Citation: Agni Hadjilouka, Konstantinos Nikolidakis, Spiros Paramithiotis, Eleftherios H. Drosinos. Effect of co-culture with enterocinogenic E. faecium on L. monocytogenes key virulence gene expression[J]. AIMS Microbiology, 2016, 2(3): 359-371. doi: 10.3934/microbiol.2016.3.359
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The activated sludge process for wastewater treatment is a biotechnology fundamental to human health and represents a model high population density ecosystem for studying microbial interactions [1]. Activated sludge is primarilycomposed of microorganisms in aggregates or flocculates (flocs), enabling low-techseparation of biomass from the bulk aqueous phase by settling under gravity. Flocs are composed of a dense microbial consortium in a matrix of extracellular polymeric substances [2] described by some as suspended biofilms or biofilms without surface association. It can be argued that human civilization is dependent on bacterial flocculate formation given the central role of this phenomenon in industrial and municipal wastewater treatmentglobally.
Despite the importance of activated sludge floc formationand function there remain significant knowledge gaps relating to how flocs form, functionand change over time. Firstly, it is unclear how floc development is initiated.Secondly, the relativecontributions of cell replication and cell recruitment to floc growth is not well characterised. Thirdly, there is no mechanistic microbiological description of the ultimatefate of a floc. Put another way, the “lifecycle” of activated sludge flocs is poorly described, possibly because heterogeneity between flocs is rife. A promising approach to addressingthese knowledge gaps is to circumvent the challenge of heterogeneity through development and interrogation of experimental models of floc formation.
Previous investigations raised the questionof whether activatedsludge flocs are in fact surface associated biofilms anchoredto particulate organic matter [3]. Activatedsludge is a well-known source of extracellular cellulase, chitinase and lipase enzymes, suggesting that particulate organic matter (cellulose, chitin and lipids) represent major substrate resources for activated sludge bacteria [4]. Cellulose is the most abundant biopolymer on Earth and the principal component of toilet paper [5]. Chitin is the second most abundant biopolymer on Earth and additionof chitin or its deacylated derivative chitosan is common practice in wastewater treatment plants to improve sludge settling properties [6]. Municipal wastewater contains lipids at concentrations between 40 and 100 µg/L representing over 30% of the chemicaloxygen demand [7,8]. Oleic acid is the most common fatty acid found in olive and sunfloweroils commonly used in domestic settings [9].
In this study we developed an experimental model of activatedsludge floc formation based on glycerol trioleate(GT) as a substrate. GT was addedto activated sludgeand observed to form small lipid spheres with which sludge biomass was tightly associated. Extracellular lipase activity was upregulated and a shift from Betaproteobacteria to Alphaproteobacteria was observed. Four bacteria associated with the GT spheres were isolated and identified and two of the isolateswere shown to produce acylated homoserinelactones (AHLs). Foundation of this experimental model enables future research into lipid based floc formationand ultimately the rationalconstruction of flocculates of desired function.
Activated sludgesamples were acquiredfrom St. Mary’s municipal wastewater treatment plant for establishment of culturesin the laboratory. The treatmentplant is located in westernSydney serving a population of app. 160000 in a catchment area of 84 km2 and discharges into the Hawkesbury-Nepean River. On a daily basis the plant processes 35 million litres of wastewater. The biological process is conductedin three stages (Anaerobic, Anoxic and Aerobic)and the sludge samplescollected for this studywere taken from the Aerobicstage.
Enrichment cultures consisted of 50 mL activated sludge in 250 mL erlenmeyerflasks incubated at 37 °C and 140 rpm. Quadruplicate cultures were treatedwith 1% glyceryl trioleate (Sigma-Aldrich T7140-10G). Samples were taken over a 25 day period, consisting of 0.5 mL for DNA extraction and 0.5 mL for lipase assays.Sterile filtered (0.22 µm Millipore) activated sludge supernatant was added back to incubations after sampling to maintain originalvolumes.
Lipid spheres were removed with tweezers and washed in phosphate buffered saline (PBS) before transferring to a glass slide. Spheres were stainedwith 5 µL SybrGreen and imaged with an Olympus Fluorescence Microscope.
DNA was extractedusing a xanthogenate-SDS based method [10]. Sludge samples were centrifuged at 16, 100 rcf for 5 min in a 1.5 mL eppendorftube before removing supernatant and pellet storage at −20 ˚C. For DNA extraction from GT spheres, semi-solid lipid droplets were removed with tweezersand washed in PBS, transferred to a 1.5 mL eppendorftube and stored at −20 ˚C. Thawed pellets were re-suspended in 0.2 mL PBS for extraction. ExtractedDNA was resuspended in molecularbiology grade water. DNA was quantifiedvia NanoDrop (NanoDrop ND-1000 Spectrophotometer) as well as Qubit (Invitrogen, Qubit 2.0 Flourometer), as per standard protocol for broad rangeDNA detection.
The PCR for denaturing gradient gel electrophoresis (DGGE) was set up with the primers 357FGC and 907R (10µM) [11] for 40 µL as follows: 20 µL EconoTaq Master mix (Lucigen), 12.48 µL molecularwater, 2 mM of each primer, 7.7mM bovine serum albumin and 5-50 ng template. The PCR protocol was the following: 2 min at 95 °C then 30 cycles of denaturingphase: 30 sec at 94°C, annealing phase: 30 sec at 54 °C, extensionphase: 1 min 30 sec at 72 °C, followed by a final extensionphase: 10 min at 72 °C and storage at 4 °C.
The DGGE was conducted using the BioRad DCode system and protocolfor 50% to 70% denaturing gradienton a 6.7% arcylamide gel. Electrophoresis was conducted at 75 V and 60 °C for 16.5 h in 1× TAE. The gel was stained with SybrGold for 20 min and imaged on a BioRad Gel Doc XR Imaging system and BioRad Image Labsoftware. Bands of interest were selected from the DGGE and excised and re-run on DGGE PCR to test their integrity before sequencing.
The sequencingPCR was set up for the 357F primer at 20 µL as follows: 1 µL BigDye terminator V3.1, 20‒50 ng PCR product, 0.32 µL primer, 3.5 µL 5× sequencing buffer and madeup to 20 µL with molecular water. The PCR protocol was the following: 26 cycles of denaturing phase: 10 sec at 96 °C, annealing phase: at 5 sec 50 °C, extension phase: 4 min at 60 °C, followed by and storage at 4 °C. The samples were cleaned up by addition of 5 µL of 125 mM EDTA and 60 µL 100% (v/v) ethanol followed by vortexing and 15 min precipitation period. The samples were then spun at 14000 rcf for 20 min and supernatant was removed. After additionof 160 µL fresh 70% ethanol, samples were spun at 14000 rcf for10 min and supernatant discarded. This step was repeated with addition of 80 µL 70% ethanol. Sanger Sequencing wasconducted by the Ramaciotti Centre for Gene Function and Analysis at the Universityof NSW, using standard ABI protocols.
The lipase assay was adapted fromChristensen et al [12]. Sludge samples (0.5 mL) were centrifuged at 16, 100 rcf for 5 min. The supernatant was removed and filter sterilised (0.22 µm Milliporefilter) to test for suspended extracellular lipase. The pellet was re-suspended via vortexing in 0.5 mL autoclavedMilliQ water with the addition of 0.1 mL Zirconium beads. The samples were bead beaten for three 45 sec cycles and centrifuged at 16.1 rcf for 5 min. The supernatant was removed and filter sterilised (0.22 µm Millipore filter) to test for EPS associatedlipase. The pellet was re-suspended via vortexing in 0.5 mL autoclaved MilliQ water to test for membrane bound lipase. Subsamples of each fraction (100 µL) were incubated with 0.9 mL of p-nitrophenyl palmitate (p-npp) containing substrate solution for one hour before reaction termination by alkaline pH inactivation of lipase with 1 mL 1 M sodium carbonate. Cleaved p-nitrophenyl groups were quantified by spectrophotometry at an absorbance of 410 nm.
Four bacterialisolates associated with GT sphereswere grown in Luria Bertanibroth overnight at 37 ˚C with shaking at 140 rpm in the presence or absence of 1% (v/v) GT. Ethyl acetate extracts (20 mL) of culturesupernatants were preparedand concentrated to 100 µL by evaporation. Extracts were assessed for the presenceof AHL like activity using an Escherichia colibased AHL monitor strain harbouring the plasmid pJBA357encoding the luxRgene and a gfp(ASV) reportergene fused to the AHL-LuxR dependentPhotobacterium fischeri luxIpromoter [13]. The fluorescent output was monitored by excitation at 485 nm and emission at 535 nm.
To establishan experimental model of floc formation based on interactions between lipids and bacteria in activated sludge, glyceryl trioleate (GT) was added to activated sludge samples at 1% v/v. On additionGT formed a transparent and hydrophobic layeron the surface of the sludge before white GT spheres formed suspended below the culturesurface within 7 days (Figure 1). The GT spheresvaried in size but decreasedin diameter untilthey were indistinguishable from the sludge after 21 days.
Figure 1. Spontaneous formationof glycerol trioleate(1% v/v) spheres upon addition to activated sludge. This image taken after 7 days aerobic incubation at 37 ˚C with shaking at 140 rpm. The lipid spheres decreased in size over time and were undetectable after 25 days.
To examine the spatial relationships betweensludge biomass and the GT spheres, samples were stained with a fluorescent DNA binding compound before imaging with epifluorescence microscopy. The fluorescent stain emits green light upon DNA binding and excitationwhile GT appears yellow and water appears black (Figure 2). Biomass associated preferentially with the GT spheresand was firmly attached as evidenced by the washing procedure applied prior to staining. This spatial distribution and cell density is reminiscent of surface associatedbiofilms and the lipid droplets with associated biomasscould be considered prototype or juvenile flocculates.
Figure 2. Epifluorescence microscopy images of sybr green stained DNA (biomass in green) associated with microscopic glycerol trioleate spheres (yellow) using 10× (A) and 40× (B) objective lenses. Filamentous, coccoid and rod shaped bacteria (red arrows) are visible as the activated sludge biomass adsorbs to the lipid droplets (blue arrows) forming ‘juvenile’ flocculates.
Lipase activitywas monitored in the sludge supernatant, in EPS and in membrane associated fractions after GT addition (Figure 3). Lipase activity was highest in membrane-associated fractions over the course of the incubationbut there was no apparent impactof GT addition. Lipase activity in EPS and supernatant fractions was more variable across replicates but there was a significant elevation in lipase activity in response to GT additionin these fractions (p < 0.05). These data suggest the presence of the GT spheres causesan increase in extracellular lipaseproduction.
Figure 3. Lipase activity in quadruplicate cultures (blue, green, yellow and orange) of glycerol trioleate treated activated sludge (left panels) and untreated activated sludge (right panels) over 25 days incubation. Sludge samples were separated into extracellular (top panels), extracellular polymeric substance (EPS) (middle panels) and membrane (bottom panels) fractions before quantifying lipase activity. Whilst significant variation is observed between biological replicates with respect to timing of emergence of lipase activity, clear evidence of upregulation of activity is observed in the extracellular fraction in response to glycerol trioleate addition.
Figure 4 shows community fingerprints of GT amended and unamended control cultures over 25 days incubation. A clear shift in bacterialcommunity composition is apparentin response to GT addition. Bands were excised, sequenced and matched with existing bacterialsequences in the NCBIdatabase (Table 1). Initiallyabundant Bacillus, Dechloromonasand Azospiralineages were replaced with Novosphingobium, Sphingomonasand Roseomonaslineages. This represented a general shift from Betaproteobacteria to Alphaproteobacteria in response to GT amendment.
Figure 4. Denaturing gradient gel electrophoresis based bacterial community analysis of untreated sludge (left panel) and glycerol trioleate treated sludge (right panel) over a 25 day incubation. A shift in bacterial community composition was observed in glycerol trioleate amended cultures. Numbered bands were sequenced (Table 1) revealing a shift from Betaproteobacteria to Alphaproteobacteria. Sphingomonas and Novosphingomonas species dominated in response to glycerol trioleate addition.
| Band Identity | Class | Closest relative (Acc. No.) |
| 1 | Bacilli | Bacillus sp. (GU271888.1) |
| 2 | Betaproteobacteria | Uncultured Dechloromonas sp. (JQ012310.1) |
| 3 | Betaproteobacteria | Azospira oryzae (KF260987) |
| 4 | Betaproteobacteria | Uncultured Dechloromonas sp. (KF003189.1) |
| 5 | Alphaproteobacteria | Novosphingobium sp. (KF544940.1) |
| 6 | Alphaproteobacteria | Novosphingobium sp. (KF544932.1) |
| 7 | Alphaproteobacteria | Sphingomonas sp. (AY521009.2) |
| 8 | Alphaproteobacteria | Sphingomonas suberifaciens (AY521009.2) |
| 9 | Alphaproteobacteria | Sphingomonas sp. (JQ928361.1) |
| 10 | Alphaproteobacteria | Roseomonas sp. (KF254767.1) |
DownLoad: CSVGT spheres were washed and plated on solid media supplemented with GT. Colonies were subcultured to isolate bacteria that use the lipid as a carbon and energy source. Four isolates were subsequently shown to utilise GT in liquid media as evidencedby growth in the presence of GT but not in its absence. SSU rRNA gene sequencing revealedthe four GT spherecolonising isolates as Betaproteobacteria Pandoraea (KF378759.1) and Achromobacter (KF378759.1) and Gammaproteobacteria Enterobacter(KF411353.1) and Pseudomonas(KC822768.1) species. Unfortunately, none of the alphaproteobacterial Novosphingobiumor Sphingomonaslineages observed in community fingerprints (Figure 4) were isolated.
The four GT sphere colonising isolateswere tested for the production of Acylated Homoserine Lactones (AHLs). AHL mediated gene expression has been linked to lipase activity in Proteobacteria [14] and therefore may play a role in lipid basedfloc formation. A LuxR based bioassay in which expression of gfp is upregulated in the presenceof AHLs was used for the detection of AHLs in culturesupernatants. The Achromobacterand Enterobacterspecies gave strong positive responsesto the LuxR based bioassay whilst the Pandoraeaand Pseudomonassp. gave a negativeresponse. Interestingly the positiveAHL bioassay responseselicited by Achromobacterand Enterobacter were enhanced in the presenceof GT and whilst the negative AHL bioassay result for Pandoraeawas unchanged by GT addition, this isolate generateda pink pigment in the presenceof GT but not in its absence.
Activated sludge flocculates underpin the global wastewater treatmentindustry but the design, production and deployment of flocs to date has involved a top-down engineering approach focused on functionrather than underlyingstructure. Here we have producedan experimental foundation for a bottom-up scientific approach to the rationale construction of wastewater treatment flocs. Specifically, five aims were addressedrelating to spatial aspects of lipid partitioning and colonisation, impactson bacterial community composition, isolation of lipid degrading bacteria and testing for AHLproduction.
The physical partitioning behaviourof lipids upon introduction to activated sludge is poorly described. As a starting point it was unclear whether the addition of GT to activated sludge would result in the formation of free phase lipid orwhether existing sludgeflocs would be coated in lipid with an absence of free phase or if multiple small lipid droplets or an emulsionwould form. This lack of information on the physical partitioning behaviour in activated sludge hampers development of an experimental model for lipid dependentfloc formation. To this end, GT was incubated in the presenceof activated sludge and direct observations were made.
During the enrichment on GT, multiple spheres formed that were suspended throughout the sludge. Objects that resemble GT spheres in oleic acid challenged anaerobic sludge have been reported previously [15]. It stands to reason that such lipid droplets representan attractive surface substratumfor colonisation by activated sludge bacteria enablingevasion of predationand ready access to a carbon and energy source. Epifluorescence microscopy was used to observe physical interactions (spatial relationships) between the activated sludge biomass and the GT spheres. The microscopy images obtained showcase the close association of biomass with the lipids.Whether the dense mass of cells observedis surrounding and growing on the lipidor whether it was embeddedwithin the lipid phase is unclearfrom the microscopy images generated.Whilst the aggregation around the droplets resemblesbiofilm formation, it is not clear if the physicochemical properties of cells and lipid droplets drive the association or if active energy dependentprocesses result in microbial colonisation of the lipiddroplets. Cells embeddedin the lipid, with no access to water, are unlikely to proliferate. Biofilm formationon lipid dropletswould ultimately generateaggregated biomass resembling a floc. The result was encouraging in that it suggestedlipid colonisation represents a realistic model system for floc formation and development.
Lipid dropletsize was observedto decrease over time until they were no longer visible. This is consistent with microbes colonising the droplets and degrading the surface substratum as a carbon and energy source. Lipase activity was also observedto increase in activated sludge samples amended with GT. An increasein lipase activitywas observed in the EPS fraction over the first three days of incubation but the greatestfold increase in lipaseactivity was observedin the culture supernatant after three days of incubationas the EPS based lipase activity decreased.It has previously been shown that lipases are weakly bound to the EPS of flocs throughhydrogen bonding [16]. The sequential appearance of lipase firstin the EPS and then in the supernatant fraction is likely due to release of lipases from the EPS.
While lipase activity was observedin the membrane fraction, there was little difference in activity when comparing GT amended treatmentsand unamended controls. The unamended controls displayed a high background level of lipase activity presumably owing to the presence of background levels of lipid in activated sludge. The decline in lipase activity observedin the EPS fraction in unamended controls is concordant with this, with the background lipid component being consumed.
It is not known which bacteriaare responsible for the degradation of lipids in activated sludge. This information is crucial in the development of an activatedsludge floc formationmodel based on lipid colonisation. The response of bacteria in activated sludge to the addition of GT was assessed using DGGE. It is evidentfrom the DGGE community analysisthat lipids have a profound impact on the microbialcommunity structure in activatedsludge. Whilst bacterial lineages enriched in the presence of GT can be implicated in lipid degradation in activatedsludge this does not represent unambiguous evidence of lipid biodegradation by these bacteria.
Based on band intensitythe most abundant bacterial lineages in the untreatedsludge samples belonged to the genera Bacillus, Dechloromonasand Azospira. Bacillusspecies have been observed in sludge previously and the sequenceretrieved here was associated with lipolytic activity in olive mill wastewater [17]. Bacillusspecies have also been associated with bioflocculation in starchwastewater treatment [18].
Dechloromonas species have been observed in membrane fouling biofilms of municipal wastewater treatment plants in Japan, with over 30% of clones belonging to this genus [19]. Azospiraare also common activatedsludge occupants [20]. Overall, the DGGE profilesof the unamended sludge community were typical of activatedsludge and did not differ greatly over the incubation period.
In sludgesamples exposed to GT the DGGE profilesover time shifteddramatically. The most abundant bacteria observed in the unamended sludge were replaced within nine days of incubation with Sphingomonas, Novosphingobiumand Roseomonasspecies. Sphingomonasspecies can be present at 5‒10% relative abundancein sludge as shown by FISH [21] and play an important role in wastewater treatment. Members of this genus degrade testosterone and sterol hormonesas well as the pollutant nonylphenol [9]. Novosphingobiumspecies have been shown to play an important role in wastewater remediation by degrading toxic dyes and estrogen [22,23]. Roseomonas species have been found at 5% relative abundance in activated sludge and are known to degrade organophosphate pesticides [24]. None of these lineageshave previously been associated with lipid consumption in wastewater treatment plants.From the data generated here bacteria belonging to the Sphingomonas, Novosphingobiumand Roseomonasgenera can be considered candidates for inclusion in an experimental systemfor investigating lipid basedfloc formation.
Four activatedsludge bacterial strains isolated directly from lipid droplets and shown to use GT as a sole carbon and energy source were identified and screened for AHL quorum sensing activity. All four isolates are Proteobacteria but only two (AchromobacterandEnterobacter) displayed AHL like activity in the LuxR basedbioassay used. This was surprising given that the two isolates that gave negative responses (Pandoraea and Pseudomonas) have close relatives known to produce AHLs and that lipase activity has only been definitively linked with AHL mediated gene transcription in Pseudomonasspecies. A soil isolate belonging to the Pandoraea genushas been shown to secrete octanoylhomoserine lactone [25]. It is possible that a more comprehensive screen for AHLs using AHL bioassays with a varietyof LuxR type receptors or a direct mass spectrometry based methodwould reveal the production of AHLs outsidethe response rangeof the LuxR protein.The role of AHLs or quorum sensingin the colonisation or biodegradation of GT spheres in activated sludgeremains to be explored.
Bacterial floc formation is essential for the remediation of domestic wastewater. This study sought to supplement the limited information available about floc formationand the impact of lipid addition on activated sludge structure as well as microbial communitycomposition. GT addition to sludge resulted in the formationof lipid spheresthat were rapidly colonisedand consumed over 25 days, which may reflect seedingand maturation phases of a flocculate “life-cycle”. During this process extracellular lipase activity increased and Sphingomonasand Novosphingomonasspecies increased in relative abundance. Lipid degrading bacteria isolated from the lipid spheres produced quorum sensing signal activity.Deciphering the nature of colonisation in a biofilm-like manner around lipid spheres extends our understanding of the mechanism of flocformation.
To interrogate this newly developed experimentalsystem for floc formation futureresearch should focus on the isolationof lipid degrading Sphingomonasand Novosphingomonas species from activated sludge, on generating pure culture flocculates based on the colonisation of lipid spheres and on describing AHL mediated gene expression systemsin lipid degrading isolates to assess whether quorum sensing regulateslipase activity in activated sludge.
Mike Manefield was supported by an Australian Research Council Future Fellowship (FT100100078).
All authorsdeclare no conflict of interest in this paper.
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Figures(3) / Tables(2)
Agni Hadjilouka, Konstantinos Nikolidakis, Spiros Paramithiotis, Eleftherios H. Drosinos. Effect of co-culture with enterocinogenic E. faecium on L. monocytogenes key virulence gene expression[J]. AIMS Microbiology, 2016, 2(3): 359-371. doi: 10.3934/microbiol.2016.3.359
| Band Identity | Class | Closest relative (Acc. No.) |
| 1 | Bacilli | Bacillus sp. (GU271888.1) |
| 2 | Betaproteobacteria | Uncultured Dechloromonas sp. (JQ012310.1) |
| 3 | Betaproteobacteria | Azospira oryzae (KF260987) |
| 4 | Betaproteobacteria | Uncultured Dechloromonas sp. (KF003189.1) |
| 5 | Alphaproteobacteria | Novosphingobium sp. (KF544940.1) |
| 6 | Alphaproteobacteria | Novosphingobium sp. (KF544932.1) |
| 7 | Alphaproteobacteria | Sphingomonas sp. (AY521009.2) |
| 8 | Alphaproteobacteria | Sphingomonas suberifaciens (AY521009.2) |
| 9 | Alphaproteobacteria | Sphingomonas sp. (JQ928361.1) |
| 10 | Alphaproteobacteria | Roseomonas sp. (KF254767.1) |
DownLoad: CSV| Band Identity | Class | Closest relative (Acc. No.) |
| 1 | Bacilli | Bacillus sp. (GU271888.1) |
| 2 | Betaproteobacteria | Uncultured Dechloromonas sp. (JQ012310.1) |
| 3 | Betaproteobacteria | Azospira oryzae (KF260987) |
| 4 | Betaproteobacteria | Uncultured Dechloromonas sp. (KF003189.1) |
| 5 | Alphaproteobacteria | Novosphingobium sp. (KF544940.1) |
| 6 | Alphaproteobacteria | Novosphingobium sp. (KF544932.1) |
| 7 | Alphaproteobacteria | Sphingomonas sp. (AY521009.2) |
| 8 | Alphaproteobacteria | Sphingomonas suberifaciens (AY521009.2) |
| 9 | Alphaproteobacteria | Sphingomonas sp. (JQ928361.1) |
| 10 | Alphaproteobacteria | Roseomonas sp. (KF254767.1) |
