Effects of Facilitated Bacterial Dispersal on the Degradation and

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Effects of facilitated bacterial dispersal on the degradation and emission of a desorbing contaminant Sally Otto, Thomas Banitz, Martin Thullner, Hauke Harms, and Lukas Y. Wick Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.6b00567 • Publication Date (Web): 19 May 2016 Downloaded from http://pubs.acs.org on May 25, 2016

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Environmental Science & Technology

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Effects of facilitated bacterial dispersal on the degradation and emission of a

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desorbing contaminant

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Sally Otto1, Thomas Banitz2, Martin Thullner1, Hauke Harms1,3, Lukas Y. Wick1*

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Microbiology, Permoserstr. 15, 04318 Leipzig, Germany

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2

UFZ - Helmholtz Centre for Environmental Research, Department of Environmental

UFZ - Helmholtz Centre for Environmental Research, Department of Ecological

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Modelling, Permoserstr. 15, 04318 Leipzig, Germany

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Deutscher Platz 5e, 04103 Leipzig

German Centre for Integrative Biodiversity Research (iDiv) Halle-Jena-Leipzig,

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Running title: Effects of facilitated bacterial dispersal

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Intended for: Environmental Science and Technology

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*

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UFZ. Department of Environmental Microbiology; Permoserstr. 15; 04318 Leipzig,

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Germany. phone: +49 341 235 1316, fax: +49 341 235 1351, e-mail: [email protected].

Corresponding author: Mailing address: Helmholtz Centre for Environmental Research -

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ACS Paragon Plus Environment

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ABSTRACT ART

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ABSTRACT

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The quantitative relationship between a compound’s availability for biological removal

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and eco-toxicity is a key issue for retrospective risk assessment and remediation

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frameworks. Here, we investigated the impact of facilitated bacterial dispersal on a model

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soil-atmosphere interface on the release, degradation and outgassing of a semi-volatile

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contaminant. We designed a laboratory microcosm with passive dosing of phenanthrene

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(PHE) to a model soil-atmosphere interface (agar surface) in the presence and absence of

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glass fibers known to facilitate the dispersal of PHE-degrading Pseudomonas fluorescens

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LP6a. We observed that glass fibers (used as a model to mimic a fungal hyphal network)

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resulted in (i) increased bacterial surface coverage, (ii) effective degradation of matrix-

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bound PHE, and (iii) substantially reduced PHE emission to locations beyond the

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contamination zone even at low bacterial surface coverage. Our data suggest that bacterial

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dispersal networks such as mycelia promote the optimized spatial arrangement of

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microbial populations to allow for effective contaminant degradation and reduction of

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potential hazard to organisms beyond a contaminated zone.

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KEYWORDS: Bacterial dispersal network, biodegradation, Pseudomonas fluorescens

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LP6a, PAH, phenanthrene, passive dosing


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INTRODUCTION

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Various environmental chemicals (contaminants) serve as substrates for microorganisms,

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while simultaneously exerting toxic effects on other organisms. Both processes are

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controlled by the chemicals’ bioavailability for the one or the other effect1. It is clear that

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the degradation of a chemical by the first group of organisms reduces the exposure for the

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second and it is generally agreed that metabolic utilization may be considered as a

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detoxification mechanism. However, these degrading organisms rely on the chemical and

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should strive to maintain an appropriate level of a chemical’s availability2. The

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quantitative relationship between the availability for biological removal and ecotoxicity is

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a key issue for any attempt to weigh extant risks against prospects of remediation3. In soil

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many organic contaminants, such as semi-volatile contaminants like polycyclic aromatic

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hydrocarbons (PAH) tend to sorb to the solid soil matrix4. Thereby the PAH may escape

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bacterial degradation and build up poorly bioavailable reservoirs5-7. Driven by

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disequilibria however, bound PAH may be released again to aqueous or gaseous soil

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phases where they may become available to soil organisms. Depending on the receiving

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organisms, this may cause toxic effects8,

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specialized bacteria able to metabolize PAH as a source of carbon and energy2. Any

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presence of PAH-degrading bacteria in the vicinity of PAH reservoirs might thus reduce

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further transport and eventual outgassing of PAH to the soil air. Moreover, PAH

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degradation, through its effect on the sorption equilibrium, would drive the desorption

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flux thus accelerating the attenuation of PAH. Efficient degradation thereby ideally

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requires a homogeneous distribution of degrading bacteria.

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The distribution of specialized bacteria and their degradation potential in the soil however

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shows a distinct spatial variability10, 11, leading to regions comprising several millimeters

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up to a few centimeters, which are devoid of degrading activity12, 13. In case of limited

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bacterial mobility patchy distribution of catabolically active microbial biomass may occur

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and/or result in biodegradation as e.g. by


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and the effectiveness of such ‘bacterial filters’ may be limited14-16. However, the effects

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of these small scale spatial heterogeneities on degradation efficiency have hardly been

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evaluated so far. Starting out from the earlier discovery that fungal mycelia facilitate

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bacterial dispersal by providing continuous paths for bacterial motility even through

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water-unsaturated soil15-18, we here hypothesize that mycelial dispersal networks may

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help distributing PAH-degrading bacteria in a way that area-covering PAH degradation

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occurs and transport of PAH including outgassing is minimized. A dense dispersal

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network thus would facilitate the formation of homogenously distributed degrader

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biomass, thus preventing contaminant outgassing from a contaminant hot spot.

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In this study, the effects of bacterial distribution in presence and absence of glass fibers

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(suitably mimicking dispersal along a fungal mycelium15,

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constantly desorbing phenanthrene (PHE) were investigated. In particular we studied (i)

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bacterial dispersal on the model surface in presence and absence of dispersal networks,

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(ii) the effects of the bacterial distribution on PHE degradation, and (iii) the role of

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bacterial distribution on PHE gas phase emission.

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) on biodegradation of

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MATERIAL AND METHODS

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Organisms and culture conditions. PHE degrading soil-borne Pseudomonas fluorescens

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LP6a20, 21 was grown at room temperature on a rotary shaker at 150 rpm in Erlenmeyer

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flasks containing 200 mL of minimal medium (MM) supplemented with 1.5 mg L-1 PHE

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crystals as the sole carbon and energy source. For microcosm experiments, 72 h old

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cultures were harvested in the exponential growth phase and washed twice in 50 mM

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potassium phosphate buffer (PB) at pH 7.2 and centrifuged at 1,000 x g for 10 min. The

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pellet was re-suspended in MM to reach an OD578 of 8 (≈ 1010 cells mL-1). Cell numbers

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were counted with a MoFlo flow cytometer (DakoCytomation, U.S.) as described

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elsewhere22. For all measurements, the instrument was adjusted using fluorescent 1-µm ACS Paragon Plus Environment

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beads. Cell counting was performed with 488 nm light of 400 mW to analyze the forward

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light scatter (FSC) and the side light scatter (SSC). Measurements of cells and reference

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microbeads were performed in triplicates.

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Sorbent. Polydimethylsiloxane (PDMS) was used as sorbent for PHE. Its qualities as

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passive dosing system are described elsewhere23. PDMS O-rings obtained from Altec

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(Order no. ORS-0793-57. Cornwall, UK) with an outer diameter of 90.7 mm, an inner

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diameter of 79.3 mm, a mass of 9.12 g (C.V. 0.5 %, n = 10) and a volume of 6.81 mL

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were used for passive dosing in biodegradation experiments. For use in the microcosm

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system, PDMS rings were cut into 1 cm long pieces with a mass of 0.356 g (C.V. 1.5 %,

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n = 20). The pieces were cleaned as described elsewhere23. Methanol was used as the

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loading solvent. PDMS pieces (57 per batch) were submerged in 75 mL of saturated PHE

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solution in methanol which had been diluted with methanol to a final volume of 500 mL

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in a closed 1-L bottle and incubated at room temperature for at least 3 days to obtain

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partitioning equilibrium. The solution was decanted, surfaces of PDMS pieces were

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wiped using lint-free tissue, and residual methanol was removed by three sequential

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washes with 200 mL of Milli-Q water. The PHE loading therefore led to a final mean

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concentration of 596 µg PHE g-1PDMS in the PDMS. For an individual experiment with a

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higher PHE load (≈ 8000 µg PHE g-1PDMS) here, PDMS pieces were submerged in 500 mL

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of saturated PHE methanol solution and treated as described above. A lower PHE load in

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the PDMS (314 µg PHE g-1PDMS) for a coverage experiment (cf. below) were adjusted by

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using 25 mL of saturated PHE methanol solution which had been diluted with methanol

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to a final volume of 500 mL.

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Calculation of the partitioning of PHE. The distribution of PHE between PDMS and

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water, i.e. agar as aqueous phase, at equilibrium can be determined experimentally and

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calculated using the dimensionless partitioning ratio:


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: =

()

(1)

()

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Here cPDMS and cwater are the PHE concentration in PDMS and water, respectively and

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: = 8404 (± 315) is the dimensionless partitioning ratio and was obtained

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experimentally in this study (cf. the SI). PHE vapor-phase concentrations were not

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directly measurable and thus calculated assuming equilibrium between the gas phase and

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the agar phase using a dimensionless partitioning ratio of ": = cair / cwater =

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0.001424. Hence, the percentage fraction of the total amount of PHE in the closed abiotic

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system, #$% present at the equilibrium in the agar can be calculated by:

#$% =

1 ( ( 1 + ": ∙ ( " + : ∙ (

(" (= 120 *+)

,

∙ 100 (2)

( (= 2 *+, "- . / + 1 *+, 012 )

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Here,

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( (= 0.76 *+), are the respective volumes. PHE concentrations in individual phases

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were calculated assuming equilibrium between the three-phases in the abiotic system and

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by using the total PHE load in the microcosm and used for the comparison between the

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experimental data in the abiotic system and the theoretically calculated value.

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Degradation and dispersal experiments. Degradation and dispersal experiments were

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conducted in microcosms with or without glass fibers as dispersal networks (Fig. 1). The

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PHE loaded PDMS allows a continuous and controlled release of the only carbon sources

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into the agar of the microcosm system as model soil atmosphere interface. The basic body

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of a microcosm consists of a glass vat (length: 3.5 cm, height: 1.0 cm, width: 1.0 cm)

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containing PHE-loaded PDMS, agar as the direct environment of the bacteria and a

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dispersal network (glass from P-D Glasseiden GmbH, Germany). Three PHE-loaded

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PDMS pieces (596 µg PHE/g PDMS) were placed at the bottom of the vat and covered with 2

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mL of minimal medium agar (MMA, 0.6% agar (w/v)). Glass fibers (approx. 300 pcs,

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length: 3.2 cm with a diameter of ≈ 8 µm) were cleaned by heating to 600 °C for 5 h.


,

and

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They were placed in parallel along the transect A-D at a distance of ≈ 0.25 cm from the

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left border of the vat in zone A and covered the whole width of the vat (Fig. 1). The

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microcosms were incubated for 18 h to allow the PHE partition between PDMS and agar.

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Then, the microcosms were inoculated with 2 µL of bacterial suspension at 0.1 cm

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distance from the glass fibers on the left-hand third of zone A (zone A, Fig. 1) to avoid

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capillary flow along them and incubated for 24 h, 96 h, 168 h (highest PHE load, cf.

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section Sorbent above), 0 h, 72 h, 120 h (medium PHE load) or 72 h (lowest PHE load),

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respectively. An agar cube (1 cm3 MMA, 2 % agar w/v) was placed at a distance of 0.3

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cm to the vat thus leaving the glass vat and the agar cube separated by an air gap (Fig. 1).

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The agar cube was used as a recipient of outgassing PHE. All microcosm setups were

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incubated in sealed standard glass Petri dishes at 25 °C and stored in a 30 L desiccator to

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minimize gaseous losses of PHE. Identical setups without glass fibers, either with or

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without bacteria were used in two independent control experiments.

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The agar cube was removed 2, 27, and 120 h after inoculation and analyzed for its PHE

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concentration by gas chromatography coupled with mass spectrometry (GC-MS, cf.

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below). As illustrated in Fig. 1, the glass vat was divided in four zones of interest (A, B,

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C and D). The surfaces of the zones were harvested individually with a spatula starting

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from zone D to a depth of about 1 mm. Due to the agar concentration used the agar allows

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for efficient bacterial dispersal on its surface only, yet restricts bacterial motility within

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the agar. The removed agar layer was then suspended in 3 mL PBS, ultravortexed for 1

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min, sonicated (with 35 kHz for 2 x 30 s with a break of 1 min)15, fixed with 2 mL 10%

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(v/v) sodium azide (NaN3) and finally analyzed for total cell number by flow cytometry

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(flow cytometry, cf. above). When glass fibers were present, they were cut at zone

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boundaries with a sharp scissor prior to sampling. The surface area of zone A was

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0.5 cm2, surface areas of zones B, C and D were 1 cm2, each. Thereafter, the remaining


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agar was removed and weighed. The PDMS pieces were removed before the PHE was

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extracted and analyzed.

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Bacterial surface coverage experiments. To estimate effects of bacterial surface

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coverage on PHE degradation and outgassing from microcosms, various initial cell

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concentrations

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(314 µg PHE/g PDMS). Suspensions of P. fluorescens LP6a calculated to result in surface

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coverages of 9%, 37% or 64%, assuming a cross-sectional area of 1.5 µm2 per cell, were

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adjusted by flow cytometry in PBS buffer and inoculated in three different regimes

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(‘heterogeneous’ or ‘homogeneous’ distribution without glass fibers or placement on

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glass fibers) on the agar surface of the microcosm system (see scheme in SI, Fig. S1).

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Microcosms were prepared and incubated for 72 h as described above but filled with 2

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mL MMA with 2 % agar to restrict bacterial movement. Heterogeneous distribution was

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achieved by spot inoculation of 3.33 µl cell suspension at each of three positions

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separated by 1 cm with the first spot placed at 0.75 cm from the left border of the glass

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vat. For homogeneous distribution, a 10 µL cell suspension was inoculated on the agar

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surface and dispersed carefully with a glass stick. For the third regime, glass fibers

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(approx. 300 pcs, length: 3.5 cm) were placed on the entire agar surface and spot

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inoculated with 10 µL cell suspension in the center of the agar surface. All setups were

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installed in triplicates along with controls without bacteria. Microcosms were harvested

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and analyzed as described above, but for the quantification of cell numbers, the whole

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agar surface was removed with a sterile spatula to a depth of about 1 mm.

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Extraction and analysis of PHE. Triplicate microcosms were harvested and their PHE

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concentration analyzed after a certain time of incubation (cf. section Degradation and

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dispersal experiments above). Briefly, agar samples were mixed during 10 min with

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approximately equal amounts of activated (5 h at 600 °C) Na2SO4 and then extracted with

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either 20 mL (agar cube) or 25 mL (agar from the vat) of a 1:3 mixture of acetone and n-

were

distributed

on

agar

containing


PHE

loaded

PDMS

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hexane containing per-deuterated PHE (PHE-d10, 10 mg mL-1 99.5 %, Dr. Ehrenstorfer,

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Germany) and acenaphthylene (ACN-d10, 10 mg mL-1 99.5 %, Dr. Ehrenstorfer,

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Germany) as extraction and injection standard, respectively25. PHE (98 % HPLC),

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methanol (99.8 %), acetone (99.8 %) and n-hexane (97.5 %) were obtained from Fluka,

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Merck, or Roth (all Germany). PHE in PDMS was extracted overnight with 8 mL of the

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solvent mixture per piece. PHE was quantified in an HP 7890 Series GC as described

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elsewhere26.

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Statistical analysis. Results of the replicate series for systems with or without transport

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networks were compared to each other and to abiotic systems, using a two-sided

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Student’s t-test to test for significant differences (p-value = 0.05).

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RESULTS

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Distribution and abundance of bacteria in the presence of a dispersal networks.

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Dispersal and growth of P. fluorescens LP6a in the presence and absence of glass fibers

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were monitored as illustrated by the mean bacterial coverage in the four zones of the

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microcosms (Fig. 2). In the absence of glass fibers (Fig. 2 left), the coverage in the

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inoculation zone A rose from initially 180 % to 870 % (i.e. formed presumed multiple

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layers of bacteria within the top ≈ 1 mm scraping depth) within 120 h, while in all other

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zones coverage increased very slowly only and remained below 33 % until the end of the

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experiment. With glass fibers (Fig. 2 right) the calculated average coverage in zone A

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reached 620 % within 120 h. In all other zones, coverages remained below 100 % but

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were at least twofold higher than in the absence of glass fibers. These differences were

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clearly significant in zones B and C after 120 h, where dense coverage (55 – 68 %) was

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found in zones B to D when glass fibers were present.

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Influence of bacterial distribution on PHE degradation. Significantly higher PHE

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removal from the PDMS pieces was detected in the presence of glass fibers (45 %) than ACS Paragon Plus Environment

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with the network-free setups (15 %; Fig. 3). Likewise 36 % (72 h) and 34 % (120 h)

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lower agar-dissolved PHE concentrations were measured in the agar of the vat in

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presence of the glass fiber networks compared to the setups without glass fibers

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(percentages relate to the theoretically calculated PHE concentration for the abiotic

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system; SI, Fig. S2).

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Influence of bacterial dispersal and abundance on PHE emission. Outgassing PHE

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was passively captured by an agar cube placed in the microcosms and analyzed over time

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to assess the PHE emission from the main body of the microcosm (Fig. 4). The partition

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equilibrium of PHE was obtained after 72 h incubation in the abiotic control (Fig. 4A). In

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agreement with PHE degradation data, the presence of LP6a bacteria reduced PHE

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outgassing by 77 % (72 h) and 88 % (120 h) relative to the calculated value. Although the

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microbial biomass in the presence of glass fibers was similar to the network-free scenario,

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no PHE accumulation in the agar cube, and hence, very limited PHE outgassing was

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detected when bacteria dispersed along glass fibers (Fig. 4A). Efficient bacterial dispersal

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along the glass fibers likewise resulted in reduced PHE emission from a PDMS

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containing a 13-fold higher PHE load (Fig. 4B). Here, ca. five- (96 h) and eight-fold

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(168 h) decreased PHE amounts were captured in the agar cubes when bacteria were able

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to disperse along glass fibers compared to the setup without glass fibers.

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Influence of cell coverage and distribution on PHE degradation. In order to further

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assess the effects of bacterial biomass and distribution on PHE degradation, in the

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bacterial surface coverage experiments nine regimes differing in their biomass

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distribution (‘heterogeneous’, ‘homogeneous’, ‘glass fibers’) and initial biomass (‘mean

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coverage’; cf. Materials and Methods) were analyzed for their effects on PHE outgassing,

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PHE degradation, and the abundance of strain LP6a after 72 h (SI, Fig. S1).

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Heterogeneous inoculation reduced PHE outgassing marginally irrespective of the

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amount of inoculated biomass (Fig. 5 top). In contrast, when the bacteria were ACS Paragon Plus Environment

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homogeneously applied to the agar surface or when glass fibers allowed their dispersal

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the outgassing of PHE was significantly decreased. Reduced outgassing went along with

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similarly efficient PHE degradation for the variants with 37% and 64% initial mean

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coverage respectively, likewise irrespective of the distribution (Fig. 5 bottom). Also here,

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no significant differences between the homogeneous distribution and the setup with glass

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fibers were observed. Interestingly, the initial biomass concentration only had marginal

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effects on the extent of PHE degradation: After 72 h of incubation, the total cell

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concentrations on the agar surface were quasi similar in all regimes independent of their

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initial abundance and distribution (SI, Fig. S3).

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DISCUSSION

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Influence of dispersal on biomass distribution and PHE biodegradation. Our study

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demonstrates that facilitated dispersal and optimized spatial arrangement of microbial

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populations enhances the degradation of constantly desorbing PHE. Efficient

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biodegradation of PHE simultaneously reduced the emission of this semi-volatile

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chemical already at surprisingly low rates of bacterial surface coverage (Fig. 4). Control

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experiments, in which biomass was ab initio inoculated at different degrees of

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homogeneity indicated that the effect of dispersal networks arises indeed from its

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influence on the cell distribution, which controls the substrate availability to individual

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cells and, consequently, their degradation rates. This can be seen from distinct

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degradation performances of differently distributed bacterial inocula despite identical

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substrate provision (Fig. 5 bottom). An important observation is that distribution of

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biomass exerted substantially different pressures on the partition equilibrium of the

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chemical; whereby a more homogeneously distributed biomass was driving PHE

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desorption by reducing PHE concentrations in the vat’s aqueous phase

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based system proved well-suited for mimicking prolonged PHE release as it is observed ACS Paragon Plus Environment

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. The PDMS-

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in PAH contaminated soil. Fungal mycelia known to facilitate bacterial dispersal15, 28, 29

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were substituted by arrangements of glass fibers for better control and longevity of the

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network as well as to prevent possible metabolic and physiological interactions between

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fungi and bacteria18. Despite the model character of this setup, we believe that the

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combination of a slow substrate release system and a receiving gas phase, separated by a

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functional system for bacterial dispersal in which bacteria upon inoculation can disperse

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and dynamically develop a biological sink for the released substrate, mimics several

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relevant features of a contaminated top soil. Our data experimentally support previous

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modelling data16,

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biodegradation rate of chemicals. Although the overall flux of PHE from the source is

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increased, aqueous concentrations of PHE available for both, degradation (SI, Fig. S2)

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and/or possible toxic exposure to non-degrader organisms are reduced. This effect is

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directly seen as reduced outgassing from the system (Fig. 4) which subsequently leads to

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a lower availability of the chemical to organisms beyond the contaminated zone. The

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higher biomass concentration behind the point of inoculation developing in microcosms

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with glass fibers indicates the positive effect of dispersal networks on the degradation

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capacity of the system in spite of the same initial substrate content. The positive effect of

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dispersal became evident despite of similar biomass production in presence of glass fibers

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in the bacterial surface coverage experiments (SI, Fig. S3), where reduced PHE loads in

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the PDMS (cf. lines 120ff) may have limited PHE bioavailability and have led to variable

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PHE allocation for bacterial growth and dispersal. Differing bacterial cell yields due to

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variable resource allocation (i.e. the division of resource uptake into fractions allocated to

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different energy-demanding processes) has been described for high metabolic costs for

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motility or flagellar synthesis 32, 33-

30, 31

, showing that network-based bacterial dispersal improves the

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Implications for the management of soil contamination. Many regulators have started

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to consider bioavailability within their retrospective risk assessment and remediation

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frameworks of organic chemicals. Such decision-making may rather depend on

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appropriate description of hazard based on the bioavailability rather than on the total-

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extractable concentration of a contaminant. Hence improved information on the emergent

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interactions between microbial dispersal, biodegradation and exposure of contaminants is

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needed to ensure appropriate protection of the environment and public health. Our results

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suggest that the presence of natural bacterial dispersal networks (e.g. provided by fungal

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mycelia or plant roots) may play an important role for the biodegradation of continuously

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released contaminants. The data furthermore show that the spatial arrangement of

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degrader biomass reflects the supply and transport characteristics of the degraded

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compounds34 provided that bacterial motility is sufficiently fast as e.g., due to dispersal

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networks. These effects of continuous structures or other biological vectors for the

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degradation capacity of bacteria in soil should also be kept in mind when designing

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bioremediation schemes3, 35, 36. Invasive management techniques such as ploughing may

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thus be detrimental for fungal helper functions37, 38. Enhanced dispersal and homogeneous

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distribution of degrader biomass may also be important for highly mobile (e.g. highly

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water soluble or quickly desorbing) contaminants, as such chemicals might easily escape

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patchy degrader populations19. Finally, our findings also may also apply for assessing the

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fate of biodegradable gases of environmental concern, such as e.g. methane evolving

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from soil and sediments39, 40.

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Acknowledgements. This work has been performed in the frame of the Helmholtz

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Association Research Programme ‘Chemicals in the Environment’ (CITE) and was

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supported by the research program Chemical Active Transport (CAT) and the Helmholtz

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Impulse and Networking Fund through the Helmholtz Interdisciplinary Graduate School ACS Paragon Plus Environment

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for Environmental Research (HIGRADE). We thank Kai-Uwe Goss for valuable

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discussions and Birgit Würz, Rita Remer and Jana Reichenbach for skilled technical help.

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We are grateful to Thomas Hübschmann for help on this work.

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Supporting Information Available

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This information is available free of charge via the Internet at http://pubs.acs.org


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(15) Kohlmeier, S.; Smits, T. H. M.; Ford, R. M.; Keel, C.; Harms, H.; Wick, L. Y. Taking the fungal highway: Mobilization of pollutant-degrading bacteria by fungi. Environ. Sci. Technol. 2005, 39 (12), 4640-4646; DOI 10.1021/Es047979zs. (16) Harms, H.; Wick, L. Y. Dispersing pollutant-degrading bacteria in contaminated soil without touching it. Eng. Life Sci. 2006, 6 (3), 252-260; DOI 10.1002/elsc.200620122s. (17) Wick, L. Y.; Remer, R.; Würz, B.; Reichenbach, J.; Braun, S.; Scharfer, F.; Harms, H. Effect of fungal hyphae on the access of bacteria to phenanthrene in soil. Environ. Sci. Technol. 2007, 41 (2), 500-505; DOI 10.1021/Es061407ss. (18) Pion, M.; Spangenberg, J. E.; Simon, A.; Bindschedler, S.; Flury, C.; Chatelain, A.; Bshary, R.; Job, D.; Junier, P. Bacterial farming by the fungus Morchella crassipes. Proc Biol Sci. 2013, 280 (1773), 2242; DOI 10.1098/rspb.2013.2242s. (19) Worrich, A.; König, S.; Miltner, A.; Banitz, T.; Centler, F.; Frank, K.; Thullner, M.; Harms, H.; Kästner, M.; Wick, L. Y. Mycelia-like Networks Increase Bacterial Dispersal, Growth and Biodegradation in a Model Ecosystem at Varying Water Potentials. Appl. Environ. Microbiol. 2016, DOI 10.1128/AEM.03901-15 s. (20) Foght, J.; Semple, K.; Westlake, D. W. S.; Blenkinsopp, S.; Sergy, G.; Wang, Z.; Fingas, M. Development of a standard bacterial consortium for laboratory efficacy testing of commercial freshwater oil spill bioremediation agents. J. Ind. Microbiol. Biotechnol. 1998, 21 (6), 322-330. (21) Bugg, T.; Foght, J. M.; Pickard, M. A.; Gray, M. R. Uptake and active efflux of polycyclic aromatic hydrocarbons by Pseudomonas fluorescens LP6a. Appl. Environ. Microbiol. 2000, 66 (12), 5387-5392. (22) Hübschmann, T.; Vogt, C.; Till, S.; Rohwerder, T.; Sand, W.; Harms, H.; Muller, S. Detection of sulfur microparticles in bacterial cultures by flow cytometry. Eng. Life Sci. 2007, 7 (4), 403-407; DOI 10.1002/elsc.200720195s. (23) Smith, K. E.; Rein, A.; Trapp, S.; Mayer, P.; Karlson, U. G. Dynamic passive dosing for studying the biotransformation of hydrophobic organic chemicals: microbial degradation as an example. Environ. Sci. Technol. 2012, 46 (9), 4852-4860; DOI 10.1021/es204050us. (24) Schwarzenbach, R. P., Gschwend, P. M., Imboden, D. M. Environmental Organic Chemistry; New Jersey, Canada, 2003. (25) Schamfuss, S.; Neu, T. R.; van der Meer, J. R.; Tecon, R.; Harms, H.; Wick, L. Y. Impact of Mycelia on the Accessibility of Fluorene to PAH-Degrading Bacteria. Environ. Sci. Technol. 2013, 47 (13), 6908-6915; DOI 10.1021/Es304378ds. (26) Gros, J.; Nabi, D.; Würz, B.; Wick, L. Y.; Brussaard, C. P. D.; Huisman, J.; van der Meer, J. R.; Reddy, C. M.; Arey, J. S. First Day of an Oil Spill on the Open Sea: Early Mass Transfers of Hydrocarbons to Air and Water. Environ. Sci. Technol. 2014, 48 (16), 9400-9411; DOI 10.1021/es502437es. (27) Dechesne, A.; Badawi, N.; Aamand, J.; Smets, B. F. Fine scale spatial variability of microbial pesticide degradation in soil: scales, controlling factors, and implications. Frontiers in Microbiology. 2014, 5 (667), DOI 10.3389/fmicb.2014.00667s. (28) Harms, H.; Schlosser, D.; Wick, L. Y. Untapped potential: exploiting fungi in bioremediation of hazardous chemicals. Nat. Rev. Microbiol. 2011, 9 (3), 177-192; DOI 10.1038/Nrmicro2519s. (29) Furuno, S.; Pazolt, K.; Rabe, C.; Neu, T. R.; Harms, H.; Wick, L. Y. Fungal mycelia allow chemotactic dispersal of polycyclic aromatic hydrocarbon-degrading bacteria in water-unsaturated systems. Environ Microbiol. 2010, 12 (6), 1391-1398; DOI 10.1111/j.1462-2920.2009.02022.xs.


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FIGURE LEGENDS

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Figure 1: Experimental setup used to assess effects of bacterial dispersal on PHE

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degradation and outgassing. A-D denote surface zones that were separately analyzed for

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bacterial biomass distribution. White lines represent the glass fibers used as model

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dispersal networks and red cylinders represent PDMS pieces used as passive dosing

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systems for continuous PHE release. An adjacent agar cube served as passive sampler for

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PHE outgassing from the vat.

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Figure 2: Calculated coverage of the agar surface of microcosms by P. fluorescens LP6a

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bacteria in % (A, B, C, D) in the absence (left) and presence (right) of glass fiber

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dispersal networks. Hundred % indicates nominal full coverage with a monolayer of

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bacteria. Values are expressed by shades of grey. Data represent averages of triplicate

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experiments. The mean measured cell concentration at a zone of interest in presence of

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glass fibers showed a statistically significant difference from the system without glass

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fibers (*) at a p-value of 0.05.

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Figure 3: PHE concentration in the PDMS in the absence and presence of glass fibers.

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Data represent averages and standard deviations of triplicate experiments for the setup

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with and without glass fibers. The abiotic control was performed in two independent

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triplicate experiments. The mean measured PHE concentration in the PDMS in presence

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of glass fibers shows a statistically significant difference from the system without glass

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fibers (a) at a p-value of 0.05.

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Figure 4: PHE concentration in the agar cube next to the glass vat from microcosms with

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PDMS containing a low PHE load (Fig 4A; system incubated for 120 h) or a high PHE

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load (Fig 4B; system incubated for 168 h). Bars represent the abiotic control (dark grey),

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the setup without glass fibers (grey) and the setup with glass fibers (grey with hatching).

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Solid line depicts calculated PHE concentrations at equilibrium in the abiotic system,


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with the dashed lines showing the 95 % confidence intervals. Data represent averages and

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standard deviations of triplicate experiments. The mean measured concentration of PHE

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in the agar cube shows statistically significant difference from the abiotic control (a) and

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a statistically significant difference from the system without glass fibers (b) at a p-value

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of 0.05.

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Figure 5: PHE concentration in the agar cube next to the vat (top) and in the PDMS

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(bottom) with the ‘heterogeneous’, ‘homogeneous’ and ‘glass fibers’ distribution regimes

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of P. fluorescens LP6a biomass after 72 h of incubation, respectively. The solid line

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depicts the mean PHE concentration of the abiotic microcosms. The dashed line depicts

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calculated PHE concentrations at equilibrium in the agar cube in the abiotic system (cf.

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equation 2; calculation of the partitioning of PHE). The mean measured concentration of

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PHE in the agar cube under different distribution regimes shows a statistically significant

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difference from the abiotic control (a) and a statistically significant difference from the

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system with a heterogeneous bacterial distribution (b) at a p-value of 0.05, respectively.


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FIGURES

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Figure 1


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Figure 2


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Figure 3


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Figure 4


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Figure 5

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Effects of Facilitated Bacterial Dispersal on the Degradation and

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