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A molecular survey of ciliates found in Shades Creek, Jefferson County, Alabama.

INTRODUCTION

Protists are important consumers in the freshwater food web (Heath et at, 2003; Marxsen, 2006), and the ciliates are key in this web as primary consumers of bacteria. However, published freshwater ciliate community data is limited In a survey using universal nuclear small subunit ribosomal DNA (SSU rDNA) primers and DNA sequencing, Slapeta et al. (2005) suggested that the diversity of freshwater eukaryotes, especially ciliates, may be much greater than previously described microscopically. In pools just south of Paris, France, they found that ciliates were the most abundant and diverse of the eukaryotes identified. Similar eukaryotic surveys (including ciliates) have been conducted on a lake in Bloomington, IN, USA (Dawson and Pace, 2002), a small river in Geneva, Switzerland (Berney et at, 2004), and a river in Spain (Amaral Zettler et al., 2002). Dopheide et al. (2008) describes the use of ciliate-specific SSU rDNA primers in the characterization of ciliates from four streams in Aukland, New Zealand. However, since ciliate survey data in the USA are still lacking and molecular methodology provides the best picture of community diversity (Slapeta et al., 2005), we describe in this report the beginning of the first detailed ciliate survey of a body of water in Alabama.

Eukaryotic surveys (including ciliates) have been conducted in various impaired environments including three sites cited above: anoxic sediments from a lake in Bloomington, IN, USA (Dawson and Pace, 2002), sediments from a small river in Geneva, Switzerland (Berney et al., 2004), and an acidic river in Spain (Amaral Zettler et al. 2002). Even within these degraded ecosystems, levels of diversity for protists remain high. Our study site, Shades Creek, is designated as an impaired body of water due to fecal coliform bacteria, sedimentation, and turbidity (USEPA, 2010). Many of the causes of impairment are related to urban land cover, and include storm sewers, construction, and impervious runoff (USEPA, 2010). Shades Creek's ecosystem instability due to the effects of urbanization makes it a stream of importance in understanding the effects of these factors on ciliate communities.

We present here the results of using various strategies that attempt to discover ciliates found in a freshwater source using two sampling techniques (tuna traps and raw-water sampling), concentration of organisms by centrifugation or filtration, then molecular biological identification. By DNA sequencing of the nuclear SSU rDNA and mitochondrial cytochrome C oxidase I (COX) gene, we identified the presence of 17 species of ciliates in upper Shades Creek and one of its small tributaries in Jefferson County, AL (Figure 1). Use of ciliate-specific primers rather than generic eukaryotic primers resulted in selection at the amplification step of ciliate DNA thus narrowing the search for these freshwater ciliates. In this report we also describe methodology for amplifying DNA from a single ciliate.

[FIGURE 1 OMITTED]

MATERIALS AND METHODS

Sample Collection

Samples were collected near Samford University from upper Shades Creek (a tributary of the Cahaba River) and one of its small tributaries that flows in a southeasterly direction through the Samford campus just east of Propst Hall. Sampling sites are indicated in Figure 1. This stretch of Shades Creek is rapidly flowing but with few rapids and has an average depth of approximately one meter. Water samples were taken from near the surface of very still pools or side waters. Tuna traps were placed at the bottom of similar shallow waters (less than approximately 20 cm deep).

Four sampling strategies, described in more detail below, were used: 1) setting tuna traps; 2) centrifugation of water samples; 3) filtration of water samples; 4) using unconcentrated raw creek water samples.

Tuna Traps. Tuna traps were constructed by placing about 4 cc of tuna in the bottom of a 50 mL disposable conical centrifuge tube and covering the opening with nylon hose material secured with a large rubber band. Traps were placed in shallow, still creek water and secured by burying the base in the soil or by covering it with rocks. Traps were collected after 2 to 4 days and examined immediately for the presence of ciliates and again after 1 to 3 days. Individual ciliates were isolated on sterile plastic petri dishes using a 10[micro]L mechanical pipettor under 30x magnification using a dissecting microscope. Cultures were established and DNA was isolated from these cultures as described below under "Ciliate Cultures." In Table 1, organisms collected by this method are indicated as "trap culture" in the column "Sampling."

Also, for DNA isolation from uncultured tuna trap water, organisms were concentrated by repeated centrifugations at 900 x g. The loose pellet was used for DNA isolation as described below. In Table 1, organisms identified by this method are indicated as "trap water" in the column "Sampling."

Centrifugation. For the second sampling technique, approximately 500 mL of water was collected from still creek water in a glass container that had been disinfected with 70% ethanol. The water sample was then heated to 70[degrees]C in a microwave to kill and immobilize organisms. Samples were then divided into twenty-four 15 mL conical centrifuge tubes and centrifuged at 6,000 x g for 4 min. The loose pellets from the 24 tubes were distributed into two 15 mL tubes and centrifuged at 6,000 x g for 4 min. One of these two loose pellets was pipetted into a 1.5 mL microfuge tube and centrifuged at 14,000 x g for 4 minutes, the supernatant discarded, then the other loose pellet pipetted into the same 1.5 mL tube and centrifuged at 14,000 x g for 4 minutes. The pellet was used for DNA isolation as described below. In Table 1, organisms identified by this method are indicated by "centri. creek" under the column "Sampling."

In some cases, tuna trap water was concentrated by centrifugation as described above.

Filtration. For the filtration method, approximately 250 mL of raw stream water was filtered through a sterile 0.22 [micro]m filter cartridge (Fisher) using a 50 mL disposable Leur-Lok syringe (Kendall), and the filtrate was discarded. The filter cartridge was opened by scoring around the perimeter where the upper and lower halves meet, then by tapping with a hammer. DNA was isolated from this filter as described below. In Table 1, organisms identified by this method are indicated as "filtration" under the column "Sampling."

Unconcentrated. In some cases, DNA was extracted directly from raw creek water. In Table 1, organisms identified by this method are indicated as "creek water" under the column "Sampling."

Ciliate Cultures

Cultures of ciliates were initiated by placing a single organism in rye tea. Rye tea was 3.5 g dry rye seed per L of water, which was boiled, cooled, then pH adjusted to 7.8-7.9 with [Na.sub.2]HP[O.sub.4]*7[H.sub.2]0. The tea was then autoclaved, cooled, adjusted to pH 6.8-7.2 with HC1, and then stored at 4[degrees]C. An aliquot of rye tea was inoculated with Klebsiella pneumoniae and cultured at room temperature overnight before use. Ciliate cultures were grown at room temperature or at 17[degrees]C in sterile 10 mL or 50 mL plastic centrifuge tubes with an initial volume of 3 to 5 mL rye tea at room temperature with the screw cap attached loosely. Cultures were fed weekly by adding about 2 mL room-temperature bacteria-inoculated rye tea. When tubes were near full, 3 to 5 mL were transferred to a new tube and the process repeated.

DNA isolation

DNA was isolated using one of the following two methods.

The Chelex Method. For the DNA isolation from a single ciliate or from 3 to 5 ciliates, a modification of the chelex method of Regensbogenova et aL (2004) was used. A single ciliate was isolated with a 10 [micro]L mechanical pipettor and washed in nuclease-free sterile water The ciliate was then transferred to 50 [micro]L of thoroughly mixed InstaGene Matrix (BioRad, 6% chelex solution), 5 [micro]L of a 49:1 dilution of LongLife Proteinase K (G-Biosciences) was added, tubes were gently mixed then incubated at 55[degrees]C for 30 min followed by heating at 98[degrees]C for 5 min. Tubes were placed on ice to cool then centrifuged at 3000 x g for 5 min. The supernatant was used immediately for PCR amplification.

In Table 1, organisms identified using this DNA isolation method are indicated as "chelex" in the column "DNA Isolation."

Wizard Genomic DNA Isolation Kit. The Wizard Genomic DNA Isolation Kit (Promega) was used as the other DNA isolation method. For isolation of DNA from tuna-trap water samples, the final supernatant described above "Centrifugation" was discarded, 600 [micro]L of Nuclei Lysis Solution (Promega) was added, and the Wizard Genomic DNA Isolation protocol was followed. In Table 1, organisms identified by this method are indicated by "Wizard" under the column "DNA Isolation." When the "Wizard" method was used for isolation of DNA from 0.22 pm filter cartridges, the filter was broken into several small pieces and placed in a 1.5 mL tube with 600 [micro]L Nuclei Lysis Solution, and the Wizard Genomic DNA Isolation protocol was followed.

PCR Amplification

DNA was amplified using the degenerate SSU rDNA (18S rDNA) primers 384F (5'-YTBGATGGTAGTGTATTGGA-3') and 1147R (5'-GACGGTATCTRATCGTCTTT-3')(Dopheide et al., 2008) or the COX primers CoxLl 1058 (5'-TGATTAGACTAGAGATGGC-3') and CoxH10176 (5'-GAAGTTTGTCAGTGTCTATCC-3')(Barth et al., 2006). These SSU primers were chosen because of their ciliate-specificity. We found that Paramecium DNA was not effectively amplified by these SSU rDNA primers, so the COX primers were also also used and proved more efficient at amplifying Paramecium DNA. PCR amplification was performed on either a MiniCycler (MJ Research) or a MiniOpticon System (BioRad) thermocycler using 33 cycles of 94[degrees]C denaturation, 55[degrees]C annealing, and 72[degrees]C replication (50[degrees]C annealing temperature was used for SSU rDNA amplification of P. multimicronucleatum). After confirmation of amplification by agarose gel electrophoresis, PCR products were cleaned up using Qiaquick spin columns (Qiagen).

Cloning and Sequencing

Cloning of PCR products was performed using the TOPO-TA pCR2.1 cloning kit (Invitrogen). PCR products were ligated into pCR 2.1 vectors and cloned into TOP-10 competent cells following the recommended Invitrogen protocol using ampicillin selection and X-gal blue/white screening. At least 10 white colonies were picked per ligation reaction. These cultures were grown in LB amp broth overnight at 35[degrees]C with shaking. In order to confirm the presence of the insert, 0.5 [micro]L, of each culture was used directly in a PCR amplification with the Ml3F(-21) and Ml3R vector primers. Agarose gel electrophoresis was used to confirm that the insert was of expected length. These cultures were streaked onto LB amp plates, which were sent to Genewiz, Plainfield, NJ for sequencing using the M13F(-21) and Ml3R vector primers. Alternatively, plasmids were isolated from transformed cells using the Quantum Prep[R] Plasmid Miniprep Kit (BioRad), and plasmids were sent to DF/HCC DNA Resource Core (Boston, MA) for sequencing.

DNA Sequence Analysis

Geneious Pro software (Drummond et at 2010) and the NCBI BLAST site (http://www.ncbi.nlm.nih.gov/BLAST/) were used for DNA sequence analysis and construction of phylogenetic trees. Alignments were performed using the Geneious Alignment tool with a 65% similarity cost matrix (5.0/4.0) and free end gaps. Neighbor Joining trees were constructed using the Tamura-Nei distance model and 1000 bootstrap replicates with a 50% support threshold.

RESULTS

We developed a protocol for the isolation of DNA from a single freshwater ciliate for PCR-amplification. The DNA isolation protocol, based on the method of Regensbogenova (2004), is summarized in Materials and Methods under "The Chelex Method."

Ciliate DNA was obtained by all collection methods and all DNA isolation procedures. The 17 ciliate species found, their collection sites, the collection method, and the DNA isolation method are indicated in Table 1. SSU rDNA sequences have been submitted to GenBank under the accession numbers shown in Table 1 and are also shown next to the sample name of each ciliate description below. All isolates were named according to the sites where organisms were collected (sites A, B, C, D, E, and F: see Figure 1). The descriptions below are listed by these assigned sample names. Table 1 lists them in order of the presumed species of each isolate. Following DNA isolation, either the COX gene or SSU rDNA was PCR-amplified, cloned, and sequenced.

1) Al (submitted to GenBank as JN232893). DNA of this ciliate was isolated using the Wizard method from centrifuged creek water samples. The SSU rDNA sequence showed 99.7% pairwise identity (616 out of 618 bases) with a published DNA sequence from Chilodonella uncinata (accession number AF300283) and 99.6% pairwise identity with another published C. uncinata sequence (AF300282).

2) A2, A3, A4, A5 (A2 submitted to GenBank as JN243999). DNA of this ciliate was isolated using the Wizard method from a centrifuged creek water sample. Three clones (A2, A3, and A4) yielded the same SSU rDNA sequence, which showed 98.6% pairwise identity (629/638 bases) with four different species of Tetrahymena (T tropicalis, EF070260; T. pyriformis, X56171, EF070254, EF070255, and M98021; T. silvana, EF070257; and T. setosa, AF364041) as well as one unidentified Tetrahymena species (EF070263). All changes were substitutions.One other clone, A5, differed from the other four by a single transition.

3) Cl, C2, C3, C4, C5, D1, D2, D3, D4, D5, E10, E12 (C1 and El2 submitted to GenBank as JN244000 and JN244004, respectively). DNA of all C site samples was isolated using the chelex method from a centrifuged pellet of tuna trap water. Four SSU rDNA clones from this sample (C1, C3, C4, C5) yielded an identical 637 base sequence. This sequence had 99.7% pairwise identity (637/639 bases) with a published DNA sequence from Colpidium campylum (X56532). The two differences are both single-base deletions in our sequence. One other clone from this site (C2) differed by one transition from the other four clones. Clones D2 and E10 were also identical to Cl, C3, C4, and C5, while D1, D3, D4, and D5 only varied by only one or two pyrimidine transitions. Although Ell also matched most closely to Colpidium campylum (X56532), it showed considerable divergence from the others (621/639 bases, 97.2% pairwise identity with several deletions) DNA of all D site samples was isolated from filtered creek water samples using the Wizard method. DNA of E10 and E12 was isolated from centrifuged tuna trap water using the Wizard method.

4) C6 (submitted to GenBank as JN244001). This large ciliate was collected from a tuna trap and has been cultured on rye grass medium. DNA was isolated from cultured C6 ciliates using the chelex method. Three clones of its COX genes yielded slightly different sequences Two clones of C6's amplified COX DNA showed 99.9% pairwise identity (766/767 bases), and one clone showed 99.7% pairwise identity (766/768 bases, 1 gap) with a published P. multimicronucleatum sequence (AM072769). The presence of multiple sequences in this P. multimicronucleatum culture is covered in the Discussion. SSU rDNA was also sequenced from this ciliate and yielded a sequence that most closely matched a published sequence from an unknown species of Paramecium (99.7% pairwise identity, 636/638 bases), but the next closest match was with P. multimicronucleatum (97.0% pairwise identity, 624/643 bases, 10 gaps). This ciliate is freely available by contacting D. A. Johnson (djohnso2@samford.edu).

5) D6 (submitted to GenBank as JN244002). DNA of this ciliate was isolated from a filtered water sample using the Wizard method. D6 SSU rDNA showed 96.1 % pairwise identity (589/613 bases, 3 gaps) with Strobilidium caudatum (AY143573).

6) El, E2, E3, E4. DNA of these ciliates was isolated from centrifuged creek water samples using the Wizard method. E2's amplified COX DNA showed 99.1% pairwise identity (838/846 bases) with a published Paramecium caudatum sequence (FJ905142). El, E3, and E4 all showed 98.9% pairwise identity (837/846) with FJ905142, but all three varied from each other by one substitution.

7) E6, E7, E8, E9 (E6 submitted to GenBank as JN244007). DNA of this ciliate was isolated from centrifuged creek water samples using the Wizard method. SSU rDNA from E6, E7, and E8 yielded an identical SSU rDNA sequence that showed 100% pairwise identity (638/638 bases) with three different species of Tetrahymena (T. lwoffi, EF070250; 7'. tropicalis, EF070259 and EF070261; and T. furgasoni, EF070247) and also matched perfectly two unidentified Tetrahymena species (AY755629 and EF070265). One other clone, E9, differed by a single base transition from the E6, E7, and E8.

8) Ell, E13 (Ell submitted to GenBank as JN244003). DNA of El 1 and E13 was isolated from centrifuged tuna trap water using the Wizard method. This ciliate's amplified SSU rDNA was identical to published sequences for three different ciliates: Hypotrichida sp. (AF508777), Pleurotricha lanceolata (AF508768 and AF164128), and Oxytricha sp. (AF164684) (643/643 bases). It is listed as "unknown ciliate" in Table 1.
Table 1, Ciliates identified from Shades Creek, Homewood, Alabama

Sample    Best BLAST Match      Sampling      DNA     Primers
                                 Method    Isolation

A1      Chilodonella         centr.      Wizard     SSU
        uncinata             creek                  rDNA

C1      Colpidium campylum   centr.      Wizard     SSU
                             creek                  rDNA

E12                          filtration  Wizard     SSU
                                                    rDNA

F1      Euplotes eurystomus  centr.      Wizard     SSU
                             creek                  rDNA

E11     Hypotrichida sp.     centr.      Wizard     SSU
                             tuna                   rDNA

        Pleurotricha
        lanceolata

        Oxytricha sp.

F2      Lembadion bullinum   centr.      Wizard     SSU
                             creek                  rDNA

F3      Levicoieps biwae     centr.      Wizard     SSU
                             creek                  rDNA

F6      Loxophyllum          centr.      Wizard     SSU
        spirellum            creek                  rDNA

E1      Paramecium caudatum  centr.      Wizard     COX
                             creek

C6      Paramecium           cultured    chelex     COX
        multimicronucleatum

                             cultured    chelex     SSU
                                                    rDNA

E17     Paramecium sp.       creek       Wizard     COX
                             water

E18     Paramecium           cultured    chelex     SSU
        tetraurelia                                 rDNA

D6      Strobiiidium         filtration  Wizard     SSU
        caudatum                                    rDNA

A2      Tetrahymena sp.      centr.      Wizard     SSU
                             creek                  rDNA

E6      Tetrahymena sp.      centr.      Wizard     SSU
                             creek                  rDNA

E14     unidentified         creek       Wizard     SSU
        ciliate              water                  rDNA

F9                           centr.      Wizard     SSU
                             creek                  rDNA

F7      unidentified         centr.      Wizard     SSU
        ciliate              creek                  rDNA

F4      Urocentrum turbo     centr.      Wizard     SSU
                             creek                  rDNA

Sample  Accession
            #

A1       JN232893

C1       JN244000

E12      JN244004

F1       JN244008

E11      JN244003

F2       JN244009

F3       JN244010

F6       JN244012

E1

C6

         JN244001

E17

E18      JN244006

D6       JN244002

A2       JN243999

E6       JN244007

E14      JN244005

F9       JN244014

F7       JN244013

F4       JN244011

"Sampling Methods" and "DNA Isolation" for ciliates are described
in the Materials and Methods. The COX gene or and SSU rDNA fragment
was amplified, cloned, and sequenced. Ciliates were identified by
comparison with published sequences using a BLAST search. "Sample"
name indicates the collection sites referred to those indicated in
Figure 1. The GenBank accession number is indicated for the SSU rDNA
sequences.


9) E14, E15, F9 (E14 and F9 submitted to GenBank as JN244005 and JN244014, respectively). El4 and EIS DNA was isolated from a creek water sample (no concentration of organisms by centrifugation or filtration) directly using the Wizard method. SSU rDNA of both most closely matched with unknown ciliates 97.3% pairwise identity with AJ810076 and AY642718 (622/639 bases, 3 gaps). The closest match with a known ciliate was with Glaucomides bromelicola (AJ810077, 97.0% pairwise identity, 620/639 bases, 3 gaps). F9 DNA was isolated from a centrifuged creek water sample using the Wizard method. Its SSU rDNA was nearly identical to AY642718 (99.8% pairwise identity, 637/638 bases), and its closest match with a known ciliate was with Glaucoma scintillans (AJ511861, 98.4% pairwise identity, 629/639 bases, 3 gaps).

10) E17. E 17 DNA was also isolated from a creek water sample (no concentration of organisms) using the Wizard method directly . El7 COX DNA showed 91.5% pairwise identity (773/845 bases, 6 gaps) with a published sequence for Paramecium caudatum (accession number FN424190). This ciliate appears to be a new species of Paramecium (see Discussion).

11) E18, E19, E20 (E18 submitted to GenBank as JN244006). This ciliate was cultured from tuna traps and has been maintained on rye grass medium. Its DNA was isolated by the chelex method, then its SSU rDNA amplified, cloned and sequenced. Two clones (E18, E19) had an identical 640 base sequence while E20 varied from these two by four separate transitions. El8 and E19's SSU rDNA had 99.8% pairwise identity (639/640 bases) with Paramecium tetraurelia (EF502045, AB252008, X03772, and AB252009).

12) F1 (submitted to GenBank as JN244008). DNA of this ciliate was isolated from centrifuged creek water samples using the Wizard method. Its SSU rDNA had 99.7% pairwise identity (718/720 bases, 1 gap) with two published sequences of Euplotes eurystomus (EF193250 and AJ310491).

13) F2 (submitted to GenBank as JN244009). DNA of this ciliate was isolated from centrifuged creek water samples using the Wizard method. Its SSU rDNA had 99.7% pairwise identity (637/639 bases) with Lembadion bullinum (AF255358). Our sequence varied from the published sequence by 2 transitions.

14) F3 (submitted to GenBank as JN244010). DNA of this ciliate was isolated from centrifuged creek water samples using the Wizard method. Its SSU rDNA had 99.0% pairwise identity (624/643 bases) with Levicoleps biwae (AB354737). Our sequence varied from the published sequence by 12 single base substitutions and 7 single base insertions or deletions.

15) F4, F5, F8 (F4 submitted to GenBank as JN244011). DNA was isolated from centrifuged creek water samples using the Wizard method. F4's SSU rDNA had 97.3% pairwise identity (637/639 bases) with Urocentrum turbo (AF255357) while F5 and F8 were identical to AF255357 (641/641 bases).

16) F6 (submitted to GenBank as JN244012). DNA of this ciliate was isolated from centrifuged creek water samples using the Wizard method. Its SSU rDNA had 99.7% pairwise identity (571/573 bases) with Loxophyllum spirellum (GU574810). This cloned rDNA fragment was considerably shorter than that of our other sequenced rDNA fragments (573 bases versus around 640 bases for the rest). Loxophyllum spirellum has been identified from a marine source (Pan et al., 2010), and the next eight closest matches on the NCBI BLAST site were also of marine origin. However, the tenth closest match (96.2% pairwise identity) is an uncultured ciliate identified in a freshwater constructed wetlands environment (Haentzsch et al., 2010).

17) F7 (submitted to GenBank as JN244013). DNA of this ciliate was isolated from a centrifuged creek water sample using the Wizard method. SSU rDNA for F7 most closely matched with unknown ciliates: FN689996 and FN689997 (96.3% pairwise identity, 554/575 bases). The closest match with a known ciliate was with Phialina salinarum (EU242508, 96.0% pairwise identity, 620/639 bases, 3 gaps).

SSU rDNA Phylogenetic Tree

Fifteen unique ciliate sequences were isolated and compared using Geneious Pro software. Published sequences were included in the tree to better illustrate interclass relationships. Eutreptia viridis (AJ532395, Eukaryota; Euglenozoa; Euglenida, Eutreptiales; Eutrptia) was used as the outgroup.

As shown in Figure 3, the 15 ciliates fall into the classes Phyllopharyngea, Litostomatea, Spirotrichea, Prostomatea, and Oligohymenophorea. Bootstrap values for inner branches are variable and support tends to increase in intermediate and outer nodes. Classes Phyllopharyngea, Litostomatea, Spirotrichea, and Prostomatea appear to be monophyletic based on their appearance and their support values, which are all above 70%. The class Oligohymenophorea does not seem to be monophyletic based on its appearance. Other publications support the notion that Oligohymenophorea may not be monophyletic (Snoeyenbos-West et al., 2004; Struder-Kypke et al., 2010).

[FIGURE 3 OMITTED]

The two clades that Oligohymenophorea occupies appear to be more specific groupings of this class. The upper Glade contains sequences from the order Peniculida. The lower Glade contains sequences from the order Hymenosyomatida, with the exception of F4, which looks to be Urocentrum turbo based on its closest match from the BLAST search results (AF255357; 99.8% pairwise identity, 100% query coverage). If this sequence is Urocentrum turbo, it should fall within the Peniculida order. It should be noted that this same species failed to group within the Oligohymenophorea class in a previous Ciliophora tree (Struder-Kypke et al., 2010). The most specific grouping within the lower Oligohymenophorea dade, with 83.2% support, appears to be the Tetrahymenia suborder based upon the taxonomy of the published sequences and all identifiable BLAST matches from our sequence data

Deeper nodes, which denote class relationships, have low support values. However, there is high support (100%) for the classes Litostomatea, Spirotrichea, Prostomatea, and Oligohymenophorea being more closely related to one another than to Phyllopharyngea. Additionally, classes Litostomatea and Spirotrichea group together with low support (55.7%); however, this relationship is suggested by several publications (Riley et al., 2001; Snoeyenbos-West el a!., 2004). Further, classes Prostomatea and Oligohymenophorea appear to be more closely related to one another than to the other classes, although the support for this grouping is once again low (5 8.8%). This relationship is also supported in several publications (Katz, 2001; Riley et aL, 2001; Snoeyenbos-West et al., 2004).

COX DNA Phylogenetic Tree

A phylogenetic tree was built with Geneious Pro software comparing all GenBank published COX P. caudatum sequences with sequences El and El7 (Figure 3). Miamiensis avidus (EU83 1213, Eukaryota; Alveolata; Ciliophora; Intramacronucleata; Oligohymenophorea; Scuticociliatia; Philasterida; Philasteridae; Miamiensis) was used as the outgroup. Tree topology for the COX DNA tree (Figure 3) is inferred to be reliable based upon the high level of branch support. All internal nodes have 70% support or above, and lower values are associated with terminal nodes. Additionally, it is important to note that the COX gene has the best resolution when comparing sequences at the genus or species level (Struder-Kypke, 2010). Thus, we can be confident that the presented relationships do not distort actual evolutionary relationships since all sequences are within the same species, with the exception of El7, which was most closely related to P. caudatum and is within the Paramecium genus.

The branching pattern shows that species El7 and one P. caudatum species, FN256283, do not assemble with the major P. caudatum grouping (Figure 3). Each species has a long branch, indicating a high degree of divergence from the rest of P. caudatum. Within the main grouping of P. caudatum, there are two clades, both of which are well supported (100% and 98.4%). Terminal nodes seem to have lower support values, which can be accounted for by the lack of variation between most P. caudatum sequences, which include many identical genotypes.

DISCUSSION

The ciliate sequences observed in this preliminary survey of upper Shades Creek indicate that we have sampled 17 different ciliate species, based upon the closest BLAST matches. Table 1 indicates that of the strategies used in this report, concentration of creek water samples followed by DNA extraction by the Wizard method was most successful.

Due to the large amount of variation between ciliate species (Struder-Kypke et al., 2010), obtaining a phylogenetic tree with high interclass support values is unlikely, even when using the well-conserved SSU rDNA gene. Although our interclass support values were low, many relationships were identified that correlate with published Ciliophora phylogenetic relationships. Most importantly, our tree allows for the observation of the relative positioning of each of our isolates with published sequences, including several sequences whose closest BLAST match was an unidentified ciliate (E11, E14, F7, F9).

Sequences E14 and F9 grouped within the lower Oligohymenophorea Glade with species that belong to the suborder Tetrahymenia, suggesting that they belong to this suborder. BLAST results suggest they may both be in the genus Glaucoma, which is supported by their grouping as shown in Figure 2. It is possible that the identities of these Tetrahymenia sequences could be reconciled by constructing a tree with published sequences from the suborder. Sequence F7 grouped within the Litostomatea Glade, suggesting that it belongs within that class. Sequence Ell was identical to several published species that fall within the class Spirotrichea, as supported by the tree.

Our observation of ciliates present in tuna traps that were allowed to age for several days indicates that, not surprisingly, this "tuna-water medium" was selective. (It is also very pungent.) That is, after several days, certain ciliates, especially a very small one and a large one resembling P. multimicronucleatum, were predominant. In fact, the culture of P. multimicronucleatum that we have maintained since September of 2009 was derived from aged tuna water. This suggests that the strategy of aging tuna traps several days might be a successful method for isolating P. multimicronucleatum.

Ciliate DNA isolated from the E17 uncultured ciliate had only 91.5% pairwise identity with any published known species (P. caudatum, FN424190). It might be assumed from this BLAST match that E17 is Paramecium but not P. caudatum. A phylogenetic tree was built with Geneious Pro software comparing all GenBank published COX P. caudatum sequences (Figure 3). The COX DNA tree shows that all of these ciliates identified as P. caudatum group together, with the exception of species FN256283, and our isolate, E17.

We have little information on how species FN256283 came to be classified as Paramecium caudatum. After running a BLAST search on this species, we discovered that its closest match showed 93.2% pairwise identity and 100% query coverage with a published P. caudatum species (accession number: FN256270). The results from this BLAST search align with the phylogenetic relationships presented in Figure 3, suggesting that species FN256283 is more closely related to the P. caudatum species than sequence El7 due to its shorter branch length and higher BLAST score. However, FN256283 appears to be more distantly related to the P. caudatum species than any other published GenBank P. caudatum COX sequence.

Additionally, the COX DNA tree contains another species we isolated, El. This uncultured ciliate showed 98.9% identity and 100% query coverage with a published P. caudatum sequence (accession number: FJ905142). El did fall within the main P. caudatum Glade in the tree.

In this project, our goal was to identify as many unique ciliates as possible. The PCR and cloning methodology employed here has the disadvantage of producing multiple clones from a single organism. Since our primary objective was to determine which species are present in these communities, this was not seen as a major problem. Also, single nucleotide differences between two strains could be due to errors during PCR amplification rather than naturally occurring diversity. However, our experience has been that Genewiz colony sequencing yields high quality, repeatable reads in the 600 to 700 by range. Therefore, when major divergence was found, we assumed it to be significant. Consequently, we believe that the COX divergence seen in C6 indicates the presence of different haplotypes in this monoculture. This raises the possibility of heteroplasmy in P. multimicronucleatum.

Eight species described in this report were represented by only one sequenced isolate and one species by only two sequenced isolates, indicating that the ciliates identified here may be an incomplete picture of the ciliate communities present in our samples. We plan to continue this investigation of these communities and would expect the list of ciliate species present in Shades Creek to grow significantly. Future studies will include other techniques, including PCR-Denaturing Gradient Gel Electrophoresis (PCR-DGGE)(van Hannen, 1998) and possibly metagenomic sequencing (using 454 pyrosequencing)(Margulies et aL., 2005), which promise to provide a more complete picture of the ciliate species comprising these freshwater communities.

ACKNOWLEDGEMENTS

We would like to thank Mary Gaines Walker, who isolated our P. multimicronucleatum strain, and Katie King for their help in this project. This research was funded by the Department of Biological and Environmental Sciences at Samford University, by a Samford Summer Undergraduate Research Fund (SURF) grant to J. Van Ausdall, and by a summer research grant from Samford's Clark Scholars Program in Computational Biology to K. Caffy.

LITERATURE CITED

Amaral Zettler, L., Gomez, F., Zettler, E., Keenan, B., Amils, R., and Sogin, M. 2002. Microbiology: eukaryotic diversity in Spain's River of Fire. Nature 417: 137.

Barth, D., Krenek, S., Fokin, S. I., and Berendonk, T. U. 2006. Intraspecific genetic variation in Paramecium revealed by mitochondrial cytochrome c oxidase I sequences. Journal of Eukaryotic Microbiology 53:20-25.

Dawson, S. and Pace, N. 2002. Novel kingdom-level eukaryotic diversity in anoxic environments. Proceedings of the National Academy of Sciences of the United States of America 99: 8324-8329.

Dopheide, A., Lear, G., Stott, R., and Lewis, G. 2008. Molecular characterization of ciliate diversity in stream biofilms. Applied and Environmental Microbiology. 74:1740-1747.

Drummond, A. J., Ashton, B., Buxton, S., Cheung, M., Cooper, A., Duran, C., Field, M., Heled, J., Kearse, M., Markowitz, S., Moir, R., Stones-Havas, S., Sturrock, S., Thierer, T., Wilson, A. 2010. Geneious v5.3, Available from http://www.geneious.com.

Haentzsch, M., Berendonk, T. U., Schlegel, M., and Bernhard, D. Molecular characterization of ciliate diversity within constructed wetlands revealed unexpected high genetic variation in rDNA sequences. Unpublished results, accession #FN826828, NCBI website (http://www.ncbi.nlm.nih.gov/).

Heath, R. T., Hwang, S. J., and Munawar, M.. 2003. A hypothesis for the assessment of the importance of microbial food web linkages in nearshore and offshore habitats of the Laurentian Great Lakes. Aquatic Ecosystems Health and Management. 6:231-239.

Katz, L. A. 2001. Evolution of nuclear dualism in ciliates: a reanalysis in light of recent molecular data. International Journal of Systematic and Evolutionary Microbiology. 51:1587-1592.

Margulies, M., Egholm, M., Altman, W. E., Attiya, S., Bader, J. S., Bemben, L. A., Berka, J., Braverman, M.S., Chen, Y. J., Chen, Z. T., Dewell, S. B., Du, L., Fierro, J. M., Gomes, X. V., Godwin, B. C., He, W., Helgesen, S., Ho, C. H., Irzyk, G. P., Jando, S. C., Alenquer, M. L. I., Jarvie, T. P., Jirage, K. B., Kim, J. B., Knight, J. R., Lanza, J. R., Leamon, J. H., Lefkowitz, S M., Lei, M., Li, J., Lohman, K. L., Lu, H., Makhijani, V. B., McDade, K. E., McKenna, M. P., Myers, E. W.. Nickerson, E., Nobile, J. R., Plant, R., Puc, B. P., Ronan, M. T., Roth, G. T., Sarkis, G. J., Simons, J. F., Simpson, J. W., Srinivasan, M., Tartaro, K. R., Tomasz, A., Vogt, K. A., Vollcmer, G. A., Wang, S. H., Wang, Y., Weiner, M. P., Yu, P. G., Begley, R. F. and Rothberg, J. M. 2005. Genome sequencing in microfabricated high-density picolitre reactors. Nature. 437:376-380.

Marxsen, J. 2006. Bacterial production in the carbon flow of a central European stream, the Breitenbach. Freshwater Biology. 51:1838-1861.

Pan, H., Gao, F., Li, J., Lin, X., Al-Farraj, S. A., and Al-Rasheid, K. A. 2010. Morphology and phylogeny of two new pleurostomatid ciliates, Epiphyllum shenzhenense n. sp. and Loxophyllum spirellum n. sp. (Protozoa, Ciliophora) from a mangrove wetland, South China. Journal of Eukaryotic Microbiology. 57: 421-428.

Regensbogenova, M., Kisidayova, S., Michalowski, T., Javorsky, P., Moon-Van Der Staay, S.Y., Moon-Van Der Staay, G. W. M., Hackstein, J. H. P., McEwan, N. R., Jouany, J. P., Newbold, C. J., Pristas, P. 2004. Rapid identification of rumen protozoa by restriction analysis of amplified SSU rRNA gene. Acta Protozoologica. 43: 219-224.

Riley, J. L. and L. A. Katz. 2001. Widespread distribution of extensive chromosomal fragmentation in ciliates. Molecular Biology and Evolution. 18:1372-1377.

Slapeta, J., Moreira, D, and Lopez-Garcia, P. 2005. The extent of protist diversity: insights from molecular ecology of freshwater eukaryotes. Proceedings of the Royal Society B 272: 2073-2081.

Snoeyenbos-West, 0. L. 0., J. Cole, A. Campbell, D. W. Coats, and L. A. Katz. 2004. Molecular phylogeny of Phyllopharyngean ciliates and their group I introns. Journal of Eukaryotic Microbiology. 51:441-450.

Strtider-Kypke, M. C., and D. H. Lynn. 2010. Comparative analysis of the mitochondrial cytochrome c oxidase subunit I (COI) gene in ciliates (Alveolata, Ciliophora) and evaluation of its suitability as a biodiversity marker. Systematics and Biodiversity 8:131-148.

U.S. Environmental Protection Agency. 2010 Waterbody Report for Shades Creek. Retrieved from http://iaspub.epa.govitmdl_waters10/attains_waterbody.

van Hannen, E.J., M.P. van Agterveld, H.J. Gons, and H.J. Laanbroek. 1998. Revealing genetic diversity of eukaryotic microorganisms in aquatic environments by denaturing gradient gel electrophoresis. Journal of Phycology. 34:206-213.

David A. Johnson, Kelsey Caffy, Jessica Van Ausdall, Kathryn Bruder-Mattson, Hunter Fleenor, and Angela Keffer

Department of Biological and Environmental Sciences, Samford University, Birmingham, AL 35229

Correspondence: David A. Johnson (djohnso2@samford.edu)
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Author:Johnson, David A.; Caffy, Kelsey; Van Ausdall, Jessica; Bruder-Mattson, Kathryn; Fleenor, Hunter; Ke
Publication:Journal of the Alabama Academy of Science
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Date:Jul 1, 2011
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