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Previous Article | Next Article 
Applied and Environmental Microbiology, March 2000, p. 1167-1174, Vol. 66, No. 3
Center for Environmental
Biotechnology,1 Department of Chemical
Engineering,2 and Department of
Microbiology and Department of Ecology and Evolutionary
Biology,4 The University of Tennessee,
Knoxville, Tennessee 37996, and Tennessee Eastman Division,
Eastman Chemical Company, Kingsport, Tennessee
376623
Received 29 October 1998/Accepted 22 October 1999
The bacterial community structure of the activated sludge from a 25 million-gal-per-day industrial wastewater treatment plant was
investigated using rRNA analysis. 16S ribosomal DNA (rDNA) libraries
were created from three sludge samples taken on different dates.
Partial rRNA gene sequences were obtained for 46 rDNA clones, and
nearly complete 16S rRNA sequences were obtained for 18 clones. Seventeen of these clones were members of the beta subdivision, and
their sequences showed high homology to sequences of known bacterial
species as well as published 16S rDNA sequences from other activated
sludge sources. Sixteen clones belonged to the alpha subdivision, 7 of
which showed similarity to Hyphomicrobium species. This
cluster was chosen for further studies due to earlier work on
Hyphomicrobium sp. strain M3 isolated from this treatment plant. A nearly full-length 16S rDNA sequence was obtained from Hyphomicrobium sp. strain M3. Phylogenetic analysis
revealed that Hyphomicrobium sp. strain M3 was 99% similar
to Hyphomicrobium denitrificans DSM 1869T in
Hyphomicrobium cluster II. Three of the cloned sequences
from the activated sludge samples also grouped with those of
Hyphomicrobium cluster II, with a 96% sequence similarity
to that of Hyphomicrobium sp. strain M3. The other four
cloned sequences from the activated sludge sample were more closely
related to those of the Hyphomicrobium cluster I organisms
(95 to 97% similarity). Whole-cell fluorescence hybridization of
microorganisms in the activated sludge with genus-specific Hyphomicrobium probe S-G-Hypho-1241-a-A-19 enhanced the
visualization of Hyphomicrobium and revealed that
Hyphomicrobium appears to be abundant both on the outside
of flocs and within the floc structure. Dot blot hybridization of
activated sludge samples from 1995 with probes designed for
Hyphomicrobium cluster I and Hyphomicrobium cluster II indicated that Hyphomicrobium cluster
II-positive 16S rRNA dominated over Hyphomicrobium cluster
I-positive 16S rRNA by 3- to 12-fold. Hyphomicrobium 16S
rRNA comprised approximately 5% of the 16S rRNA in the activated sludge.
Activated sludge, a common
biological treatment method for both municipal and industrial
wastewater, represents a complex microbial community. Due to intricate
interactions within the microbial community, process control of
wastewater treatment plants can be difficult. Population shifts within
the microbial community may result from changes in the plant operating
conditions and cause sludge quality problems such as poor sludge
settling, compaction, and dewatering (22). The application
of molecular analysis to activated sludge is of considerable interest
as a means for determining the microbial diversity and robustness,
identifying populations associated with process upsets, and developing
probes for diagnosing, monitoring, and controlling activated-sludge
problems (5, 7, 8, 16, 27, 33, 37). Some operational
problems with activated sludge can often be detected microscopically at
the microorganism level. For example, poor sludge settling due to filamentous bulking is due to excessive growth of bacterial filaments (e.g., Sphaerotilus natans, Microthrix
parvicella, Hyphomicrobium spp., Thiothrix
nivea, etc.) (19). Diagnosis and correction of such
activated-sludge problems require the correct identification of the
responsible microbial population(s) and institution of appropriate
process changes to select for or against specific organisms.
A potential for considerable variation between the activated sludge of
industrial and that of municipal wastewater treatment exists due to the
differences in chemical composition of the treated waste streams. In
this study, the microbial community from an activated sludge system
used to treat wastewater from chemical manufacturing processes was
examined. This differs from municipal wastewater in that it is much
higher in its total organic carbon load, comprising mostly simple
organic acids and alcohols, with little fibrous or complex carbon
sources present. One microorganism of particular interest routinely
monitored in this wastewater treatment system is Hyphomicrobium.
Hyphomicrobium spp. have been reported in both sewage treatment
plants and adjacent waters (15, 17, 20). Recently, the 16S
ribosomal DNA (rDNA) sequences for seven species of the genus
Hyphomicrobium were published (30). These species
fall into two distinct phylogenetic clusters indistinguishable by
morphological characteristics alone. The maintenance of
Hyphomicrobium in this industrial activated sludge is
important due to its ability to degrade C-1 compounds such as methanol,
which is found in the influent wastewater (18, 20). However,
hypertrophic growth (i.e., hyphal elongation) of
Hyphomicrobium can lead to poor sludge settling and
compaction (A. J. Meyers and C. D. Meyers, Abstr. 86th Annu.
Meet. Am. Soc. Microbiol. 1986, abstr. N-93, 1986). Therefore, the
ability to differentiate and reliably monitor Hyphomicrobium levels in the flocs is fundamental for optimal control and operation of
the wastewater treatment plant.
Other researchers (5, 33, 37) have proposed that a
combination of approaches is needed to understand the basic microbial community structure of activated sludge. These methods include the
construction and analysis of 16S rDNA libraries, hybridization with
rRNA-targeted oligonucleotides, and comparison of those results with a
specific focus on the presence of Hyphomicrobium-like strains.
Samples.
Activated sludge samples were collected from the
Eastman Chemical Company wastewater treatment plant (Tennessee Eastman
Division, Kingsport, Tenn.). The influent wastewater contains primarily low-molecular-weight organic acids (e.g., acetic acid, propionic acid,
n-butyric acid) and short-chain alcohols such as methanol, ethanol, and isopropanol (10). The plant includes four
parallel trains, each with three aeration basins in series, and is
operated in a modified step-feed flow configuration. The relevant
operating characteristics for the 3 months in which samples were taken
are provided in Table 1.
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Quantification of Hyphomicrobium Populations in
Activated Sludge from an Industrial Wastewater Treatment System as
Determined by 16S rRNA Analysis
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
TABLE 1.
Performance summary for the wastewater treatment plant
during the 3 months of sampling
Isolation and growth of Hyphomicrobium strains. Hyphomicrobium sp. strain M3 was previously isolated and identified from this activated sludge during an episode of poor settling due to the abundance of its elongated hyphae protruding from the flocs (Meyers and Meyers, Abstr. 86th Annu. Meet. Am. Soc. Microbiol. 1986). Isolation of the strain was on a liquid medium consisting of 1.36 g of KH2PO4, 0.5 g of (NH4)2SO4, 0.4 g of KNO3, 0.01 g of CaCl2 · 2H2O, 0.0031 g of MnSO4 · 4H2O, 2.13 g of Na2HPO4, 0.2 g of MgSO4 · 7H2O, 0.005 g of FeSO4 · 7H2O, and 0.0025 g of Na2MoO4 · 2H2O/liter with a final pH of 7.2 (1) and supplemented with 10 ml of methanol/liter under anoxic conditions at 30°C. After sequential enrichment of the strain on media with methanol, the strain was routinely cultivated aerobically at 30°C using the above-described medium without KNO3 and supplemented with 10 g of methylamine hydrochloride/liter. Isolation in pure culture was accomplished on this medium solidified with 15 g of agar/liter. Hyphomicrobium sp. strain M3 has been deposited in the American Type Culture Collection as ATCC 202122.
Strain M3 was identified as Hyphomicrobium based on morphological and physiological characteristics. In pure culture, this gram-negative, oval-shaped bacterium showed the diagnostic "mother cell," with its monopolar budding giving rise to a "daughter cell" at the terminus of a thin, thread-like hypha (prostheca) (18). Hyphomicrobium sp. strain M3 is mesophilic (growth temperature range of 10 to 35°C; optimum, 30°C), is relatively slow growing (doubling time on methanol at 30°C is 9.1 h), is facultatively anaerobic, and has hyphal lengths of 3 to 5 µm (Meyers and Meyers, Abstr. 86th Annu. Meet. Am. Soc. Microbiol. 1986). This particular strain can utilize several C1 compounds (e.g., methanol, methylamine HCl, formic acid, methylurea, dimethylamine, and formamide) but not others (e.g., methane, formaldehyde, dimethyl ether, and dimethyl sulfoxide), qualifying it as a restrictive type of methyltroph. Selective C2 compounds such as acetic acid, ethanol, and ethylamine support its growth, while higher alcohols (e.g., n-isopropanol and n-butanol) and organic acids (e.g., propionic, n-butyric, succinic, and citric), sugars (e.g., glucose and sucrose), and other compounds (e.g., ethylene glycol and acetone) fail to support growth. A rather wide range of nitrogen sources are employed by Hyphomicrobium sp. strain M3; these include ammonium chloride, sodium nitrate, sodium nitrite, sodium azide, oxamic acid, and various amines (e.g., methylamine, dimethylamine, and glucosamine) and amides (e.g., formamide, acetamide, and N,N-dimethylformamide). Amino acids do not serve as either N or C sources for the organism. As with other Hyphomicrobium isolates, strain M3 accumulates intracellular reserves of poly-
-hydroxybutyric acid, engages in rosette formation, and produces a pellicle in quiescent liquid culture.
Extraction of DNA from sludge.
DNA was extracted from this
sludge by following a modified method of Ogram et al. (28).
Sludge samples (volumes ranging from 20 to 150 ml) were centrifuged at
5,500 × g at 10°C for 10 min. The supernatant
fractions were discarded, and the pellets were resuspended in 50 ml of
0.12 M Na2PO4 (pH 8.0)-2.5 ml of 5% (wt/vol)
sodium dodecyl sulfate. These samples were incubated at 70°C for
1 h with periodic inversions by hand. Five grams of 0.1-mm-diameter glass beads was added to the samples, and the samples
were blended for two 2-min bursts separated by a 1-min rest period. The
samples were recovered and centrifuged again at 5,500 × g for 25 min at 10°C; the resulting supernatants were collected
and stored at 4°C. The pellets were resuspended in 25 ml of 0.12 M
Na2PO4 and incubated at 70°C for 20 min with
periodic inversions by hand. After centrifugation at 5,500 × g for 25 min at 10°C, the supernatants were pooled with the
previously collected supernatants. Precipitation for >2 h at 
20°C
was performed by the addition of 0.1 volume of 2 M sodium acetate
(NaOAc) solution and 0.8 volumes of isopropanol. The precipitates were
then centrifuged at 11,500 × g at 4°C for 30 min and
dried under vacuum at 100°C. Excess salts were eliminated by
dialysis using Spectra dialysis tubing (molecular weight cutoff of
6,000 to 8,000) overnight against 10 mM Tris-HCl-1 mM EDTA buffer (pH
8.0) (TE buffer). Dialyzed samples were extracted with Tris-saturated
phenol followed by extraction with chloroform-isoamyl alcohol (24:1
[vol/vol]). The recovered aqueous phases were precipitated at
20°C for 2 h by the addition of 0.1 volume of 2 M NaOAc and 2 volumes of absolute ethanol. The final precipitation product was
centrifuged at 11,500 × g for 30 min at 4°C and
dried under vacuum at 100°C. The samples were resuspended in 1 ml
of sterile TE buffer and stored at 20°C. RNA was removed from the
samples by treatment with 5 µl of DNAse-free RNase (Boehringer
Mannheim Corp., Indianapolis, Ind.) at 37°C for 1 h.
PCR amplification and cloning. Libraries of 16S rDNA were constructed by PCR amplification of the target genes from DNA extracted from sludge and Hyphomicrobium sp. strain M3, followed by cloning into pCRII (TA cloning kit; Invitrogen, Carlsbad, Calif.). PCR amplification of the 16S rDNA was performed using the eubacterial primers 27f and 1525r or 1492r (24). The PCR mixture consisted of 2 µl of DNA template, 2.5 µl of each primer (5 ng/µl), 10 µl of 10× PCR buffer (GeneAmp PCR reagent kit; Perkin-Elmer Corp., Norwalk, Conn.), 2 µl each of a 200 µM solution of dATP, dCTP, dGTP, and dTTP, and 75 µl of sterile water. Each reaction mixture was overlaid with filter-sterilized mineral oil. The reaction mixtures were heated to 100°C, followed by the addition of 0.5 µl of Taq polymerase (5 U/µl; Gibco BRL, Gaithersburg, Md.). PCR amplification was performed for 38 cycles using 94°C for 1 min, 60°C for 1 min, and 72°C for 1 min. For each set of PCR amplifications, a control reaction without template was performed to check the kit and solution purity.
Amplified DNA was cloned into pCRII plasmid, and the ligation mixture was transformed into OneShot competent Escherichia coli cells (Invitrogen Corp.) by following manufacturer protocols. Colonies containing plasmid inserts were identified by blue/white color selection on Luria-Bertani (LB) plates with X-Gal (5-bromo-4-chloro-3-indolyl--D-galactopyranoside). Alkaline lysis plasmid preparations were made from cultures grown in 100 ml of LB broth with either ampicillin (100 µg/ml) or kanamycin (50 µg/ml). Inserts were verified by restriction digestion of the
plasmids with EcoRI.
DNA sequencing and analysis.
Plasmids containing 16S rDNA
inserts were sequenced by Retrogen, Inc. (San Diego, Calif.), or the
Molecular Biology Resource Facility at the University of Tennessee
(Knoxville) using an ABI PRISM dye terminator cycle sequencing kit with
AmpliTaq DNA polymerase (protocol P/N 402078, revision A)
and an Applied Biosystems 373 DNA sequencer (Perkin-Elmer, Foster City,
Calif.). Greater than 400 bp were sequenced for each insert using
single primer extension with the 1492r primer or 27f primer
(24). Nearly full-length 16S rDNA sequences were obtained
for Hyphomicrobium sp. strain M3, and 18 16S rDNA clones
from the sludge were obtained using the additional primers 27f, 530f,
907r, and 926f (24), M13f (40), and M13r located on the
pCRII plasmid.
|
Whole-cell hybridizations. Whole-cell hybridizations were performed using fluorescent 16S rRNA probes by following published methods (4, 6, 34) with the exceptions that Igepal CA-630 was substituted for Nonidet P-40 and an additional 5-min sonication step in an ice-water bath was used after fixation in paraformaldehyde. Prior to sonication, sludge samples were resuspended in 10 mM EDTA to enhance the permeability of the flocs and probe penetration. Hybridization experiments were performed with 5'-end-labeled fluorescein oligonucleotide probes obtained from Genosys (The Woodlands, Tex.). The control strains used to determine the concentration of formamide needed at 37°C to discriminate zero, one, and two mismatches using probe S-G-Hypho-1241-a-A-19 included H. vulgare ATCC 27500 (no mismatches), Hyphomicrobium sp. strain M3 (no mismatches), Zoogloea ramigera ATCC 19623 (two mismatches), and Nitrobacter winogradskyi ATCC 14123 (one mismatch). The optimum concentration of formamide for hybridization was determined using 20, 30, 40, and 60% (vol/vol) formamide concentrations. Both H. vulgare and Hyphomicrobium sp. strain M3 were positive with all formamide concentrations tested. Z. ramigera was positive at 20 and 30% (vol/vol) formamide and had weak fluorescence at 40 and 60% (vol/vol) formamide. N. winogradskyi was positive at 20 and 30% (vol/vol) formamide and negative at 40 and 60% (vol/vol) formamide. Based on these results, hybridization with this probe was carried out at 40% (vol/vol) formamide at 37°C. Slides were visualized using a Zeiss Axioskop with a Zeiss 100-W epifluorescence illuminator and Zeiss fluorescein isothiocyanate filter 31001. Pictures were obtained with a Zeiss (optronics) ZVS-3C75DE three-chip video camera and a TCX Frame Grabber board or Sony CVP-M3 video printer.
Extraction of RNA from sludge samples and slot blot
hybridizations.
RNA was isolated from approximately 2 g of
sludge solids or 100 ml from pure cultures by following previously
published methods (14). Final RNA pellets were resuspended
in 100 µl of diethyl pyrocarbonate-treated water and stored at
80°C. RNA dot blots were prepared as outlined by Sambrook et al.
(32). RNA amounts on the dot blots were 1 µg and 500, 250, 100, 50, and 10 ng for control samples, and 0.1-ml aliquots were used
for sludge samples. Samples were vacuum blotted onto
0.45-µm-pore-size Biotrans membranes (ICN, Irvine, Calif.) using a
Bio-Rad dot blot apparatus and baked for 1 h at 80°C. The blots
were pretreated for at least 2 h in a hybridization solution
(11). Probes were end labeled with [
-32P]ATP by following the protocol described by Life
Technologies (Gaithersburg, Md.) and purified using Nuc-Trap push
columns (Stratagene, La Jolla, Calif.). Stringency experiments were
conducted with four control strains with zero to six probe mismatches
as shown in Table 2. The optimal hybridization temperature was
determined by hybridizing blots containing the control strains with the
probes at temperatures from 42 to 65°C. Optimal hybridization was
achieved at 50°C for S-S-HyphoC1-648-a-A-20, 52°C for
S-S-HyphoCII-654-a-A-18, and 65°C for S-G-Hypho-1241-a-A-19.
Hybridizations with universal probe 1390 were carried out at 45°C
(38). Blots were hybridized overnight and were washed at the
same hybridization temperature with 1× SSC (0.15 M NaCl plus 0.015 M
sodium citrate)-0.1% sodium dodecyl sulfate solution two times for 15 min each. The RNA levels were quantified using a Storm 840 PhosphorImager (Molecular Dynamics, Sunnyvale, Calif.). The number of
nanograms of RNA hybridizing in sludge samples with each probe was
determined by regression analysis of standard curves (nanograms of
total 16S rRNA versus signal intensity) generated from probe
hybridization with the positive-control strains. The percentage of the
total RNA probing positive in each sample was calculated as
[(nanograms of specific RNA)/(nanograms of universal 16S rRNA)]
· 100. The percentage of the total 16S rRNA probing positive in
sludge samples for each month, except January, was the average of
results from three or four samples.
Nucleotide sequence accession numbers. The 16S rDNA clone sequences were deposited in GenBank at the National Center for Biotechnology Information under accession no. AF097766 to AF097829, and the 16S rDNA sequence from Hyphomicrobium sp. strain M3 received accession no. AF098790.
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RESULTS |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Analysis of 16S rDNA sequences from activated-sludge samples.
Preliminary analysis based on sequences using the 1492r or 27f primers
was performed on 67 16S rDNA clones from three different sludge samples
taken from the same industrial wastewater treatment plant. Individual
clones were given numbers based on the month and year of the sludge
sample and an individual clone number. Clones from the January 1995 sample begin with 195, those from the February 1995 sample begin with
295, and those from the April 1995 sample begin with 495. All clones
were subjected to CHIMERA_CHECK analysis through the Ribosomal Database
Project. Three of the clones were considered potential chimeras and
were not further analyzed. Preliminary work to classify the clones into
known subdivisions was performed using the SIMILARITY_RANK program of
the Ribosomal Database Project and BLAST analysis of all sequences in
GenBank. None of the sequences were identical to previously isolated
sequences. Sequences of the majority of strains exhibited 90 to 95%
similarity to 16S rRNA sequences in GenBank. Four clones had sequences
that were less than 90% similar to published 16S rRNA sequences in GenBank. The sequences of several of these clones showed high similarity to sequences obtained from a phosphate-removing sludge in a
sequencing batch reactor (SBR) (8). These included clones 2951, 2952, 2958, and 29523 from the
Holophaga/Acidobacterium cluster with 95% similarity to
SBR1078 and SBR10103 and 1959 in an undescribed cluster of
Cytophaga with 96% similarity to UNSBR1093. Seven clones
also showed a 95 to 96% sequence homology with beta subdivision clones
from municipal activated sludge (33). Based on these
identifications, the cloned 16S rDNAs were assigned to Eubacteria groups and Proteobacteria subdivisions
(Table 3). In this analysis, population
shifts were seen among the 3 months, with alpha and beta type bacteria
dominating in January and April. The February clones differed from the
January and April clones in that alpha subdivision clones were absent
and the majority of clones belonged to the
Holophaga/Acidobacterium group.
|
Analysis of alpha subdivision clones.
More than 1,300 bp of
16S rDNA sequences was determined for Hyphomicrobium sp.
strain M3 and for 12 clones which fell into the alpha subdivision. A
phylogenetic tree based on evolutionary distance and neighbor joining
was constructed with 950 bp of 16S rDNA sequences from three
Pedomicrobium spp., nine Hyphomicrobium spp., and
three other members of the alpha subdivision (Fig.
1). In this phylogenetic analysis, known
Hyphomicrobium spp. grouped into two clusters as described
by Rainey et al. (30) and the Pedomicrobium spp.
also fell into a distinctive phylogenetic group. The 16S rDNA sequence
from Hyphomicrobium sp. strain M3 was 99.6% similar to that
of Hyphomicrobium denitrificans DSM 1869T in
Hyphomicrobium cluster II. Three of the clone sequences
(4953, 49512, and 49518) also grouped with cluster II and were 96.2 to 96.5% similar to those of Hyphomicrobium sp. strain M3 and
H. denitrificans DSM 1869T. Another two clones
(1951 and 1956) were most closely related to Hyphomicrobium
cluster I containing the H. vulgare species and were 96.6 to
96.8% similar to H. vulgare MC-750. Clones 49519 and 49520 also grouped most closely to the Hyphomicrobium cluster I
(i.e., 94.9 to 95.5% similarity) but were only 94.3 and 95.1% similar
to clones 1951 and 1956, respectively, and may represent a different
cluster within the genus Hyphomicrobium.
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Whole-cell hybridizations in activated sludge. General Hyphomicrobium probe S-G-Hypho-1241-a-A-19 was designed to hybridize to all Hyphomicrobium strains based on sequence alignments between the 495 16S rDNA sequences and that of H. vulgare. This probe was intended to be degenerate in one position (G/C) to include H. vulgare, 49520, 49519, 4953, 49518, and 49512. Isolation and additional sequence analysis indicated that the probe would also hybridize to Hyphomicrobium sp. strain M3 and to the 16S rDNA cloned sequences 1951 and 1956. The specificity of the probe was initially checked using PROBE_CHECK from the Ribosomal Database Project. At that time the only strains which showed no mismatches were H. vulgare and Hyphomicrobium-like organism US-353. A few other alpha subdivision bacteria, including Rhizobium and Sphingomonas species, showed one or two mismatches to the probe. Most of the sludge 16S rDNA sequences from Hyphomicrobium-type clones were complementary to the probe with the G nucleotide, whereas sequences from H. vulgare and Hyphomicrobium-like sp. strain US-353 were complementary to the probe with the C nucleotide.
At this industrial wastewater treatment plant, detection and quantification of Hyphomicrobium levels in the sludge have been based on morphology and a rating system using conventional microscopic techniques. When phase-contrast illumination on either simply stained or Gram-stained slide preparations was used, individual Hyphomicrobium cells were indistinguishable from other microbial cells within the flocs; indeed, only those cells whose hyphae protruded from the floc periphery were readily identifiable as Hyphomicrobium. However, with the aid of the fluorescently labeled S-G-Hypho-1241-a-A-19 probe, clear visualization of Hyphomicrobium embedded within the floc structure by fluorescence microscopic examination was made possible (Fig. 2). Although the distinguishing hypha and tip structure are not seen on the inside of the floc as they are on the outside, the cells had the characteristic ovoid morphology seen at the tips of the cells on the outside of the flocs.
|
Characterization and quantification of Hyphomicrobium
in activated sludge samples.
Based on the sequence alignments used
to create the phylogenetic tree in Fig. 1, probes were designed for
Hyphomicrobium cluster I and cluster II. H. vulgare MC-750 was used as the control strain for cluster I, and
Hyphomicrobium sp. strain M3 was used as the control strain
for cluster II (Table 2). These two probes also had higher numbers of
mismatches to other known alpha subdivision sequences than
S-G-Hypho-1241-a-A-19 and should allow better quantification of
Hyphomicrobium populations (Table 2). RNA was extracted from 48 sludge samples collected from the wastewater treatment plant in
1995. The RNA was hybridized with the general Hyphomicrobium probe (S-G-Hypho-1241-a-A-19) and the cluster I
(S-S-HyphoC1-648-a-A-20) and cluster II (S-S-HyphoCII-654-a-A-18)
probes. The results from these analyses were arithmetically averaged by
monthly periods (Fig. 3). The calculated
values for total Hyphomicrobium-positive 16S rRNA using
probe S-G-Hypho-1241-a-A-1 and for cluster II using probe
S-S-HyphoCII-654-a-A-18 were similar for all months and ranged from 3 to 12% of the total 16S rRNA. The average percentage of
Hyphomicrobium-positive 16S rRNA for the year was 4.8% ± 2.3%. The percentage of cluster I-positive 16S rRNA was lower and
ranged from 0.2 to 1.5% of the total 16S rRNA.
Hyphomicrobium-specific 16S rRNA was present throughout the
year and peaked in March to April 1995. These results are in agreement
with the visual microscopic ratings, which ranged from 2 (some) to 5 (abundant). The highest visual rating for Hyphomicrobium was
in April, with an average score of 4.6.
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Sequence analysis of 16S rRNA is a useful tool for studying the basic microbial community structure of activated sludge. For comparative purposes, ribosomal sequence data from a variety of activated sludge sources would be beneficial. In addition, to determine how operational parameters affect microbial communities, information on the plant operating conditions, as well as on sludge and effluent quality at the time of sampling is also needed. In this study, some plant operation data are provided (Table 1), although it is still unclear how process changes between January, February, and April affected the microbial community structure. The February sample was taken during a time period in which the sludge was experiencing poor dewatering due to zoogloeal infestation (23), whereas in April sludge dewatering had returned to normal conditions.
Libraries of 16S rDNA were constructed from three samples to relate potential shifts in the microbial populations to changes in operational parameters. While none of the clones were completely homologous to sequences found in GenBank, the majority of January (195) and April (495) clone sequences could be classified into well-known groups. Due to the increasing number of 16S rDNA sequences in GenBank, comparisons among diverse activated-sludge communities could be made. As found in other activated-sludge samples (8, 33, 37), the January and April samples had a large number of beta subdivision clones whose sequences had high similarity to published sequences. The microbial community in this same industrial sludge also had an abundance of alpha subdivision 16S rDNA sequences (16 of 64). Seven of these sequences were similar to the hyphal budding bacterium Hyphomicrobium or Pedomicrobium. Also of interest was the high similarity (96%) of the 49531 and 1959 clones to UNSBR1093 from a phosphate-removing bioreactor (8). These sequences have low similarity to other sequences in GenBank and may represent undescribed organisms in the activated sludge. Fourteen clones from February were most closely related to clones of soil bacteria (21, 25) which aggregate into cluster C of the recently described phylum Holophaga/Acidobacterium (25). Organisms belonging to this phylum appear to be widely distributed in soils, sediment, and activated sludge. The organisms in cluster C of this phylum have not been cultured, and their metabolic function in the activated sludge is unclear at this time.
The potential for biases in the construction of 16S rDNA libraries includes inefficient cell lysis, DNA recovery, and PCR amplification and cloning (13, 35). Therefore, the 16S rDNA sequences obtained in this present study may represent some of the members in the community but may not be all-inclusive or reflect the frequency of individuals in the community. For example, the February 16S rDNA sequences were very different from those sequences obtained from the January and April samples. It is unclear whether the 16S rDNA sequences obtained in the library from the February sample reflect changes in the microbial community during that time period or whether they are simply the result of bias in nucleic acid extraction efficiency and library formation. The fact that Hyphomicrobium type clones were not isolated in the February sludge library, even though the sludge samples probed positive with the general Hyphomicrobium probe (S-G-Hypho-1241-a-A-1) and the cluster II probe (S-S-HyphoCII-654-a-A-18) at approximately 3%, suggests that this library may be biased.
The presence of Hyphomicrobium in this industrial activated sludge was corroborated by its morphology using light microscopy, by isolation and identification of Hyphomicrobium sp. strain M3, and genetically by probe analysis. Hyphomicrobium sp. strain M3 was isolated from this sludge over 10 years, prior to the inception of this study (Meyers and Meyers, Abstr. 86th Annu. Meet. Am. Soc. Microbiol. 1986). Surprisingly, Hyphomicrobium sp. strain M3 showed a slightly higher sequence similarity to the cultured strain H. denitrificans DSM 1869T than to Hyphomicrobium clone isolates (99 versus 96%). This may represent a slight shift in the 16S rDNA sequence of the Hyphomicrobium cluster II population in this sludge or a bias that resulted from isolation and cultivation of Hyphomicrobium sp. strain M3. The placement of several cloned library sequences and strain M3 in the Hyphomicrobium cluster II was verified by phylogenetic analysis of 16S rDNA sequences. These analyses were in good agreement with the taxonomic placement of Pedomicrobium and Hyphomicrobium (12). Physiologically and morphologically, strain M3 belongs in the Hyphomicrobium genus because of its ability to utilize the C1 compounds (e.g., methanol and methylamine) and the formation of the characteristic mother cells with hyphae (18). Monopolar hyphal budding is the major morphological feature distinguishing Hyphomicrobium and Pedomicrobium (18). Hyphomicrobium sp. strain M3 has been shown to utilize monopolar budding as its mode of reproduction, based on light microscopic examination of pure cultures of the organism, thereby sustaining its genus identification.
A probe created to detect the Hyphomicrobium group was designed using the 16S rDNA library and was used to detect Hyphomicrobium spp. in activated-sludge samples. The distinctive morphology of Hyphomicrobium spp. allowed for the verification of the S-G-Hypho-1241-a-A-19 probe by whole-cell hybridization with activated sludge from the wastewater treatment plant. The fluorescein-labeled S-G-Hypho-1241-a-A-19 probe in conjunction with fluorescence microscopy provided a better resolution of Hyphomicrobium both within the structure and around the periphery of the flocs than that obtained with conventional light microscopy.
In this study, three cloned Hyphomicrobium sequences (4953, 49512, and 49518) fit into Hyphomicrobium cluster II with a high level of confidence. For purposes of probe creation, two other pairs of clones (1951 and 1956, and 49519 and 49520) were grouped with Hyphomicrobium cluster I although they were only 94 to 95% similar to each other and may represent different clusters within Hyphomicrobium. The grouping of these sequences together allowed H. vulgare MC-750 to be used as a control strain for hybridization studies. Hybridizations of samples taken from almost 1 year of operation indicated that the Hyphomicrobium cluster II organisms dominated over the Hyphomicrobium cluster I organisms by 3- to 12-fold. Holm et al. (17) suggest that morphologically and nutritionally similar isolates of Hyphomicrobium may show a high level of genetic diversity and that Hyphomicrobium populations may vary seasonally. In this wastewater treatment plant the Hyphomicrobium populations showed genetic diversity but did not vary seasonally. Gliesche and Fesefeldt (15) also did not see seasonal variations in Hyphomicrobium DNA/DNA hybridization group HG 27 in activated sludge. It would be interesting to determine whether plant upsets change the dominant Hyphomicrobium population.
In addition to validation of the molecular analysis methods by using more-conventional cultivation methods and microscopy, comparative information across a wide spectrum of activated sludge plants and operating conditions is needed. The isolation and cultivation of bacterial strains from sludge provide information on the roles and the niches of particular organisms in the community, whereas community analysis and probing methods are valuable in verifying that the isolated organisms are actually present in significant numbers within the community.
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ACKNOWLEDGMENTS |
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This work was supported by a research grant from Eastman Chemical Company, Tennessee Eastman Division, Kingsport, and WMREI at the University of Tennessee.
We gratefully acknowledge the assistance of Claudia Werner for cultivation of strains and Neil Quigley at the Molecular Biology Resource Facility (UTK) for 16S rDNA sequencing. Thanks also to Janet Hensley (Eastman Chemical Company) for preparation of media and successful resuscitation of Hyphomicrobium sp. strain M3 cultures.
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FOOTNOTES |
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* Corresponding author. Mailing address: Center for Environmental Biotechnology, The University of Tennessee, 676 Dabney Hall, Knoxville, TN 37996. Phone: (865) 974-8080. Fax: (865) 974-8086. E-mail: alayton{at}utk.edu.
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REFERENCES |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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