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Aquatic Toxicology 161 (2015) 253–266

Contents lists available at ScienceDirect

Aquatic Toxicology

j ourna l ho me pa ge: www.elsev ier .com/ locate /aquatox

esilience and recovery: The effect of triclosan exposure timinguring development, on the structure and function of river biofilmommunities

.R. Lawrence a,∗, E. Topp b, M.J. Waiser a, V. Tumber a, J. Roy a, G.D.W. Swerhone a,. Leavitt c, A. Paule d, D.R. Korber e

Environment Canada, 11 Innovation Blvd., Saskatoon, SK S7N 3H5, CanadaAgriculture and Agri-Food Canada, London, ON, CanadaUniversity of Regina, Regina, SK, CanadaGlobal Institute for Water Security, University of Saskatchewan, Saskatoon, SK, CanadaFood and Bioproduct Sciences, University of Saskatchewan, Saskatoon, SK, Canada

r t i c l e i n f o

rticle history:eceived 15 October 2014eceived in revised form 22 January 2015ccepted 20 February 2015vailable online 21 February 2015

eywords:esilienceecoveryiofilmommunityriclosanototorque

a b s t r a c t

Triclosan (TCS) is a ubiquitous antibacterial agent found in soaps, scrubs, and consumer products. Thereis limited information on hazardous effects of TCS in the environment. Here, rotating annular reactorswere used to cultivate river biofilm communities exposed to 1.8 �g l−1 TCS with the timing and durationof exposure and recovery during development varied. Two major treatment regimens were employed: (i)biofilm development for 2, 4 or 6 weeks prior to TCS exposure and (ii) exposure of biofilms to TCS for 2, 4or 6 weeks followed by recovery. Biofilms not exposed to TCS were used as a reference condition. Commu-nities cultivated without and then exposed to TCS all exhibited reductions in algal biomass and significant(p < 0.05) reductions in cyanobacterial biomass. No significant effects were observed on bacterial biomass.CLSM imaging of biofilms at 8 weeks revealed unique endpoints in terms of community architecture.Community composition was altered by any exposure to TCS, as indicated by significant shifts in dena-turing gradient gel electrophoresis fingerprints and exopolymer composition relative to the reference.Bacterial, algal and cyanobacterial components initially exposed to TCS were significantly different fromthose TCS-free at time zero. Pigment analyses suggested that significant changes in composition of algaland cyanobacterial populations occurred with TCS exposure. Bacterial thymidine incorporation rateswere reduced by TCS exposure and carbon utilization spectra shifted in terms substrate metabolism.

Direct counts of protozoans indicated that TCS was suppressive, whereas micrometazoan populationswere, in some instances, stimulated. These results indicate that even a relatively brief exposure of a riverbiofilm community to relatively low levels of TCS alters both the trajectory and final community struc-ture. Although some evidence of recovery was observed, removal of TCS did not result in a return to theunexposed reference condition.

Crown Copyright © 2015 Published by Elsevier B.V. All rights reserved.

. Introduction

Triclosan (2,4,4-trichloro-2-hydroxydiphenylether) (TCS) is aigh-use, broad-spectrum antimicrobial compound, and it has been

etected in sewage waste waters and sludges at concentrationss high as 1000–8000 ng l−1 and in a variety of aquatic habitats,ncluding rivers and streams (Halden and Paull, 2005). Recorded

∗ Corresponding author. Tel.: +1 306 975 5789; fax: +1 306 975 5143.E-mail address: john.lawrence@ec.gc.ca (J.R. Lawrence).

ttp://dx.doi.org/10.1016/j.aquatox.2015.02.012166-445X/Crown Copyright © 2015 Published by Elsevier B.V. All rights reserved.

levels of triclosan range from 0.05 to 0.15 �g l−1 (Okumura andNishikawa, 1996) while others report a concentration rangebetween 0.027 and 2.7 �g l−1 in waste water treatment plant efflu-ents (Chalew and Halden, 2009). General toxicity of TCS to otherbiota is possible due to its action as a reactive chemical or anon-polar narcotic (Verhaar et al., 1992); thus, toxicity concernsextend beyond aquatic bacteria. For example, evidence suggests

that environmentally-relevant concentrations of triclosan inducechanges in hormone-mediated metamorphosis in the North Amer-ican bullfrog (Veldhoen et al., 2006). It is also an inhibitor ofenoyl–acyl carrier protein reductase (ENR), an enzyme which is

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nvolved in bacterial lipid biosynthesis (Adolfsson-Erici et al., 2002),uggesting links to antibiotic resistance.

Despite evidence for TCS’s toxicological effects, particularly forlgae (Wilson et al., 2003), its potential bioaccumulative behaviorAdolfsson-Erici et al., 2002; Balmer et al., 2004) and demon-trated presence in the aquatic environment (Halden and Paull,005), questions still remain regarding the ecological implica-ions, particularly at environmentally-relevant concentrations. Toddress these questions, we examined the response to TCS of com-lex microbial communities i.e., river biofilms, which are highlyiverse, abundant, ubiquitous, stationary, reliably-exposed to thetress, and are integral components of a functional ecosystem.ommunity-level testing in ecotoxicology can be a useful tool dueo its relatively high ecological realism and reliability (Porsbringt al., 2007; Sabater et al., 2007). Several studies have addressedriclosan’s impacts on complex aquatic communities (Drury et al.,013; Franz et al., 2008; Lawrence et al., 2009; Ricart et al., 2010;roia et al., 2011; Nietch et al., 2013). Lawrence et al. (2009)eported that TCS exposure at 10 �g l−1 resulted in significanteductions in algal biomass, carbon utilization, changes in bacterialommunity composition and alteration of the community struc-ure. Exposures to effluents, however, may occur intermittently,r in pulses (Ellis 2007), therefore, pulsed or time-variable expo-ures are also of considerable interest (Proia et al., 2011; Reinertt al., 2002; Zafar et al., 2012). Proia et al. (2011) used an exposureecovery design to assess river biofilm community responses to TCS.urthermore, impacts are influenced by concentration, length ofxposure and developmental phase of biofilm communities. Thus,t is essential, to also consider community resistance and resilienceo relevant concentrations of pollutants. Such information couldrovide a more complete understanding of the potential impactsf these stressors to the ecosystem as a whole (Proia et al., 2011).

n the present study, effects of 1.8 �g l−1 of TCS on river biofilmommunities were studied with time-variable durations of estab-ishment, exposure and recovery. The main hypothesis was thatCS would cause changes in the structure and function of biofilmsfter variable terms of development, exposure and recovery, andhat these effects and degree of recovery would differ dependentpon the timing of exposure during biofilm development. Basedn established toxicological effects of TCS, we would predict sig-ificant negative impacts on photosynthetic components of theommunity. However, there is little basis for prediction regardinghe trajectory of recovery.

. Materials and methods

.1. Microcosm operation

Experimental exposures were performed in rotating annulareactors (see Lawrence et al., 2000, 2004). Natural river waterSouth Saskatchewan River, Saskatoon, SK, Canada) was used asnoculum and a source of carbon and nutrients. TCS was addedirectly to individual reactors using a peristaltic pump. Nutrient

evels were assessed as described by Chénier et al. (2003) andhe reactors were maintained at 21 ± 2 ◦C under continuous illu-

ination and rotating at 150 rpm. Water was pumped throughhe clear glass reactors at a rate of 500 ml per day (one reactorolume) using a multichannel peristaltic pump (Watson Marlow,

ilmington, MA). Replicated (n = 3) experimental treatments con-isted of two basic TCS (1.8 �g l−1) exposure regimes: (i) whereCS was added from time = 0 and after 2, 4, or 6 weeks replaced

y river water alone (T2, T4, and T6) or continuously exposedo triclosan (T8) or, (ii) where river water alone was added fromime = 0 and replaced by river water plus TCS after 2, 4, 6 weeksR2, R4, and R6) of growth. Triplicate reactors that received river

ology 161 (2015) 253–266

water alone (R8) acted as the reference condition or control forall other treatments. TCS concentration used was several orders ofmagnitude below medical-industrial application levels (0.2–1.0%)and within the range reported for aquatic environments (i.e., Ricartet al., 2010; Nietch et al., 2013). After 8 weeks of biofilm develop-ment, the twelve, 1 × 10 cm polycarbonate coupons were removedfrom replicate reactors for immediate analysis (confocal laser scan-ning microscopy (CLSM) microscopic analyses of bacterial, algal,cyanobacterial and exopolymer composition and biomass, carbonutilization, thymidine incorporation, TCS degradation, pigments,and dry weight) and also frozen at −80 ◦C and stored for subsequentDNA extraction and molecular analyses.

2.2. Confocal laser scanning microscopy (CLSM) and imageanalysis

Microscopy studies were carried out with an MRC 1024 confo-cal laser scanning microscope (Bio-Rad Hemel Hempstead, UnitedKingdom) attached to a Microphot SA microscope (Nikon, Tokyo,Japan). The following water-immersible lenses were used: 63×/0.9NA (Zeiss, Jena, Germany), 40×/0.55 NA and 10×/0.35 NA (Nikon,Japan). Replicate slides from all replicate reactors were cut into1 cm2 pieces and mounted in small Petri dishes using Dow Corn-ing #3140 acid-free silicone (WPI, Inc., Sarasota, FL) and thenstained and analyzed. Bacteria were stained with the fluorescentnucleic acid stain (SYTO 9) (excitation wavelength, 488 nm [ex488]; emission wavelength, [em 522/32]); a lectin probe (Triticumvulgaris-TRITC [tetramethyl rhodamine isothiocyanate]; ex 568,em 605/32) was used as a general stain to visualize exopolymer,and autofluorescence (ex 647, em 680/32) was used to detectalgal and cyanobacterial cells (Neu et al., 2004). The CLSM opti-cal thin sections obtained as described above at a 5 �m sectioninginterval in each of the three channels allowed determination ofbacterial cell area (biomass), exopolymer biomass, cyanobacte-rial biomass, and total photosynthetic biomass at various opticalsectioning depths. Biofilm thickness was determined using pro-jections of the optical thin section stacks. NIH Image version 1.61(http://rsb.info.nih.gov/nih-image/), with macros written for semi-automated quantification as described in Manz et al. (1999), wasused for image analyses. Three color red–green–blue projections(cells stained red, polymer stained green, and algal autofluores-cence stained blue) of the biofilms were computed for visualizationpurposes.

2.3. Exopolymer analyses

Lectins labeled with either fluorescein isothiocyanate or TRITC(Sigma, St. Louis, MO) or Cy5 (Research Organics, Cleveland, OH)were applied for exopolymer analyses. The lectins T. vulgaris (�(1,4)N-acetyl glucosamine, N-acetyl neuraminic acid), Arachis hypogaea(terminal �-galactose, N-acetyl galactosamine (associated withalgal–cyanobacterial polymers)), Canavalia ensiformis (�-linkedmannose or glucose residues), Glycine max (terminal �- or �-linkedN-acetylgalactosamine (associated with algal–cyanobacterial poly-mers)), and Ulex europaeus (�-l-fucose) were used alone or incombination for in situ analyses of polymer composition. Staining,imaging, image analyses and calculations of lectin binding volumeswere carried out using the equations of Neu et al. (2001).

2.4. Protozoan and micrometazoan enumeration

Protozoa and micrometazoa (rotifers, nematodes) were enu-merated weekly by counting manually on triplicate 2 cm2

subsamples using phase contrast on an Olympus BH-2 microscope

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ith an MS plan 10×, NA 0.03, objective lens. The entire 2 cm2

ubsample was examined at each time period.

.5. Carbon utilization

Carbon utilization spectra were determined for biofilm samplessing commercial Eco-plates (Biolog, Hayward, CA) as describedreviously (Lawrence et al., 2004, 2009). Biofilm was removed fromhe coupons using a sterile spatula, samples were homogenized byortexing (30 s), diluted in sterile river water 10−4 ml and inoc-lated in triplicate into the Eco-plates. Plates were incubated at2 ± 2 C for 7 days prior to reading analyses with a Biolog-96 welllate reader and manufacturers software as described by the man-facturer.

.6. Chemical analyses

Reference materials as well as influent and effluent from theeactors were collected, placed on ice and submitted to the Albertannovates Analytical Facility (Vegreville, AB, Canada) for extractionnd determination of TCS levels. A liquid/liquid (dichloromethane)xtraction method was used and extracts analysed on an Ion Trap,arian GC/MS (Santa Clara, CA).

.7. Radioisotope analyses

TCS mineralization was assessed as the percentage of 14CO2roduced from universally-labelled (UL) ring of [14C] triclosan (spe-ific activity 0.1 mCi mmol−1 added at 100 �g l−1 and 36,000 dpm).hymidine analysis: 4 replicates of 5 cm2 of biofilm on theolycarbonate coupons were put into 3–100 mm × 15 mm sterileolystyrene petri plates containing 4.88 ml of filter sterilized riverater and 1 plate with added formaldehyde to kill the biofilm.

25 ml of a [methyl]-3H thymidine solution was added to bringhe concentration of the thymidine to 20 nM and the plates werencubated for 30 min. Then the biofilm was killed with formalde-yde scraped off the coupons with a silicone scraper, transferredo a 50 ml conical bottom polypropylene centrifuge tube and ana-yzed according to the procedure of Robarts and Wicks (1989). All

egative controls were killed with formaldehyde at 4% final con-entration. All activity levels were measured by liquid scintillationpectrometry (Tri-Carb 2100TR, Packard Instruments, Downersrove, IL).

ig. 1. Representative CLSM photomicrographs of microbial communities that experiencCS was added from time = 0 and after 2, 4, or 6 weeks replaced by river water alone (T2,

iver water plus TCS after 2, 4, and 6 weeks (R2, R4, and R6). Bacteria (green), Triticum vuFor interpretation of the references to color in this figure legend, the reader is referred to

ology 161 (2015) 253–266 255

2.8. Pigment analyses

Algal and bacterial pigments were extracted, filtered and driedunder N2 gas following the procedures of Leavitt and Hodgson(2001). Briefly, lipid-soluble pigments were extracted from aknown surface area (cm2) of biofilm by soaking in a mixture of ace-tone:methanol:water (80:15:5, by volume) for 24 h in darkness andunder an inert N2 atmosphere at 4 ◦C. Pigment concentrations werequantified by reversed-phase high performance liquid chromatog-raphy (RP-HPLC). Specifically, carotenoid, chlorophyll (Chl), andpigment-derivative concentrations were quantified using an Ali-gent 1100HPLC system following the reversed-phase procedure ofLeavitt and Hodgson (2001). The Agilent 1100 system was equippedwith a C-18 column (5-�m particle size; 10-cm length), and anAgilent model 1100 photodiode array spectrophotometer (435-nmdetection wavelength). An internal reference standard (3.2 mg l−1)of Sudan II (Sigma Chemical Corp., St. Louis, MO) was injected ineach sample.

2.9. Molecular analyses

2.9.1. Total community DNA extractionFor each treatment bioreactor, a frozen (−80 ◦C) polycarbon-

ate strip was aseptically cut (2 cm2) and transferred to a 50 mlpolypropylene tube (Falcon, Becton Dickinson, Franklin Lanes, NJ).Cells and associated materials from the frozen biofilm sampleswere removed from the polycarbonate strip with a sterile metalscraper. Total DNA was extracted using the FastDNA spin kit forsoil (Bio101 systems Qbiogene, Carlsbad, CA) according to manu-facturer’s instructions.

2.9.2. PCR amplificationThe bacterial 16S rRNA gene was amplified using “universal”

primers to perform DGGE. The primer consensus sequence forwardwas 5′- CCT ACG GGA GGC AGC AG -3′, preceded by a GC clampfor DGGE (not for sequencing) = CGC CCG CCG CGC CCC GCG CCCGGC CCG CCG CCC CCG CCC G (40 nt) and reverse, 5′- CCG TCA ATTCMT TTG AGT TT-3′ position (length) 341–357 (17 nt) and 907–926(20 nt), respectively, and the amplified PCR fragment size was 586

base pairs (Muyzer et al., 1993; Muyzer and Ramsing, 1995). PCRamplification was conducted in a 25 �l reaction volume contain-ing 1 �l of DNA template, 10 pmol of each appropriate primer asdescribed by Muyzer et al. (1993) and Muyzer and Ramsing (1995),

ed no TCS exposure (R8) versus continuous TCS-treatment (T8), and those whereT4, and T6), or where river water alone was supplied from time = 0 and replaced bylgaris-TRITC lectin binding polymer (red), photosynthetic biomass (blue/magenta).

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exopolymeric composition. Detailed analyses of replicate commu-nities (n = 3) and CLSM image stacks (n = 15) were carried out toestablish the quantitative distribution of each component. ANOVAresults indicated that TCS had no significant effect (p < 0.05) on bac-

Fig. 2. (A) Results of ANOVA (p < 0.05) using the results of bacterial, cyanobacterialand algae biomass determined by digital image analyses (ANOVA bars followed by

56 J.R. Lawrence et al. / Aquati

.25 U Taq DNA polymerase (New England Biolabs Ipswich, MA),× PCR buffer, 2.5 mM MgCl2, and 200 �M dNTPs. A touchdownCR program using the PTC-200 thermocycler (MJ Research, Inc.,altham, MA) consisted of an initial denaturation step of 94 ◦C

or 5 min, followed by 10 cycles of denaturation at 94 ◦C for 1 min,nnealing at 66 ◦C (decreasing in each cycle by 1 ◦C) for 1 min andn elongation step of 72 ◦C for 1 min. Following these steps, another0 cycles of 95 ◦C for 1 min, annealing at 56 ◦C for 1 min, and elonga-ion at 72 ◦C for 1 min, with a final elongation step of 72 ◦C for 7 min,ere performed. The correctly-sized PCR product was verified by

lectrophoresis on a 1.5% w/v agarose gel in 1.0× Tris-acetate-EDTATAE) buffer (40 mM Tris, 20 mM acetic acid and 1 mM EDTA) for.0 h at 100 V. Gels were stained using ethidium bromide and docu-ented using the AlphaImager 3300 gel documentation and image

nalysis system (Alpha Innotech Corporation, San Leandro, CA).

.9.3. Denaturing gradient gel electrophoresis analysis (DGGE)After the specificity and size of the amplified products were

hecked on agarose gels, the PCR product was separated by DGGEMuyzer et al., 1993; Muyzer and Ramsing, 1995) using an IngenyhorU2 system (Ingeny, Leiden, The Netherlands). Aliquots (20 �l)f PCR product were mixed with 4 �l of loading dye buffer andesolved on a 6% (w/v) polyacrylamide gel in 1.0× TAE buffer usingenaturing gradients from 45 to 65% (100% denaturant contains 7 Mrea and 40% deionized formamide). DGGE was carried out at 40 V

or 10 min, and then 100 V for 18 h at 60 ◦C. After electrophoresis,he gel was stained with SYBR Green I (1:10000 dilution; Molecularrobes, Eugene, OR) for 15 min with gentle agitation and pho-ographed using the AlphaImager 3300 gel documentation andmage analysis system (Alpha Innotech Corporation, San Leandro,A).

.10. Experimental design and statistical analyses

The experimental design included the following reference treat-ent: no exposure to TCS, for the entire 8 week development period

R8). River biofilms were allowed to either develop in the absencef TCS for 2, 4 or 6 weeks (designated “R”, R2, R4, and R6) with cor-esponding periods of exposure 6, 4, or 2 weeks or to develop withnitial TCS exposure for 2, 4, 6 or 8 weeks (designated “T” T2, T4,6, and T8) with corresponding periods of recovery for 6, 4, 2, or 0eeks. Each treatment had 3 identical replicate reactors randomly

ssigned to it on the reactor bench (experimental replications).ach analysis was done on subsamples from randomly selectediofilm coupons from among the 12 identical coupons from eacheplicate reactor (n = 3) at the end of the 8 week experiment. CLSMmaging was done at 5 locations across a transect on a 1 cm2 piecef the biofilm coupon from each replicate reactor. Subsamplingor other analyses, protozoan counts, thymidine incorporation,CS mineralization and carbon utilization, was also carried outsing randomly selected subsamples from among the 12 identi-al coupons from each replicate reactor (n = 3). Analysis of varianceas used to detect significant differences among sample means

t p < 0.05 using the commercial package, MiniTab (State College,A). Band detecting, matching and processing of DGGE gels wereompleted with the GelCompare II software 4.6 (Applied Maths,otrijk, Belgium). Fingerprint data based on presence absence wasrocessed by generating a band-matching table (Boon et al., 2002).he binary data was exported and compared by principal compo-ent analysis (PCA) with PRIMER v6 software (PrimerE-Ltd., Lutton,K). Statistical analyses of PCA scores generated from the first twoxes were run using the analysis of similarity (ANOSIM) test based

n Euclidean distance, with PRIMERr v6 software (Clarke 1993).he inclusion of DGGE ladders allowed GelCompare II to normalizehe position of bands in all of the lanes under examination. PRIMERersion 6 was also used to perform PCA and cluster analyses on

ology 161 (2015) 253–266

other data sets obtained from the analyses i.e., carbon utilization,biomass, lectin binding, etc.

3. Results

3.1. Total community composition

Examination of confocal laser micrographs shown in Fig. 1 indi-cated that each community, regardless of its exposure regime,appeared unique in terms of algal, cyanobacterial, bacterial and

the same letter are not significantly different p < 0.05), (B) cluster analyses of thebacterial, algal, cyanobacterial biomass data, and (C) results of PCA analyses, showingfundamental groupings based on algal, bacterial and cyanobacterial biomass andwhich elements are the major drivers of the separation. Circles indicate Euclideandistance.

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erial biomass. Significant (p < 0.05, ANOVA) impacts on algal andyanobacterial biomass, however, were detected (Fig. 2A). Clusternalyses of algal–cyanobacterial–bacterial biomass data sets con-rmed a fundamental separation of communities based on whether

hey were exposed to TCS at time = 0 (“T”), or subsequent to a TCS-ree establishment interval (“R”) (Fig. 2B). Further PCA analysesFig. 2C) indicated that the drivers of these differences were changesn algal biomass relative to the reference community, while changesn cyanobacterial biomass contributed to separations in the otherxposure regimes. In the case of phototrophs, the most significantegative effects were evident in communities that initially devel-ped in the absence of TCS (R series). Increasing photosyntheticiomass (algae plus cyanobacteria) in the longer-term 6 and 8 weekontinuous TCS exposures was observed perhaps consistent withdaptation (Fig. 2A).

.2. Algal–cyanobacterial biomass

Confocal imaging of biofilms developing without TCS all showed decline in both algal and cyanobacterial biomass when subse-uently exposed to the chemical (Fig. 2A). The only significant algaliomass reduction (ANOVA p < 0.05) however, was detected in theommunity only exposed to TCS for the last two weeks of develop-ent (R6).

Those receiving TCS for 4 or 6 weeks (and then having thetress removed for 4 or 2 weeks) conversely showed a trendf increasing algal biomass consistent with recovery. Of biofilms

nitially-exposed to TCS (T8 and T6) algal biomass was not sig-ificantly different from the untreated reference (Fig. 2A). Thoseeceiving TCS for 2 or 4 weeks prior to switching however, hadignificantly lower algal biomass than the reference community.lthough algal biomass tended to increase with TCS exposure timemong initially-treated biofilms (T series); in no case did photo-

ynthetic biomass increase to the same level as in the referenceommunity. Although based on the increase in total photosyntheticiomass (algae plus cyanobacteria) in T6 and T8 there was a trendo development of an adapted photosynthetic community after 6–8

ig. 3. Results of pigment analyses for microbial communities exposed to variable dureference community (R8) = 0, (B) results of cluster analyses showing the similarity of comundamental grouping based on pigment concentrations and the major drivers of the sep

ology 161 (2015) 253–266 257

weeks of TCS exposure. This adaptation is evident in the clusteranalyses shown in Fig. 2B.

Cyanobacterial biomass in biofilms subsequently exposed to TCS(R series) was less than, and significantly different (p < 0.05) from,the reference community in all cases (Fig. 2A). In contrast, when TCSwas added from time = 0, cyanobacterial biomass was not signifi-cantly different (p < 0.05) in any case, from the unexposed referencecommunity (Fig. 2A). For cyanobacteria, there was also a trendto biomass recovery with increasing TCS exposure time (Fig. 2A).Overall, these results indicate that exposure at any time duringdevelopment will significantly affect the photosynthetic biomassand initiate selection of a more resistant or tolerant community.

3.3. Photosynthetic community structure

Pigment analyses (Fig. 3A–C) indicated that thealgal–cyanobacterial community was significantly (p < 0.05)different compared to unexposed reference (R8) and 8 week TCSexposure (T8) communities. Interestingly, particular changes inChl-a and fucoxanthin were associated with longer TCS exposures(Fig. 3A). As shown in Fig. 3B, communities receiving TCS postbiofilm-establishment for 4 and 6 weeks (R2 and R4) were moresimilar to biofilms receiving TCS for 8 weeks. In contrast, thosebiofilms receiving TCS for 2, 4 or 6 weeks before shifting to TCS-freewater were more similar to the reference community (R8), as werelate TCS-exposed biofilms receiving only two weeks TCS (T2)in the final development phase. This observation suggests thatalgal–cyanobacterial community structure (but not biomass) mayrecover from exposure in as little as 2 weeks, but when developedin the absence of TCS may resist significant change in communitystructure for more than 2 weeks, although (see above) biomassmay be affected. Generally, any TCS exposure had clear effects onphotosynthetic community structure.

3.4. Bacterial biomass and activity

In no case did TCS exposure result in a significant (p < 0.05)decline in bacterial biomass (CLSM imaging and digital image anal-

ations of TCS. (A) Illustrates the change in pigment concentration relative to themunities to the reference condition and each other, and (C) PCA analyses showing

aration. Circles indicate Euclidean distance.

258 J.R. Lawrence et al. / Aquatic Toxic

Table 1Results of thymidine incorporation assays.

Treatment % Thymidinea

R8 100b*

R6 76abR4 64aR2 62abT2 57aT4 63aT6 57aT8 70a

* Values followed by the same letter are not significantly different (p < 0.05A

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the major driver of this is their use of polymers. Whereas, in keep-ing with plots showing the deviation from the reference condition

F(

NOVA) (n = 3).a % relative to reference R8 = 100%.

ses (Figs. 1 and 2A)) or biofilm dry weight measures (data nothown). Thymidine incorporation (Table 1) however, suggestedhat although not always significantly different (p < 0.05) any TCSxposure resulted in a decline in bacterial community metabolicotential relative to the unexposed reference. When TCS exposureccurred during establishment (T series), reduction in thymidine

ncorporation was sustained throughout the experimental periodith no significant (p < 0.05) indication of recovery.

ig. 4. Results of carbon utilization assays using the Biolog Ecoplate system showing shiftA) polymers, (B) amino acids, (C) amines, and (D) others. Asterix indicates significantly d

ology 161 (2015) 253–266

3.5. Carbon substrate utilization

Significant effects of TCS on community metabolism weredetected using carbon utilization spectra. Generally, there was adifference in response between T series and R series regardless ofsubstrate. As indicated by examining shifts in utilization relativeto the reference (R8), TCS exposure resulted in either significantdepression or enhancement of substrate use (Fig. 4A–D). For exam-ple, use of polymers and selected amino acids were significantlyincreased in communities initially exposed to TCS (Fig. 4A and C).Conversely, TCS exposure suppressed utilization of carboxylic acids(Fig. 4B) amines and phosphates + esters (Fig. 4D). Further, in com-munities subsequently treated with TCS (R series), the 6 weeks TCSexposure (R2) appeared to shift polymer utilization patterns (i.e.,Tween 40/80) (Fig. 4A).

Carbon utilization analyses based on those sources showingdeviation from the reference condition indicated that communi-ties that experienced a recovery phase after exposure (T2, T4, andT6) were similar (Fig. 5A and B). As shown by the PCA analyses

(Fig. 4 A–D), those that were subsequently exposed to TCS for aninterval <6 weeks after development clustered based on their use

s in utilization of specific substrates classes relative to the reference community = 0,ifferent from reference community (R8), p < 0.05.

J.R. Lawrence et al. / Aquatic Toxicology 161 (2015) 253–266 259

Fig. 5. Results of carbon utilization assays using the Biolog Ecoplate system showing (A) cluster analyses using selected carbon sources (polymers = pol, amino acids = aa,amines = am, carbohydrates = cbh. carboxylic acids = cbc, esters = es, phosphates = ph) shown to respond to TCS treatment (see Fig. 4) and (B) PCA analyses using selectedcarbon sources. Circles indicate Euclidean distance.

Fig. 6. Results of fluorescent lectin binding analyses of major sugar residues in the exopolymers of the treatment communities. Bars with different letters are significantlydifferent from the reference community and other treatments (p < 0.05, ANOVA) ANOVA bars labelled with the same letter are not significantly different p < 0.05.

2 c Toxicology 161 (2015) 253–266

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Fig. 7. Results of fluorescent lectin binding analyses of the exopolymers of the treat-ment communities; (A) illustrates of the change in lectin binding volumes relativeto the reference community = 0, (B) presents results of cluster analyses showing thesimilarity of communities to the reference condition and each other, and (C) shows

60 J.R. Lawrence et al. / Aquati

f amines (Fig. 5 A and B). Phosphates and carboxylic acids (Fig. 5B)lso contributed to these separations. Interestingly, those biofilmseceiving TCS either for a long term or not at all grouped similarly,riven in part by their use of esters, carbohydrates and amines. Noecovery to the reference condition was observed.

.6. Exopolymer composition

Since EPS composition is dependent upon the types of produc-ng organisms present lectin binding can be used to assess microbialommunity structure. Biofilm communities initially grown withoutCS (R2, R4, and R6) were all significantly different from the unex-osed reference community based on EPS composition (p < 0.05)Fig. 6). Within this set of late TCS exposure regimes however,lthough communities with either 2 or 4 weeks of TCS-free growthR2 and R4) were not significantly different, EPS in communi-ies with 6 weeks TCS-free growth (R6) was significantly differentrom all others. Exposure of communities to TCS for the 8 or 6

eeks period (T6, T8) resulted in similar EPS pattern/structurep < 0.05); interestingly, T8 was also not significantly different fromiofilms grown for 6 weeks TCS free before exposure to TCS forhe final two weeks (R6). This observation was consistent withhe demonstrated effect on algal–cyanobacteria community struc-ure, as shown above. The pairs of 4 weeks TCS on–off/4 weeks TCSff–on communities (T4 and R4), as well as both cases of 6 weeksCS on–off/2 weeks TCS off–on, (T6 and R2) were not significantlyifferent from each other (p < 0.05) (Fig. 6). Relative to referenceonditions, Fig. 7A showed a consistent decrease in the amounts oferminal �-galactose, N-acetyl galactosamine residues (associatedith algal–cyanobacterial polymers) and increases in �-l-fucose

esidues, more in keeping with bacterial EPS. These pairings werelso seen in the outcomes of the PCA analyses (Fig. 7C) which sug-ests that terminal �-galactose, N-acetyl galactosamine residuesnd terminal �- or �-linked N-acetyl galactosamine residues wereajor drivers of differences separating the 8 week reference com-unity from the 6 week- and 8 week-TCS initially-exposed biofilms

T6 and T8). Indeed, any exposure to TCS resulted in a decreasen these two residues relative to the reference community, anbservation entirely consistent with their being associated withlgal/cyanobacterial exopolymers. In contrast, �-linked mannoser glucose residues and �-l-fucose increased when TCS exposureccurred regardless of duration, in keeping with their bindingffinities for bacterial polymers. These changes which occurredithin two weeks of exposure are reflective of shifts in the com-unity structure and diversity (both photosynthetic and bacterial).

nterestingly, the symmetrical T and R series 2-4-6 week exposuresesult in similar communities based on lectin binding analyses.ecovery to the reference condition was not observed in any case.

.7. Bacterial community structure (DGGE)

DGGE analyses were used to assess the bacterial communitytructure and how it varied with treatment. DGGE profiles clearlyhowed differences in banding patterns between treatments (dataot shown). Analyses of these patterns of the profiles (as observed

or other parameters) supported the clear distinction of commu-ities initially receiving TCS exposure (T series) versus subsequentxposure (R series) of established biofilm communities. PCA anal-ses of DGGE data (Fig. 8) further illustrates this outcome. ANOSIMnalyses, where R = 0.78 and p < 0.001 confirmed a significant dif-erence between communities initially exposed to TCS (T) andhose subsequently exposed (R). The community exposed to TCS

or eight weeks, as well as the 2, 4, and 6 week TCS (T series)xposures, were significantly (p < 0.001) different from all com-unities that were initially TCS-free (R series). The community

nitially-exposed to TCS for two weeks (T2) was not significantly

PCA analyses where fundamental groupings based on exopolymer concentrationsand the major drivers of the separation. Asterix indicates significantly different fromreference (R8) p < 0.05, Circles indicate Euclidean distance.

different from its TCS-initial T4, T6, and T8 weeks exposure coun-terparts; conversely, the community initially-exposed to TCS for4 weeks (T4) was significantly different from that exposed to TCSfor 6 weeks prior to switching (T6), or continuously exposed to

TCS for 8 weeks (T8). By this measure, the community exposed toTCS for 6 weeks prior to switching was not significantly (p < 0.001)different from 8 full weeks of TCS exposure. Within communitiesnot initially exposed to TCS (R series), the R2 biofilms (2 weeks

J.R. Lawrence et al. / Aquatic Toxicology 161 (2015) 253–266 261

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xposed to TCS and those subsequently exposed. Circles indicate percent similarity

CS-free growth, 6 weeks TCS exposure) were significantly dif-erent from their counterparts, while the R4 community was notignificantly different from the community that developed 6 weeksCS free followed by a final 2 weeks of TCS exposure (R6). A signif-

cant difference (p < 0.001) however, was detected between the R6nd the unexposed reference community (R8). Average numbers ofands detected in the replicate treatments were as follows, T2/11.6,4/11.0, T6/8, T8/10.6, versus R2/15.7, R4/14.0, R6/13.7, R8/13.7.hus, initial exposure to triclosan (T-series) during developmentroduced reductions in total band number detected.

The results suggest that recruitment to the attachment surfacend biofilm in the presence of TCS resulted in a substantially-ifferent bacterial community. This community was quite resistanto change within the 8 week experimental cycle since the 2 weekCS initial exposure community (T2) was not significantly differ-nt from the T4-, T6- and T8-exposed communities. Apparently,ommunities that initially develop in the absence of TCS were notignificantly different from each other, with the exception of theairwise comparison of the 6 week TCS-free (R6) and the unexposedR8) reference community. Based on the DGGE analysis it appearshat the initial bacterial community recruited from the planktonichase is relatively-resistant to change.

.8. Protozoa and micrometazoa

Microscopic examination of the developing microbial commu-ities allowed regular (weekly) monitoring of the predatory orrazer populations in each treatment. Addition of TCS after anstablishment phase generally appeared to stimulate micrometa-oan populations, although the impact diminished with increasedxposure time (Fig. 9A–D). In contrast, when TCS was present dur-ng establishment there was evidence of recovery when the initialCS exposure was limited to two weeks (T2), with clear negative

mpacts for TCS 4 weeks on/4 weeks off (T4) and TCS 8 week expo-ures (Fig. 9B and D). Protozoa were most abundant in the referenceommunity and TCS exposure resulted in negative impacts on pro-ozoan populations in all treatments relative to the control (Fig. 9And C). These observations suggest effects of reduced grazing inten-ity on the microbial community structure as a consequence of anyCS exposure, however no significant differences (p < 0.05) wereetected.

.9. Degradation of 14C labeled triclosan

The various treatments had no effect on the degradation of 14Cabeled TCS by the biofilm communities (data not shown).

8 and p < 0.001 confirmed a significant difference between communities initially

4. Discussion

In the current experimental study, triclosan addition created adisturbance with defined timing, frequency and duration. Relativeto the life cycle of microbial community members, the approachused here may be considered a pulse disturbance, although thisis subject to some discussion (see Shade et al., 2012). As notedby Victoria and Gómez (2010), in ecotoxicology it is increasinglyrecognized that elucidating how communities respond to distur-bances, whether they exhibit resistance or resilience, is criticalto understanding potential hazards. Aquatic biofilm communi-ties drive many biogeochemical cycles and trophic relationships,respond to change over short time frames, and are amenableto the use of multiple types of analyses. These characteris-tics make them highly-relevant and suitable for assessment ofresistance–resilience phenomena in aquatic ecosystems (Lawrenceet al., 2009; Proia et al., 2011, 2012; Sabater et al., 2007). Previ-ous resilience experiments used a variety of community cultivationapproaches: artificial channels, mesocosms, as well as in situ cul-tivation and transplantation to create scenarios of exposure andrecovery where return to reference composition and function ismonitored using various parameters (Begon et al., 2006; Baho et al.,2012; Proia et al., 2011; Zafar et al., 2012). In the current studythe experimental set up used rotating annular reactors (Lawrenceet al., 2000) where there was continuous flow-through of riverwater with metered addition of TCS to achieve a controlled expo-sure. This flow-through approach eliminates concerns regardingdevelopment of toxic degradation products such as dioxins, whichmay be generated by photodegradation in the reactor (Ricart et al.,2010). The selection of 1.8 �g l−1 TCS as the exposure concentra-tion was based on exceeding the no observed effects concentration(NOEC) of 0.21 �g l−1 for bacteria (Ricart et al., 2010) and in therange reported to cause effects on algal communities (from 0.015to 1.5 �g l−1; Wilson et al., 2003) and with reported environmentalconcentrations for TCS (0.027–2.7 �g l−1; Ricart et al., 2010).

TCS effects on a microbial community may occur both directly(toxicity) and indirectly (ecological interactions), as indicatedpreviously (Nietch et al., 2013; Lawrence et al., 2009). Hypo-thetically, longer exposures may cause greater indirect effects onthe microbial community through effects on bacterial-phototrophinteractions and grazing, as seen in this study (see below). It iswell-established that TCS is a broad spectrum antimicrobial active

against both Gram-positive and Gram-negative bacteria, specifi-cally it inhibits the enoyl–acyl carrier protein reductase enzyme(ENR), which is involved in bacterial membrane lipid biosynthesis(Adolfsson-Erici et al., 2002; McMurry et al., 1998; Levy et al., 1999).

262 J.R. Lawrence et al. / Aquatic Toxicology 161 (2015) 253–266

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ig. 9. Results of protozoan and metazoan analyses by treatment; (A and B) shows thhe change in numbers of grazers relative to the reference community = R8.

hen a compound has a specific target that is common to a num-er of organisms it may have toxicity at concentrations much lowerhan predicted (Verhaar et al., 1992; Orvos et al., 2002). An examplef such a target is the FabI pathway which is involved in plant, algal,rotozoan and bacterial lipid biosynthesis (Tatarazako et al., 2004;hite et al., 2005). TCS exhibits a broad range of impacts with NOEC

evels for aquatic species that include: algae 0.69 �g l−1, aquaticlants 62.5 �g l−1, fish 100 �g l−1, acute invertebrates 6 �g l−1, andhronic invertebrates 40 �g l−1 (Reiss et al., 2002).

.1. Response of phototrophs

TCS appears to be phytotoxic, as shown in mixed communitiesWilson et al., 2003), and in a number of pure culture toxicity assaysTatarazako et al., 2004). Ricart et al. (2010) used a measure of pho-osynthetic efficiency to determine a NOEC for the algal communityf 0.42 �g l−1. Generally, algae are considered the most sensitiverganisms to TCS (Dann and Hontela, 2011). Cyanobacteria, such asnabaena flos-aquae, are also sensitive with a NOEC of 0.67 �g l−1.

n contrast, Nietch et al. (2013) reported that based on enumerationf identified taxa and Chl-a determinations, stimulation occurredt low doses (0.1 �g l−1 TCS) relative to the control but suppressiont 5 and 10 �g l−1 doses over an extended exposure. For example,

yanobacteria were stimulated at <1 �g l−1, although there wereostly undetectable effects at 5 and 10 �g l−1. Findings here and

hose of Lawrence et al. (2009), suggest that some cyanobacteriaay be tolerant or adapt to TCS exposure, such tolerance resulting

ulative counts for protozoa and metazoa on a weekly basis, and (C and D) illustrates

in recovery of cyanobacterial biomass as previously indicated byNietch et al. (2013). Drury et al. (2013) treated artificial streamsto attain an initial TCS sediment concentration of 17 �g g−1. Theyreported a dramatic increase in relative abundance of cyanobacte-rial sequences, suggesting greater resistance to TCS than algae andpotential dominance with attendant ecological implications. Thesevariable results are may be explained by several factors includ-ing: initial community composition and habitat (light, nutrients,substrata) and sediment/water chemistry (Nietch et al., 2013).

In an experiment evaluating resilience of periphyton commu-nities, Proia et al. (2011) grew river biofilms in mesocosms for4-weeks and then subjected them to 48-h of short pulses ofTCS at nominal concentrations of 60 �g l−1. Following exposure,the water was replaced and communities sampled after 1 and 2weeks recovery. Monitoring included extracellular enzyme activ-ities, Phosphorus (P)-uptake, photosynthetic parameters, Chl-a,live:dead ratios for bacteria and diatoms, total bacterial numbersas well as diatom numbers and taxonomy. Proia et al. (2011) notedthat TCS increased diatom mortality at 1 week post-exposure. Bac-terial mortality also increased but returned to reference levelsafter 1 week. TCS apparently compromized the cellular integrityof Spirogyra sp., and significantly inhibited P-uptake. TCS evidentlyaffected both algal and bacterial parameters however, these param-

eters returned to reference levels within two weeks of exposure.Despite measurement of different parameters by Proia et al. (2011),apparent recovery intervals (circa 2 weeks) for algal/cyanobacterialbiomass and some bacterial parameters were similar to those

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bserved in the rototorque study reported here although the expo-ure level was much lower (1.8 versus 60 �g l−1) and the time scaleor effects and recovery longer. The data reported here suggestshat with the longer time scale, recruitment of tolerant speciesccurred in the initial phases of biofilm development, thereby cre-ting a more tolerant community than when TCS was not initially

selective agent. Lawrence et al. (2009) indicated that there waso significant reduction in cyanobacterial biomass at 10 �g l−1 TCS,

result consistent with selection-recruitment of tolerant species.verall, our results suggest that TCS exposure at any time dur-

ng development will negatively affect photosynthetic biomass andlter community composition. The most significant effects in termsf reduced photosynthetic biomass however, were observed whenstablished algal–cyanobacterial communities were exposed toCS. Observations suggest that after TCS exposure photosyntheticiomass will not be as extensive, and that based on exopolymernd pigment analyses, community composition will be altered.ignificant effects were detected after only 2 weeks exposure inn unadapted community (R6) in these rototorque experiments.dapted or exposed communities however, may accumulate simi-

ar photosynthetic biomass, and when TCS exposure is limited to 2eeks, will exhibit similar composition to the reference condition

T2). Recovery in terms of biomass appeared to occur within thexperimental time frame i.e., 6–8 weeks in adapted communitiesT series). While community composition (based on pigment data)uggested that established communities with no TCS exposure mayesist change for 2 weeks, those exposed to TCS for two weeks mayhift toward reference status within 6 weeks (T2). These effects onhototrophs (algae and cyanobacteria) will typically have a range ofegative effects on algal–bacteria linkages e.g., competition, withinhe community (Haak and McFeters, 1982a,b,b), further influencingevelopment and energy and carbon flows.

.2. Response of the bacterial community

At low concentrations, triclosan inhibits the growth of a wideange of bacteria, and acts as a bactericide at higher concentrationsGomez et al., 2005). Nietch et al. (2013) found that at 1.0 �g l−1

r less, TCS stimulated periphyton while at 5 and 10 �g l−1, itignificantly lowered bacterial and cyanobacterial cell densities.onsistent with this, inhibition of litter processing at concentra-ions of 1–10 �g l−1 was observed. Our data suggest that whileacterial biomass, as measured by digital imaging, may not changeignificantly (p < 0.05), reduction in thymidine incorporation andhanges in carbon utilization indicate effects on bacterial commu-ity metabolism. When TCS was introduced after an initial TCS freeevelopment period (R series), there was a decline in thymidine

ncorporation rate with increasing exposure. TCS application dur-ng biofilm development (T2, T4, T6, and T8) significantly (p < 0.05)educed thymidine incorporation relative to the reference (R8).ur results show changes in carbon utilization in terms of generalctivity and specific substrate utilization (polymers and amines).hese observations suggest that from a metabolic perspective, com-unities receiving TCS for a long term, or not at all, were more

imilar. Despite differences in composition, these communitiesay become functionally similar. The clustering/PCA results (Fig. 5)

or initial R-series versus T-series TCS exposure suggest that theature of the initial community recruited at time = 0 persists whenhe stress is either applied or relieved.

For biofilms developing in experimental channels, other authorsRicart et al., 2010) have reported, that TCS increased bacterial mor-ality with a NOEC of 0.21 �g l−1. Proia et al. (2011) applied a single

ose of TCS, recording increased bacterial mortality but with recov-ry within 1 week of application. In contrast, Drury et al. (2013)eported that even 8 �g g−1 sediment of TCS did not affect bacterialell abundance or community respiration rates over 34 days. They

ology 161 (2015) 253–266 263

suggested that binding of TCS to sediment may have limited theexposure, or that their sampling regime missed the response. Orvoset al. (2002) reported increased bacterial mortality upon exposure,but as with Proia et al. (2011), a return to baseline occurred withinone week.

Virtually all of our measured parameters indicated that expo-sure to TCS during development (T) results in different effectscompared to exposure of the established community (R). DGGEanalyses suggested that both bacterial community types are resis-tant, i.e., that the initial community recruited to the surface tendsnot to change significantly over the experiment. This observationis in keeping with the fact that the community with only a final 2week exposure to TCS (an unadapted community (R6)), was mostsignificantly affected. The DGGE results pointing to a change incommunity structure are also consistent with both toxic and selec-tive TCS effects. The findings of Drury et al. (2013) (8 �g g−1) andLubarsky et al. (2012) (2–100 �g l−1) also show that chronic TCSexposure resulted in changes in bacterial community composi-tion and a loss of diversity. Such changes may be due to selectiverecruitment or inhibition of sensitive species during biofilm devel-opment. In fact, Schreiber and Szewzyk (2008) reported that evennanogram per liter of antimicrobials significantly affected therecruitment of bacteria to the attached community. In contrast,Lubarsky et al. (2012) suggested that while surface recruitment wasnot affected by TCS, biofilm development was inhibited by increas-ing TCS levels. Morin et al. (2010a), utilized in situ incubations toexamine effects of pesticides used in vineyards and reported thatbiofilm dry weight, ash free dry weight, and Chl-a suggested thatbiofilms translocated to contaminant free sites recovered withintwo months. Diatom diversity indices however, suggested that aone month recovery period did not allow return to reference condi-tions. Although communities did not recover to reference conditionthey exhibited “recovery potential” (Morin et al., 2010a). Findingsreported by Morin et al. (2010b) studying effects of TCS were consis-tent with Dorigo et al. (unpublished data) who used PCR–DGGE tofind that neither eukaryotic nor prokaryotic community structurecould be restored after TCS exposure, within a 1–2 month recoveryperiod. Similarly, we found that a number of measures indicatedno recovery, even though changes in biomass, carbon utilizationand pigment composition were suggestive of recovery. This wasdespite the fact that in the current study, the exposure level wasmuch lower, chronic rather than acute, and the time scale for effectsand recovery longer than in many studies.

4.3. Response of the protozoa/micrometazoa

Although the effects of grazers on biofilm development areimportant, few studies have assessed impacts of stress on this com-munity compartment. Nietch et al. (2013) included “consumers”in their examination of TCS impacts on stream periphyton. In thisstudy effects on nematodes appeared to be related to reducedresource availability rather than direct TCS toxicity. They did how-ever, find that ostracods were significantly- and directly-affectedby TCS, and estimated a lowest observed effects level of 0.5 �g l−1

(Nietch et al., 2013). This observation is in keeping with a direct TCSeffect on Rotifera at 0.5 �g l−1 reported by Lawrence et al. (2009).In general, the NOEC TCS concentrations for invertebrates basedon single species testing are much greater than 4 �g l−1 (Lyndallet al., 2010). In the present study, micrometazoans (rotifers, nema-todes) appeared to be stimulated by TCS when added to establishedbiofilms (R) but not when added during initial biofilm development(T) – there appears to be no clear explanation for this effect given

that the toxicity of TCS to invertebrates occurs at high levels i.e.,40 �g l−1 (Reiss et al., 2002). Few assessments of TCS effects onprotozoan grazers have been carried out. Lawrence et al. (2009)reported that there were no detectable effects of TCS at 0.5, 5, and

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64 J.R. Lawrence et al. / Aquati

0 �g l−1 on Euplotes sp., Dileptus sp., Blepharisma sp., Stentor sp.,pirostomum sp., Euglena sp., and Paramecium sp. Further, no sig-ificant impacts on the protozoa or micrometazoa numbers were

ound when the community developed in the presence of 10 �g l−1

CS. Miyoshi et al. (2003) however, reported that TCS was toxico Paramecium trichium and Paramecium caudatum at concentra-ions of 2.58 �M (about 750 �g l−1) and 1.64 �M (about 476 �g l−1),espectively. In the current study there were clear negative effectsn protozoan grazers relative to the reference community. Theumber of protozoa in treatments receiving longer duration TCSxposure (6–8 week) suggests that tolerant populations may beecruited over time, in keeping with the no effects observations ofawrence et al. (2009). Due to their role(s) as ecosystem engineersJones et al., 1994), changes in protozoan and micrometazoan num-ers or types may have significant “knock-on” effects. Protozoa play

critical role in ecosystem processes, influencing carbon cycling asell as nutrient availability to higher trophic levels (Zöllner et al.,

009). Furthermore, they can be highly selective, thereby affectingommunity composition (Glucksman et al., 2010).

.4. Lack of recovery

The lack of recovery to reference conditions when exposure times increased has been attributed to a number of causes. For example,t may be linked to experimental design, length of time pre- orost- exposure, immigration processes, age of biofilm during thexposure, sensitivity of parameters measured (see below). It alsoepends on the definition of recovery: is it functional or structural?

n the literature lack of recovery has been linked with retentionf contaminants in the biofilm that prevents complete removal ofhe stress even though it is no longer present in the bulk liquidhase (Ivorra et al., 1999). For example, Arini et al. (2012) usedrtificial streams to assess biofilm recovery following Cd and Znxposure. After 56 days, full recovery was not observed, and lackf recovery was attributed in part, to retention of the metals inhe biofilm matrix. Although a possible factor in the current study,4C–triclosan studies have shown that only four to seven percentf the radioactive TCS was recovered sorbed to organic materialr retained by the biofilm (Lawrence et al., 2009) and similar tobservations were made during the current study. Therefore, thisactor is unlikely to have contributed to the lack of recovery in theurrent experiments.

.5. What measurements are best?

The application of a wide variety of measurements has gainedonsiderable acceptance for assessing microbial community struc-ure and activity. Generally, these methods may be divided intohree classes: (i) those based on estimates of biomass (ash free dryeight, dry weight, biovolume, and Chl-a), (ii) activity (respiration,

hymidine and leucine incorporation, photosynthesis, enzymes,nd carbon utilization), and (iii) structure-diversity (DGGE, nexteneration sequencing (NGS), pigments, and fatty acid methylsters (FAME)). Here, we have used several biomass measures,ncluding biovolume, biofilm dry weights, biofilm thickness, as wells specific biomass measurements for bacteria, cyanobacteria andlgae. While the general biomass measures appeared relativelynsensitive measures of effects and recovery, the specific mea-ures related to algal/cyanobacterial and bacterial biomass, wereevealing with regard to community response to TCS exposurend recovery. Arini et al. (2012) reported that diversity indices

nd biovolumes measured from biofilms under decontaminationonditions did show the beginnings of recovery. Activity measure-ents include thymidine incorporation and compound-specific

stimates based on labeled substrates, as well as commonly-used

ology 161 (2015) 253–266

Biolog Ecoplate. Although not applied here, monitoring of spe-cific enzymes has also been successfully applied. In the currentstudy, thymidine incorporation appeared sensitive to TCS effects,while carbon utilization spectra (Biolog) revealed shifts in theuse of specific substrates or classes. Measures of diversity maybe based on microscopic examination, as is commonly performedfor diatom assemblages where taxonomy, function and evidenceof stress may be interpreted or from chemical- (lipid, EPS, pig-ments, fatty acid methyl esters), molecular- or genomic-basedapproaches. Here, we utilized pigment analyses, fluorescent lectinbinding assays and DGGE, all of which were sensitive indicatorsof changes in algal and bacterial community structure. A com-bination of structural and functional information at population,community and ecosystem levels may provide realistic results ofthe effects of different disturbances on fluvial ecosystems, moredetailed analyses are necessary on the intrinsic mechanisms oper-ating in the biofilm for a better interpretation of response relativeto disturbance (Victoria and Gómez, 2010). From the perspectiveof ecosystem services, both functional and structural recovery arenecessary to ensure diversity, function and maintain resilience.In general it appears that higher resolution measurements basedon activity and diversity are the most sensitive measures. Indeedapplication of high-throughput DNA sequencing for taxonomicand transcriptomic analyses of community structure and activ-ity responses may be particularly revealing (Yergeau et al., 2010)and amenable for use as a part of resistance–resilience–recoverystudies.

5. Conclusions

We applied various measures to assess the impacts of time-variable exposures to the broad-spectrum antimicrobial, TCSin a microbial community. Community structure was alteredby any exposure to TCS; clear impacts were detected inalgal–cyanobacterial abundance and diversity, while similar effectswere seen for protozoan grazers. Biofilm bacteria were markedlyaffected in terms of diversity and metabolic activity (i.e., thymi-dine/carbon utilization). Results presented here indicate that evenrelatively brief exposures of a river biofilm community to low-levels of TCS alter both the trajectory and endpoint of development,resulting in significantly changed community structure and func-tion. Although we recognize that microbial communities may beunique products of their developmental history sensu Matthewset al. (1996) in this instance the final outcome is likely influencedby balance between selection of resistant populations when TCSis present during development and the dominance of sensitivespecies in those initially developing TCS free. While evidence wasobtained for recovery and adaptation by the community, a returnto reference conditions was not confirmed in any treatment. Thus,TCS presents a risk in aquatic ecosystems and may impact microbialcommunity structure and function in developing and establishedcommunities.

Acknowledgements

This research was supported by grants from Health Canada andEnvironment Canada. AP was supported by the Global Institute forWater Security, University of Saskatchewan.

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