Introduction

Rhamnolipids are

surface-active molecules produced by Pseudomonas

aeruginosa and are essential

biotechnological products with a wide range of applications in many

areas, e.g. cosmetics (emulsifiers), food industry (food formulation

ingredients) [25], biomedicine (due to their antiadhesive and

antimicrobial properties) [31], agriculture (due to their antimicrobial and

antifungal effects) and bioremediation (removal of toxic heavy metals from soils) [14, 24,

29].

Soil washing

Soil washing with

rhamnolipids is one of the most often proposed potential strategies for

reducing heavy metal toxicity to indigenous soil bacteria (for review see

[24]), and it is postulated that the removal of metal toxicity includes complexation

of heavy metals by rhamnolipids [26].

Bioremediation

Although for remediation

of sites polluted with heavy metals, both the potential of rhamnolipids to

remove heavy metals from soils as well as the possible toxic effects of

rhamnolipids to soil microorganisms should be taken into account. Existing data

about the interactions between rhamnolipids−metal complexes and soil

(micro)organisms are relatively rare and controversial. Stacey et al. [35]

postulated that rhamnolipids from neutral lipophilic complexes with cationic

metal ions and enhance the absorption of zinc by plant roots. Conversely,

Al-Tahhan et al. Rhamnolipids reduced the cell surface charge of Gram-negative

bacteria resulting in increased hydrophobicity and, thus, reduced cadmium

uptake.

Similarly, several

studies have shown a reduction in heavy metal toxicity to bacteria in the

presence of rhamnolipids [20, 33]. On the other hand, Shin et al. [34] showed

that the addition of 240 mg l-1 rhamnolipids for an in situ remediations

inhibited the phenanthrene degrading bacteria referring to the potential toxic

effect of biosurfactants. Indeed, rhamnolipids have been shown to exhibit

powerful antibacterial, antifungal and algicidal activities [8, 37]. Thus, to

be used for in situ bioremediation, preparations of rhamnolipids

should be thoroughly characterized not only concerning their remediation

efficiency but also for potential toxic effects pathogenicity.

Objectives of the study

The main aim of this

study was to characterize the rhamnolipids produced by Pseudomonas aeruginosa for their inherent toxicity to Gram-negative (Vibrio fischeri, Pseudomonas fluorescens, P.

aeruginosa, Escherichia coli) and

Gram-positive (Bacillus subtilis) bacteria and their potential to decrease Cd

bioavailability and remove Cd- toxicity in aqueous media and soils.

Materials and Methods

1. Materials

CdCl2 (>98%) was

obtained from Sigma, Tween 80 from Serva, components of growth media were

either from LabM or Sigma, L-rhamnose, orcinol and 1-N- phenylnaphthylamine

(1-NPN) were from Sigma-Aldrich. Rhamnolipids were purified from the culture

broth of Pseudomonas aeruginosa. A sandy soil (initial concentration of Cd 0.17 mg

kg-1) spiked with CdCl2 (1.5, 15, 150, 1500 or 15 000 mg of Cd kg-1) as

previously reported [11] was used for the bioavailability studies. Before

spiking, the soil was characterized in a certified laboratory and had the

following properties: 10.6% of clay, 10.6% of silt, 72.8% of sand, 5.7% of

organic matter; 39 g·kg-1 of CaCO3, 3.59 g·kg-1 of N, 0.62 g·kg-1 of P, 0.17 mg

kg-1 of Cd; with 2.3 cmol+kg-1 of CEC (cation exchange capacity) and pH of 7.3.

Deionized water was used throughout the study.

2.

Characterization of the rhamnolipids-producing bacterium Pseudomonas aeruginosa

In the current study, the 16S rRNA gene of P. aeruginosa DS10-129 was sequenced by DSMZ (German Collection of

Microorganisms and Cell Cultures; and the partial sequence of the 16S rRNA gene

was deposited in the EMBL nucleotide sequence database with accession number AM419153.

The series was compared with those available in EMBL and NCBI databases using

the program BLAST to find its closest homologues. The similarity matrix was

constructed by pairwise analysis of validated (most completed) sequences using

the corrections computed by the Kimura’s 2-parameter model [18]. A phylogenetic

tree was created with the partial 16S rRNA gene sequences using a multiple

sequence alignment software CLUSTALW [10] by the Neighbour-Joining method [32]

and illustrated with the TreeView program. The resulting hierarchical

clustering tree was “pruned” to save space, and the closest

relatives were retained.

3. Isolation, purification and characterization of rhamnolipids

Rhamnolipids were

isolated from P. aeruginosa DS10-129 cell-free supernatant. P. aeruginosa was grown on the mineral medium containing (per l

water) 20 g of glycerol, 0.7 g of KH2PO4, 2 g of Na2HPO4, 0.4 g of

MgSO4•7H2O, 0.01 g of CaCl2 and

0.001 g of FeSO4•H2O and 1% of Yeast Extract. The presence of the rhamnolipids

was verified by thin-layer chromatography on silica gel 60 F254, according

to Matsufuji et al., [21]. Rhamnolipids were extracted from the cell-free supernatant

of the 4-day bacterial culture by centrifugation at 8000 g for 20 minutes at 4°C

and the subsequent filtration through a sterile filter (pore size 2 µm). The

cell-free culture was acidified to pH 2 with 2 M H2SO4, and the precipitated

rhamnolipids were extracted with an equal volume of 2:1

dichloromethane/methanol (liquid-liquid extraction) [38]. The organic phase was dried with

anhydrous Na2SO4 to remove excess water and evaporated on a rotary evaporator

(Buchi) at 60–70 °C to yield rhamnolipids. The rhamnolipids were dissolved in

0.05 M NaHCO3. The concentration of rhamnolipids was determined

using the orcinol assay [6] by mixing 100 µl of diluted solution of

rhamnolipids (purified with liquid-liquid extraction) with 900 µl of freshly prepared 0.19%

orcinol solution in 53% H2SO4. The mixture was heated at 80oC for 30 min and its absorbance

was measured spectrophotometrically at 421 nm. The concentration

of rhamnolipids was calculated according to L-rhamnose standard curve (0 to 50 mg

l-1) and by multiplying the result with a coefficient of 3.4 obtained from the

correlation of pure rhamnolipids/rhamnose [3]. The critical micelle concentration (CMC) was

determined by measuring the surface tension of serial dilutions of the

rhamnolipids [17]. Fourier Transform Infrared spectrophotometer (FTIR) Perkin Elmer 100

series was used to determine the molecular structure of the rhamnolipids. The

cell-free supernatant was acidified to pH 2 by adding drops of 2M Sulphuric acid to

precipitate the rhamnolipids. The precipitated rhamnolipids were extracted with an

equal volume of 2:1 dichloromethane/methanol. The organic phase was

dried with anhydrous Sodium Sulphate (Na2SO4) and evaporated on a rotary

evaporator (Buchi, Rota vapour R-200 Germany) set at 60-70°C. Approximately 2-5mg of the

concentrated rhamnolipids were analysed with the FTIR spectrophotometer.

4. Luminescent bacterial strains for toxicity and bioavailability assays

The luminescent bacterial strains used for toxicity and bioavailability studies are listed in Table 1. Constitutively luminescent bacterial strains, both natural (Vibrio

fischeri) and recombinant strains

were used to evaluate the toxicity of the rhamnolipids using bioluminescence

inhibition as a toxicity endpoint. Recombinant luminescent Cd- inducible sensor

bacteria were used to study the modulatory effect of rhamnolipids on

availability of Cd to bacteria. All recombinant bioluminescent bacterial

strains, except Pseudomonas

aeruginosa DS10-129

(pDNcadRPcadAlux) were constructed previously (Table 1). P. aeruginosa DS10-129 (pDNcadRPcadAlux) was initially constructed

as Cd-inducible strain by electroporating a 14,525 bp plasmid pDNcadRPcadAlux

[13], which contains bioluminescence-encoding luxCDABE genes

under the control of Cd response elements: Cd-regulated promoter (promoter of cadA)

and a Cd-binding regulatory protein (CadR), into P. aeruginosa DS10-129 competent cells [3]. Bacteria were plated

onto LB agar (10 g of tryptone, 5 g of yeast extract and 5 g of NaCl per 1 l of

deionised / distilled? water) containing 50 mg l-1 of tetracycline and the

plasmid-containing colonies were selected by luminescence. During further

experiments P. aeruginosa DS10-129 (pDNcadRPcadAlux) failed to be induced with

Cd (maximum induction below the limit of detection), mostly due to its high

background luminescence. However, the high bioluminescence level favoured the

use of this strain as a model organism for general toxicity testing. 17.

5. Analysis of toxicity and Cd bioavailability

Luminescent bacterial strains were either rehydrated from lyophilised culture (Vibrio fischeri) obtained from Aboatox, Turku, Finland or

cultivated. V. fischeri was reconstituted in heavy metal MOPS medium (HMM

medium) supplemented with 2% NaCl at room temperature for 1 h. The HMM medium

contained (per l of deionised / distilled? water): 8.4 g of MOPS buffer, 0.4 g of

glucose, 0.22 g of glycerol-2- phosphate, 3.7 g of KCl, 0.54 g of NH4Cl, 0.06g of

MgSO4, 0.162 mg of FeCl3. All recombinant bacteria were cultivated freshly by

growing the cultures overnight in 3 ml of LB medium [44] supplemented

with appropriate antibiotics as in Table 1. The overnight culture was diluted

1:50 with 10–50 ml of HMM medium, grown until OD600 of 0.3 and then diluted to

OD600 of ~0.1 prior to test.

5.1. Toxicity testing

To measure toxicity CdCl2 or CdCl2-rhamnolipids mixtures were analyzed by measuring the inhibition of bioluminescence of constitutively luminescent bacterial strains (Table 1). In addition, Pseudomonas aeruginosa DS10- 129(pDNcadRPcadAlux) was used to assess the

inhibitory effect of rhamnolipids. The effect of rhamnolipids on viability of

bacteria was evaluated by plating the treated bacteria on solidified growth

medium (see below). Dilutions of CdCl2 (0.1 – 100 mg l- 1 as final

concentrations), rhamnolipids (5 – 200 mg l-1 as final concentrations) or Cd-

rhamnolipids mixture (the final concentration of rhamnolipids was 50 mg l-1 or

a concentration reducing the light output of bacteria by 20%, indicated below)

were prepared in HMM medium or HMM medium supplemented with 2% NaCl (in case of V. fischeri).

For toxicity assessment, luminescence inhibition assay was performed in 96-well

microplates essentially as in Mortimer et al. [23]. Briefly, 100 ml of the

diluted test compound(s) was pipetted into 96-well microplate and 100 ml of the

bacterial suspension was automatically dispensed into the wells. Bacteria were

incubated at 20°C (V. fischeri) or 30°C (recombinant luminescent bacteria) and the

luminescence was continuously recorded during the first 30 seconds of exposure

and once after 30 minutes of incubation using Fluoroskan Ascent FL plate

luminometer (ThermoLabsystems). Inhibition of bacterial bioluminescence by the

tested compounds/mixtures was calculated as percentage of the unaffected

control (HMM medium or HMM supplemented with 2% NaCl, respectively). 30-s

and 30- min EC50 and EC20 values (the concentration of chemical which reduces

the light output of bacteria by 50 or 20% after the respective exposure times)

were calculated by linear regression from dose-response curves of the studied

compounds. Measurements were performed in three independent assays. Viability

of bacteria was assessed after their exposure to 100 mg l-1 of rhamnolipids by

plating the bacteria onto LB agar plates containing appropriate antibiotics.

Plates were incubated for 24 h at 30°C after which colony forming units (CFU)

were counted.

5.2. Bioavailability

testing

Availability of Cd (with or without rhamnolipids) to Cd-sensor bacteria (Table 1) both in aqueous environment and in soil-water suspension was analysed as described by Bondarenko et al. [4]. CdCl2 dilutions at final concentrations of 0.01-10 mg l-1 were prepared by rotating soil:water (1:10) aqueous suspensions of Cd-spiked soils at room temperature for 24 h. Non-spiked soil was used as a control for soil assays and deionised / distilled? water served as a control for CdCl2 dilutions. Rhamnolipids were added to CdCl2 dilutions or Cd-spiked soil suspensions to the final concentrations of 10, 20 and 40 mg l-1 . Samples (100 µl) were added to 100 µl of the sensor bacterial culture in HMM medium in 96-well microplates and incubated at 30oC for 2 h. Luminescence was measured with Fluoroskan Ascent FL plate luminometer. Induction of luminescence of sensor bacteria by Cd was calculated as follows:

Induction = LS/LB, where

LS is luminescence in the sample (CdCl2, CdCl2-rhamnolipids mixture, soil-

water suspension or its mixture with rhamnolipids) and LB is the background

luminescence (bacteria in HMM medium added to water or unspiked soil). The

concentration of Cd in the sample causing induction of bioluminescence twice

above the background value was defined as minimal inducing concentration

(defined also as limit of determination (LOD) by Ivask et al. [13]).

Bioavailability analyses were performed in three independent assays. 243

6. Analysis of membrane permeability

The enhancement in permeability of Escherichia coli MC1061(pDNlux) cell membranes by rhamnolipids was

measured by the uptake of a hydrophobic probe 1- N-phenylnaphthylamine (1-NPN)

as described by Helander et al. [12]. As compared to hydrophilic environments,

the fluorescence of 1-NPN is significantly enhanced in hydrophobic environments

(e.g. membrane phospholipids), rendering it a suitable dye

to probe outer membrane

integrity of Gram-negative bacteria [9]. Briefly, 50 µl of 40 µM 1-NPN dye and

50 µl of the surfactants (rhamnolipids or non-ionic chemical surfactant Tween

80 serving as a positive control) in 5 mM HEPES buffer (pH 7.2) were pipetted

into black microplates. 5 mM HEPES buffer was used as a negative control. 100

µl of bacterial suspension in 5 mM HEPES buffer were automatically dispensed

into each well and the fluorescence was immediately measured (Fluoroskan Ascent

FL plate luminometer; excitation/emission filters 350/460 nm). The final

concentrations of both surfactants in the test were 10, 40 and 100 mg l-1. The

1-NPN cell uptake factor was calculated as a ratio of fluorescence values of

the bacterial suspension in the presence and absence of surfactants.