Bioremediation of PAH-contaminated soil with fungi – From laboratory to field scale

International Biodeterioration & Biodegradation 86 (2014) 238e247

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Bioremediation of PAH-contaminated soil with fungi e From laboratory to field scale Erika Winquist a, *, Katarina Björklöf b, Eija Schultz b, Markus Räsänen a, Kalle Salonen a, Festus Anasonye c, Tomás Cajthaml d, Kari T. Steffen c, Kirsten S. Jørgensen b, Marja Tuomela c a

Department of Biotechnology and Chemical Technology, Aalto University School of Chemical Technology, P.O. Box 16100, FI-00076 Aalto, Finland Finnish Environment Institute, P.O. Box 140, FI-00251 Helsinki, Finland Department of Food and Environmental Sciences, University of Helsinki, P.O. Box 56, FI-00014 Helsinki, Finland d  ská 1083, CZ-14220 Prague 4, Laboratory of Environmental Biotechnology, Institute of Microbiology, Academy of Sciences of the Czech Republic, Viden Czech Republic b c

a r t i c l e i n f o

a b s t r a c t

Article history: Received 15 May 2013 Received in revised form 16 September 2013 Accepted 17 September 2013 Available online 13 November 2013

The purpose of this study was to develop a fungal bioremediation method that could be used for soils heavily contaminated with persistent organic compounds, such as polyaromatic hydrocarbons (PAHs). Sawmill soil, contaminated with PAHs, was mixed with composted green waste (1:1) and incubated with or without fungal inoculum. The treatments were performed at the laboratory and field scales. In the laboratory scale treatment (starting concentration 3500 mg kg
1, sum of 16 PAH) the high molecular weight PAHs were degraded significantly more in the fungal-inoculated microcosms than in the uninoculated ones. In the microcosms inoculated with Phanerochaete velutina, 96% of 4-ring PAHs and 39% of 5- and 6-ring PAHs were removed in three months. In the uninoculated microcosms, 55% of 4-ring PAHs and only 7% of 5- and 6-ring PAHs were degraded. However, during the field scale (2 t) experiment at lower starting concentration (1400 mg kg
1, sum of 16 PAH) the % degradation was similar in both the P. velutina-inoculated and the uninoculated treatments: 94% of the 16 PAHs were degraded in three months. In the field scale experiment the copy number of gram-positive bacteria PAH-ring hydroxylating dioxygenase genes was found to increase 1000 fold, indicating that bacterial PAH degradation also played an important role. Ó 2013 Elsevier Ltd. All rights reserved.

Keywords: Bioremediation Fungi Phanerochaete velutina Polyaromatic hydrocarbons Creosote Contaminated soil

1. Introduction Polyaromatic hydrocarbons (PAHs) are resistant to microbial degradation because of their low water solubility and complex structure with two or more fused benzene rings (Antizar-Ladislao et al., 2004). Since many PAHs are toxic and carcinogenic, their accumulation in the environment is of great concern. A typical source of PAH contamination in soil is coal-tar creosote, which was commonly used to preserve and waterproof crossties and power line poles. Particularly in Finland, former sawmill sites constitute a major problem. In addition to PAHs, such soils may contain chlorophenols, polychlorinated dibenzo-p-dioxins and -furans (PCDD/F), and heavy metals (Kitunen et al., 1987). Bacteria degrade PAH compounds by an assimilative process where they gain carbon and energy for the growth, which typically * Corresponding author. Tel.: þ358 50 573 1529. E-mail address: erika.winquist@aalto.fi (E. Winquist). 0964-8305/$ e see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.ibiod.2013.09.012

leads to mineralization of the compound (Kästner et al., 1994; Haderlein et al., 2006). Bacteria generally use intracellular dioxygenase enzymes for the degradation of PAHs (Johnsen et al., 2005). The aerobic bacterial degradation of PAH compounds is initiated by an oxygenation of the ring structure to form a cis-dihydrodiol followed by a dehydrogenation to form dihydroxylated intermediates. The enzymatic pathway of the PAH degradation and the genes encoding for the corresponding enzymes are well known. A wide array of both gram-positive and gram-negative bacteria are known to degrade PAHs, and their genes are somewhat different (Cebron et al., 2008). Gram-negative bacteria, such as Burkholderia (b-proteobacteria), can easily degrade 2- or 3-ring PAHs, whereas grampositive bacteria, such as Mycobacterium, are more efficient in degrading higher ring PAHs (Johnsen et al., 2005). Ligninolytic fungi are also able to degrade PAHs and they might even be more efficient degraders than bacteria (Davis et al., 1993). When growing on wood or litter, fungi degrade lignin with extracellular oxidizing enzymes in a co-metabolic process (Hatakka,

E. Winquist et al. / International Biodeterioration & Biodegradation 86 (2014) 238e247

2001). These ligninolytic enzymes are non-specific and function through radical reactions. In addition to lignin, they are able to degrade compounds with structural similarities to lignin, such as many xenobiotic organic chemicals, e.g. PAHs (Tuomela and Hatakka, 2011). Fungal ligninolytic enzymes might have an important role in the initial attack on high molecular weight PAHs in soil (Sack et al., 1997; Harms et al., 2011). Since ligninolytic enzymes are extracellular, they are able to diffuse effectively to the highly immobile high molecular weight PAHs. The resulting metabolites are more water soluble, and thus more bioavailable. The formed compounds can be substrates for many bacteria (Sack et al., 1997), but they may also be further degraded by fungal intracellular enzymes, such as cytochrome P-450 monooxygenase (Pozdnyakova, 2012). When the degradation of several PAHs by Irpex lacteus was studied, structures of some of the metabolites suggested involvement of both ligninolytic enzymes and cytochrome P-450 monooxygenase (Cajthaml et al., 2002a, 2006). In addition to mineralization, a significant fraction of PAHs is incorporated into humic substances during bioremediation (Kästner et al., 1994, 1999). Oil-contaminated soil can efficiently be remediated by composting (Jørgensen et al., 2000). Often these soils also contain PAH compounds. Small molecular weight PAHs containing two or three benzene rings are degraded during a well optimized composting process (Antizar-Ladislao et al., 2004). Unlike petroleum oil, creosote oil mainly consists of PAHs. Various strategies for bioremediation of PAH-contaminated soil have been investigated, such as amendment with various types of compost (Semple et al., 2001; Antizar-Ladislao et al., 2004) and bioaugmentation with various fungal species (Pozdnyakova, 2012). Scientific research supports the basis of high diversity bioremediation and has shown that fungal inoculum positively compliments the bacterial communities in soil and in many ways stimulates contaminant degradation  (Kohlmeier et al., 2005; Snajdr et al., 2011; Furuno et al., 2012; Lladó et al., 2012). However, only few studies have reported up-scaling of fungal treatment (Steffen and Tuomela, 2010). The purpose of this study was to develop a bioremediation method that could be used for soils heavily contaminated with persistent organic compounds, such as high molecular weight PAHs. Bioremediation offers a sustainable alternative to other treatment methods, such as thermal treatment and bitumen stabilization (Khan et al., 2004). Bioremediation requires much less energy than thermal treatment. Furthermore, raw material for bitumen stabilization is derived from petroleum and the solution cannot be considered as final because the contaminants are not degraded but only bound to the matrix. The Regulation No 850/ 2004 given by the European Community (2004) declares: “The persistent organic pollutant content in waste is to be destroyed or irreversibly transformed into substances that do not exhibit similar characteristics, unless other operations are environmentally preferable.” Therefore, treatment methods which actually destroy the contaminants should always be the first choice. For a bioremediation process to be successful the microorganisms, or their enzymes, need to be in physical contact with the organic contaminant. Both properties of the soil and the type of the contaminant determine bioavailability and bioaccessibility (Harms, 2011). Bioavailability represents the fraction that is uptaken by the cells, and can cause toxic effects, or can be biodegraded by intracellular mechanisms. The term bioaccessibility, often also called environmental availability, considers the fraction that is potentially available for biota in soils. From the risk assessment point of view this phenomenon is more important than the total concentration, because toxic effects can be attributed to a  carová et al., 2013). contaminant only when it is accessible (Cvan This is also true for biodegradation phenomena that include the

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action of extracellular degradative enzymes, which do not require penetration into the cells and are often overlooked (Leonardi et al., 2007; Covino et al., 2010). Particularly, high molecular weight PAHs have low water solubility and high solid/water distribution ratio. In aged contaminated soil, they might not be bioaccessible because of sorption (adsorption to solid surfaces, absorption in organic matter), physical entrapment (immobilization in micropores) and covalent bonding (oxidative coupling to phenolic compounds) (Kästner et al., 1999). This work examines the addition of both fungal inoculum and composted green waste to a PAH-contaminated sawmill soil in two environments: a laboratory and a field scale. The aged creosote contaminated soil, used in both treatment scales, contained considerable amounts of fluoranthene (35% of 16 PAHs) and pyrene (18% of 16 PAHs), which both contain four benzene rings. Even some five- to six-ring PAHs were present and the total amount of 16 PAHs (listed by the United States Environmental Protection Agency) was high (6000 mg kg
1). We studied the capability of selected ligninolytic fungi to grow in the soil, produce extracellular oxidizing enzymes, and degrade PAH compounds. The addition of composted green waste improved the structure of soil for better aeration and water holding capacity, and provided organic matter as a carbon and energy source for fungi (Semple et al., 2001). To determine the role of bacterial PAH degradation in the treatments, we quantified PAH degradation genes by quantitative PCR. Furthermore, to predict the bioaccessible fraction of PAHs in soil,  sek, supercritical fluid extraction (SFE) was used (Cajthaml and Sa 2005). The biodegradation results were compared with the bioaccessible fraction. Since some of the metabolites formed during bioremediation of the PAH compounds might be more toxic than the parent compounds (Lundstedt et al., 2007), the soil was also tested for ecotoxicity before and after the treatment. 2. Materials and methods 2.1. Soils and soil properties Heavily PAH-contaminated soil (6000 mg kg
1) from a former sawmill was used in the experiments (Table 1). A stock pile of the same origin was used for the field scale experiment, after storage of the excavated soil outdoors for one year. Commercially available composted green waste (Helsinki Region Environmental Services Authority, Finland) was used to dilute the PAH-contaminated soil. Dry matter of the soil was determined by drying the fresh soil at 105  C for 16 h and organic matter content as loss on ignition (mass % of dm) after 4 h at 550  C. Soil pH was measured in 1 mol l
1 KCl solution with a suspension ratio 1:2.5 (w/v). PAH concentration was measured as a sum of 16 PAH compounds namely: naphthalene, acenaphthylene, acenaphthene, fluorene, phenanthrene, anthracene, fluoranthene, pyrene, benzo(a)anthracene, chrysene, benzo(b)fluoranthene, benzo(k)

Table 1 Properties of soils used in the laboratory experiments. Soil

Dry matter (%)

PAH-contaminated soil Diluted PAH-soil (1:1) Composted green waste

93  2a

a b

Organic matter (% dm)

pH

PAH concentration, sum of 16 PAHs (mg kg
1)

4

5.8

6000

76  2

10

6.6

3500

59  2

16

7.4

ndb

Average value of three replicates  SD. nd ¼ not determined.

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fluoranthene, benzo(a)pyrene, indeno(1,2,3-cd)pyrene, dibenzo(a,h)anthracene and benzo(g,h,i)perylene. For quantitative PAH determination, the soil samples were extracted using ultrasonic method according to US EPA method 3550C. Approximately 8 g of soil (wet weight) were placed in an ultrasonic bath in tightly closed flask with 30 ml of acetone for 30 min. Afterwards, the solvent contents of the extraction vessels were centrifuged (5000 g, 10 min). Aliquots (1 ml) were transferred to 2 ml auto sampler vials and 100 ml of deuterated internal-standard solution was added (naphthalene-D8, acenaphthene-D10, phenanthrene-D10, chrysene-D12, perylene-D13) to each sample. The samples were then analysed using gas chromatography-mass spectrometry system (5977A Series GC/MSD, Agilent, USA). The samples were analysed using split/splitless injection method following the standard ISO 18287:2006 (International Standardization Organization, 2006) (SGS Inspection Services Oy, Finland). The individual PAHs were monitored with single-ion monitoring method (SIM) using specific ions of the respective PAH representatives. The dry weights of the extracted soil samples were determined according to the standard ISO 11465:1993 (International Standardization Organization, 1993). In addition to PAHs, heavy metals (As, Cd, Cr, Cu, Ni, Pb, Zn, Hg) were analysed from the soil (SGS Inspection Services Oy, Finland). The concentration of arsenic (28 mg kg
1) was slightly higher than natural concentrations (0.1e25 mg kg
1) in Finnish soils, but the concentrations of the other heavy metals were at naturally occurring levels (results not shown). 2.2. Bioaccessibility of PAH compounds The PAH bioaccessibility analysis was performed using a SFE extractor (Labio a.s., Czech Republic). The soil samples (1 g) were placed at the bottom of a 2 ml extraction cell and the dead volume was filled with sodium sulphate. The samples were extracted by a downward stream of carbon dioxide at 50  C and 200 bar. The flow rate was set by a manual restrictor to 1 ml min
1. The PAH analytes were collected on octadecyl silica at
20  C and eluted with 12 ml of acetone at 20  C. The extractions were carried out in three parallel runs and the compounds were collected after 5, 10, 20, 40, 60, 80, 120, 160 and 200 min intervals. The acetone extracts were analysed by HPLC (Waters 2695) equipped with a diode array (Waters 2996) and a fluorescence detectors (Waters 2475) (Covino et al., 2010). The desorption kinetic curves of the individual PAHs were modelled with an empirical two-site model (Williamson et al., 1998):

with extensive screening of 147 fungal strains (Valentin et al., 2009), ten strains were selected for screening experiments: Agrocybe dura FBCC478 (MW71-2), Agrocybe praecox FBCC587 (K163), Gymnopilus luteofolius FBCC466 (X9), Irpex lacteus FBCC1012 (CCBAS617), Mycena galericulata FBCC598 (K175), Phanerochaete velutina FBCC941 (T244), Physisporinus rivulosus FBCC939 (T241i), Stropharia aeruginosa FBCC521 (K47), Stropharia rugosoannulata FBCC475 (11372) and Trametes ochracea FBCC1011 (HAM9) (old strain number in parenthesis). The strains were maintained on malt extract agar plates at þ4  C and sub-cultured at least every 3 months. 2.4. Laboratory experiments 2.4.1. Fungal inoculum grown in liquid Liquid inoculum was prepared by cutting four (5 mm  5 mm) agar plugs from malt extract agar plates. These were extruded through a syringe to the sterile liquid medium in 500 ml Erlenmeyer flasks containing 200 ml malt extract (2% w/v). Fungi were cultivated for 6e7 days at 25  C with continuous agitation (100e150 rpm). 2.4.2. Fungal inoculum grown on bark Scots pine (Pinus sylvestris) bark is a very promising substrate for fungal inoculum cultivation (Valentin et al., 2010). It is a by-product from pulp, paper and timber production. Pine bark is composed mainly of lignocellulose and it provides nutrients to fungi. It is also a selective growth substrate, since it contains phenolic extractives with antimicrobial properties. The fungi were cultivated on bark in high density polyethene (HDPE) plastic bags. Bark was first soaked in water overnight and then drained. Wet bark, 1 kg in each plastic bag, was autoclaved (30 min, 121  C) and the cooled bark was inoculated with 200 ml of liquid inoculum. The bags were incubated at room temperature (21  C) and aerated continuously with moisturized air (1 l min
1). The incubation time varied from three to five weeks until the surface of the bark was fully covered with fungal mycelium. The average dry matter and organic matter contents of the “ready to use” bark inoculum were 30% and 95% (Alén, 2011) respectively. To study the enzyme activity of the fungal inoculum, cultivations were performed in small plastic jars (100 g of wet bark in each plastic jar) in similar conditions as in the 1 kg scale. Liquid inoculum (20 ml) was homogenized at 17500 rpm for 10 s (UltraTurrax T25, IKA-Werke, Germany) and mixed with 100 g of wet bark. One jar represented one sample. Ten samples (with three replicates) were taken for enzyme activity measurements, from 7 to 42 days after inoculation, with three to four days interval.

St ¼ F S0 expð
k1 tÞ þ ð1
FÞS0 expð
k2 tÞ; where St is the pollutant concentration remaining in the soil after extraction using SFE at time t; F is the fraction of rapidly released chemical; S0 is the original concentration of the pollutant in the soil (the total concentration extracted by Accelerated Solvent Extractor, ASE); and k1 and k2 are the first-order rate constants. The parameter F (Fast fraction) represents bioaccessible fraction of each PAH and it was averaged from the three independent analyses. The value ranges from 0 to 1, where 1 represents 100% bioaccessibility. The amount after exhaustive analytical accelerated solvent extraction was considered as the total concentration   (100%) of each PAH (Cvan carová et al., 2013). 2.3. Fungal strains Fungal strains were obtained from the Fungal Biotechnology Culture Collection (FBCC) of the Department of Food and Environmental Sciences, University of Helsinki. Based on an earlier study

2.4.3. Enzyme activity analysis Manganese peroxidase (MnP) and laccase activities were determined from bark with fungal growth. Twenty-five grams of fresh bark was suspended in 50 ml of 50 mM sodium phosphate buffer (pH 6.5), and the suspension was mixed continuously (33 rpm) on a roller mixer for 1 h at room temperature. Afterwards, it was filtered through a nylon cloth, and the filtrate was centrifuged at 4300  g for 15 min. Enzyme activity was determined from the supernatant. Laccase activity was determined with 2,20 -azinobis(3-ethylbenzthiazoline-6-sulfonate) (ABTS) as the substrate, according to the method of Niku-Paavola et al. (1988). The measurements were carried out in 25 mM sodium succinate buffer (pH 4.5) with a spectrophotometer (Shimadzu UV-2100, Japan) at 436 nm (ε436 ¼ 29300 M-1 cm
1). MnP activity was measured at 270 nm by following the formation of Mn(III)-malonate complexes as described by Wariishi et al. (1992) and modified by Hofrichter et al. (1998) (ε270 ¼ 11590 M-1 cm
1). The activities are expressed as mU g-1 dm of bark.

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2.4.4. Screening of fungal strains The screening experiments were done with five different growth media: 1) PAH-contaminated soil, 2) diluted PAHcontaminated soil (1:1), 3) composted green waste (Table 1), 4) sandy soil mainly contaminated with oil hydrocarbons and 5) slightly contaminated organic soil. In these two last soils the concentration of oil hydrocarbons (C5eC40) was 30,000 mg kg
1 and PAHs 140 mg kg
1 in sandy soil, and 2300 mg kg
1 and 220 mg kg
1 respectively in organic soil (not included in Table 1). A Petri-dish was filled with non-sterile soil, moistened if needed, and 2.5 g of pine bark with fungal mycelium was placed on top of the soil. Fungal growth was examined visually during 30 days of incubation at room temperature in dark. Moisture content of the soil was kept constant by adding water when needed. 2.4.5. Degradation experiments in aerated bottles None of the screened fungi showed visible growth in undiluted PAH-contaminated soil. Thus, in further experiments PAHcontaminated soil was mixed with composted green waste (1:1, on wet mass basis) and sieved through 5 mm mesh prior to use for laboratory experiments (starting concentration 3500 mg kg
1). Two-litre glass bottles were filled with a layer of expanded clay (150 g) at the bottom, followed by layers of moist soil (400 g), fungal inoculum growing on bark (100 g), and a top layer of moist soil (400 g). Control bottles contained only expanded clay (150 g) and soil (800 g) without fungal inoculum. All bottles were aerated with moist air (0.5 l min
1). The inlet air tube was placed to the expanded clay layer at the bottom, so that air would be spread evenly and move up through all layers. Carbon dioxide (CO2) was measured on-line from the exhaust air by CO2 analyser (custom made instrument based on IR absorbance measurement, eLabs Oy Engineering, Finland). CO2 production during the experiment was calculated by subtracting inlet air CO2 concentration from exhaust CO2 concentration, and after that calculating the cumulative CO2 production, when the aeration rate was kept constant. Since the amount of total organic matter (including bark) affects the CO2 evolvement, the CO2 production was presented as g per 100 g of organic matter (loss of ignition).

241

90  C for 1 h, the cabinet was then allowed to cool overnight, and steaming was repeated the next day. This type of heat treatment is called tyndallisation. The first steaming will kill the growing cells, but not the spores. On the second day the spores, that survived the first steaming, have germinated into growing cells. These cells will be killed by the second steaming. The bark (3.5 kg) in each cultivation basket was inoculated with 400 ml of liquid inoculum (P. velutina) and the fungus was cultivated for five weeks at room temperature (21  C). Moist air was circulated into the cultivation cabinet (30 l min
1). At the end of the cultivation, the surface of the bark was fully covered with fungal mycelium. 2.5.2. Field scale experiment in aerated soil piles PAH-contaminated soil was mixed with composted green waste (1:1) on wet weight basis (starting concentration 1400 mg kg
1). Altogether four tons of soil-compost mixture were divided into two piles. The fungal inoculum (70 kg) was added to one pile in two layers: the lower layer contained 42 kg of inoculum and the upper 28 kg of inoculum. The other pile was left without inoculum. The length of one pile was 2.0 m and the maximum height was 0.6 m. Both piles were aerated with moist air (70 l min
1) and covered with a tarpaulin to avoid drying of the soil. During incubation, data loggers measured temperature eight times a day inside and outside the piles. The field scale experiment was carried out at a soil treatment facility in a non-heated hall with a concrete floor. 2.5.3. Sampling and analyses Two parallel composite soil samples were taken with an Edelman auger (Eijkelkamp Agrisearch Equipment BV, the Netherlands) once a month from both piles. One composite sample consisted of four sub-samples taken at 0.3 m depth. The sub-samples were mixed together and sieved through 8 mm mesh. Samples were analysed for dry matter, organic matter, pH and PAH concentration. In addition, the P. velutina ITS region DNA copy number, the bacterial 16S rDNA copy number and quantification of dioxygenase genes involved in PAH degradation were determined. The first and last samples were run for ecotoxicological tests. 2.6. Microbiological analyses

2.5. Field scale experiment 2.5.1. Fungal inoculum for the field scale experiment Production of fungal inoculum on bark was scaled up for the field experiment (Steffen and Tuomela, 2010). Two cabinets were built from stainless steel sheets and 12 plastic dishwasher baskets (length 50 cm, width 50 cm, height 10 cm) were placed inside both cabinets as shelves. The capacity of one basket was 3.5 kg of wet bark, thus the total capacity of two cabinets was 84 kg. Moist pine bark was surface sterilized with steam, which was conducted into the cultivation cabinet from the bottom and led through the baskets. The temperature of the cabinet was kept over

The total amount of indigenous bacteria was estimated by analysing the number of 16S rDNA genes in the soil sample. All prokaryotic cells contain on average four copies of this gene. The PAH degradation potential by gram-positive and gram-negative bacteria was estimated by analysing the number of ring hydroxylating dioxygenase (RHD) genes of both bacterial classes (Cebron et al., 2008). The growth of P. velutina was also evaluated by a DNA quantification method using specific primers designed for the  ITS region of this fungus (Snajdr et al., 2011). Since there are usually more than one ITS region in a fungal cell, the cell number of P. velutina is not directly comparable with the amount of ITS regions

Table 2 Sequences, annealing temperatures (Ta) and temperatures for measurements of fluorescence (FimT) for the primers used in gene quantification for this study. Target

Primer

Sequence 50 -3’a

Ta ( C)

FimT ( C)

Reference

P. velutina ITS region

PvF PvR PAH-GNf PAH-GNr PAH-GPf PAH-GPr Eub338 Eub518

AAC GCA CCT TGC GCT CCC T CTT CAC GAC CAC GGC GCA GA GAG ATG CAT ACC ACG TKG GTT GGA AGC TGT TGT TCG GGA AGA YWG TGC MGT T CGG CGC CGA CAA YTT YGT NGG GGG GAA CAC GGT GCC RTG DAT RAA ACT CCT ACG GGA GGC AGC AG ATT ACC GCG GCT GCT GG

69, 1 min

75

 Snajdr et al., 2011

57

80

Cebron et al., 2008

54

80

Cebron et al., 2008

53

80

Lane 1991 Muyzer et al., 1993

PAH genes in gram-negatives PAH genes in gram-positives Bacterial 16S rDNA a

K ¼ G or T; Y ¼ C or T; W ¼ A or T, M ¼ A or C, R ¼ A or G, N ¼ A or C or G or T, D ¼ G or A or T.

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in a sample. However, comparisons can be made how the P. velutina ITS region DNA copy number is changing over time. P. velutina ITS region, bacterial 16S rDNA, and PAH-ring hydroxylating dioxygenase (RHD) gene copy numbers were estimated using quantitative PCR (qPCR) methods (Table 2). Quantification was done by comparing the fluorescence of samples to standard samples containing known amounts of the same genes cloned into TOPO TA plasmids using cloning kits (Invitrogen). For qPCR the Maxima SYBR Green/ROX qPCR 2 master mix was used (Fermentas, Thermo Fisher Scientific), except for 16S qPCR where QuantiTect SYBR Green PCR 2 master mix was used (Qiagen). All the analyses were performed in 25 ml reaction volumes using an Applied Bioscience 7300 Real Time PCR System (CA, USA). The concentration of the primers was 0.3 mM, except for P. velutina qPCR 0.6 mM. In P. velutina qPCR, bovine serum albumin (BSA) was used at a concentration of 0.45 mg ml
1. The first qPCR step was always a denaturation step (95  C for 10 min), after which the following cycle was repeated forty times: 95  C for 15 s, annealing at primer specific temperatures for 30 s, elongation at 72  C for 30 s, and measurement of fluorescence at qPCR specific FimT temperatures. Controls without added template were routinely used as negative controls. 2.7. Ecotoxicological tests 2.7.1. Germination tests Garden cress (Lepidium sativum) and red clover (Trifolium pratense) were used for germination tests. The method is a modification of ISO 17126:2008 (International Standardization Organization, 2008d) and ISO 11269-2:2008 (International Standardization Organization, 2008c). Three replicate samples and 6 controls were tested using 40 seeds for one Petri-dish (Ø14 cm). One hundred grams of experimental soil, or clean sand (particle size 0.3e0.9 mm) for controls, was placed on the dish and ultrapure water was added to achieve approximately 85% of water holding capacity. Seeds were placed on the moist test material, covered with 90 g of quartz sand (particle size 0.7e1.2 mm) and incubated at 20  2  C. The total time of incubation was 5 d for the garden cress and 6 d for the red clover. Incubation was started with a 48 h dark period and continued with light/dark cycle (16/8 h, 4300  430 lux) until the end of the incubation. At the end of the test, the number of emerged seedlings was counted. The result was calculated as a percentage of inhibition in comparison with the germination of the seeds in controls. 2.7.2. Test with earthworms Earthworm (Eisenia fetida) tests were performed as a modification of the standards ISO 11268-1:2008 and ISO 11268-2:2008 (International Standardization Organization, 2008a,b). The acute and reproduction tests were combined in a single protocol: survival was recorded after 4 weeks’ exposure, the adult earthworms were then removed, and the test was continued for additional 4 weeks to determine the effects on reproduction. A mixture of equal volumes of artificial soil and commercial garden soil was used as the control soil in earthworm tests. Water was added to achieve 50% of water holding capacity before adding the earthworms. Four replicate samples, containing 300 g soil and 10 earthworms per vessel, were prepared for the tests. The number of the juveniles was counted at the end of the incubation at 20  C. 3. Results 3.1. Laboratory experiments The capability of ten selected fungal strains to grow in the soil, compete against indigenous micro-organisms, and tolerate the

Fig. 1. Total number of indigenous bacteria (16S rDNA copy numbers) and PAH-ring hydroxylating dioxygenase genes involved in PAH degradation in gram-positive and gram-negative bacteria in soil in the laboratory experiment (SDs are shown with error bars). PAH-contaminated soil diluted with composted green waste (1:1) was incubated 3 months with or without fungal inoculum.

contaminants was screened. Based on the screening, P. velutina, S. rugosoannulata and G. luteofolius were selected for laboratory degradation experiments (results not shown). None of the fungi showed visible growth in the original PAH-contaminated soil at the PAH concentration of 6000 mg kg
1. Further screening was performed in PAH-contaminated soil, which was diluted with composted green waste (1:1). This soil was still partially toxic to most of the fungi, only P. velutina showed maximum visible growth in the diluted PAH-contaminated soil. Oil-contaminated soil inhibited the growth of all the experimental fungi. However, minor growth was observed with the selected fungal species on plates with oilcontaminated soil. The activities of laccase and MnP were determined from the fungal inoculum grown on bark. The laccase activities were very low (<100 mU g-1 dm, results not shown). On the contrary, P. velutina and S. rugosoannulata produced high MnP activities. The maximum MnP activities were 2300  1100 mU g-1 dm for P. velutina (after 21 d of cultivation), 900  800 mU g-1 dm for S. rugosoannulata (after 35 d of cultivation) and 140  90 mU g-1 dm for G. luteofolius (after 24 d of cultivation). The enzyme production appeared together with the fungal growth. The incubation time varied from three to five weeks, after which the surface of the bark was fully covered with fungal mycelium. With G. luteofolius and P. velutina the fungal inoculum was ready to use after three weeks and with S. rugosoannulata after five weeks of the solid state cultivation. PAH-contaminated soil, and particularly composted green waste, contained indigenous micro-organisms. The total amount of bacteria was nearly the same in all samples (Fig. 1). When the fungal inoculum was added to the soil, the PAH degradation potential by gram-positive bacteria slightly increased and by gramnegative bacteria slightly decreased. During microbial growth, bark and soil organic matter are degraded and CO2 is produced. The CO2 production tripled in bottles with fungal inoculum, when compared to the control without inoculum (Fig. 2). Treatment with S. rugosoannulata inoculum produced in 3 months approximately 10% more CO2 than the other fungal treatments. Also in a previous study S. rugosoannulata produced more CO2 than P. velutina and G. luteofolius when growing in soil (Winquist et al., 2009). When the fungal inoculum was added to the soil, 95e96% of total PAHs were degraded (Fig. 2). PAH compounds were degraded even in the control bottle without the fungal inoculum to less than one third of the original concentration. However, the degradation of individual PAH compounds showed that 4-ring or even larger

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Fig. 2. CO2 production per 100 g of organic matter (light columns) and PAH concentration in soil (dark columns) before incubation (untreated soil) and after 3 months incubation in the laboratory experiment (average value of three replicates, SDs are shown with error bars). PAH-contaminated soil diluted with composted green waste (1:1) was incubated 3 months with or without fungal inoculum.

PAH molecules were degraded significantly more when the fungal inoculum was used than by indigenous microbes alone (Table 3). The highest degradation of high molecular weight PAHs was reached when P. velutina inoculum was added. This treatment (P. velutina together with indigenous microbes) was able to degrade 96% of 4-ring PAHs and 39% of 5- and 6-ring PAHs. In the control bottles 55% of 4-ring PAHs and only 7% of 5- and 6-ring PAHs were degraded. When the biodegradation results with the addition of P. velutina inoculum were compared to the bioaccessible fraction, it could be noticed that they were very close to each other. Thus, it can be concluded that the whole bioaccessible fraction was degraded by P. velutina together with indigenous microbes. In control soil approximately half of the bioaccessible 4-ring PAHs and most of the bioaccessible 5- and 6-ring PAHs were still present after three months incubation. 3.2. Field scale experiment The field experiment was performed during the summer season in Finland (from the beginning of June until the end of October). During this time, the temperature inside the soil piles and the

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experiment hall varied between 6  C and 23  C, and between 2  C and 30  C, respectively. Aeration with moist air worked well and covering the piles helped to keep suitable moisture in the soil. Dry matter (66e71%) and organic matter (13e16% of dm) of the two piles varied only little. The pH decreased slightly during the experiment in both piles: in the fungal treatment pile from 7.2 to 6.8 and in the control pile from 7.3 to 7.0. In the field experiment 94% of total PAHs were degraded in both piles already in three months although the field experiment was continued for five months (Fig. 3). PAH compounds were degraded in both piles equally: 92% of 3-ring PAHs, 95% of 4-ring PAHs and 33% of 5- and 6ring PAHs. Bark adsorbed 0.2% of PAHs. Fungal mycelium was clearly visible on the surface of the pile after three months of incubation. The visual growth corresponded well with the P. velutina ITS region DNA copy number (Fig. 4), which was approximately thousand-fold higher in most of the samples taken from the fungal treatment pile than from the control pile. The total amount of indigenous bacteria was estimated with bacterial 16S rDNA copy number. It was constant during the field experiment in both piles. The copy number was equivalent to laboratory experiments (1010 copies g-1 dm). In addition, dioxygenase genes involved in PAH degradation were measured. Although the amount of dioxygenase genes in the two piles (inoculum versus no inoculum) did not vary, the number of dioxygenase genes in gramnegative and gram-positive bacteria differed greatly (Fig. 5). While the copy number of gram-negative bacteria was almost the same during the whole field experiment, the copy number of grampositive bacteria increased thousand fold (from 105 to 108 copies g-1 dm) during the first month of the experiment. The toxicity of soil was determined by survival of earthworms and inhibition of seed germination (Table 4). In the beginning of the field experiment the soil diluted with composted green waste was still very toxic and all earthworms died within three days from the beginning of the incubation. After the field experiment, however, acute toxicity had almost disappeared and reproduction in treated soil was even higher than in the control soil. Moreover, the toxicity of the soil in the beginning of the field experiment inhibited the seed germination even though the effect was not as strong as with the earthworms. The toxicity of the soil decreased during the field experiment also as measured by seed germination (Table 4).

Table 3 PAH concentrations, degradation and bioaccessibility in laboratory experiments with PAH-contaminated soil diluted with composted green waste (1:1). PAH compound (No. of rings) Naphthalene (2) Acenaphtylene (3) Acenaphthene (3) Fluorene (3) Phenanthrene (3) Anthracene (3) Fluoranthene (4) Pyrene (4) Benzo(a)anthracene (4) Chrysene (4) Benzo(b)fluoranthene (5) Benzo(k)fluoranthene (5) Benzo(a)pyrene (5) Dibenzo(a,h)anthracene (5) Indeno(1,2,3-cd)pyrene (6) Benzo(g,h,i)perylene (6) Sum of 16 PAHs a b c

Untreated soila Conc. (mg kg <0.2 10 297 208 735 62 1279 720 53 59 15 13 8.9 1.5 4.5 2.7 3469  140b


1

Aerated soila dm)

Bioaccess. Fraction (%) c

nf 96 nf 96 96 95 95 95 84 83 30 44 nf nf 54 43

Conc. (mg kg <0.2 5.0 9.8 4.6 4.8 8.0 345 535 31 39 15 11 8.7 1.6 4.0 2.5 1025  4

P. velutina treatmenta
1

dm)

Degradation (%)

Conc. (mg kg
1 dm)

Degradation (%)

51 97 98 99 87 73 26 42 34 1 15 3 0 12 10

<0.2 3.5 0.4 0.8 0.7 5.6 22 41 6.6 14 10 6.8 5.9 1.1 2.6 1.4 123  11

65 100 100 100 91 98 94 88 76 33 47 34 30 41 47

Untreated soil: no incubation, Aerated soil: incubation time 3 months, P. velutina treatment: incubation time 3 months. Average value of three replicates  SD. nf ¼ not fitting the model.

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Fig. 3. The PAH concentration during the field experiment with or without fungal inoculum (average value of two replicates, the range of variations are shown with error bars). PAH-contaminated soil diluted with composted green waste (1:1) was used.

Fig. 5. Amount of dioxygenase genes involved in PAH degradation in gram-negative and gram-positive bacteria in soil during the field experiment with or without fungal inoculum (SD: 0.02e0.42 log, not shown in the figure). PAH-contaminated soil diluted with composted green waste (1:1) was used.

4. Discussion The most efficient bioremediation fungus in our study was P. velutina. The treatment, where P. velutina inoculum was added to the soil, was able to degrade the whole bioaccessible fraction of the PAH compounds in the laboratory experiment: 96% of total PAHs were degraded in three months. Covino et al. (2010) determined bioaccessible fraction by SFE as in our work, and observed that treatment with Pleurotus ostreatus degraded 80% of total PAHs in non-sterile creosote contaminated soil, which was approximately the whole bioaccessible fraction (82%) (Table 5). In our experiment, PAH compounds were degraded also in the control soil without the fungal inoculum by indigenous micro-organisms. The addition of composted green waste resulted in a high number of indigenous bacteria as well as genes involved in PAH degradation from grampositive and gram-negative bacteria. In the laboratory scale, almost all 3-ring PAHs and approximately half of the 4-ring PAHs were degraded, but the amount of 5- and 6-ring PAHs remained nearly the same. High molecular weight PAHs are not easily degraded by bacteria, because the solubility in water decreases with the increase in molecular mass. In the field, P. velutina grew extensively into the fungal treatment pile, yet the total amount of indigenous bacteria was found to be high both in the fungal treatment pile and in the control pile. The

dilution of contaminated soil with composted green waste influenced the microbial activity in three ways: 1) PAH concentration was diluted and the soil was less toxic, 2) soil composition was changed to favour microbial growth and 3) the composted green waste itself acted as a strong microbial inoculant. Approximately 94% of total PAHs were degraded in both piles in three months. The use of fungal inoculum did not increase the degradation of PAHs as in the laboratory experiments. Possible reason for this was a lower PAH concentration before the treatment in field scale (1400 mg kg
1) compared to that in laboratory scale (3500 mg kg
1). Since the soil was less toxic, the indigenous microbial activity was higher and bioremediation was enhanced. More work is needed to find out when the addition of fungal mycelium is beneficial. During the field experiment the number of dioxygenase genes in gram-positive bacteria grew thousand fold (from 105 to 108 copies g-1 dm) in both piles. Therefore, it could be concluded that in the field experiment with composted green waste, grampositive bacteria most likely had a significant role in PAH degradation. In an earlier study, where bacteria were isolated from PAHcontaminated soil, most species that could utilize PAHs as sole carbon source were gram-positive actinobacteria (Kästner et al., 1994). Also gram-positive mycobacteria are known to be able to efficiently degrade particularly 4-ring PAHs (López et al., 2008). Composted green waste most likely contained also indigenous fungi, although their presence was not directly quantified. However, P. velutina DNA copy number in the control pile without the fungal inoculum was elevated, indicating that the primers were not completely specific and that fungi, related to P. velutina, were

Table 4 Ecotoxicity tests with earthworms, garden cress and red clover, samples taken before (start) or after (inoculum/no inoculum) the field experiment with PAHcontaminated soil mixed with composted green waste (1:1).

Fig. 4. The P. velutina ITS region DNA copy number in soil during the field experiment with and without fungal inoculum (detection level is shown as a black line, SDs are marked with error bars). PAH-contaminated soil diluted with composted green waste (1:1) was used.

Soil

Acute death rate of earthworms (%)

Reproduction of earthworms

Inhibition with garden cress (%)

Inhibition with red clover (%)

Control soil (clean) Start No inoculum (5 months) Inoculum (5 months)

0

218  41a

0

0

100 3

0 312  32

24 0

19 2

0

343  34

8

12

a

Average value of four replicates  SD.

E. Winquist et al. / International Biodeterioration & Biodegradation 86 (2014) 238e247 Table 5 Comparison of laboratory scale remediation with fungi of PAH-contaminated nonsterile soil from various wood treatment plants.

PAH concentration before the treatment (mg kg
1) Fungus Working scale (g) Degradation (bioaccessible): Total PAHs (%) 3-ring PAHs (%) 4-ring PAHs (%) 5-/6-ring PAHs (%) Incubation time (d) a

Eggen 1999

Covino et al., 2010

This work

1900

2300

3500

Pleurotus ostreatus 3000

Pleurotus ostreatus 25

Phanerochaete velutina 800

86 89 87 48 49

80 99 71 51 60

(82)a (96) (77) (28)

96 99 96 39 90

(94) (96) (94) (39)

bioaccessible fraction in parenthesis after the degraded fraction.

present. Fungi affect PAH degradation also with other mechanisms than direct metabolism. Saprotrophic fungi decompose organic matter in soil with the help of extracellular hydrolysing enzymes. Since small molecular mass nutrients are released outside the cell they are available for both their producers and other soil micro organisms. Snajdr et al. (2011) found in their studies a positive correlation between bacterial and fungal biomass. Fungi also increase the bioavailability of immobile and unevenly distributed PAHs for bacterial degradation both by increasing bacterial mobility and by transporting PAH compounds within fungal mycelia. Bacteria may use liquid films around fungal hyphae for bridging airfilled pores and penetrating soil aggregates (Kohlmeier et al., 2005). Fungi may also take up and actively translocate PAHs through their hyphae via cytoplasmic streaming (Furuno et al., 2012). Since we observed no difference between the fungal treatment and the control treatment, we compared our results with composted green waste (control pile, without inoculum) to two other reported composting experiments in field scale (Table 6). There are two approaches how to utilize compost in bioremediation: 1) adding primary compost ingredients or 2) adding mature compost to contaminated soil (Semple et al., 2001). When the primary compost ingredients, e.g. wheat straw and chicken manure, are mixed with the contaminated soil an increase in temperature and amount of thermophilic micro-organisms follows. When mature compost is added to contaminated soil almost no increase in

Table 6 Comparison of field scale experiments where compost was used for bioremediation of PAH-contaminated soil.

Origin of soil Compost Soil to compost dilution ratio Concentration after dilution (mg kg
1) Working scale (kg) Degradation: Total PAHs (%) 3-ring PAHs (%) 4-ring PAHs (%) 5-/6-ring PAHs (%) Incubation time (d)

Cajthaml et al., 2002b

 sek et al., 2003 Sa

This work

Tar producing plant Mushroom compost 1:4

Manufactured gas plant Mushroom compost 1:4

Sawmill Composted green waste 1:1

1800

600

1400

500

1000

2000

64 80 65 54 142

69 88 61 57 154

94 92 95 33 92

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temperature follows, but the indigenous micro-organisms are activated by more favourable growth conditions. Cajthaml et al.  sek et al. (2003) applied mushroom compost, a (2002b) and Sa “ready to use” compost mix without fungal inoculum (Table 6). After mixing compost to soil an actual composting process started with a fast increase in temperature. These experiments can be compared to our control pile as we incubated soil with mature green waste compost, but without a major increase in temperature. We obtained higher degradation of total PAHs with lower dilution rate. However, the comparison is difficult since properties of both the soil and the contaminants affect the bioremediation process. All the soils in these experiments were from a different origin: from a tar producing plant (Cajthaml et al., 2002b), a manufactured gas  sek et al., 2003) and a sawmill (this work). Nevertheless, plant (Sa also the bioremediation process has an important effect on the PAH degradation. Haderlein et al. (2006) noticed that pyrene degradation was more efficient in ambient than at high (55  C) temperature. At ambient temperature a higher diversity of micro-organisms are present, and proteobacteria, actinobacteria, and fungi can work in co-operation (Antizar-Ladislao et al., 2004). In addition, higher pyrene mineralization was reached with mature compost than with fresh compost (Haderlein et al., 2006). According to the ecotoxicity tests, a dramatic improvement in the soil quality occurred during the field scale treatment. Survival and reproduction of earthworms demonstrated that the soil was suitable as a habitat for these animals after treatment, while the soil before treatment was acutely toxic. This is in agreement with re  sults of Cvan carova et al. (2013) who documented that earthworms are very sensitive towards bioaccessible PAHs. Seed germination test was not as good indicator as earthworm test, but the trend was the same. This is not surprising, since it is well known that germination tests are not as sensitive to contaminants as are several soil invertebrates (Dorn et al., 1998; Saterbak et al., 1999). The effects on plants are important factors in the assessment of soil quality or remediation efficacy, and for making decision on the possible use of the remediated material. When the soil is intended for landscaping, several plant species should be tested, and plant growth tests would be of additional value. We demonstrated 94% degradation of PAHs in a field scale experiment with composted green waste and with or without fungal inoculum. The residual concentration of total 16 PAHs was 80e100 mg kg
1. Even with an effective bioremediation process some residual contamination is left in the soil, but this is comparable to the bioaccessible fraction of PAHs in soil (Reichenberg et al., 2010). However, the actual risk of exposure is more important than the total concentration. The evaluation for reuse of this soil, for example in various landscaping, should be based on ecotoxicological tests and bioaccessibility analysis. Acknowledgements The authors thank Petr Baldrian (Institute of Microbiology, Academy of Sciences of the Czech Republic) for his co-operation in the development of a DNA identification method for P. velutina. We also thank Laura Häkkinen, Ilse Heiskanen and Minna Sepponen (Finnish Environment Institute) for technical assistance with the DNA methods and ecotoxicological analysis, Siiri Viljanen (Aalto University School of Chemical Technology) for assistance with the enzyme activity measurements, and Seppo Jääskeläinen and Pekka Koivulaakso (Aalto University School of Chemical Technology) for taking part in designing and building the aeration system used in the field scale experiment. This research was funded by the Finnish Funding Agency for Technology and Innovation (TEKES) through SYMBIO program and by the companies who took part in the program: Ekokem-Palvelu Oy, Suomen

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Erityisjäte Oy (former Niska & Nyyssönen Oy), Soilrem Oy, Mzymes Oy, Ramboll Finland Oy, the Finnish Defence Forces, and the City of Helsinki. Particularly, we thank Riina Rantsi from Suomen Erityisjäte Oy for offering us a place to perform field scale studies. The work was also supported by Competence Center TE01020218 of the Czech Technology Agency. References Alén, R., 2011. Structure and chemical composition of biomass feedstocks. In: Alén, R. (Ed.), Biorefining of Forest Resources. Bookwell Oy, Porvoo, Finland, pp. 18e54. Antizar-Ladislao, B., Lopez-Real, J.M., Beck, A.J., 2004. Bioremediation of polycyclic aromatic hydrocarbon (PAH)-contaminated waste using composting approaches. Crit. Rev. Environ. Sci. Technol. 34, 249e289.  sek, V., 2005. Application of supercritical fluid extraction (SFE) to Cajthaml, T., Sa predict bioremediation efficacy of long-term composting of PAH-contaminated soil. Environ. Sci. Technol. 39, 8448e8452.  sek, V., Popp, P., 2002a. 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