benzene in both anaerobic and aerobic environments (Singh, Sar and Bennett, 2009). Nonetheless the species isolated was not able to grow in 1000 ppm nitrobenzene.

The Actinobacteria isolated belonged to the genera Micrococcus and Dermacoccus. Strains from the genus Micrococcus can use nitrobenzene as a sole source of carbon nitrogen and energy under aerobic conditions (Zheng et al., 2009) and in this study the isolated species could grow in nitrobenzene 1000 ppm. Characteristics from the genus Dermacoccus were already mentioned above.

The phyla present in all the CW beds in all the conditions are: Firmicutes and Actinobacteria.

The phylum Ascomycota and Bacteroidetes are only present in L5, and the predominant phyla is Firmicutes (42.7%) followed by the phylum Ascomycota (21.4%), Proteobacteria and Actinobacteria (14.3%) and finally the phyla Bacteroidetes (7.1%).

In L3 with 0 cm depth the predominant phyla is Firmicutes (40%) followed by Proteobacteria and Actinobacteria (30%).

In the L3 with 10 cm depth the predominant phyla is Actinobacteria (66.7%) followed by Firmicutes (33.3%).

The CW L5 is the one that show most isolated microorganisms. Implying that in LECA® a greater diversity of microorganism is found.

Since the microorganisms were grown and isolated from the rhizosphere of a Phragmites sp. of two CWs treating benzene derivative compounds, it was tested if the microorganisms could grow with nitrobenzene being the sole source of carbon and nitrogen. The isolated bacteria that grew individually in MM with 1000 ppm of nitrobenzene were then identified according to a comparison to other sequences deposited in GenBank, using the Blast tool, and with that identification, a research was done in order to find previous work relating the genus and/or species with

32 nitrobenzene. The isolates 2 and 16 whose identification was Acinetobacter radioresistens and Acinetobacter sp. respectively, have a strain with two catechol 1,2-dioxygenase genes characterized (Caposio et al., 2002), which is an enzyme that is involved in the oxidative pathway of degradation of nitrobenzene. The isolates 10 and 12 identified as Lysinibacillus sp. and Lysinibacillus sphaericus respectively, have a strain that can catalyse the reduction of nitrocompounds to their respective amines (Misal et al., 2014). The isolates 20 and 21 both identified as Bacillus sp. can degrade nitrobenzene (Li et al., 2014). For the isolated 22 identified as Cohnella sp. there was no information relating this genus with nitrobenzene. The isolate 18 identified as Microbacterium sp. has no information about the relation of this genus and nitrobenzene. The isolates 23 and 27 were identified as being Dermacoccus sp. any information was found relating this genus with nitrobenzene. Isolates 17 and 28 both identified as Staphylococcus sp. which have nitroreductases identified (Oliveira et al., 2010), that are enzymes involved in the reductive pathway of the degradation of nitrobenzene. The isolates 26 and 30 were identified as Micrococcus sp., that have strains that can use nitrobenzene as sole source of carbon nitrogen and energy under aerobic conditions (Zheng et al., 2009). As observed in this study all of these isolates were able to grow individually using nitrobenzene being the sole source of carbon and nitrogen, and upon some research in the literature and to date none of them was described as being able to use nitrobenzene as sole source of carbon and nitrogen, with the exception of the genus Micrococcus that already was described as capable of doing it. From all isolated bacteria from: L3 at the surface, the isolates 16, 17, 18, 20, 21, 22, and 23 (70% of the isolated bacteria in this conditions), L3 at 10 cm depth the isolates 26 ,27, 28 and 30 (67% of the isolated bacteria in this conditions) and L5 the isolates 4, 10, 12 (21% of the isolated bacteria in this conditions) grew in MM with 1000 ppm nitrobenzene.

The rest of the other isolated bacteria, could not grow individually in the MM with 1000 ppm of nitrobenzene, but when they were put together in the same medium the results evidenced bacterial growth. Even thought it was not possible to clarify if all of them grew or in one was more abundant than other. Nonetheless the obtained results show how important a community is for microorganism’s survival and/or to degrade xenobiotics. It is also shown that they live together with each other in symbiosis and that is why in the environment they are found together as a community and not individually. These may be one of the reasons why some microorganisms are not cultivable in the lab.

Since the genes required for degradation of aromatic compounds have been found to be encoded on plasmids in different bacteria (Park and Kim, 2000; Roldán et al., 2008) a study was conducted in order to check whether there were plasmids in the isolated bacteria. For this purpose, plasmids were extracted as described in materials and methods, using a kit able to extract bacterial artificial chromosomes and cosmids up to 52 kb, although no plasmids were detected. The methodology used was designed for plasmids of medium size and one cannot exclude the possibility that plasmids of larger size could be present in the bacterial cells. Future work should address this issue, by using other methods for the extraction of plasmids of larger size, as well as of

33 megaplasmids. In agreement with this consideration is the fact that Park and Kim, (2000) et al have reported the isolation of a 59.1Kb plasmid encoding a nitrobenzene reductase.

Nitrobenzene can be degraded by one of two major pathways, an oxidative pathway, where the catechol 2,3-dioxygenase enzyme activity participates, and a reductive pathway, where the nitrobenzene nitroreductases are present (Arora and Bae, 2014). To gain further clues on the pathways that are used by bacteria isolated from the rhizosphere of Phragmites sp. enzyme assays were conducted to detect the nitroreductase and catechol 2,3-dioxygenase activities. The protein content of the crude extract obtained from the biomass previously grown in 10 petri dishes was quite low (< 700 µg/mL). Since the extract was not purified the results from the enzyme activity detected or not detected are just indicative.

For nitroreductase assay no activity could be detected. The experimental conditions used in this work were based on methods previously published by Xiao et al. (2006), Yang et al. (2016) and Yanto et al. (2010), although these authors have used purified protein.

In the catechol 2,3-dioxygenase assay the specific activity obtained was 5.265 × 10-5 U/mg, a low specific activity. Nishino and Spain (1995) reported catechol 2,3-dioxygenase specific activities of 3.04 U/mg. The discrepancy between results from Nishino and Spain and those reported in this work might result from the distinct methods used for protein extract preparation, since in our study the extracts were obtained from solid medium. The colonies harvested were cultivated for 3 days and the extracts were used in the day after their preparation. Nishino and Spain (1995) also reported that the extracts loose their activity when not used immediately after preparation.Anyway the obtained results suggest that nitrobenzene bacterial removal occurs by the oxidative pathway and not by the reductive pathway, at least under the used conditions and for the bacterial isolates used to prepare the crude extracts.

To gain further insights into the pathway used by the bacterial isolates to metabolize nitrobenzene, an alternative method than could have been used was the measurement of ammonia or nitrite released. As stated by authors Nishino and Spain (1995), the release of ammonia is indicative of the usage of the reductive pathway, while nitrite release is directly related to the usage of the oxidative pathway.

5. Conclusion and Future work

To better understand the microbial community composition and diversity present in the rhizosphere of Phragmites sp. from two CWs, the microorganisms were cultivated in solid medium and identified based on the amplification of the 16S rRNA gene or the ITS region. The results showed that 14 of the cultivated bacteria could grow in the presence of nitrobenzene, as sole source of carbon and nitrogen at a concentration of 1000 ppm. Of these 14 bacterial isolates, 13 have not been described yet as capable of using nitrobenzene as sole source of carbon and nitrogen. Results from this work also show that microorganisms that individually could not grow in nitrobenzene at 1000 ppm, when grown together in community they were able to grow.

The studied CWs are complex wastewater treatment systems with a diverse microbial community, which is stimulated by the roots and rhizomes by the release of oxygen and roots exudates (Stottmeister et al., 2003).

In conclusion, available species-specific data about the microbial communities is scarce and is difficult to make any general conclusions about the dynamic of the microbial community in wetlands or how the removal process functions. This knowledge is expected to lead to the creation of starter cultures for wastewater treatment. It is also important that obtained sequences (such as 16S rDNA or other functional gene) in different studies are deposited in available databases with the proper amount of background information, to facilitate knowledge accumulation of these microbiomes and a better understanding of the complex processes in these systems.

In future work, metagenomics should be performed in order to gain further insights into the composition and structure of the microbial communities thriving in CWs. In particular, data from metagenomics might contribute to better understand the possible routes of nitrobenzene degradation/metabolism. Although results from this study suggest that the preferred pathway of degradation of nitrobenzene is the oxidative, a thorough investigation should be performed using the isolates able to grow in nitrobenzene at a concentration of 1000 ppm.

The combination between the knowledge gained from this work and future metagenomic analysis will contribute to a better understanding of what happens in the reactive zone of the rhizosphere and will help the improvement of design and operations of CWs, making them as efficient as possible in wastewater treatment.

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