Department of Civil and Environmental Engineering,
University of Illinois at Urbana-Champaign, Urbana, Illinois
61801,1 and
Department of Civil,
Environmental and Architectural Engineering, University of
Colorado, Boulder, Colorado 803092
Previous studies have shown the predominance of mycolic
acid-containing filamentous actinomycetes (mycolata) in foam layers in
activated sludge systems. Gordona (formerly
Nocardia) amarae often is considered the major
representative of this group in activated sludge foam. In this study,
small-subunit rRNA genes of four G. amarae strains were
sequenced, and the resulting sequences were compared to the sequence of
G. amarae type strain SE-6. Comparative sequence analysis
showed that the five strains used represent two lines of evolutionary
descent; group 1 consists of strains NM23 and ASAC1, and group 2 contains strains SE-6, SE-102, and ASF3. The following three
oligonucleotide probes were designed: a species-specific probe for
G. amarae, a probe specific for group 1, and a probe
targeting group 2. The probes were characterized by dissociation
temperature and specificity studies, and the species-specific probe was
evaluated for use in fluorescent in situ hybridizations. By using the
group-specific probes, it was possible to place additional G. amarae isolates in their respective groups. The probes were used
along with previously designed probes in membrane hybridizations to
determine the abundance of G. amarae, group 1, group 2, bacterial, mycolata, and Gordona rRNAs in samples obtained
from foaming activated sludge systems in California, Illinois, and
Wisconsin. The target groups were present in significantly greater
concentrations in activated sludge foam than in mixed liquor and
persisted in anaerobic digesters. Hybridization results indicated that
the presence of certain G. amarae strains may be regional
or treatment plant specific and that previously uncharacterized
G. amarae strains may be present in some systems.
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INTRODUCTION |
Activated sludge treatment plants in
different parts of the world with widely varying wastewater
characteristics and operating conditions have experienced foaming
episodes in their aeration basins and anaerobic digesters (9, 12,
30, 38, 45). Measures that have been used to control foaming have
not produced consistent results; consequently, the adoption of control
mechanisms remains empirical (39). Foaming can lead to
numerous problems, such as reduction in effluent quality, severe
operational difficulties in anaerobic digesters, and clogging of gas
collection systems (29). The recent isolation of a
pathogenic Nocardia sp. from activated sludge foam
(44) suggests that foaming also may represent a health
concern because of the possible spread of pathogens in windblown scum
(8).
Mycolic acid-containing actinomycetes or mycolata (14) often
have been isolated from foams and are considered the main cause of foam
stabilization in many activated sludge systems throughout the world.
The phylogeny of the mycolata recently has been elucidated by using
small-subunit (SSU) rRNA sequence analyses. Goodfellow and coworkers
distinguished two suprageneric lineages and proposed that the mycolata
should be grouped into the following two families: the family
Corynebacteriaceae, which encompasses the genera
Corynebacterium, Dietza, and
Turicella; and the family Mycobacteriaceae,
containing the genera Gordona, Mycobacterium,
Nocardia, Rhodococcus, Tsukamurella, and Skermania (13, 14). Stackebrandt et al.
(40) proposed a new suborder within the order
Actinomycetales, the suborder Corynebacterineae,
which would include the families Nocardiaceae, Gordoniaceae, Mycobacteriaceae,
Dietziaceae, Tsukamurellaceae, and
Corynebacteriaceae. Some members of these groups, including Gordona (Nocardia) amarae and
"Microthrix parvicella," an unclassified actinomycete
(11), commonly have been cited as predominant organisms in
activated sludge foams (9, 12, 25, 38). Stackebrandt et al.
(40) also proposed that the genus Gordona should
be renamed Gordonia based on correct etymology.
In most studies in which it was determined that G. amarae
was a major microorganism in activated sludge foam, morphology- and
physiology-based identification methods were used. These methods are
inadequate for describing microbial diversity in complex environments, such as activated sludge (37, 47). Filament identification techniques (20, 23), which are widely used in activated
sludge microbiology, are hampered by the morphological similarities of actinomycetes (7) and the transitions that these organisms undergo depending on their growth stages (hyphal to coccal stages and
vice versa) (39). Biases presented by culture-based methods and the recent finding that G. amarae is one of the easiest
actinomycetes to isolate and maintain in pure culture (24)
make quantification of this organism with culture-based techniques
unreliable. Therefore, previous reports of G. amarae
abundance in foam samples may have been inadequate. These observations
raise new questions concerning the significance of this species in
foaming.
Oligonucleotide probes that target rRNA may provide more reliable
characterizations of microbial community structure (4, 5,
42). We previously designed a probe for G. amarae
based on a limited number of sequences, probe S-S-G.am-0192-a-A-18
(probe nomenclature is based on the nomenclature of the Oligonucleotide Probe Database [2]) (17). However, G. amarae quantification was impaired after it was determined that
the probe targeted only a few of the known G. amarae
strains. As a result, assessment of the significance of G. amarae in foaming was not possible.
In this study, we report interstrain heterogeneity in the SSU ribosomal
DNA (rDNA) sequences of G. amarae, which explains the
variable responses of various G. amarae strains with the
previously designed species-specific probe (17). We describe
the development and application of a probe for G. amarae
that targets all available strains, as well as two probes that
distinguish the interstrain sequence variability in this species.
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MATERIALS AND METHODS |
Organisms, culture techniques, and nucleic acid extractions.
The following seven strains of G. amarae were used in this
study: SE-6 (= ATCC 27808) (type strain), SE-102 (= ATCC 27809), SE-149B (= ATCC 27810), NM23, ASAC1, RBI, and ASF3. Strains SE-6, SE-102, and SE-149B were isolated by plate dilution by Lechevalier and
Lechevalier (25) from activated sludge foams in Andover, Fla. (Miami, Fla.), Bordentown Township, N.J., and Jamaica Bay, N.Y.,
respectively. Strain ASAC1 was obtained by micromanipulation from mixed
liquor from the Sacramento, Calif., Regional Water Quality Plant by
L. L. Blackall and H. Ho; strain RBI was isolated by plate
dilution from mixed liquor from the Richmond, Calif., wastewater
treatment plant by M. Richard (46); and strain ASF3 was
isolated by micromanipulation from mixed liquor from the San Francisco,
Calif., Southeast Water Pollution Control Plant. Strain NM23 was
obtained from an activated sludge system in Queensland, Australia, by
L. L. Blackall by micromanipulation (9). A variety of
organisms not targeted by the probes designed in this study served as
controls in specificity studies and fluorescent in situ hybridization
(FISH) studies (see below).
The strains were grown in yeast extract-glucose broth (10 g of tryptone
per liter, 7 g of NaCl per liter, 2.5 g of yeast extract per
liter, 1 g of dextrose per liter; pH 7.0) for 3 to 5 days at
37°C with constant shaking. Strain ASF3 was grown on the mineral salts medium of Stanier et al. (43) after it was found that this organism did not grow well in yeast extract-glucose broth. The
cells were harvested by centrifugation, and the cell pellets were
immediately frozen on dry ice and kept at 
80°C.
Genomic DNA was extracted by using a microwave protocol for
gram-positive bacteria. This method involved washing of the cell pellets with TE buffer (10 mM Tris [pH 8.0], 10 mM EDTA),
resuspension in TE buffer and 10% sodium dodecyl sulfate (SDS), and
incubation at 65°C for 30 min to lyse the cells. The lysate was
centrifuged, the tubes were heated in a microwave oven (1,000 W) twice
for 1 min, and the resulting pellets were dissolved in TE buffer. DNA
was extracted with chloroform-isoamyl alcohol-phenol (24:1:25, vol/vol/vol), precipitated with ethanol (overnight, 20°C), and centrifuged, and the resulting pellets were resuspended in water. DNA
concentrations were determined spectrophotometrically by assuming that
1 mg of DNA per ml was equivalent to 20 optical density units at a
wavelength of 260 nm. DNA quality was assessed by gel electropheresis on 1% agarose gels, and bands were viewed with a charge-coupled device
camera (Bioimaging Technologies, Elburn, Ill.).
RNA was extracted by a low-pH hot-phenol method (41). The
quality of the extracted RNA was evaluated by polyacrylamide gel electrophoresis (1), and the extracted RNA was quantified
with the BIT Image Analysis software (Bioimaging Technologies).
Amplification and cloning of SSU rDNA.
The SSU rDNA was
amplified by PCR with primers S-D-Bact-0011-a-S-17 and
S-D-Bact-1492-a-A-21 by using the following protocol: one cycle
consisting of 4 min at 94°C, 1 min at 50°C, and 1.5 min at 72°C;
28 cycles consisting of 1 min at 92°C, 1 min at 50°C, and 5 min at
72°C; and temporary storage at 4°C. The sizes and quantities of the
PCR products were evaluated by agarose gel electrophoresis, and the PCR
products were quantified with the BIT Image Analysis software.
Cloning and transformation were carried out by using instructions
provided with an Invitrogen TA cloning kit, version B (Invitrogen Corporation, San Diego, Calif.). Transformants were screened further by
preparing plasmids and performing restriction enzyme digestions (36).
Sequencing of the SSU rDNA.
Cell pellets were prepared from
overnight growth of transformed Escherichia coli cells on
Luria-Bertani broth, and sequencing was performed at the University of
Illinois Biotechnology Center Sequencing Laboratory with four primers
specific for the bacterial domain, S-*-Bact-0343-a-A-15
(TACGGGAGGCAGCAG), S-*-Bact-1100-a-S-16 (AGGGTTGCGCTCGTTG), S-*-Bact-0519-a-S-18
(GTATTACCGCGGCTGCTG), and S-*-Bact-0907-a-A-20
(AAACTCAAATGAATTGACGG) (19), as well as the
M13(-20) forward and M13 reverse primers (Invitrogen Corporation). The
sequences obtained for strains SE-102, ASAC1, ASF3, and NM23 were
assembled and edited by using the Fragment Assembly program of the
Genetics Computer Group software, version 9.0, Wisconsin Package, and
the Sequencher program (Gene Codes Corp., Ann Arbor, Mich.).
Phylogenetic analyses were done with programs available in the PHYLIP
package (version 3.5) (21), such as the distance and
maximum-likelihood methods.
Design and characterization of oligonucleotide probes.
Based
on the SSU rDNA sequences obtained for strains NM23, ASAC1, ASF3, and
SE-102 and the sequence of type strain SE-6 available from the
Ribosomal Database Project (RDP) (26), oligonucleotide probes were designed to target the G. amarae strains. The
newly designed oligonucleotide probes, their target groups, and their sequences are shown in Table 1 and Fig.
1. Additional probes used in this study
also are listed in Table 1. The oligonucleotide probes were synthesized
with a DNA synthesizer (Applied Biosystems, Foster City, Calif.) at the
University of Illinois Biotechnology Center DNA Synthesis Laboratory
and were purified with an oligonucleotide purification cartridge
(Applied Biosystems).
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FIG. 1.
Td studies for probes
S-S-G.am-0205-a-A-19, S-*-G.am1-0439-a-A-19, and S-*-G.am2-0439-a-A-19.
Adjacent to the probe dissociation results are the SSU rRNA sequences
of target and nontarget species and probe sequences. The top SSU rRNA
sequence in each list of sequences is the target sequence. Dots in the
sequences below this sequence indicate identical nucleotides, and
replacement nucleotides are nucleotides that differ from the
nucleotides in the target sequences.
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To determine the optimal hybridization wash temperatures, dissociation
temperature (Td) studies were performed
(17, 50) by using RNAs from a variety of target and
nontarget organisms (Fig. 1). Oligonucleotide probes were 5' end
labeled with [
-32P]ATP (ICN Radiochemicals, Irvine,
Calif.) (32) and were purified with a Nensorb 20 cartridge
(DuPont Corp., Boston, Mass.) (42). Prehybridization,
hybridization, washing, and scintillation counting were carried out as
described by de los Reyes et al. (17). The temperature at
which 50% of the hybridized probe washed off was determined to be the
Td of the probe.
The specificities of the probes were assessed experimentally by
hybridizing them with RNAs from 41 target and nontarget organisms (Fig.
2). The final wash of the hybridization
membranes was performed at the previously determined
Td, the membranes were exposed to PhosphorImager
screens (Molecular Dynamics, Sunnyvale, Calif.), and hybridization
signals were viewed and quantified with a PhosphorImager (Molecular
Dynamics) (17).
Characterization of environmental samples.
The environmental
samples used and the wastewater treatment plants from which they were
obtained are listed in Table 2. All of
the plants experienced foaming at the time of sampling or had a history
of recurring foaming. The samples were stored in 50-ml conical
centrifuge tubes (Corning Costar Corp., Cambridge, Mass.), immediately
put on ice, and brought or mailed overnight to the laboratory. Aliquots
were centrifuged, and cell pellets were frozen in ethanol with dry ice
and stored at 80°C until they were used for RNA extraction. RNA was
extracted by a low-pH hot-phenol extraction method (41).
Quantification of extracted RNA and hybridizations with radioactive
oligonucleotide probes were performed as previously described (17). All hybridization membranes were prepared in
duplicate; for each set of membranes, one membrane was hybridized with
a universal probe, and the other membrane was hybridized with a specific probe (Table 1). The hybridization responses of triplicate applications of RNAs obtained from samples and of a dilution series of
RNA obtained from a pure culture (target organism) were used to
determine the relative concentrations of target SSU rRNAs in the
samples. Hybridization results were expressed in terms of the relative
SSU rRNA concentrations of target groups, which were calculated by
dividing the amount of rRNA (in nanograms) targeted by a specific probe
by the amount of rRNA (in nanograms) bound to the universal probe.
FISH and microscopy.
A tetramethyl rhodamine (TRITC)-labeled
version of the species-specific probe S-S-G.am-0205-a-A-19 was obtained
from Genosys Biotechnologies, Inc. (The Woodlands, Tex.). Pure cultures
and wastewater treatment plant samples were fixed in 4%
paraformaldehyde for 1 min at room temperature (17). FISH
were performed on SuperCured heavy Teflon-coated slides (Cel-Line
Associates Inc., Newfield, N.J.) as previously described
(17), with minor modifications. The hybridization buffer
consisted of 30% formamide, 0.9 M NaCl, 0.1 M Tris (pH 7.2), and 0.1%
SDS. Hybridizations were conducted in saturated humidity chambers at
46°C for 2 h. The slides were rinsed once and immersed in 50 ml
of wash solution (80 mM NaCl, 0.05 M sodium phosphate buffer [pH
7.2], 0.1% SDS, 5 mM EDTA) at 48°C for 20 min. These hybridization
and wash conditions were found to be optimal for in situ hybridization
with this probe. In some cases, dual staining with
4',6-diamidino-2-phenylindole dihydrochloride (DAPI) was performed
(17). The slides were then washed with ice-cold water, air
dried, and mounted in Citifluor (UKC Chemical Laboratory, Canterbury,
United Kingdom).
The slides were visualized with an epifluorescence microscope
(Axioskop; Carl Zeiss, Oberkochen, Germany) fitted with filter sets for
DAPI and TRITC (Chroma Technology Corp., Brattleboro, Vt.)
(17). Images were captured with a charge-coupled device camera (Photometrics Ltd., Tucson, Ariz.) and IPLab Spectrum image analysis software (Signal Analytics, Vienna, Va.). Images were imported
to Adobe Photoshop 3.0 (Adobe, Seattle, Wash.) for printing.
Nucleotide sequence accession numbers.
Nucleotide sequences
have been deposited in the GenBank database under accession no.
AF020329 for ASAC1, accession no. AF020330 for SE-102, accession no.
AF020331 for ASF3, and accession no. AF020332 for NM23.
| |
RESULTS AND DISCUSSION |
Sequence analysis.
Complete SSU rDNA sequences for strains
NM23, ASAC1, ASF3, and SE-102 and partial sequences for RBI and SE-149B
were obtained. Alignment with sequences deposited in the RDP and
GenBank-EMBL databases showed that the sequences of all of the strains
were most similar to the G. amarae SE-6 sequence (accession
no. X80635). An analysis to determine the presence of chimeras produced
negative results for all sequences.
A comparison of the full-length sequences and similarity values
obtained from the Pileup program of the Genetics Computer Group
software indicated that strains ASAC1 and NM23 were more similar to
each other than to the other strains, while strains ASF3, SE-102, and
SE-6 clustered together (data not shown). This apparent grouping of the
G. amarae strains into two clusters was supported by the
results of phylogenetic analyses in which we used maximum-likelihood
and distance-based neighbor-joining methods, which produced similar
phylogenetic trees. Figure 3 shows the phylogenetic tree obtained with a neighbor-joining-unweighted pair
group method with arithmetic averages (distances were calculated by
using a Kimura two-parameter distance setting), which shows the
distinct branching of the strains into two groups; strains NM23 and
ASAC1 cluster together in one group (group 1), and strains SE-102,
ASF3, and SE-6 cluster together in another group (group 2). The partial
sequences of strains SE-149B and RBI indicated that these organisms
belong to group 1, and this was confirmed by the results of probe
specificity and Td studies (see below). The
evolutionary relationships among mycolata reflected in the phylogenetic
tree are consistent with the evolutionary relationships observed by
Chun et al. (14).
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FIG. 3.
Phylogenetic tree showing the positions of
representative mycolata, inferred from a comparison of SSU rRNA
sequences. The tree was constructed by using a neighbor-joining method.
The bar represents 5 estimated changes per 100 nucleotides. The
oligonucleotide probes are shown with their respective target groups.
Probes I, II, and III were designed in this study; probes IV and V have
been described previously (17).
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Intraspecific variation in SSU rDNA sequences has been attributed to
sequencing errors, misidentification of strains, and rRNA operon
heterogeneity (16, 27, 28, 31, 35). The interstrain
variability observed among the sequences obtained in this study (0.5 to
1%) is higher than the estimated 0.1% random sequencing error for
sequences deposited in GenBank (15). Moreover, alignment of
the sequences showed that the strains were properly classified as
G. amarae. Interoperon differences cannot be ruled out as a
potential source of the interstrain variability observed in this study.
This can be tested if intrastrain variability is compared with
interstrain variability (31). Even though we did not
evaluate this possibility through additional sequence analyses, the
results of probe specificity studies indicated that it is unlikely (see
below).
Design of oligonucleotide probes.
The following three
oligonucleotide probes were designed and characterized based on the
results of the sequence analysis: S-S-G.am-0205-a-A-19, which is
species specific for G. amarae; S-*-G.am1-0439-a-A-19, which
targets group 1 G. amarae strains; and
S-*-G.am2-0439-a-A-19, which is specific for group 2 strains (Fig. 1
and 3). The specificities of the probes were initially assessed by
performing a Basic Local Alignment Search Tool (BLAST) in GenBank
(3) and by using the CHECK_PROBE program provided by the RDP
(26). The species-specific probe perfectly matched all
G. amarae strains for which sequences were available and was an improvement over the previously designed G. amarae probe
(17), which was found to have two mismatches with the group
1 strains. Probe S-S-G.am-0205-a-A-19 had at least two consecutive
mismatches (at positions 216 and 217 [Escherichia coli
numbering]) with the SSU rRNA sequences of nontarget species.
Probe S-*-G.am1-0439-a-A-19 perfectly matched all group 1 strains. A
few closely related nontarget species (Gordona terrae, Gordona rubropertincta, and Gordona bronchialis)
had one mismatch. Other nontarget organisms had at least two
mismatches, while group 2 strains had five mismatches. Similarly, probe
S-*-G.am2-0439-a-A-19 perfectly matched all group 2 strains, had at
least three mismatches with nontarget organisms, and had five
mismatches with group 1 strains.
Other factors considered during probe design were G+C content, probe
length, and the presence of self-complementary regions. The G+C content
and probe length were adjusted as much as possible to obtain similar
theoretical Td values. This is important when multiple probes are used in simultaneous membrane or in situ
hybridizations (34). No regions of internal complementarity
were present in S-S-G.am-0205-a-A-19 and S-*-G.am1-0439-a-A-19, whereas
probe S-*-G.am2-0439-a-A-19 contained a 4-base region of
self-complementarity, but this internal complementarity region likely
did not affect the hybridization response, as discussed previously
(34).
Optimization of wash temperatures.
The
Td values were experimentally determined for
each probe and were used as posthybridization wash temperatures to
ensure dissociation of duplexes with one or more mismatches
(42). Figure 1 shows the Td curves
obtained for the three probes and the sequence alignments for the
probes and the target and nontarget organisms used in the
Td studies.
Strains SE-149B (group 1) and SE-102 (group 2) had overlapping
Td curves (Td, 53.5°C),
indicating that they had identical responses with the species-specific
probe S-S-G.am-0205-a-A-19. G. terrae and
Mycobacterium vaccae, both of which had two mismatches with
this probe, had Td values between 42 and 45°C.
The group 1 probe S-*-G.am1-0439-a-A-19 had a Td
of 65°C for group 1 strain SE-149B and lower
Td values for the following nontarget organisms:
57°C for G. terrae (one mismatch), 53°C for Rhodococcus equi (three mismatches), and 52.5°C for group
2 strain SE-102 (five mismatches). These results indicate that the
group 1 probe can be used to distinguish a target strain from a
nontarget strain with a single nucleotide mismatch when a
posthybridization wash temperature of approximately 65°C is used.
The Td values for group 2 strains SE-6 and
SE-102 were 61 and 63°C, respectively, for group 2 probe
S-*-G.am2-0439-a-A-19. Corynebacterium renale (three
mismatches) had a Td of 51°C, while the group
1 strains SE-149B and NM23 (five mismatches) exhibited Td values between 50 and 51°C.
Probe specificity studies.
A diverse selection of RNAs
obtained from target and nontarget organisms (Fig. 2) were hybridized
with the oligonucleotide probes to assess their specificities. Figure
4 shows the hybridization membranes used
for this specificity study. Membrane b was hybridized with a universal
probe (S-*-Univ-1390-a-A-18 [50]), which targeted all
RNA samples applied to the membrane. Variations in the signal intensities on this membrane were likely due to differences in the
amounts of RNA applied. Membrane c shows the results of hybridization with the species-specific probe; this probe targeted all seven G. amarae strains, including strains SE-149B and RBI, for which only
partial sequences were available. The variations in signal intensity
for the G. amarae strains were generally consistent with the
variations observed with the universal probe. The specificity results
for the group 1 probe (membrane d) show that this probe hybridized with
all group 1 strains, including strains SE-149B and RBI. We used a wash
temperature of 66°C for this probe; this temperature was 1°C higher
than the experimentally derived Td, which
resulted in minimal nonspecific binding for closely related nontarget
organisms. For example, the hybridization responses for G. terrae and M. vaccae RNAs comprised only 2.6 and 3.1%, respectively, of the signal for the target group 1 RNAs. Membrane e
shows that the group 2 probe hybridized only with group 2 strains. The
wash temperature that resulted in strong hybridization responses for
the target strains and negligible signals for nontarget organisms was
61°C. Overall, the specificity studies showed that the three probes
were specific to their target organisms, which was consistent with the
results of database searches.
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FIG. 4.
Results of the probe specificity study. Membrane
hybridization results were analyzed with a PhosphorImager and were
scanned and printed with Adobe Photoshop 3.0. (a) Template. Names of
individual organisms are shown in Fig. 2. (b) Probe
S-*-Univ-1390-a-A-18. (c) Probe S-S-G.am-0205-a-A-19. (d) Probe
S-*-G.am1-0439-a-A-19. (e) Probe S-*-G.am2-0439-a-A-19.
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Characterization of environmental samples.
The probes designed
in this study, as well as probes for the genus Gordona, the
mycolata, and the bacterial domain (Table 1), were used to determine
whether G. amarae strains were involved in foaming problems
experienced in six full-scale wastewater treatment plants in Illinois,
California, and Wisconsin (Table 2). Hybridization results for samples
from these treatment plants are summarized in Table
3. Samples were obtained from activated
sludge foam and mixed liquor from the Illinois and Wisconsin treatment
plants. Because three of the California plants have high-purity
oxygen-activated sludge systems, samples were taken from the return
activated sludge (RAS) lines. The hybridization results for the
Illinois plant revealed similar rRNA levels for all target groups for
RAS and mixed liquor (data not shown), suggesting that RAS data can be used to estimate target group abundance in mixed liquor. In addition to
the activated sludge samples, anaerobic digester sludge samples were
obtained from the Illinois and California plants, and an anaerobic
digester foam sample was taken at the Illinois plant.
The hybridization results for the Urbana, Ill., plant indicated that
activated sludge foam had significantly higher rRNA levels than mixed
liquor for the bacterial domain, mycolata, the genus Gordona, G. amarae, and group 1 strains. This
finding supports the results of previous studies in which researchers
observed a greater abundance of filamentous actinomycetes in activated sludge foam than in mixed liquor (12, 49).
Gordona strains appeared to be the only mycolata
representatives present in this activated sludge system since the
relative rRNA levels obtained with the Gordona-specific
probe were slightly higher than the levels obtained with the myclolata
probe in foam and mixed liquor (Table 3). Furthermore, a significant
percentage of the Gordona strains were G. amarae
strains (65% for activated sludge foam, 51% for mixed liquor), the
majority of which were group 1 strains (55% for activated sludge foam,
71% for mixed liquor). Group 2 strains probably were not present in
this system since the hybridization response was close to or below the
detection limit (Table 3). Thus, it appears that G. amarae
group 1 strains were involved in the foaming problem observed in the
Urbana treatment plant. However, since group 1 strains contributed only
approximately one-half of the G. amarae rRNA in foam,
previously uncharacterized G. amarae strains likely were
present in this system.
Results obtained with samples from a foaming anaerobic digester at the
Urbana treatment plant indicate that the relative rRNA levels for
mycolata, the genus Gordona, and G. amarae were
higher in anaerobic digester sludge than in activated sludge mixed
liquor. This implies that there was a transfer of mycolata from the
activated sludge basin to the anaerobic digester through
waste-activated sludge feeding. These findings also suggest that rRNAs
from strictly aerobic filamentous actinomycetes persisted under
anaerobic conditions. The survival of significant quantities of
metabolically competent Gordona (Nocardia)
filaments in anaerobic digesters has previously been observed by
Hernandez and Jenkins (22). In continuous-flow digestion
experiments, these authors observed only 37% filament reduction after
a 14-day retention time and noted that 60% of the filaments retained
their ability to respire, as judged by 2-(p-iodophenyl)-3-(p-nitrophenyl)-S-phenyltetrazolium
chloride reduction, after 14 days of anaerobic digestion. A discrepancy between ribosome content and metabolic activity has also been observed
for a methanogenic reactor which was amended with sulfate (33); the methanogenic activity ceased immediately after
sulfate was added (as indicated by a cessation of methane production), but the methanogen rRNA levels decreased at a much lower rate. Thus, in
accordance with previous observations obtained with oligonucleotide probes (33, 48), our results suggest that rRNA levels may be
poor indicators of reduced metabolic activity in environmental samples.
However, foaming is a physical phenomenon caused by the entrapment of
gas within a film of liquid and stabilized by the presence of
filamentous microorganisms with hydrophobic cell walls (23).
Even when filamentous actinomycetes are metabolically inactive, their
presence can contribute to foam formation and stabilization. Therefore,
the fact that cellular rRNA is present for considerable time periods
even if organisms are metabolically inactive might make it possible to
develop an early-warning system for foaming based on hybridizations
with rRNA-targeted probes.
In previous studies (22, 45) workers have observed selective
accumulation of mycolata strains in anaerobic digester foam, and these
findings contradict our findings that mycolata rRNA is more abundant in
anaerobic digester sludge than in anaerobic digester foam (Table 3). In
these studies, however, the researchers used conventional and
immunofluorescent stains, which are not dependent on cellular
metabolism for visualization. The rRNA content of mycolata cells in
anaerobic digester foam may be low enough to account for this
discrepancy. This is because the partitioning of digester foam from
digesting sludge may lead to longer retention times for digester foam.
Increased retention times in anaerobic digesters significantly increase
the degradation of Gordona filaments and reduce their
viability (22). It is also possible that biogas production
in the Urbana anaerobic digester resulted in a level of selective
accumulation of mycolata strains in anaerobic digester foam lower than
the level of selective accumulation caused by aeration in activated
sludge basins. In any case, since foaming in the Urbana anaerobic
digester appeared to be linked to the presence of mycolata, control
strategies should take into consideration the recycling of filamentous
actinomycetes; the seeding of anaerobic digesters with activated sludge
foam can cause foaming problems in anaerobic digesters, and recycling
of anaerobic digester supernatant to activated sludge systems may
result in reseeding of the aeration basin.
The mycolata, Gordona, and G. amarae rRNA levels
in the RAS of the San Francisco and Richmond plants were significantly
higher than those in the mixed liquor of the Urbana plant (Table 3), suggesting that filamentous actinomycetes should also be very abundant
in the activated sludge foam of these California plants. Almost all of
the mycolata rRNA was Gordona rRNA, and the majority of the
Gordona rRNA was G. amarae rRNA in these plants.
On the other hand, the rRNA levels in the Oakland and Sacramento plants were significantly lower than those in the Urbana treatment system, suggesting that filamentous microorganisms not targeted by our probes
were responsible for foaming. Gordona spp. other than
G. amarae and members of genera other than the genus
Gordona constituted a significant fraction of the mycolata.
This further supported the observation that G. amarae was
not contributing to foaming problems in the Oakland and Sacramento
plants.
The combined abundance of group 1 and group 2 rRNAs approximated the
abundance of G. amarae rRNA in Sacramento and Oakland samples. For the San Francisco and Richmond samples, the group probes
targeted 56 to 79 and 44 to 53%, respectively, of the G. amarae strains. These results indicate that novel strains of
G. amarae that are not targeted by the group probes may have
been present in the San Francisco and Richmond plants. Hybridization data obtained with the group probes also demonstrated the predominance of G. amarae strains in sites from which they were isolated
originally. For example, group 2 strains were more abundant in San
Francisco samples, from which strain ASF3 (group 2) was originally
isolated, and the group 1 strain rRNA levels were higher in the
Sacramento and Richmond treatment plants, the sites of isolation of
strains ASAC1 and RBI (group 1), respectively. Since some of the
strains were isolated by micromanipulation techniques, these results
indicate that micromanipulation can be a reliable tool for isolating
predominant strains from environmental samples, as suggested by Soddell
and Seviour (39).
Hybridization results obtained with samples from a milk-processing
wastewater treatment plant in Wisconsin, which was experiencing severe
foaming at the time of sampling, showed that the mycolata, Gordona, and G. amarae rRNA levels were only
slightly higher in activated sludge foam than in mixed liquor (Table
3), suggesting that mycolata did not selectively accumulate in foam. In
addition, the levels of these organisms in foam were significantly
lower than the levels in the foam of the Urbana plant. The genus
Gordona was found to be the predominant genus within the
mycolata, but G. amarae rRNA constituted less than one-half
of the Gordona rRNA. Group 1 rRNA contributed approximately
one-third of the G. amarae rRNA, while group 2 strains were
absent.
Since mycolata apparently were not responsible for foaming in this
milk-processing wastewater treatment plant, surfactants present in the
wastewater or filamentous microorganisms not targeted by our probes may
have caused the foaming problem. For example, it is possible that
"Microthrix parvicella," a common constituent of
activated sludge foam (8, 12, 30), was involved in foaming in this plant. Using the recently published SSU rRNA sequence of
"Microthrix parvicella" (10), we determined
that none of our probes target this organism.
FISH.
The results of membrane hybridizations indicate the
contributions of organisms involved in filamentous foaming to total
activity, as reflected by rRNA levels. However, it is important to
develop a method that relates the abundance of filamentous
microorganisms (not only rRNA levels) to the severity of foaming. FISH
has better potential to accomplish this goal. Therefore, we addressed
problems with permeabilizing cell walls of gram-positive bacteria for
probe accessibility in a previous study (17). We
demonstrated that fixation with 4% paraformaldehyde for very short
time periods (1 min) rendered most mycolata permeable for labeled
probes (17). Schuppler et al. (37b) recently
demonstrated that ethanol fixation combined with mutanolysin incubation
after dehydration also resulted in excellent FISH results for mycolata.
We also developed a method for quantifying members of the genus
Gordona on a mass basis in foaming activated sludge systems
and anaerobic digesters by using FISH (17, 18). Additional
studies are necessary to relate this and other quantitative approaches
to foaming potential.
To demonstrate the potential of FISH for such studies, we used a
TRITC-labeled version of the G. amarae-specific probe
S-S-G.am-0205-a-A-19. The optimum formamide concentration in the
hybridization buffer was determined to be 30%, and the optimum NaCl
concentration in the corresponding wash solution was found to be 80 mM.
Figures 5a and b show the specificity of
the probe when it was hybridized with a mixture of cultures of R. rhodochrous and G. amarae SE-102. Figure 5a shows the
mixed culture stained with DAPI, while Fig. 5b shows the same image
field viewed with a TRITC filter set. A comparison of Fig. 5a and b
indicates that G. amarae can be distinguished clearly from
nontarget cells at the hybridization stringency used. Figures 5c and d
show the application of FISH to a RAS sample from the San Francisco
treatment plant. Figure 5c shows the phase-contrast image, while Fig.
5d shows the same image field when a TRITC filter set was used. These
images demonstrate the ability of FISH to distinguish target cells
inside and extending from activated sludge flocs without regard for
morphology. The formation of G. amarae filamentous clusters
was observed, which confirmed previous observations (17). In
addition, we observed very low levels of nonspecific probe binding and
activated sludge floc autofluorescence. This is particularly
encouraging for future FISH studies with this probe.
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FIG. 5.
Phase-contrast and epifluorescence images after FISH of
mixed cultures and activated sludge. Bar = 10 µm. (a and b)
Mixture of cultures of R. rhodochrous and G. amarae SE-102 hybridized with TRITC-labeled probe
S-S-G.am-0205-a-A-19 and dual stained with DAPI. Panels a and b show
the same image field, as viewed by epifluorescence microscopy, prepared
with the DAPI and TRITC filter sets, respectively. (c and d) RAS sample
from San Francisco Southeast Water Pollution Control Plant hybridized
with probe S-S-G.am-0205-a-A-19 labeled with TRITC. Panels c and d show
the same image field, as viewed by phase-contrast microscopy and
epifluorescence microscopy, respectively.
|
|
Conclusions.
In this study, we developed oligonucleotide
probes for G. amarae and two subgroups within this species.
The probes allowed identification and quantification of G. amarae strains often implicated in foaming activated sludge and
anaerobic digester systems by membrane hybridization and FISH. Use of a
nested set of probes permitted a quantitative assessment of G. amarae strains in foaming samples, which is necessary when the
importance of this species in foaming is evaluated. The quantities and
types of G. amarae rRNA were found to vary in different
treatment plants. Future studies on the analysis of foaming occurrences
should benefit from the use of phylogenetic probes specific for
foam-related microorganisms, such as the probes designed in this study.
In particular, correlating FISH and membrane hybridization results to
changes in foaming potential could provide threshold values which could
be used to predict foaming incidents.
We thank the following personnel of the wastewater treatment
plants for providing samples: Tim Bachman, Urbana-Champaign Sanitary District; Paul Pitt, City and County of San Francisco; David Fretias, Oakland East Bay Municipal Utilities District; William Louis, City of
Sacramento Regional Wastewater Treatment Plant; and the operators of
the Richmond Wastewater Treatment Plant.
This research was supported by grant BES 9410476 from the U.S. National
Science Foundation.
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Abstract
Introduction
