APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Aug. 2009, p. 5261–5272 Vol. 75, No. 160099-2240/09/$08.00�0 doi:10.1128/AEM.00412-09Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Relative Diversity and Community Structure of Ciliates in StreamBiofilms According to Molecular and Microscopy Methods�†

Andrew Dopheide,1 Gavin Lear,1 Rebecca Stott,2 and Gillian Lewis1*School of Biological Sciences, University of Auckland, 3a Symonds Street, Auckland, New Zealand,1 and National Institute for

Water and Atmospheric Research, P.O. Box 11-115, Hamilton, New Zealand2

Received 18 February 2009/Accepted 18 June 2009

Ciliates are an important component of aquatic ecosystems, acting as predators of bacteria and protozoa andproviding nutrition for organisms at higher trophic levels. Understanding of the diversity and ecological roleof ciliates in stream biofilms is limited, however. Ciliate diversity in biofilm samples from four streams subjectto different impacts by human activity was assessed using microscopy and terminal restriction fragment lengthpolymorphism (T-RFLP) analysis of 18S rRNA sequences. Analysis of 3� and 5� terminal fragments yieldedvery similar estimates of ciliate diversity. The diversity detected using microscopy was consistently lower thanthat suggested by T-RFLP analysis, indicating the existence of genetic diversity not apparent by morphologicalexamination. Microscopy and T-RFLP analyses revealed similar relative trends in diversity between differentstreams, with the lowest level of biofilm-associated ciliate diversity found in samples from the least-impactedstream and the highest diversity in samples from moderately to highly impacted streams. Multivariate analysisprovided evidence of significantly different ciliate communities in biofilm samples from different streams andseasons, particularly between a highly degraded urban stream and less impacted streams. Microscopy andT-RFLP data both suggested the existence of widely distributed, resilient biofilm-associated ciliates as well asciliate taxa restricted to sites with particular environmental conditions, with cosmopolitan taxa being moreabundant than those with restricted distributions. Differences between ciliate assemblages were associated withwater quality characteristics typical of urban stream degradation and may be related to factors such asnutrient availability and macroinvertebrate communities. Microscopic and molecular techniques were consid-ered to be useful complementary approaches for investigation of biofilm ciliate communities.

Heterotrophic microeukaryotes such as ciliates are thoughtto be of considerable importance in aquatic ecosystems, as theyare major predators of bacteria and constitute a nutritionalresource for other protozoa, invertebrates, and probably fishlarvae (9, 22, 36, 52, 62, 63, 71). In addition, protozoan bacte-rivory contributes to enhanced decomposition of leaf detri-tus—a vital nutrient resource in streams—by increasing turn-over of bacterial populations through predation (57). It is notwell understood, however, how ciliate diversity and communitystructure in streams are affected by changing environmentalconditions, or how ciliate communities affect other streambiota and processes. The effects of various physical, chemical,and biological factors on freshwater protozoan communitieshave been considered by a number of studies, but most of thesehave focused upon planktonic organisms in lentic habitats (forexample, see references 2, 11, and 44). However, the complexmicrobial communities in biofilms have been recognized asimportant contributors to critical ecological processes, such asauxotrophic primary production, nitrogen fixation, and nutri-ent cycling, and may underpin the function of stream foodwebs (31, 45, 61). The few studies which have investigatedbenthic habitats in lotic systems have found evidence of the

existence of diverse communities of abundant ciliates (3, 20,56) and shifts in community structure in response to ecophysi-ological parameters (30, 42, 43). With one exception, however,these investigations were based on aquatic sediments, and theorganisms within epilithic biofilms have continued to receivelittle attention.

Most studies of ciliate diversity and ecology have utilizedmicroscopy-based methods of identification (for example, seereferences 3 and 56), as ciliate cells are relatively large andmorphologically diverse. Such methods demand a high level oftaxonomic expertise, however, and are difficult and time-con-suming—for example, many ciliates are fragile and fast mov-ing, and they often require difficult fixing and staining proto-cols for reliable identification. Molecular biological tools offerthe possibility of more accurate and efficient methods for pro-tozoan study and may provide a useful complement to tradi-tional approaches (12, 18, 28, 65), yet we know of only a fewmolecular studies of environmental ciliate diversity (18, 20,37). A series of recent investigations used culture-independentanalysis of 18S rRNA gene sequences to reveal the existence ofdiverse microeukaryote communities in assorted marine, an-oxic, and extreme environments (40, 48, 66, 69, 70, 72). Fur-thermore, a growing body of evidence suggests the existence ofsignificant genetic diversity among various ciliate taxa whichhas escaped detection by microscopy (14, 18, 23, 34, 60, 64, 78),pointing to the potential for molecular techniques to generatenew insights into ciliate diversity and ecology, and suggesting aneed for comparison of the effectiveness of these differenttechniques in environmental samples.

Terminal restriction fragment length polymorphism (T-RFLP)

* Corresponding author. Mailing address: School of Biological Sci-ences, University of Auckland, Private Bag 92019, Auckland, NewZealand. Phone: 64 (9) 373 7599. Fax: 64 (9) 373 7416. E-mail: gd.lewis@auckland.ac.nz.

† Supplemental material for this article may be found at http://aem.asm.org/.

� Published ahead of print on 26 June 2009.

5261

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

analysis provides an efficient, inexpensive, and semiquantita-tive means for comparing microbial molecular diversity be-tween different samples and has been widely used to investi-gate bacterial communities, although only a few studies haveapplied T-RFLP methods to the analysis of microeukaryotediversity (6, 16, 17). In this study, ciliate diversity and commu-nity structure were investigated in biofilm samples fromstreams representing a range of levels of anthropogenic deg-radation, with the objective of testing the null hypothesis thathuman impacts have no effect upon this important heterotro-phic component of stream ecosystems. To achieve this, ciliate-targeted PCR primers were used in conjunction with T-RFLPand multivariate statistical analyses. Additionally, ciliate diver-sity measures obtained using molecular techniques were com-pared with those derived from microscopy-based methods inorder to assess the relative effectiveness of these approaches.

MATERIALS AND METHODS

Sampling sites. Biofilm samples were collected from each of four differentlyimpacted streams in Auckland, New Zealand. Site 1 (Cascade Stream) is alargely unimpacted stream, located in an undeveloped native forest catchment(36°53�32� S, 174°31�07� E). Site 2 (Stoney Creek) is mildly impacted, located ina partially developed native forest catchment with nearby houses and roads(36°54�24� S, 174°34�06� E). Site 2 is a lower-order tributary of site 3 (OpanukuStream), which is proximate to rural agricultural development and is moderatelyimpacted (36°53�42� S, 174°35�44� E). Site 4 (Pakuranga Stream) is located in ahighly developed urban catchment (36°53�50� S, 174°54�21� E) and is highlyimpacted. Sites 1, 2, and 3 all have natural stony substrates, while site 4 consistsof a concrete channel at the sampling location.

Sites 1 and 3 are ranked as having the best and fifth-best water quality,respectively, of 25 streams throughout the Auckland region based on monthlymonitoring between 1995 and 2005; three locations in the site 4 stream catch-ment are ranked in the worst five (4, 5). Physical and chemical attributes of thestreams are presented in Table 1.

Sample collection. Quantitative methods for sampling biofilm material andassociated protozoa from submerged surfaces were developed. There are noclearly established protocols for sampling protozoa associated with epilithicbiofilms in lotic systems, and for this reason two methods were tested in thisstudy. For both methods, stream biofilm was collected from substrate surfaceswhile submerged, to avoid the potential loss of material upon removal of stones

from the water column (29). The first method involved the use of sterile Speci-Sponges (Nasco, Fort Atkinson, WI) to thoroughly swab submerged surfaces(rocks or concrete channel) within a 55-cm2 area defined by a circular neoprenetemplate. Dislodged biofilm material was then squeezed from the collectingsponges into sterile Whirl-Pak bags (Nasco).

The second biofilm collection method involved a syringe sampler based ondevices recommended for subsurface sampling of epilithic periphyton (1, 39, 54,67). The syringe sampler, illustrated in Fig. S1 in the supplemental material,consisted of a 60-ml syringe with its end removed to create a wide opening anda toothbrush head glued to the end of the syringe plunger. A rubber ring wasattached to the end of the syringe to seal the sampler against the rock surface andto minimize the loss of dislodged material due to water currents. Biofilm materialwas removed from a 4.91-cm2 area by pressing down and rotating the syringeplunger. Loosened material was drawn up into a 10-ml collection syringe at-tached to the base of the larger syringe with plastic tubing. Samples were thendecanted into sterile Whirl-Pak bags (Nasco).

Biofilm sampling was carried out during January (summer), May (autumn),August (winter), and November (spring) of 2005. On each sampling occasion,biofilm material was collected from two 20 m reaches of each stream. In general,the exposed surfaces of 4 to 10 randomly selected rocks (220 to 550 cm2 in total)were sampled using the sponge method, and the surfaces of 10 rocks (about 50cm2 in total) were sampled using the syringe method, from within each 20-mreach of each stream. Similarly, 10 samples were collected from each of two 20-mreaches along the concrete channel at site 4, using each sampling method. The 10samples obtained using each method at each sample point were combined, givinga total of four composite samples per stream (one sponge sample and one syringesample from each of two sampling points in each stream). Samples were chilledon ice for transport.

Assessment of ciliate diversity by microscopy-based analysis. Samples werestored at 4°C and analyzed within 4 to 10 hours of collection. For the enumer-ation of ciliates, subsamples of 1 ml were transferred to a Sedgewick Rafter celland scanned at a magnification of �25 to generate preliminary lists of taxa.Subsamples were then examined at magnifications of �200 to �630. Due to thelow density of biofilm material and associated ciliates, concentration of samplesfrom site 1 prior to examination was typically required, as follows: 25- to 100-mlsamples were concentrated by filtering through 25-�m nylon mesh and back-washing the retentate into a graduated 15-ml tube using filtered water (typicalfinal volume, �3 ml). Aliquots (1 ml) were then transferred to a SedgewickRafter cell and examined at magnifications of �200 to �630. Ciliate cells wereidentified to at least genus level, where possible, using criteria described intaxonomic keys (25, 53). Photographs were used to ensure that identificationswere consistent. The relative abundance of different taxa was scored on a scaleof 1 to 8, corresponding to approximate abundances of 1 to 5, 5 to 10, 10 to 15,15 to 20, 20 to 50, 50 to 100, 100 to 200, and over 200 cells per ml, respectively.

TABLE 1. Physical and chemical characteristics of streams included in this study throughout 2005a

MeasurementResult for:

Site 1 Site 2e Site 3 Site 4

Catchment area (ha)b 233 375 2,652 275Land useb (%) (native, forestry, agriculture, urban) 100, 0, 0, 0 98.9, 0, 1.1, 0 55, 2, 25.4, 17.6 1, 0.3, 0, 98.7Stream widthc (m) 5.8 4.1 6.3 0.7Water depthc (cm) 11 (10–15) 20 (15–24) 23 (19–28) 13 (14–18)Water velocityc (ms�1) 0.31 (0.15–0.61) 0.46 (0.18–0.6) 0.71 (0.54–0.88) 0.4 (0.15–0.63)Temperaturec (°C) 13.6 (11.9–14.4) 13.8 (10.9–16.2) 14.2 (10.8–17.2) 18.1 (14.0–25.5)pHc 7.6 (7.2–7.9) 7.3 (7.0–7.6) 7.5 (7.2–7.6) 7.5 (7.3–7.8)Turbidityd (NTU) 3.8 (1.1–13.0) 7.7 (1.9–18.0) 11.9 (3.8–46.9)Dissolved oxygend (gm�3) 9.9 (8.8–11.1) 9.4 (7.3–10.8) 8.9 (5.8–12.3)Conductivityd (�S cm�2) 166.4 (134.7–187.9) 142.2 (124.2–168.1) 311.3 (208.9–411.3)Ammoniacal nitrogend (gm�3) 0.01 (0.01–0.02) 0.03 (0.01–0.05) 0.09 (0.03–0.18)Nitrate/nitrited (gm�3) 0.01 (0.00–0.03) 0.2 (0.01–0.83) 0.60 (0.13–1.63)Total Kjeldahl nitrogend (gm�3) 0.31 (0.21–0.93) 0.33 (0.21–1.14) 0.61 (0.20–1.80)Total nitrogend (gm�3) 0.22 (0.20–0.92) 0.41 (0.20–0.84) 0.96 (0.42–2.24)Dissolved reactive phosphorusd (gm�3) 0.021 (0.013–0.031) 0.016 (0.010–0.024) 0.021 (0.017–0.045)Total phosphorusd (gm�3) 0.032 (0.023–0.046) 0.045 (0.024–0.064) 0.100 (0.044–0.336)

a Values are means and ranges. NTU, nephelometric turbidity units.b Data were generated using the Land Cover database 2 (Terralink International Ltd., Wellington, New Zealand).c Data were recorded in this study.d Data are from reference 5 and are unavailable for site 2.e Site 2 is an upstream tributary of site 3. It is presumed that water quality at site 2 is the same as or better than that at site 3.

5262 DOPHEIDE ET AL. APPL. ENVIRON. MICROBIOL.

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

DNA extraction and PCR amplification. Subsamples (30 ml) of each combinedbiofilm sample were transferred to preweighed sterile 35-ml centrifuge tubes andcentrifuged at 6,000 � g for 10 min at 4°C. Supernatants were removed andpellets resuspended in 15% glycerol to achieve final concentrations of 100 to 200mg biofilm ml�1. Samples were then frozen at �80°C until required.

DNA was extracted from biofilm samples as previously described (20). Fol-lowing extraction, the concentration of DNA in each extract was assessed usinga Quant-iT PicoGreen double-stranded-DNA kit (Invitrogen, Auckland, NewZealand) according to the manufacturer’s directions, in combination with absor-bance measurements at 260 nm using a Nanodrop ND-1000 spectrophotometer(Thermo Fisher Scientific, Waltham, MA), and electrophoresis on 1% agarosegels stained with Sybr Safe (Invitrogen). Based on the combined results of theseprocedures, the concentration of DNA in extracts was standardized to approxi-mately 20 ng �l�1 before use as templates in PCRs.

PCR primers 384F (5�-YTB GAT GGT AGT GTA TTG GA-3�) (20) and1147R (5�-GAC GGT ATC TRA TCG TCT TT-3�) (20), targeting an �700-bpfragment of the ciliate 18S rRNA gene, were labeled at their 5� termini with6-carboxyhexachlorofluorescein (HEX) and 6-carboxyfluorescein (FAM) fluoro-phores (Invitrogen), respectively. DNA was amplified from biofilm extracts usingthese primers in 50-�l PCR mixes (25 �l GoTaq Green master mix [Promega, InVitro Technologies, Auckland, New Zealand], 0.5 �M forward and reverseprimers, 0.4% BSA [Invitrogen] and 2 �l of template DNA). The following PCRprotocol was used: initial incubation for 5 min at 94°C, then 30 amplificationcycles of 45 s at 94°C, 60 s at 55°C and 90 s at 72°C, followed by a final extensionstep of 7 min at 72°C. PCR products were purified using a Purelink PCRpurification kit (Invitrogen) according to the manufacturer’s instructions. Theconcentration of fluorescently labeled PCR products was determined by absor-bance measurements at 260 nm using a Nanodrop ND-1000 spectrophotometer.

T-RFLP analysis. T-RFLP analysis is a semiquantitative molecular finger-printing technique which provides an efficient method of comparing populations.Fluorescently labeled PCR products are digested with one or more restrictionenzymes, resulting in the production of fluorescently labeled terminal fragments,the length (in base pairs) and abundance of which can be automatically detected.This results in the generation of profiles in which the number of peaks indicatesthe number of different terminal fragments present, while the height and area ofpeaks indicate their relative abundance. As terminal fragment length variesacross taxa, these data can provide a profile of community structure within eachsample.

For T-RFLP analysis in this study, the DNA concentration in each purifiedPCR product was adjusted to 20 ng �l�1. PCR products were digested with therestriction endonucleases HaeIII and RsaI (Invitrogen) in 10-�l reaction mix-tures, incubated overnight at 37°C. Each digestion reaction mixture contained 1U of each enzyme, 1 �l of reaction buffer (Invitrogen), and approximately 175 ngof purified amplicon. Digested samples were electrophoresed alongside a sizestandard with markers at 20-bp intervals up to 1,200 bp (LIZ1200; AppliedBiosystems, Melbourne, Australia). Terminal restriction fragments were de-tected using a 3130XL genetic analyzer (Applied Biosystems). This resulted ingeneration of peak profiles representing the abundance of HEX- and FAM-labeled terminal fragments, which were analyzed using GeneMapper 4.0 (Ap-plied Biosystems), which automatically calculates the number, height, and area ofpeaks and their corresponding fragment lengths (in base pairs). Profiles from tworuns of each sample were compared to check for consistency, and any inconsis-tent results were discarded. T-RFLP peaks with a height of �50 relative fluo-rescence units and a length of less than 10 bp were excluded from analysis toeliminate background interference. Each remaining peak was presumed to rep-resent a different terminal restriction fragment and hence a different ciliate-derived 18S rRNA gene sequence. Peaks representing terminal fragments inexcess of about 650 bp in length were assumed to represent PCR products whichwere not cut during restriction digestion.

The length (in base pairs) and area of HEX- and FAM-labeled peaks in eachT-RFLP profile were imported into Microsoft Excel. Peak positions wererounded to the nearest whole number, and the overall area of each profile wasstandardized to 1, to ensure comparability between samples.

Statistical analyses. Analysis of variance (ANOVA) was used to test forsignificant differences among numbers of ciliate taxa detected in samples fromdifferent streams in each month. Significant ANOVA results were further ana-lyzed with post-hoc Tukey-Kramer honestly significant difference multiple com-parison tests. Differences between numbers of taxa in samples obtained using thetwo different sampling methods and according to the two different analysismethods were investigated using t tests. These analyses were carried out in JMP7.0 (SAS Institute Inc., Cary, NC). As each different PCR product can beexpected to produce both a HEX-labeled fragment (primer 384F) and a FAM-labeled fragment (primer 1147R), the numbers of HEX-labeled and FAM-

labeled T-RFLP peaks were averaged to provide a composite diversity estimatefor each sample.

For multivariate analysis, data from both HEX-labeled and FAM-labeledT-RFLP peaks were combined into a single data set for each sample and sub-jected to a square-root transformation to moderate the influence of large peaksin subsequent analyses. Relative abundance data obtained by microscopywere not transformed. Multivariate analyses were carried out in Primer v6.1.6(Primer-E Ltd., Plymouth, United Kingdom). Bray-Curtis similarity between allpairs of biofilm samples was calculated. Similarities and differences among ciliatecommunities were visualyzed by nonmetric multidimensional scaling (MDS),which clusters samples with higher levels of pairwise similarity more closely thansamples with lower pairwise similarity. Analysis of similarities (ANOSIM) wasused to test the null hypotheses of no significant differences between ciliatecommunities from different streams, seasons, and sampling methods. ANOSIMcompares within-group similarity and between-group similarity; R values around0 indicate that within-group and between-group similarities are the same, whileR values approaching 1 indicate that samples within groups are more similar toeach other than to samples from different groups, allowing the null hypothesis tobe rejected (15).

RESULTS

Sampling methods. The effectiveness of sponge- and sy-ringe-based biofilm sampling methods for detecting ciliateswas compared, but there was no clear evidence of greaterefficacy of either method in terms of ciliate diversity detected,according to either microscopy or T-RFLP. Little evidence forsignificant differences in ciliate community structure was foundbetween sponge-derived and syringe-derived samples accord-ing to ANOSIM (Table 2), and furthermore, sponge- and sy-ringe-derived samples from the same stream and sampling datewere typically grouped very closely in MDS plots, suggestingvery similar composition of the communities sampled by eachmethod. Sponge- and syringe-derived samples from each siteand date were therefore pooled for subsequent analyses.

Ciliate diversity in stream biofilm samples (microscopy andT-RFLP methods). Our methodology targeted a standard areaof substrate for all samples, and our diversity results representthe number of taxa detected per area sampled. However, bio-film biomass was particularly sparse at site 1, especially inwinter (G. Lewis and S. Tsai, unpublished data). Similarly, thedensity of ciliates at site 1 was generally very low, necessitatingconcentration of these samples for analysis. Even after con-centration, the number of ciliate cells detected and identifiedin site 1 samples was low, and this may have affected the levelof diversity detected.

The number of ciliate taxa detected in stream biofilm sam-ples using microscopy-based methods ranged from 0, for sam-

TABLE 2. Comparison of ciliate diversity in sponge and syringesamples according to microscopy and T-RFLP

analysis (ANOSIM)

Month of sponge-syringesample comparison

Differenceaccording

to microscopy

Differenceaccording

to T-RFLP

Global Rstatistic P Global R

statistic P

Januarya 0.438 0.07May 0.031 0.48 �0.106 0.67August 0.167 0.44 �0.086 0.59Novembera �0.313 1.00

a Molecular analysis of January sponge and November syringe samples wasunsuccessful.

VOL. 75, 2009 DIVERSITY OF CILIATES IN STREAM BIOFILMS 5263

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

ples from site 1 in May and August, to 17, for a site 4 samplefrom May (Fig. 1). Site 1 typically had the fewest biofilm-associated ciliate taxa, while samples from site 3 and site 4contained the highest numbers of ciliate taxa. Significant dif-ferences were detected among microscopy results from Mayand November (ANOVA; P � 0.05) but not from January orAugust.

The number of different peaks present in a T-RFLP profileis assumed to reflect the number of different ciliate 18S rRNAgene sequences—and therefore the number of ciliate taxa—present in the stream biofilm sample. The number of T-RFLPpeaks detected ranged from 5 (site 1, November) to 61 (site 3,January) (Fig. 1). Significant differences were detected in allmonths (ANOVA; P � 0.005). The pattern of ciliate diversityacross different streams according to T-RFLP analysis wasbroadly similar to that derived from microscopic investigations,with the lowest ciliate diversity being detected in the mostpristine stream and higher diversity at the more impacted sites.

Overall, in each stream the average number of peaks de-

tected in T-RFLP profiles exceeded the average number ofciliate taxa detected by microscopy (t test; P � 0.0001) (Fig. 2).In total, 183 different HEX-labeled terminal fragments and191 different FAM-labeled terminal fragments were detectedamong all samples, compared with 68 different ciliate taxaidentified by microscopy.

Differences in ciliate diversity and community structurebetween streams and seasons according to microscopy andT-RFLP. Microscopy-based analysis of biofilm samples foundevidence of ciliate taxa common to multiple stream environ-ments as well as many taxa restricted to individual sites (Fig. 3).Taxa common to all four streams were detected only in Janu-ary and November, although these taxa were always present inat least one stream throughout the year. In January, a greaternumber of unique taxa were found in site 2 than in site 3. In allother months, taxa unique to site 3 and site 4 together ac-counted for the majority of taxa detected, although these twostreams also had generally higher overall levels of diversitythan the other sites. Taxa unique to site 4 were particularly

FIG. 1. Numbers of ciliate taxa detected by microscopy and numbers of ciliate 18S rRNA gene sequences indicated by T-RFLP analysis ofstream biofilm samples. The numbers on the x axis denote samples from two different reaches within each stream. Each bar shows the mean ofcounts from two samples (microscopy data) or the mean of HEX-labeled and FAM-labeled terminal fragment counts from two samples (T-RFLPdata). Error bars show one standard deviation. Significant differences were found between streams during each month according to T-RFLPanalysis and in May and November according to microscopy data (ANOVA; P � 0.05). Within each month, samples not linked by the same letter(A to C) are significantly different (Tukey-Kramer honestly significant difference; P 0.05). �, samples from site 1 were typically concentrated dueto low biomass and ciliate abundance; there was insufficient biofilm biomass at site 1 in August for T-RFLP analysis.

5264 DOPHEIDE ET AL. APPL. ENVIRON. MICROBIOL.

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

frequent in August. In November, taxa unique to site 2 werenot detected, and taxa unique to site 1 were evident only inMay. Fewer than one in five of the taxa unique to particularstreams were detected on more than one sampling date.

Overall, only 7% of ciliate taxa identified throughout theyear using microscopy were common to all four streams (Fig.3). Forty-four percent of the detected taxa were each found inonly one stream, most commonly site 3 or site 4, and no taxawere unique to the least-impacted stream, site 1.

According to microscopy, ciliates common to all four streamswere generally small species from Oligohymenophorea orPhyllopharyngea, such as Glaucoma spp., Trochilia spp., orCyclidium spp. These ubiquitous species were generally moreabundant in the more impacted streams. Several unidentifiedhypotrichs were typically characteristic of site 2 biofilm sam-ples, while taxa unique to site 3 included Actinobolina spp.,Aspidisca lynceus, and sessile peritrich species such as Epistylisspp. and Vorticella spp. A large number of taxa were found onlyat site 4, including Spirostomum spp., Stylonychia spp., Euplotesspp., Strombilidium spp. and predatory taxa, including Mono-dinium spp. and several species of Litonotus.

Visual inspection of T-RFLP profiles shows that some peaksare present in profiles from multiple streams (although thesepeaks are often markedly different in magnitude), while otherpeaks appear to be unique to particular stream biofilms (Fig. 4).Overall, 17% of the different T-RFLP peaks detected throughoutthe year were found in all four streams. Forty-five percent of theT-RFLP peaks occurred in only one stream (Fig. 3), consistentwith the microscopy data. The proportion unique to each streamranged from 5% (site 1) to 16% (site 3).

The proportions of T-RFLP peaks found in different streamsshowed a higher degree of consistency between months thanthe microscopy data (Fig. 3). Peaks unique to each of the fourstreams were detected in all months except August, when bio-film biomass at site 1 was insufficient for molecular analysis.Compared with the microscopy data, the proportion of T-RFLP peaks unique to site 3 and site 4 together accounted forless of the total diversity detected, except for in January sam-ples. Peaks unique to site 1 were more frequently detected,however, as were peaks unique to site 2 in November samples.As for the microscopy data, T-RFLP peaks unique to site 4were most frequent in August samples, although the propor-tion of peaks unique to site 3 was highest in January.

The T-RFLP peaks found in all four streams throughout theyear (17% of all peaks) together accounted for 75% of totalprofile area, indicating that the corresponding terminal frag-ments were relatively abundant (Fig. 5). Conversely, the peaksthat were each detected in only one stream (45% of all peaks)accounted for less than 6% of the total T-RFLP profile area,and as for the microscopy data, few of these unique T-RFLPpeaks were detected on more than one sampling date. These

FIG. 2. Average numbers of ciliate taxa detected by microscopy(white bars) and average numbers of ciliate 18S rRNA gene sequencesindicated by T-RFLP analysis (gray bars) in stream biofilm samplesthroughout 1 year. Each bar shows the mean of counts from eightsamples (microscopy data) or the mean of HEX-labeled and FAM-labeled terminal fragment counts from eight samples (T-RFLP data).Error bars show one standard deviation. The average number of T-RFLP peaks detected significantly exceeded the average number oftaxa identified by microscopy in all cases (t test; P � 0.0001).

FIG. 3. Comparison of diversities of ciliate taxa occurring in biofilms from different stream environments, according to microscopy and T-RFLPanalysis. �, there were no T-RFLP data for site 1 in August due to insufficient biofilm biomass for analysis.

VOL. 75, 2009 DIVERSITY OF CILIATES IN STREAM BIOFILMS 5265

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

findings suggest the existence of populations of abundant andcosmopolitan ciliate taxa in stream biofilms from different en-vironments, together with rarer taxa, which had low abundanceand restricted spatial and temporal distributions.

Microscopy relative abundance data and T-RFLP peak areadata were used to generate nonmetric MDS plots in whicheach data point represents the assemblage of ciliate taxa orT-RFLP peaks detected in one sample. The proximity of the

FIG. 4. T-RFLP profiles derived from stream biofilm samples in November 2005 using ciliate-targeted PCR primers. Sizes of gray and blackpeaks, respectively, indicate the abundance of HEX-labeled and FAM-labeled terminal restriction fragments.

FIG. 5. Abundance and diversity of ciliates in stream biofilms according to T-RFLP profiles. Bars represent the combined area of T-RFLPpeaks found in profiles from individual streams only and peaks found in profiles from multiple streams, as a proportion of total profile area foreach sampling date. Numbers of peaks contributing to each bar are indicated. �, there were no T-RFLP data from site 1 for August.

5266 DOPHEIDE ET AL. APPL. ENVIRON. MICROBIOL.

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

data points to each other reflects the relative similarity of theirciliate assemblages. MDS plots based on microscopy data showseparation of samples from highly impacted site 4 and moder-ately impacted site 3 from each other and from samples fromless impacted site 2 and site 1 (Fig. 6). Similarly, MDS plotsbased on T-RFLP data show site 4 samples forming a clearlyseparated group, while site 3 and site 2 samples form overlappingclusters and site 1 samples are scattered widely. Temporal pat-terns are less clear, with little discernible grouping of data pointsby sampling month according to microscopy data. According toT-RFLP data, November samples and May samples from site 1are more dispersed than January and August samples.

ANOSIM analysis of microscopy and T-RFLP data providedevidence of significant differences between ciliate communitiesin different streams and between samples from different seasons(Table 3). For the microscopy data, significant and generally largedifferences were found between ciliate assemblages from allstreams, and moderate differences between assemblages fromdifferent sampling dates. Similarly, significant differences werefound between ciliate assemblages from all streams accordingto T-RFLP analysis, with the exception of site 1 and site 2. Thelargest differences according to T-RFLP were found betweensite 4 and both site 2 and site 3, while the largest differencesaccording to microscopy were found between site 1 and site 3and between site 3 and site 4. The largest difference betweensampling months according to T-RFLP analysis was betweenJanuary and August, but these months showed relatively littledifference according to microscopy. Significant differenceswere not detected between January and May or between Jan-uary and November T-RFLP results.

Links between environmental data and ciliate assemblageMDS data. Links between ciliate community assemblage dataand environmental trends can be visualized as bubbles overlaidon MDS ordination plots, with the size of bubbles representing

the magnitude of environmental parameters at sites and datescorresponding to biofilm sampling occasions. The resulting fig-ures suggest that observed MDS ordination patterns are associ-ated with a combination of environmental factors (Fig. 7). Sepa-ration of site 4 samples from others appears to be associated withfactors typically associated with urban stream degradation, suchas higher levels of nitrogenous compounds and lower levels ofdissolved oxygen, in addition to very low levels of forest cover.Turbidity, temperature, and phosphorus levels are similarly ele-vated at site 4 on certain sampling occasions, while water velocityand pH do not follow this trend. Conversely, site 1 samples areassociated with high native forest cover, low levels of nitrogenouscompounds, total phosphorus and turbidity, and elevated oxygen.Grouping of site 3 samples in a cluster adjacent to site 4 samplesappears to be related to intermediate levels of nitrogenous com-pounds and total phosphorus, although pH and levels of dissolvedreactive phosphorus are generally lower than for site 1 samples.

DISCUSSION

Ciliate diversity according to microscopy and T-RFLP anal-ysis. Ciliates have been considered very amenable to micro-scopic study due to their high level of morphological diversityand relatively large size. However, it has been suggested that acurrent list of described ciliate morphospecies may contain 5 to10 times as many biological species (13, 24, 26). Furthermore,ciliate morphospecies have been shown to include organismswith clearly different ecophysiological characteristics (75). Thissuggests that the morphospecies concept may substantially un-derestimate ciliate species diversity and ecosystem complexity(26, 75). Morphologically identical but genetically, physiolog-ically, or biochemically divergent ciliates are likely to occupyseparate ecological niches, and measurement of this functionalciliate diversity is relevant to ecological studies. While molec-

FIG. 6. MDS grouping of ciliate assemblages in stream biofilm samples based on microscopy data (left; two-dimensional stress 0.15) andT-RFLP profiles (right; two-dimensional stress 0.19).

VOL. 75, 2009 DIVERSITY OF CILIATES IN STREAM BIOFILMS 5267

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

ular techniques have recently contributed to great insights intoprotistan diversity in various inaccessible and extreme environ-ments, few studies have applied these techniques to specificphyla, such as Ciliophora.

This investigation found that the number of ciliate taxa sug-gested by T-RFLP analysis was more than double that indi-cated by microscopy. This suggests the existence of a significantcomponent of genetic ciliate diversity in stream biofilms whichis not evident upon microscopic examination of morphology. Asimilar finding was made in a recent study of oligotrich ciliatediversity in seawater, with diversity according to molecularanalysis being about 10-fold higher than that according tomorphological observations (18). In the present study, ciliateswere identified using simple light microscopy methods, andalthough most species can be detected at relatively low mag-nification, it is possible that some organisms may have beenoverlooked due to small size or inconspicuousness, or due totheir being present in encysted form. Silver staining proceduresand electron microscopy can improve taxonomic discrimina-tion of ciliate taxa based on morphological and morphometri-

cal analysis. However, even when these more complex methodshave been used, ciliate diversity based on molecular analysishas still been found to exceed diversity according to morphol-ogy (34). A growing number of studies provide evidence ofcryptic molecular diversity exceeding apparent morphologicaldiversity in various ciliate taxa, including Carchesium (78),Cyclidium (23), Halteria (34), Oxytricha (60), Strombidium (34),Tetrahymena (41, 49, 64), and Zoothamnium (14). The consis-tency of these findings suggests little reason to expect anydifference in the great majority of ciliate taxa which have notyet been subjected to genetic analysis. Cyclidium and rRNAgene sequences closely matching those of Oxytricha, Tetrahy-mena, and Zoothamnium have all previously been detected inthese Auckland streams (20) and therefore may have contrib-uted to the cryptic genetic diversity detected using T-RFLP inthis study.

Although our molecular analysis indicates a high level ofgenetic diversity underlying ciliate morphospecies, the PCRprimers used in this study, while highly ciliate specific, are notperfectly so (20). It is possible that a limited number of non-ciliate sequences and resulting terminal restriction fragmentsmay be represented in our results. However, this effect may beoutweighed by the tendency of T-RFLP analysis to underesti-mate diversity of closely related taxa, due to conservation ofrestriction sites and consequent generation of terminal frag-ments of identical length (16). Additionally, ciliate taxa withindistinguishable 18S rRNA gene sequences may be discrimi-nated by examination of other genes (41), suggesting that thesequences targeted in this study may not provide completeresolution of different species. It thus seems likely that theassessments of ciliate diversity provided by T-RFLP analyses inthis study are conservative.

Limitations and complementarity of methods. Although T-RFLP analysis is an efficient method of obtaining and compar-ing microbial genetic diversity data, interpretation of T-RFLPinformation, in isolation, remains challenging. T-RFLP analy-sis lacks a straightforward means of reliably assigning taxo-nomic identities to observed peaks in complex samples, partic-ularly for groups of organisms for which availability of DNAsequence data is limited, such as ciliates. The value of diversitymeasures derived from T-RFLP analyses has been questioned,on the bases that different organisms may contribute to singleT-RFLP peaks, different restriction enzymes will produce dif-ferent results, and the use of thresholds to eliminate back-ground noise from T-RFLP profiles means that terminal frag-ments (and organisms) of low abundance will be excluded fromthe resulting analysis (10). Multivariate statistical analysis ofT-RFLP data is considered reliable, however, with conclusionsbeing little affected by the exclusion of minor T-RFLP peaks orthe choice of restriction enzyme (8, 77). T-RFLP is thus auseful method for comparing complex microbial communitystructures.

Microscopy-based analysis of morphology does permit iden-tification of ciliates, subject to sufficient taxonomic expertisebeing available, and can allow the classification of ciliates intofunctional categories, such as feeding groups, which can beused to examine the ecological role of protozoa (55). The levelof taxonomic resolution used can affect whether significantdifferences between protozoan communities will be detected,with identification of protozoa to taxonomic levels higher than

TABLE 3. ANOSIM comparison of ciliate assemblages in samplesfrom different streams and sampling dates according to

microscopy and T-RFLP data

Comparison R Pa

Stream comparisonMicroscopy datab

Global 0.719 0.001*Site 1-Site 2 0.414 0.001*Site 1-Site 3 0.988 0.001*Site 1-Site 4 0.798 0.001*Site 2-Site 3 0.748 0.001*Site 2-Site 4 0.621 0.001*Site 3-Site 4 0.842 0.001*

T-RFLP datac

Global 0.49 0.001*Site 1-Site 2 0.269 0.102Site 1-Site 3 0.497 0.01*Site 1-Site 4 0.508 0.006*Site 2-Site 3 0.332 0.037*Site 2-Site 4 0.769 0.002*Site 3-Site 4 0.678 0.001*

Sampling date comparisonMicroscopy datab

Global 0.383 0.001*January-May 0.487 0.002*January-August 0.284 0.011*January-November 0.469 0.001*May-August 0.257 0.011*May-November 0.428 0.001*August-November 0.418 0.001*

T-RFLP datac

Global 0.395 0.001*January-May 0.201 0.105January-August 0.722 0.002*January-November 0.125 0.222May-August 0.502 0.001*May-November 0.433 0.006*August-November 0.376 0.020*

a �, statistically significant result (P � 0.05).b n 13 to 16 samples per stream or month.c n 8 to 12 samples per stream and 8 to 15 samples per month.

5268 DOPHEIDE ET AL. APPL. ENVIRON. MICROBIOL.

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

genus being less effective in discriminating surface-associatedprotozoan communities (35). Microscopy does have the advan-tage of quantitative power—cells can be counted, albeittediously—which may be lacking in PCR-based assays. Clearly,morphological identification is possible only within the con-straints of the morphospecies concept, which has recognizedlimitations (26).

In this study, both T-RFLP and microscopy-based analysesshowed broadly similar overall trends of diversity in the differ-ent streams, and both methods produced evidence of signifi-cant differences between ciliate communities in differently im-pacted stream biofilms. It seems, therefore, that T-RFLP andmicroscopic analyses may be considered complementary meth-ods, the former providing a robust and efficient method forcomparing ciliate community structure, and the latter allowingattribution of differences between microbial assemblagesand systems to particular ciliate taxa or functional groups. Ofcourse, group-targeted PCR primers such as those used in thisstudy can be used for cloning and sequencing, which—if se-

quence and morphological data have been reconciled—doesallow reliable identification of microbes in environmental sam-ples, thus avoiding one of the limitations of T-RFLP. Fluores-cence in situ hybridization-based techniques offer a usefulmeans of linking molecular sequence data with microscopy-derived morphological information (68).

Further studies combining molecular and microscopic meth-ods are necessary for the expansion of currently limited se-quence database coverage of microeukaryotes (21). Assumingthat the availability of microeukaryote DNA sequence infor-mation will improve, it seems probable that in future, molec-ular identification methods may prove more straightforwardand accurate than morphology-based methods. However, it islikely that combined approaches may prove more informativethan either molecular or microscopy-based techniques alone(68, 73). Molecular profiling methods allow efficient and robustcomparisons of community structure, while identification anddescription of taxa using sequencing, microscopy, and fluores-cence in situ hybridization-based techniques can provide addi-

FIG. 7. MDS grouping of ciliate assemblages in stream biofilm samples, based on T-RFLP analysis, showing magnitude of various environ-mental parameters associated with samples (two-dimensional stress 0.19). Bubble sizes are scaled to reflect the ranges of values indicated inTable 1. Samples for which environmental data were unavailable were omitted.

VOL. 75, 2009 DIVERSITY OF CILIATES IN STREAM BIOFILMS 5269

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

tional insights and links to ecological, phenotypic, and physio-logical information and may allow pinpointing of ecologicallyimportant organisms.

Differences between biofilm ciliate communities in differentstreams. Both microscopy and T-RFLP analysis methods haveprovided clear evidence of differences in ciliate assemblages inbiofilms from differently impacted streams. These differencescan be associated with environmental parameters typical ofurban stream degradation, suggesting that our null hypothesis,that human impacts have no effect upon biofilm-associatedciliate communities, can be rejected.

The fewest ciliate taxa were found at site 1, a relativelypristine stream, while the most ciliate taxa were detected insamples from moderately impacted site 3 and highly impactedsite 4. This trend of greater diversity at the more impacted sitesseems contrary to the generally accepted tendency for ecolog-ical perturbation to lead to a simplification of communitystructure. A number of contrasting biotic and abiotic factorssuggest possible reasons for this finding. Site 1 and site 2 arecharacterized by little exposure to sunlight or anthropogenicpollutants. Site 3 and site 4, in contrast, are exposed to elevatednitrogen loads, derived from nearby areas of agricultural andurban land use, respectively. Reduced density and height ofriparian vegetation expose site 2 and site 3 to more sunlightthan site 1, and site 4 receives virtually no shade whatsoever.Nutrient enrichment and sunlight have been shown to promoteperiphyton growth in lotic systems (27, 32, 74), suggesting thatthe more impacted sites are likely to have communities ofmore abundant phototrophic organisms than the less impactedstreams. This is consistent with observations of increased bio-film biomass and more diverse bacterial and algal (especiallydiatom) communities at sites 3 and 4 (Lewis and Tsai, unpub-lished). The three less impacted sites in this study also receivesignificant amounts of allochthonous debris from surroundingvegetation, while site 4 does not, suggesting a difference in therelative importance of detritus-derived nutrients in thesestreams. Elevated nutrient availability can increase benthicciliate abundance in rivers and streams (19, 50, 58) and mayaffect ciliate abundance, biomass, and community compositionin lentic habitats (2, 33, 51, 76). It seems possible that theabundant biofilms in the more impacted streams in this studymay provide more resources and a wider variety of feedingniches for heterotrophic protozoan organisms. The greatervariety and abundance of bacterivorous, algivorous, and pred-atory ciliates detected in samples from the two more impactedsites is consistent with this suggestion.

Site 1 is home to diverse and abundant benthic macroinver-tebrates, while site 4 has a macrobenthic invertebrate fauna ofvery low diversity, consisting almost entirely of chironomidlarvae (38). Biofilms at site 1 may therefore be subjected tovery different grazing pressures than biofilms at site 4, which islikely to further affect the nutrient resources available in thesestreams. Macroinvertebrates may also negatively affect proto-zoa through predation (51, 71). In addition, macroinverte-brates consume meiofauna, such as rotifers (59), which mayalso predate upon protozoa (47). Studies of the effects oftop-down predation pressures on ciliates in lakes and pondshave had mixed results (2, 51, 76). There is very little informa-tion available on the nature of trophic interactions betweenciliates and invertebrates in stream biofilms, although one

study found evidence of invertebrate predation and/or compe-tition negatively affecting biofilm-associated ciliates (46). Nev-ertheless, it can be speculated that the homogeneity of theinvertebrate community at site 4 may mean that ciliates of onlycertain types and sizes are subjected to predation pressures,resulting in the selective proliferation of nontarget taxa. Incontrast, the diverse invertebrates at site 1 may represent abroad competitive and predatory factor, perhaps contributingto the lower abundance and diversity of ciliates at this site.

Different catchment land uses may cause development ofdifferent biofilm-associated ciliate assemblages by favoring tol-erant taxa while eliminating sensitive organisms. Being sur-rounded by extensive urban development, site 4 is probablyexposed to various pollutants in addition to increased levels ofnitrogenous compounds and lower levels of dissolved oxygen.Furthermore, the artificial substrate in site 4 may lack refugesfor flow-sensitive or light-sensitive organisms. A previous in-vestigation suggested that site 4 may be home to fewer sessileperitrich taxa and a higher frequency of predatory ciliates suchas Litonotus spp. and Loxophyllum spp. than the other streamsinvestigated in this study (20). Similarly, in this study, preda-tory ciliates such as Monodinium spp. and Litonotus spp. weredetected by microscopy-based analysis only in the site 4 bio-film. These, and other characteristic taxa identified in site 4,are typically found in -� mesosaprobic or polysaprobic wa-ters, indicating that they can tolerate reasonably high organicloads and hence more heavily polluted environments (25). Thissuggests that physicochemical conditions in site 4 influence thedevelopment of a very different ciliate community than those inthe less impacted streams included in this study. How thisdifferent ciliate community affects the ecological processes andinteractions occurring in this stream awaits further investiga-tion.

Conclusion. Ciliates and other protozoa are major predatorsof bacteria and provide an important trophic link in aquatichabitats, such as stream biofilms (52). Understanding proto-zoan community diversity and abundance is therefore impor-tant for gaining insights into the function of these hot spots ofmicrobial activity, which contribute substantially to ecosystemprocesses in streams (7). Both molecular and microscopy-based analyses provided evidence of diverse biofilm-associatedciliate communities, with greater diversity being seen in themore impacted streams and with significant differences be-tween ciliate assemblages in streams in different states of deg-radation being observed. These observations may be related toa variety of differences in environmental parameters charac-teristic of urban stream degradation, such as elevated nutrientand sunlight availability, as well as different assemblages ofautotrophic biofilm organisms and communities of benthicmacroinvertebrates. The discrepancy between numbers of taxasuggested by T-RFLP analysis and those obtained by micros-copy-based analysis in this study adds to evidence that ciliatediversity has been underestimated by traditional microscopicapproaches. Microscopic analysis allowed identification of cil-iate taxa characteristic of different stream environments, how-ever. Future application of these complementary techniqueswill improve our understanding of the causes and effects ofstream degradation at the microbial level, leading to develop-ment of more effective stream monitoring and remediationstrategies.

5270 DOPHEIDE ET AL. APPL. ENVIRON. MICROBIOL.

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

ACKNOWLEDGMENTS

We thank the anonymous reviewers for their helpful contributions tothe manuscript.

Funding for this research was provided by the New Zealand Foun-dation for Research, Science and Technology Public Good ScienceFund UOA306.

REFERENCES

1. Aloi, J. E. 1990. A critical review of recent freshwater periphyton fieldmethods. Can. J. Fish. Aquat. Sci. 47:656–670.

2. Andrushchyshyn, O. P., A. K. Magnusson, and D. D. Williams. 2006. Re-sponses of intermittent pond ciliate populations and communities to in situbottom-up and top-down manipulations. Aquat. Microb. Ecol. 42:293–310.

3. Andrushchyshyn, O. P., K. P. Wilson, and D. D. Williams. 2007. Ciliatecommunities in shallow groundwater: seasonal and spatial characteristics.Freshwater Biol. 52:1745–1761.

4. Auckland Regional Council. 2007. River water quality: state and trends inthe Auckland region. Auckland Regional Council technical publication 336.Auckland Regional Council, Auckland, New Zealand.

5. Auckland Regional Council. 2007. State of the environment monitoring:rivers & streams water quality data report 2005. Auckland Regional Counciltechnical publication 327. Auckland Regional Council, Auckland, New Zea-land.

6. Baldwin, A. J., J. A. Moss, J. D. Pakulski, P. Catala, F. Joux, and W. H.Jeffrey. 2005. Microbial diversity in a Pacific Ocean transect from the Arcticto Antarctic circles. Aquat. Microb. Ecol. 41:91–102.

7. Battin, T. J., L. A. Kaplan, J. D. Newbold, and C. M. E. Hansen. 2003.Contributions of microbial biofilms to ecosystem processes in stream meso-cosms. Nature 426:439–442.

8. Bennett, L. T., S. Kasel, and J. Tibbits. 2008. Non-parametric multivariatecomparisons of soil fungal composition: sensitivity to thresholds and indica-tions of structural redundancy in T-RFLP data. Soil Biol. Biochem. 40:1601–1611.

9. Berninger, U. G., B. J. Finlay, and P. Kuuppo-Leinikki. 1991. Protozoancontrol of bacterial abundances in freshwater. Limnol. Oceanogr. 36:139–147.

10. Blackwood, C. B., D. Hudleston, D. R. Zak, and J. S. Buyer. 2007. Interpret-ing ecological diversity indices applied to terminal restriction fragmentlength polymorphism data: Insights from simulated microbial communities.Appl. Environ. Microbiol. 73:5276–5283.

11. Burns, C. W., and L. M. Galbraith. 2007. Relating planktonic microbial foodweb structure in lentic freshwater ecosystems to water quality and land use.J. Plankton Res. 29:127–139.

12. Caron, D. A., P. D. Countway, and M. V. Brown. 2004. The growing contri-bution of molecular biology and immunology to protistan ecology: molecularsignatures as ecological tools. J. Euk. Microbiol. 51:38–48.

13. Chao, A., P. C. Li, S. Agatha, and W. Foissner. 2006. A statistical approachto estimate soil ciliate diversity and distribution based on data from fivecontinents. Oikos 114:479–493.

14. Clamp, J. C., and D. Williams. 2006. A molecular phylogenetic investigationof Zoothamnium (Ciliophora, Peritrichia, Sessilida). J. Euk. Microbiol. 53:494–498.

15. Clarke, K. R. 1993. Non-parametric multivariate analyses of changes incommunity structure. Aust. J. Ecol. 18:117–143.

16. Countway, P. D., R. J. Gast, P. Savai, and D. A. Caron. 2005. Protistandiversity estimates based on 18S rDNA from seawater incubations in theWestern North Atlantic. J. Euk. Microbiol. 52:95–106.

17. Díez, B., C. Pedros-Alio, T. L. Marsh, and R. Massana. 2001. Application ofdenaturing gradient gel electrophoresis (DGGE) to study the diversity ofmarine picoeukaryotic assemblages and comparison of DGGE with othermolecular techniques. Appl. Environ. Microbiol. 67:2942–2951.

18. Doherty, M., B. A. Costas, G. B. McManus, and L. A. Katz. 2007. Culture-independent assessment of planktonic ciliate diversity in coastal northwestAtlantic waters. Aquat. Microb. Ecol. 48:141–154.

19. Domenech, R., A. Gaudes, J. C. Lopez-Doval, H. Salvado, and I. Munoz.2006. Effects of short-term nutrient addition on microfauna density in aMediterranean stream. Hydrobiologia 568:207–215.

20. Dopheide, A., G. Lear, R. Stott, and G. Lewis. 2008. Molecular character-ization of ciliate diversity in stream biofilms. Appl. Environ. Microbiol.74:1740–1747.

21. Duff, R. J., H. Ball, and P. J. Lavrentyev. 2008. Application of combinedmorphological-molecular approaches to the identification of planktonic pro-tists from environmental samples. J. Euk. Microbiol. 55:306–312.

22. Finlay, B. J., and G. F. Esteban. 1998. Freshwater protozoa: biodiversity andecological function. Biodiversity Conservation 7:1163–1186.

23. Finlay, B. J., G. F. Esteban, S. Brown, T. Fenchell, and K. Hoef-Emden.2006. Multiple cosmopolitan ecotypes within a microbial eukaryote mor-phospecies. Protist 157:377–390.

24. Foissner, W. 2006. Biogeography and dispersal of micro-organisms: a reviewemphasizing protists. Acta Protozool. 45:111–136.

25. Foissner, W., and H. Berger. 1996. A user-friendly guide to the ciliates(Protozoa, Ciliophora) commonly used by hydrobiologists as bioindicators inrivers, lakes, and waste waters, with notes on their ecology. Freshw. Biol.35:375–482.

26. Foissner, W., A. Chao, and L. A. Katz. 2008. Diversity and geographicdistribution of ciliates (Protista: Ciliophora). Biodiversity Conservation 17:345–363.

27. Francoeur, S. N. 2001. Meta-analysis of lotic nutrient amendment experi-ments: detecting and quantifying subtle responses. J. N. Am. Benthol. Soc.20:358–368.

28. Fried, J., R. Psenner, W. Ludwig, and K. H. Schleifer. 2002. Improvement ofciliate identification and quantification: a new protocol for fluorescence insitu hybridization (FISH) in combination with silver stain techniques. Syst.Appl. Microbiol. 25:555–571.

29. Fukuda, M., A. Ashida, Y. Tomaru, and S. I. Nakano. 2004. An improvedmethod for collecting heterotrophic microorganisms living on pebbles instreams. Limnology 5:41–46.

30. Gucker, B., and H. Fischer. 2003. Flagellate and ciliate distribution in sed-iments of a lowland river: relationships with environmental gradients andbacteria. Aquat. Microb. Ecol. 31:67–76.

31. Hall, R. O., Jr., and J. L. Meyer. 1998. The trophic significance of bacteriain a detritus-based stream food web. Ecol. Lett. 79:1995–2012.

32. Hillebrand, H. 2002. Top-down versus bottom-up control of autotrophicbiomass—a meta-analysis on experiments with periphyton. J. N. Am.Benthol. Soc. 21:349–369.

33. Hillebrand, H., M. Kahlert, A. L. Haglund, U. G. Berninger, S. Nagel, andS. Wickham. 2002. Control of microbenthic communities by grazing andnutrient supply. Ecology 83:2205–2219.

34. Katz, L. A., G. B. McManus, O. L. O. Snoeyenbos-West, A. Griffin, K. Pirog,B. Costas, and W. Foissner. 2005. Reframing the ‘everything is everywhere’debate: evidence for high gene flow and diversity in ciliate morphospecies.Aquat. Microb. Ecol. 41:55–65.

35. Kepner, R. L., Jr., and J. R. Pratt. 1996. Characterization of surface-asso-ciated protozoan communities in a Lake Erie coastal wetland (Old WomanCreek, Ohio). J. Great Lakes Res. 22:63–76.

36. Kneitel, J. M., and J. M. Chase. 2004. Disturbance, predator, and resourceinteractions alter container community composition. Ecology 85:2088–2093.

37. Lara, E., C. Berney, H. Harms, and A. Chatzinotas. 2007. Cultivation-independent analysis reveals a shift in ciliate 18S rRNA gene diversity in apolycyclic aromatic hydrocarbon-polluted soil. FEMS Microbiol. Ecol. 62:365–373.

38. Lear, G., I. K. G. Boothroyd, S. J. Turner, K. Roberts, and G. D. Lewis. 2009.A comparison of bacteria and benthic invertebrates as indicators of ecolog-ical health in streams. Freshw. Biol. 54:1532–1543.

39. Loeb, S. L. 1981. An in situ method for measuring the primary productivityand standing crop of the epilithic periphyton community in lentic systems.Limnol. Oceanogr. 26:394–399.

40. Lopez-García, P., F. Rodríguez-Valera, C. Pedros-Alio, and D. Moreira.2001. Unexpected diversity of small eukaryotes in deep-sea Antarctic plank-ton. Nature 409:603–607.

41. Lynn, D. H., and M. C. Struder-Kypke. 2006. Species of Tetrahymena iden-tical by small subunit rRNA gene sequences are discriminated by mitochon-drial cytochrome c oxidase I gene sequences. J. Euk. Microbiol. 53:385–387.

42. Madoni, P., and N. Bassanini. 1999. Longitudinal changes in the ciliatedprotozoa communities along a fluvial system polluted by organic matter. Eur.J. Protistol. 35:391–402.

43. Madoni, P., and S. Braghiroli. 2007. Changes in the ciliate assemblage alonga fluvial system related to physical, chemical and geomorphological charac-teristics. Eur. J. Protistol. 43:67–75.

44. Madoni, P., and F. Sartore. 2003. Long-term changes in the structure ofciliate communities in a small isolated pond. Italian J. Zool. 70:313–320.

45. Marxsen, J. 2006. Bacterial production in the carbon flow of a centralEuropean stream, the Breitenbach. Freshw. Biol. 51:1838–1861.

46. McCormick, P. V. 1991. Lotic protistan herbivore selectivity and its potentialimpact on benthic algal assemblages. J. N. Am. Benthol. Soc. 10:238–250.

47. McCormick, P. V., and J. Cairns, Jr. 1991. Effects of micrometazoa on theprotistan assemblage of a littoral food web. Freshw. Biol. 26:111–119.

48. Moon-van der Staay, S. Y., R. De Wachter, and D. Vaulot. 2001. Oceanic 18SrDNA sequences from picoplankton reveal unsuspected eukaryotic diversity.Nature 409:607–610.

49. Nanney, D. L., C. Park, R. Preparata, and E. M. Simon. 1998. Comparisonof sequence differences in a variable 23S rRNA domain among sets of crypticspecies of ciliated protozoa. J. Euk. Microbiol. 45:91–100.

50. Norf, H., H. Arndt, and M. Weitere. 2009. Responses of biofilm-dwellingciliate communities to planktonic and benthic resource enrichment. Microb.Ecol. 57:687–700.

51. Pace, M. L., and E. Funke. 1991. Regulation of planktonic microbial com-munities by nutrients and herbivores. Ecology 72:904–914.

52. Parry, J. D. 2004. Protozoan grazing of freshwater biofilms. Adv. Appl.Microbiol. 54:167–196.

53. Patterson, D. J. 1998. Free-living freshwater protozoa. Manson PublishingLtd., London, United Kingdom.

VOL. 75, 2009 DIVERSITY OF CILIATES IN STREAM BIOFILMS 5271

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from

54. Persson, J., H. Hansson, and I. Renberg. 2006. The Stone Brusher, a newsampler for submerged epilithic material in shallow streams and lakes. Hy-drobiologia 560:385–392.

55. Pratt, J. R., and J. Cairns, Jr. 1985. Functional groups in the protozoa: rolesin differing ecosystems. J. Protozool. 32:415–423.

56. Reiss, J., and J. M. Schmid-Araya. 2008. Existing in plenty: abundance,biomass and diversity of ciliates and meiofauna in small streams. Freshw.Biol. 53:652–668.

57. Ribblett, S. G., M. A. Palmer, and D. W. Coats. 2005. The importance ofbacterivorous protists in the decomposition of stream leaf litter. FreshwaterBiol. 50:516–526.

58. Rublee, P. A., and A. Partusch-Talley. 1995. Microfaunal response to fertil-ization of an Arctic tundra stream. Freshw. Biol. 34:81–90.

59. Schmid-Araya, J. M., and P. E. Schmid. 2000. Trophic relationships: Inte-grating meiofauna into a realistic benthic food web. Freshw. Biol. 44:149–163.

60. Schmidt, S. L., D. Bernhard, M. Schlegel, and W. Foissner. 2007. Phylogenyof the Stichotrichia (Ciliophora; Spirotrichea) reconstructed with nuclearsmall subunit rRNA gene sequences: discrepancies and accordances withmorphological data. J. Euk. Microbiol. 54:201–209.

61. Sheldon, F., and K. F. Walker. 1997. Changes in biofilms induced by flowregulation could explain extinctions of aquatic snails in the lower RiverMurray, Australia. Hydrobiologia 347:97–108.

62. Sherr, E. B., and B. F. Sherr. 1994. Bacterivory and herbivory: key roles ofphagotrophic protists in pelagic food webs. Microb. Ecol. 28:223–235.

63. Sherr, E. B., and B. F. Sherr. 2002. Significance of predation by protists inaquatic microbial food webs. Antonie Leeuwenhoek Int. J. Gen. Mol. Mi-crobiol. 81:293–308.

64. Simon, E. M., D. L. Nanney, and F. P. Doerder. 2008. The “Tetrahymenapyriformis” complex of cryptic species. Biodiversity Conservation 17:365–380.

65. Sims, G. P., R. Aitken, and A. Rogerson. 2002. Identification and phyloge-netic analysis of morphologically similar naked amoebae using small subunitribosomal RNA. J. Euk. Microbiol. 49:478–484.

66. Slapeta, J., D. Moreira, and P. Lopez-Garcia. 2005. The extent of protistdiversity: Insights from molecular ecology of freshwater eukaryotes. Proc. R.Soc. Lond. Ser. B 272:2073–2081.

67. Stockner, J. G., and F. A. J. Armstrong. 1971. Periphyton of the experimentallakes area, northwestern Ontario. J. Fish. Res. Board Canada 28:215–229.

68. Stoeck, T., S. S. Epstein, and W. H. Fowle. 2003. Methodology of protistandiscovery: from rRNA detection to quality scanning electron microscopeimages. Appl. Environ. Microbiol. 69:6856–6863.

69. Stoeck, T., B. Hayward, S. S. Epstein, G. T. Taylor, and R. Varela. 2006. Amultiple PCR-primer approach to access the microeukaryotic diversity inenvironmental samples. Protist 157:31–43.

70. Stoeck, T., G. T. Taylor, and S. S. Epstein. 2003. Novel eukaryotes from thepermanently anoxic Cariaco Basin (Caribbean Sea). Appl. Environ. Micro-biol. 69:5656–5663.

71. Stoecker, D. K., and J. McDowell Capuzzo. 1990. Predation on protozoa: itsimportance to zooplankton. J. Plankton Res. 12:891–908.

72. Takishita, K., M. Tsuchiya, M. Kawato, K. Oguri, H. Kitazato, and T.Maruyama. 2007. Genetic diversity of microbial eukaryotes in anoxic sedi-ment of the saline meromictic Lake Namako-ike (Japan): on the detection ofanaerobic or anoxic-tolerant lineages of eukaryotes. Protist 158:51–64.

73. Vaerewijck, M. J. M., K. Sabbe, J. Bare, and K. Houf. 2008. Microscopic andmolecular studies of the diversity of free-living protozoa in meat-cuttingplants. Appl. Environ. Microbiol. 74:5741–5749.

74. Von Schiller, D., E. Martí, J. L. Riera, and F. Sabater. 2007. Effects ofnutrients and light on periphyton biomass and nitrogen uptake in Mediter-ranean streams with contrasting land uses. Freshw. Biol. 52:891–906.

75. Weisse, T., and S. Rammer. 2006. Pronounced ecophysiological clonal dif-ferences of two common freshwater ciliates, Coleps spetai (Prostomatida)and Rimostrombidium lacustris (Oligotrichida), challenge the morphospeciesconcept. J. Plankton Res. 28:55–63.

76. Wickham, S. A., S. Nagel, and H. Hillebrand. 2004. Control of epibenthicciliate communities by grazers and nutrients. Aquat. Microb. Ecol. 35:153–162.

77. Zhang, R., V. Thiyagarajan, and P. Y. Qian. 2008. Evaluation of terminal-restriction fragment length polymorphism analysis in contrasting marineenvironments. FEMS Microbiol. Ecol. 65:169–178.

78. Zhang, W. J., J. Yang, Y. H. Yu, S. W. Shu, and Y. F. Shen. 2006. Populationgenetic structure of Carchesium polypinum (Ciliophora: Peritrichia) in fourChinese lakes inferred from ISSR fingerprinting: high diversity but lowdifferentiation. J. Euk. Microbiol. 53:358–363.

5272 DOPHEIDE ET AL. APPL. ENVIRON. MICROBIOL.

on May 29, 2020 by guest

http://aem.asm

.org/D

ownloaded from