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APPLIED AND ENVIRONMENTAL MICROBIOLOGY,
0099-2240/97/$04.0010
Mar. 1997, p. 1107–1117 Vol. 63, No. 3
Copyright q 1997, American Society for Microbiology
Group-Specific Small-Subunit rRNA Hybridization Probes To
Characterize Filamentous Foaming in Activated Sludge Systems
FRANCIS L. DE LOS REYES, WOLFGANG RITTER,† AND LUTGARDE RASKIN*
Environmental Engineering and Science, Department of Civil Engineering,
University of Illinois at Urbana-Champaign, Urbana, Illinois 61801
Received 24 October 1996/Accepted 23 December 1996
Foaming in activated sludge systems is characterized by the formation of a thick, chocolate brown-colored
scum that floats on the surface of aeration basins and secondary clarifiers. These viscous foams have been
associated with the presence of filamentous mycolic acid-containing actinomycetes. To aid in evaluating the
microbial representation in foam, we developed and characterized group-, genus-, and species-specific oligo-
nucleotide probes targeting the small subunit rRNA of the Mycobacterium complex, Gordona spp., and Gordona
(Nocardia) amarae, respectively. The use of a universal base analog, 5-nitroindole, in oligonucleotide probe
design was evaluated by comparing the characteristics of two different versions of the Mycobacterium complex
probe. The temperature of dissociation of each probe was determined. Probe specificity studies with a diverse
collection of 67 target and nontarget rRNAs demonstrated the specificity of the probes to the target groups.
Whole-cell hybridizations with fluorescein- and rhodamine-labeled probes were performed with pure cultures
of various members of the Mycobacterium complex as well as with environmental samples from a full-scale
activated sludge plant which experienced foaming. Quantitative membrane hybridizations with activated
sludge and anaerobic digester foam showed that 15.0 to 18.3% of the total small-subunit rRNAs could be
attributed to members of the Mycobacterium complex, of which a vast majority consisted of Gordona rRNA.
Several G. amarae strains made up only a very small percentage of the Gordona strains present. We demon-
strated that group-specific rRNA probes are useful tools for the in situ monitoring and identification of
filamentous bacteria in activated sludge systems.
The success of the activated sludge process in treating in-
dustrial and domestic wastewater hinges crucially on the effi-
cient separation of microbial biomass (i.e., activated sludge or
mixed liquor) from the effluent stream. Filamentous foaming,
which manifests itself as a “viscous, stable, chocolate-colored”
scum layer (17, 41) on the surface of activated sludge aeration
basins and secondary clarifiers, prevents this separation, thus
reducing effluent quality. In addition, foaming may represent a
public health concern because of the possible spread of patho-
gens in windblown scum (8). Foaming problems are wide-
spread: recent surveys reported that 66% of 114 U.S. activated
sludge plants had been affected by foaming (29), and 51% of
129 activated sludge plants in eastern Australia had a foam
problem at the time of the survey (37). In addition, foaming
problems have been reported for activated sludge plants in
France, South Africa, Japan, Germany, Switzerland, and En-
gland (10, 17, 41).
Foaming has been associated with the presence of mycolic
acid-containing actinomycetes belonging to the family Nocar-
diaceae, as well as other filamentous organisms. In particular,
the actinomycete Gordona amarae (formerly Nocardia amarae
[18, 34]) often has been implicated by microscopic and physi-
ological analyses as the dominant species in foam (21). This
has led to the term “Nocardia foam” or “nocardioform foam,”
despite the isolation from foam of several genera, including
Nocardia, Rhodococcus, Gordona, Tsukamurella, and Mycobac-
terium, as well as other gram-positive bacteria such as Micro-
thrix parvicella (9, 38–40).
The taxonomy of mycolic acid-containing actinomycetes has
undergone tremendous changes in the light of modern phylo-
genetic methods (12, 36). At present, the actinomycetes that
have cell walls of chemotype IV and contain mycolic acids are
classified in the Mycobacterium complex, a suprageneric group
that forms a distinct phyletic line (34) and contains the genera
Corynebacterium, Tsukamurella, Mycobacterium, Gordona, Rho-
dococcus, and Nocardia (24, 34). The name “enlarged Coryne-
bacterium-Mycobacterium-Nocardia (CMN)” has also been
used for this group (31, 34).
Since many of these filamentous actinomycetes are morpho-
logically similar, reliance on traditional identification and enu-
meration techniques has contributed to the difficulty in pre-
scribing solutions to foaming problems. Discrepancies between
results obtained with culture-dependent methods and total di-
rect microscopic counts for aquatic environments have long
been recognized (45). More recently, biases introduced by tra-
ditional methods have been pointed out in the context of re-
lating operational problems in activated sludge systems to spe-
cific causative microbial populations (25, 41, 48). Identification
and classification based on cultivation and morphology are
especially challenging for actinomycetes because of their low
growth rates, complex life cycles, and spore patterns (42).
Hernandez et al. (16) moved beyond morphological and
culture-based methods for identifying filamentous actinomyce-
tes by using fluorescently labeled antibody probes to identify
Gordona (Nocardia) strains in activated sludge and anaerobic
digesters. While this immunofluorescent technique is an im-
provement over conventional methods, the production of an-
tibodies still requires cultivation of target microorganisms. In
contrast, the use of rRNA-targeted oligonucleotide hybridiza-
tion probes allows for the identification and enumeration of
* Corresponding author. Mailing address: Department of Civil En-
gineering, University of Illinois at Urbana-Champaign, 3221 Newmark
Civil Engineering Laboratory, 205 N. Mathews, Urbana, IL 61801.
Phone: (217) 333-6964. Fax: (217) 333-6968 or -9464. E-mail: lraskin
@uiuc.edu.
† Present address: Dresdner Str. 69, 80993 Munich, Germany.
1107
microorganisms in complex environments without prior culti-
vation (14). Another advantage of oligonucleotide probes is
their ability to target organisms at different phylogenetic levels.
Thus, by using such probes, it may become possible to resolve
conflicting reports (40, 41) about the causative organisms of
nocardioform foaming at the genus or species level. The iden-
tification and in situ detection of microbial populations in
activated sludge with fluorescently labeled oligonucleotide
probes have been reported previously for the subclasses of the
proteobacteria (48), gram-negative filamentous bacteria (47),
Acinetobacter species (49), ammonia-oxidizing bacteria (27, 50,
51), and nitrite-oxidizing bacteria (27, 51).
In this study, we report on the development and character-
ization of group-, genus-, and species-specific small subunit
(SSU) rRNA-targeted oligonucleotide probes for the CMN
group and demonstrate the usefulness of these probes in char-
acterizing filamentous foaming in full-scale activated sludge
systems.
MATERIALS AND METHODS
Organisms, culture conditions, and nucleic acid extraction. The organisms
used in this study are listed in Table 1. Most organisms were obtained from the
American Type Culture Collection (ATCC [Rockville, Md.]). Additional strains
were obtained from various collections at the University of Illinois, Urbana, and
the University of California, Berkeley. Proteobacteria were grown in liquid
tryptone-yeast extract medium (10 g of Bacto-tryptone,5gofyeast extract, and
7 g of NaCl per liter [pH 7.0]). Gram-positive bacteria with a high DNA G1C
content were grown in TGY medium (5 g of Bacto-tryptone,1gofglucose, 3 g
of yeast extract per liter [pH 7.0]) or yeast-glucose medium (5 g of glucose and
3 g of yeast extract per liter [pH 7.0]). Other strains were grown as recommended
by ATCC. The organisms were grown to mid-log phase, and nucleic acids were
extracted from cell pellets by a phenol-chloroform-isoamyl alcohol extraction
procedure (44). The concentrations of the nucleic acids were measured spectro-
photometrically, assuming that 1 mg of RNA per ml is equal to 20 optical density
units at A
260
. The quality of extracted RNA was evaluated by polyacrylamide gel
electrophoresis (1). For in situ hybridizations, cells were harvested in mid-log
phase and fixed as described below.
Environmental samples. Grab samples were taken from the foam and mixed
liquor in the aeration basins and from anaerobic digester foam at the Urbana-
Champaign Sanitary District Northeast Treatment Plant (Urbana, Ill.). For
membrane hybridizations, 50-ml samples were collected and immediately trans-
ported to the laboratory for processing and storage. Nucleic acids were extracted
by a phenol-chloroform-isoamyl alcohol extraction procedure (44). For fluores-
cent in situ hybridizations (FISH), samples were immediately fixed in absolute
ethanol (final concentration, 50% [vol/vol]) or in 4% paraformaldehyde (PFA) as
described below.
Oligonucleotide probes. Oligonucleotide probes were designed by comparing
the SSU rRNA sequences of 32 organisms of the Mycobacterium complex, which
were obtained from the Ribosomal Database Project (RDP) (24). The oligonu-
cleotide probes used, their target groups, and the SSU rRNA target sites are
listed in Table 2. In addition, a universal probe, S-*-Univ-1390-a-A-18 (53),
complementary to the SSU rRNA of most known organisms, and probe S-D-
Bact-0338-a-A-18 (4), complementary to the SSU rRNA of bacteria, were used.
The probes were synthesized on a DNA synthesizer (Applied Biosystems, Foster
City, Calif.) at the University of Illinois Biotechnology Center’s Genetic Engi-
neering Facility. A universal base analog, 5-nitroindole (N
5
; Glen Research,
Sterling, Va.) (22), was incorporated during the synthesis of one of the probes
(S-*-Myb-0736-b-A-22). Oligonucleotides used for FISH were synthesized with
an aminolinker at the 59 end (Aminolink 2; Applied Biosystems) and were
coupled to tetramethylrhodamine isothiocyanate (TRITC) or fluorescein isothio-
cyanate (FITC) (Molecular Probes, Eugene, Oreg.) as previously described (5,
30). A reaction mixture was set up with the following ingredients: 350 mlof
oligonucleotide (approximately 200 mg of DNA), 120 ml of sodium carbonate
buffer at pH 9.2, 75 ml of fluorochrome dissolved in dimethylformamide (10
mg/ml), and 55 ml of double-distilled H
2
O. The reaction mixture was incubated
in the dark at room temperature for 12 to 16 h. Free fluorochrome was separated
from the oligonucleotide by running the mixture through a Sephadex G-25
DNA-grade NAP10 column (Pharmacia Biotech, Piscataway, N.J.). Unlabeled
oligonucleotide was separated from labeled oligonucleotide with an oligonucle-
otide purification cartridge (OPC; Applied Biosystems) according to the manu-
facturer’s protocol. Purified probes were stored in the dark at 2208C. Oligonu-
cleotides used for membrane hybridizations were purified with the OPC and 59
end labeled with
32
P with T4 polynucleotide kinase (Promega Corporation,
Madison, Wis.) and [g-
32
P]ATP (ICN Radiochemicals, Irvine, Calif.) (32).
Cell fixation, FISH, and microscopy. To determine the best fixation method
for the mycolic acid-containing actinomycetes, the following fixation treatments
were evaluated: 4% PFA for 1 min, 4% PFA for 2 h, 50% absolute ethanol for
2 h, and 50% absolute ethanol for 12 to 16 h. Three volumes of fresh (prepared
within 24 h of use) PFA fixative (33) or 50% absolute ethanol was applied to
cells. After fixation, samples were washed with phosphate-buffered saline (PBS)
buffer (130 mM NaCl, 10 mM sodium phosphate [pH 7.2]) and resuspended in
1:1 (vol/vol) PBS-ethanol. Fixed cells were stored at 2208C. Teflon-coated slides
(Cel-Line Associates, Newfield, N.J.) were cleaned in ethanolic KOH (10%
KOH in ethanol) and coated with gelatin [0.1% gelatin, 0.01% CrK(SO
4
)
2
z
12H
2
O]. Fixed cells were applied to the wells of the slide. The slides were
air-dried and dehydrated by serial immersion in 50, 80, and 96% ethanol (3 min
each). Subsequently, 9 ml of hybridization solution was mixed with 1 ml (25 ng)
of FITC- or TRITC-labeled probe and applied to each well. The slides were
incubated at 378C in a moisture chamber (5) for at least 2 h. After hybridization,
the slides were rinsed once with prewarmed (378C) wash solution and then
washed for 20 min at 378C. The hybridization and wash solution consisted of
0.1% sodium dodecyl sulfate (SDS), 20 mM Tris-HCl, and X M NaCl (where X
varied depending on the probe, as described below). The slides were dipped in
a cold solution of 49,6-diamidino-2-phenylindole (DAPI; Sigma, St. Louis, Mo.)
and then rinsed with cold wash solution corresponding to 0% formamide (FA)
(0.9 M NaCl in wash solution). The DAPI solution consisted of 0.1 M Tris-HCl
(pH 7.2), 0.9 M NaCl, and DAPI (final concentration of 6.26 mg/ml). The slides
were mounted in Citifluor (Citifluor Ltd., London, United Kingdom) and visu-
alized with an epifluorescence microscope (Axioskop; Carl Zeiss, Germany)
fitted with filter sets (Chroma Tech. Corp., Brattleboro, Vt.) for TRITC (filter set
41002), FITC (filter set 41001), and DAPI (filter set 31000) and a Zeiss 50-W
high-pressure bulb. Images were captured with a liquid-cooled charge coupled
device (CCD) camera (Photometrics MXC200L, class 2 Kodak KAF 1400 CCD,
1,317-by-1,035 pixel array, 6.8-mm pixel size; Photometrics Ltd., Tucson, Az.)
controlled by a PowerPC 7100 (Apple Computer, Inc., Cupertino, Calif.). Ex-
posure times were varied from 0.05 to 0.2 s for epifluorescence and phase-
contrast images. To provide a comparative measure of fluorescence, a constant
exposure time of 0.1 s was used for image capture to compare fixation treat-
ments. Images were imported to Adobe Photoshop 3.0 (Adobe, Seattle, Wash.)
for printing.
Probe-conferred fluorescence was quantified with IPLab Spectrum image
analysis software (Signal Analytics, Vienna, Va.), with semiautomatic image
thresholding. Background signals were similarly quantified and subtracted from
the mean pixel value for each image. For quantification, at least 100 cells for each
hybridization and wash condition were used. The optimal hybridization strin-
gency for the different probes was determined by quantification of probe-con-
ferred fluorescence to target and nontarget reference cells with different NaCl
concentrations in the hybridization buffer and wash solution (50).
Determination of optimal wash temperatures. Organisms with zero, one, or
two mismatches in the probe target region of their SSU rRNA, as determined
from the RDP database, were used in temperature of dissociation (T
d
) studies.
Nucleic acid samples (30 ng) were slot blotted in triplicate to nylon membranes
(Magna Charge; Micron Separation, Inc., Westboro, Mass.). Baked membranes
were prehybridized for2hat408C and hybridized overnight at 408C (32).
Subsequently, the membranes were washed in 100 ml of 1% SDS–13 SSC (0.15
M NaCl, 0.015 M sodium citrate) twice for1hat408C. The membranes were
then cut into separate individual hybridized blots. Each blot was washed in 3 ml
of 1% SDS–13 SSC for 10 min at the first temperature (308C). The blot was then
removed and transferred to a new scintillation vial with a wash solution at the
next higher temperature, and washes were continued at increasing temperatures
until a temperature of 808C was reached (total of 16 temperatures). The amount
of probe released at each wash temperature was quantified by liquid scintillation
counting with a model 1600CA liquid scintillation analyzer (Packard Instru-
ments, Downers Grove, Ill.).
Probe specificity studies. The specificity of the oligonucleotide probes was
examined by membrane hybridizations (32) with nucleic acids from 67 organisms,
representing a broad spectrum of phylogenetic diversity (Table 1). The previ-
ously determined T
d
for each probe was used as the final wash temperature for
the membranes. Hybridization signals were quantified by storage phosphor anal-
ysis with a 400 series PhosphorImager and ImageQuant software package (Mo-
lecular Dynamics, Sunnyvale, Calif.).
Quantitative membrane hybridization. The oligonucleotide probes were used
to quantify the abundances of the microbial groups (group, genus, and species)
in activated sludge and anaerobic digester systems. Denatured RNA samples and
dilution series of pure culture G. amarae SE102 RNA were applied in triplicate
to Magna Charge nylon membranes (33). These membranes were hybridized
with universal and specific probes, and the resulting hybridization responses were
used to determine the relative concentration of target SSU rRNA in the samples
(33, 53).
RESULTS AND DISCUSSION
Probe design. The design of the hybridization probes was
based on the phylogeny of the mycolic acid-containing actino-
mycetes inferred by SSU rRNA sequence analysis from RDP.
The mycolic acid-containing actinomycetes under the Myco-
bacterium complex are divided into members of the families
1108 DE LOS REYES ET AL. APPL.ENVIRON.MICROBIOL.
Nocardiaceae (genera Gordona, Nocardia, Rhodococcus, Tsuka-
murella, and Corynebacterium [15]) and Mycobacteriaceae (ge-
nus Mycobacterium). A phylogenetic tree with representative
organisms from each genus is shown in Fig. 1. Since many
studies have implicated G. amarae as the dominant foam-
causing organism (17, 21, 41), we developed a probe targeting
a G. amarae-specific region of the SSU rRNA. In addition, we
designed a genus-specific probe for Gordona and two Myco-
bacterium complex-specific probes. The groups of actinomyce-
tes targeted by these probes are shown in Fig. 1, and probe
sequences are given in Fig. 2. Probe design also involved
a check of SSU rRNA sequences for nontarget group com-
plementarity, with the CHECK_PROBE program provided
by RDP (24) and the Basic Local Alignment Search Tool
(BLAST) network service (3).
The Mycobacterium complex probe (S-*-Myb-0736-a-A-22),
targeting the SSU rRNA of Gordona, Rhodococcus, Nocardia,
Tsukamurella, Corynebacterium, and Mycobacterium, perfectly
matches all Gordona spp. This probe has one A:C mismatch
with the target sequences of 9 Rhodococcus spp., 3 Corynebac-
terium spp., and 78 Mycobacterium spp. and one A:A mismatch
with the target sequences of 3 Nocardia spp. at position 743
(Escherichia coli numbering). Target species with two mis-
matches in their target sequences include 1 Rhodococcus sp., 1
Corynebacterium sp., 2 Tsukamurella spp., and 11 Mycobacte-
rium spp. For all of these organisms, one of the two mis-
matches occurs in position 743. All nontarget species (non-
members of the Mycobacterium complex) have at least two
mismatches with the probe. We designed a new probe (S-*-
Myb-0736-b-A-22) by incorporating a new universal base ana-
log, N
5
, at position 743 in order to reduce the effect of this
mismatch. Loakes and Brown (22) showed that N
5
was supe-
rior to other base analogs in providing higher duplex stability
and behaving indiscriminately towards four of the five natural
bases. This new probe was characterized along with the origi-
nal probe to evaluate the applicability of N
5
for improving
probe design. Thus, probe S-*-Myb-0736-b-A-22 perfectly
matches most organisms of the Mycobacterium complex and
has only one mismatch with a few species in this group.
The Gordona-specific probe S-G-Gor-0596-a-A-22 perfectly
matches most Gordona spp. (including G. amarae), but it has
one G:G mismatch with G. terrae (ATCC 25594) at position
613 (E. coli numbering) and a possible mismatch at position
605 (base not reported for this position) with G. bronchialis.
Nontarget organisms have at least two mismatches in their
target sequence with this probe.
Probe S-S-G.am-0192-a-A-18 was designed to be species
specific for G. amarae. At the time of probe design, only one
strain of G. amarae, SE6, was available in the database. The
probe has no mismatches with G. amarae SE6, and nontarget
organisms have at least two mismatches (e.g., Tsukamurella
paurometabolum).
Optimization of wash temperatures. The use of oligonucle-
otide probes for accurate identification and quantification of
target organisms relies on the specific duplex formation be-
tween probe and target rRNA. Especially in complex environ-
ments such as activated sludge, this specificity must be ensured
not only by comparison with existing rRNA sequence data-
bases, but also by optimization of experimental hybridization
conditions. An important parameter in this regard is the T
d
,
defined as the temperature at which 50% of the duplex remains
intact during a specified washing period (46). Use of the ex-
perimentally determined T
d
for perfect duplexes (perfect
match between probe and target) as the posthybridization
wash temperature generally ensures that duplexes with one or
more mismatches are dissociated (51). Table 2 provides a sum-
TABLE 1. List of organisms used in the specificity study
Position
a
Organism
A1 ................................... Rattus norvegicus
A2 ................................... Dictyostelium discoideum
A3 ................................... Methanobrevibacter sp.
A4 ................................... Methanogenium cariaci
A5 ................................... Methanosarcina acetivorans
A6 ................................... Chloroflexus aurantiacus ATCC 29366
A7 ................................... Cytophaga lytica
A8 ................................... Weeksella virosa ATCC 43766
A9 ................................... Flavobacterium ferrugineum
A10 ................................. Bacteroides thetaiotaomicron
A11 ................................. Azospirillum brasilense ATCC 29145
A12 ................................. Caulobacter sp. strain MCS10
A13 ................................. Rhizobium meliloti
A14 ................................. Chromobacterium violaceum ATCC 12472
A15 ................................. Alcaligenes eutrophus
A16 ................................. Alcaligenes faecalis ATCC 8750
A17 ................................. Oxalobacter formigenes
A18 ................................. Nitrosomonas sp.
A19 ................................. Chromatium vinosum
A20 ................................. Legionella pneumophila
A21 ................................. Oceanospirillum linum
A22 ................................. Pseudomonas aeruginosa
B1.................................... Vibrio harveyi
B2.................................... Aeromonas hydrophila
B3.................................... Escherichia coli K-12
B4.................................... Proteus vulgaris
B5.................................... Desulfovibrio desulfuricans
B6.................................... Megasphaera elsdenii
B7.................................... Sporohalobacter lortetii
B8.................................... Mycoplasma neurolyticum
B9.................................... Lactobacillus casei
B10.................................. Listeria monocytogenes
B11.................................. Staphylococcus aureus
B12.................................. Bacillus subtilis
B13.................................. Brevibacterium linens ATCC 9172
B14.................................. Micrococcus luteus ATCC 4698
B15.................................. Micrococcus roseus ATCC 534
B16.................................. Amycolata autotrophica ATCC 19727
B17.................................. Saccharopolyspora rectivirgula ATCC 33515
B18.................................. Mycobacterium komossense ATCC 33013
B19.................................. Mycobacterium fortuitum ATCC 6841
B20.................................. Mycobacterium smegmatis ATCC 19420
B21.................................. Mycobacterium vaccae ATCC 15483
B22.................................. Tsukamurella paurometabolum ATCC 8368
C1.................................... Corynebacterium xerosis ATCC 373
C2.................................... Corynebacterium renale ATCC 19412
C3.................................... Nocardia asteroides
C4.................................... Nocardia otitidiscaviarum ATCC 14629
C5.................................... Nocardia camea ATCC 6847
C6.................................... Nocardia paratuberculosis ATCC 23826
C7.................................... Nocardia transvalensis ATCC 6865
C8.................................... Nocardia nova ATCC 33726
C9.................................... Nocardia brasiliensis
C10.................................. Dietzia maris ATCC 35013
C11.................................. Rhodococcus rhodochrous ATCC 13808
C12.................................. Rhodococcus equi
C13.................................. Rhodococcus rhodnii ATCC 35071
C14.................................. Rhodococcus coprophilus ATCC 29080
C15.................................. Gordona terrae ATCC 25594
C16.................................. Gordona rubropertinctus ATCC 14352
C17.................................. Gordona aichiensis ATCC 66611
C18.................................. Gordona sputi ATCC 29627
C19.................................. Gordona amarae SE149B
C20.................................. Gordona amarae NM23
C21.................................. Gordona amarae SE102
C22.................................. Gordona amarae SE6 ATCC 27808
C23.................................. Gordona amarae RBI
a
Location on the hybridization membrane (Fig. 3).
VOL. 63, 1997 rRNA HYBRIDIZATION PROBES FOR ACTIVATED SLUDGE FOAMING 1109
mary of the experimentally determined T
d
values for the four
probes characterized in this study.
Probe S-*-Myb-0736-a-A-22 has a T
d
value of 55 to 578C for
three G. amarae strains, SE149B, SE102, and SE6 (Fig. 2a). G.
amarae NM23 has a lower T
d
of 51.58C. This lower T
d
suggests
a possible mismatch in the target region. We currently are
investigating this possibility by SSU rDNA sequencing of the
different strains of G. amarae. As expected, T. pauro-
metabolum, which has two mismatches with this probe, has a
lower T
d
of 488C. The relatively large spread in T
d
values
decreased for the N
5
-substituted probe, S-*-Myb-0736-b-A-22
(Fig. 2b). The three strains of G. amarae have T
d
values of 50.5
to 51.58C. The T
d
for G. amarae NM23 is still slightly lower at
498C, while T. paurometabolum has the lowest T
d
at 488C.
Thus, substitution with N
5
improved the probe by decreasing
the spread in T
d
values from 98C (57 to 488C) to 3.58C (51.5 to
488C). Furthermore, substitution with N
5
decreased the T
d
values for the organisms with no mismatches in the target
region. These results are consistent with previous studies (22)
and ongoing studies in our laboratory which have demon-
strated the effectiveness of N
5
as a “universal” nucleotide in
probe and primer design (28).
Probe S-G-Gor-0596-a-A-22 showed excellent potential as a
genus-specific probe, with a very small T
d
spread of 0.48C (54.2
to 53.88C). G. terrae (ATCC 25594), which has one mismatch
with this probe at position 613 (based on RDP), has T
d
values
similar to those of the other Gordona species. The species-
specific probe S-S-G.am-0192-a-A-18 exhibits two distinct T
d
ranges. Two G. amarae strains, SE6 and SE102, have high T
d
values (65 to 678C), while two other strains, SE149B and
FIG. 1. Phylogenetic tree showing representative mycolic acid-containing actinomycetes, inferred from comparison of the SSU rRNA sequences. The tree was
constructed by the neighbor-joining method (35). The bar at the bottom represents 5 estimated changes per 100 nucleotides. The oligonucleotide probes are shown
with their respective target groups.
TABLE 2. Probe names, target groups, target sites, optimized experimental hybridization conditions,
and signal intensities of nontarget signals
Probe
c
Target group
Target site
(SSU rRNA positions
[E. coli numbering])
Exptl T
d
(8C)
% Formamide
in FISH
Avg nontarget signal
intensity (% of
target signal
intensity)
a
Range (% of
target signal
intensity)
b
S-*-Myb-0736-a-A-22 Mycobacterium complex 0736–0757 56 30 0.15 0.08–0.56
S-*-Myb-0736-b-A-22 Mycobacterium complex 0736–0757 51 30 0.44 0.17–0.94
S-G-Gor-0596-a-A-22 Gordona 0596–0617 54 20 0.56 0.12–5.60
S-S-G.am-0192-a-A-18 G. amarae 0192–0209 66 30 0.83 0.29–5.34
a
Average nontarget signal intensity, % 5 100 3 (average signal intensity for nontarget)/(average signal intensity for target).
b
Lower range, % 5 100 3 (lowest signal intensity for nontarget)/(average signal intensity for target); upper range, % 5 100 3 (highest signal intensity for
nontarget)/(average signal intensity for target).
c
Probe nomenclature has been standardized according to the oligonucleotide probe database (2).
1110 DE LOS REYES ET AL. APPL.ENVIRON.MICROBIOL.
FIG. 2. T
d
studies for probes S-*-Myb-0736-a-A-22, S-*-Myb-0736-b-A-22, S-G-Gor-0596-a-A-22, and S-S-G.am-0192-a-A-18. Adjacent to the probe dissociation
results are SSU rRNA sequences of target and nontarget species and probe sequences. The top SSU rRNA sequences for each list of organisms are the target sequences.
Periods in the succeeding sequences signify identical nucleotides, and replacement nucleotides indicate differences in the target sequences at the respective positions.
Nocardia otitidis., N. otitidiscaviarum.
VOL. 63, 1997 rRNA HYBRIDIZATION PROBES FOR ACTIVATED SLUDGE FOAMING 1111
NM23, as well as T. paurometabolum, have T
d
values ranging
from 48 to 52.58C. These results suggest that G. amarae
SE149B and NM23 have one or more mismatches with probe
S-S-G.am-0192-a-A-18. We currently are evaluating this by
sequencing the SSU rDNA of the various G. amarae strains.
Specificity studies. Table 1 lists the organisms that were
used in the specificity study in the order that they were applied
to the hybridization membranes, and Fig. 3a shows a schematic
representation of the same membrane template with phyloge-
netic group names. Figure 3b presents the hybridization re-
sponse with probe S-*-Univ-1390-a-A-18. Since this probe tar-
gets almost all known organisms, all 67 RNAs resulted in a
positive hybridization response. The lower response for some
positions can be attributed to smaller amounts of RNA applied
to the membranes. Hybridization with probe S-*-Myb-0736-a-
A-22 resulted in a positive signal for all members of the My-
cobacterium complex, except for Nocardia paratuberculosis
(C6) and Nocardia brasiliensis (C9) (Fig. 3c). Additional hy-
bridizations showed that both organisms bind weakly to this
probe (data not shown). However, the number of mismatches
of the sequence of N. paratuberculosis to the probe cannot be
determined, because the SSU rRNA sequence is not available.
For N. brasiliensis, analysis of its sequence shows that it has one
A:A mismatch at position 743. The N
5
-substituted probe (Fig.
3d) resulted in a more uniform response for all members of the
Mycobacterium complex compared to the original version of
the probe. Thus, the positive effect of using a universal base
analog in the degenerate base position was confirmed in this
study. The genus-specific probe (Fig. 3e) was specific for all
Gordona spp., while the species-specific probe (Fig. 3f) tar-
geted only G. amarae SE102 and SE6. These results confirmed
results obtained during the T
d
studies. In general, the four
probes hybridized strongly to nucleic acids from target organ-
isms but not to those of nontarget organisms. Quantification of
hybridization signals with a PhosphorImager showed that the
highest response for nontarget SSU rRNA was 5.6% of the
response for target SSU rRNA. On average, the nontarget
response was only 0.5% of the target response (Table 2).
Effects of cell fixation. A crucial issue in FISH is the fixation
of cell samples. The ideal fixative should not only preserve
cellular morphology, but should also maximize the diffusion of
probe throughout the cytoplasmic matrix. The most common
fixation methods for FISH (i.e., treatment with PFA or with
ethanol and formaldehyde) have been found to be effective for
a wide variety of cells. However, it has been reported that many
gram-positive bacteria may be more difficult to permeabilize
with these common fixatives (6, 23, 43).
Some actinomycetes, including Gordona, Mycobacterium,
Nocardia, Rhodococcus, and Tsukamurella, have cell envelopes
that contain straight-chain, saturated, and unsaturated fatty
acids and tuberculostearic acid in intimate association with
high-molecular-mass 3-hydroxy 2-alkyl branched-chain fatty
acids (mycolic acids) (23, 26). It has been suggested that the
presence of these mycolic acids in the cell envelopes results in
increased hydrophobicity and thus leads to difficulties in per-
meabilization with PFA (7, 23).
Lawrence and Singer (19) showed that fixation with PFA for
1 min was effective for in situ hybridization of chicken embry-
onic muscle cultures and that cellular retention and hybridiza-
tion were not improved by increasing fixation times up to 15
min. Macnaughton et al. (23) performed successful hybridiza-
tions with mycolic acid-containing actinomycetes with an acid
hydrolysis fixation (HCl treatment). However, this fixation pro-
cedure did not work for Gordona bronchialis, Mycobacterium
fortuitum, N. brasiliensis, Rhodococcus erythropolis, Rhodococ-
cus fascians, Rhodococcus rhodochrous, and T. paurometab-
FIG. 3. Probe specificity study. Membrane hybridization results were analyzed with a PhosphorImager and were scanned and printed with Adobe Photoshop 3.0
(Adobe, Seattle, Wash.). (a) Template (names of individual organisms are given in Table 1); (b) probe S-*-Univ-1390-a-A-18; (c) probe S-*-Myb-0736-a-A-22; (d) probe
S-*-Myb-0736-b-A-22; (e) probe S-G-Gor-0596-a-A-22; (f) probe S-S-G.am-0192-a-A-18.
1112 DE LOS REYES ET AL. APPL.ENVIRON.MICROBIOL.
olum. Thus, they concluded that the optimum fixation proce-
dure was strain dependent.
To assess this potential problem, we evaluated four fixation
treatments with G. amarae SE102, G. amarae NM23, T. pau-
rometabolum, R. rhodochrous, and Mycobacterium smegmatis
cells. Bacterial probe S-D-Bact-0338-a-A-18, labeled with
TRITC, was used in this experiment. The fixation treatments
included PFA fixation for 1 min and 2 h and ethanol fixation
for 2 h and 12 to 16 h. The shorter fixation time with PFA was
hypothesized to be a good compromise to allow retention of
cell morphology while avoiding excessive cross-linking of the
proteins in the cell wall by PFA. Figures 4 and 5 show that
treatment with 4% PFA for 1 min renders G. amarae, R.
rhodochrous, and M. smegmatis permeable for labeled probes.
T. paurometabolum responds best to overnight ethanol fixation.
The finding that T. paurometabolum is not permeabilized by
PFA is consistent with the results from a previous study (23).
The differential results obtained with different species indicate
that it may be difficult to find a single method for permeabi-
lizing all cells. This observation may be especially problematic
in quantitative studies of complex environmental samples. In
population characterization studies, this problem could be
minimized by side-by-side fixations and use of normalization of
cell counts obtained with specific probes to cell counts ob-
tained with a universal probe. Based on the results of these
fixation studies, we decided to use 4% PFA fixation for 1 min
for further probe characterization studies and the FISH of
environmental samples.
Optimization of FISH conditions. To determine the optimal
hybridization conditions for FISH, probe-conferred fluores-
cence was quantified for eight different hybridization condi-
tions, representing 0 to 70% FA in the hybridization and wash
solutions. Increasing the FA concentration by 1% has an ap-
proximate equivalent effect of increasing the T
d
by 0.78C (43).
Thus, various stringencies can be simulated with different FA
concentrations. In this study, equivalent FA concentrations
were achieved by changing the NaCl concentration in the hy-
bridization and wash solutions, based on empirical formulas
for estimating T
d
values (43). Representative results of these
studies are shown for probe S-S-G.am-0192-a-A-18 in Fig. 6. In
general, higher FA concentrations resulted in lower signal in-
tensities (mean pixel value), and signals for target species were
consistently higher than those for nontarget species. The op-
timum FA concentrations were determined to be 30% for both
Mycobacterium complex probes, 20% for the genus-specific
probe, and 30% for the species-specific probe (Table 2 and Fig.
6).
FISH of environmental samples. In situ images of foam
obtained from the Urbana activated sludge and anaerobic di-
gester systems taken after FISH with probes for Gordona and
G. amarae are shown in Fig. 7. Initial FISH studies showed that
activated sludge samples contained organic matter and cells
which autofluoresce; in addition, out-of-focus fluorescence in
thick flocs hampered image clarity. Thus, the fluorescent dyes
and filter sets that we used, TRITC and FITC, tended to
impart some red (TRITC) or green (FITC) even to nontarget
cells. This uncertainty can lead to subjectivity in cell identifi-
cation and can only be resolved by quantification of the fluo-
rescence of all cells in activated sludge. In the absence of a
valid quantitative measure, we developed a qualitative method
to improve image visualization. This involved inclusion of the
dye DAPI in FISH studies with TRITC- and FITC-labeled
probes. DAPI, which binds to DNA, has a maximum emission
at 460 nm (blue), TRITC has a maximum emission at 610 nm
(red-orange), and FITC has maximum emission at 535 nm
(green).
Figures 7a and b present images of activated sludge foam
hybridized with probe S-G-Gor-0596-a-A-22 (labeled with
TRITC) and counterstained with DAPI. Figure 7a shows the
phase-contrast image, while Fig. 7b presents a superimposition
of the same image field obtained with TRITC and DAPI filter
sets. By this approach, target organisms theoretically should
show as a combination of blue and red (purple [TRITC and
DAPI]), and nontarget cells should be blue (DAPI only). How-
ever, because of the autofluorescence and out-of-plane fluo-
rescence mentioned above, some red showed up in suspected
nontarget cells. We found that by optimizing the exposure time
with the TRITC filter, the distinction between target and non-
target organisms could be improved. Thus, the greater inten-
sity of TRITC in target cells resulted in a red color for mem-
bers of the genus Gordona, thereby clearly showing the target
organisms, while nontarget cells were purple. This technique
thus allowed for detection of target organisms despite the
presence of autofluorescing organic matter and limited spatial
resolution of conventional epifluorescence microscopy. The
phase-contrast image showed that several different types of
filaments were present in the activated sludge foam. The flu-
orescence image demonstrated that the Gordona probe de-
tected some filament types, but not others, enabling us to
distinguish members of the genus Gordona from other filamen-
tous types. This is particularly significant, because the mor-
phology (e.g., length and branching of filaments and fragmen-
tation into rods and cocci) of filamentous organisms is affected
by different growth rates (20, 41).
We also demonstrated this approach with anaerobic digester
foam. Samples were hybridized with probe S-S-G.am-0192-a-
FIG. 4. Effects of fixation treatment on G. amarae. (a) G. amarae SE102 fixed
with PFA for 1 min. (b) G. amarae SE102 fixed with ethanol for 12 h. (c) G.
amarae NM23 fixed with PFA for 1 min. (d) G. amarae NM23 fixed with ethanol
for 12 h. Bar, 10 mm.
VOL. 63, 1997 rRNA HYBRIDIZATION PROBES FOR ACTIVATED SLUDGE FOAMING 1113
A-18 (labeled with FITC) counterstained with DAPI. Figure 7c
shows the phase-contrast image, and Fig. 7d is a superimposi-
tion of images obtained with FITC and DAPI filter sets. As
described above, autofluorescence in thick flocs can be circum-
vented by optimizing exposure time for the target organisms.
The G. amarae cells thus show up as green, while nontarget
organisms are light blue. Again, it was possible to identify G.
amarae cells without regard for morphology, because both
filaments and rods were detected. These images show the pres-
ence of G. amarae in anaerobic digester foam and demonstrate
the potential of FISH to study the role of filamentous foaming
in anaerobic digesters. In addition to the advantages of FISH
discussed above, the demonstration that FISH can be used to
study filaments in anaerobic digesters may be of particular
practical significance. It has been suggested that Nocardia
(Gordona) filaments lose their Gram-staining property in high-
rate anaerobic digesters (16), which makes study of them with
traditional staining methods especially problematic.
The abovementioned difficulties with FISH of activated
sludge and anaerobic digester samples made microscopic anal-
ysis difficult. Out-of-plane fluorescence, typical for conven-
tional epifluorescence microscopy, and very thick flocs, which
led to overlaying of nontarget bacteria in the same microscopic
field, added to the lack of image clarity. Filamentous bacteria
were observed to cluster together in situ, forming thick fluo-
rescent “balls” and bundles, which were difficult to analyze by
FIG. 5. Effects of fixation treatment on R. rhodochrous, M. smegmatis, and T. paurometabolum.(atod)M. smegmatis;(etoh)R. rhodochrous;(itol)
T. paurometabolum. (a, e, and i) One-minute PFA fixation; (b, f, and j) 2-h PFA fixation; (c, g, and k) 2-h ethanol fixation; (d, h, and l) 12-h ethanol fixation.
FIG. 6. Optimization of hybridization and wash conditions for FISH with
probe S-S-G.am-0192-a-A-18.
1114 DE LOS REYES ET AL. APPL.ENVIRON.MICROBIOL.
two-dimensional epifluorescence microscopy. Thus, it was eas-
ier to visualize target organisms at floc edges rather than within
flocs. Although beyond the scope of this paper, some possibil-
ities for minimizing these problems include the use of confocal
laser scanning microscopy, which would minimize out-of-focus
fluorescence due to the three-dimensional nature of flocs, and
the use of competitor probes, which would minimize nonspe-
cific binding within the floc. Various sample preparation tech-
niques must also be explored to reduce the floc density and to
separate filaments.
Quantitative membrane hybridization. The group-, genus-,
and species-specific probes also were used to characterize the
microbial community structure of foam and mixed liquor from
the Urbana activated sludge and anaerobic digester systems,
which experienced foaming problems at the time of sampling.
The abundances of different microbial groups were expressed
as a percentage of the total SSU rRNA in the sample (Table
3).
The N
5
-substituted Mycobacterium complex probe consis-
tently detected higher fractions of target organisms than the
original probe, demonstrating the improved detection of all
target organisms. The results show that a relatively high per-
centage (15.0% 6 2.7%) of the rRNA in activated sludge foam
can be attributed to members of the Mycobacterium complex,
of which the vast majority consisted of Gordona rRNA. rRNAs
from G. amarae SE6 and SE102 made up only small fractions
of the Gordona rRNA present (the stringency of the species-
specific probe was controlled to target G. amarae SE6 and
FIG. 7. Phase-contrast and epifluorescence images of FISH of environmental samples. (a and b) Activated sludge foam; (c and d) anaerobic digester foam.
Magnification, 3630. Bar, 10 mm. (a and b) Phase-contrast and epifluorescence micrographs, respectively, with S-G-Gor-0596-a-A-22 labeled with TRITC and dual
staining with DAPI; (c and d) phase-contrast and epifluorescence micrographs, respectively, with S-S-G.am-0192-a-A-18 labeled with FITC and dual staining with
DAPI.
TABLE 3. Quantitative hybridization results, expressed as percentages of total SSU rRNA in samples
Target group and probe
% of total SSU rRNA in sample (SD)
Activated sludge Anaerobic digester
Foam Mixed liquor Foam Sludge
Mycobacterium complex
S-*-Myb-0736-a-A-22 12.9 (2.2) 7.0 (2.6) 15.6 (2.1) 3.0 (1.6)
S-*-Myb-0736-b-A-22 15.0 (2.7) 8.8 (3.1) 18.3 (2.6) 3.8 (2.0)
Gordona, S-G-Gor-0596-a-A-22 12.6 (2.0) 8.6 (2.4) 14.6 (1.9) 3.6 (1.4)
G. amarae SE6 and SE102, S-S-G.am-0192-a-A-18 0.8 (0.8) 0.6 (1.0) 0.8 (1.2) 0.5 (0.5)
V
OL. 63, 1997 rRNA HYBRIDIZATION PROBES FOR ACTIVATED SLUDGE FOAMING 1115
SE102 only [Fig. 2d]). The relative abundance of Mycobacte-
rium complex rRNA in anaerobic digester foam was even
higher than that in activated sludge foam (18.3% 6 2.6%).
However, as expected, the percentage in activated sludge
mixed liquor (8.8% 6 3.1%) was higher than that in anaerobic
digester sludge (3.8% 6 2.0%). As for activated sludge foam,
most of the Mycobacterium complex rRNA in activated sludge
mixed liquor, anaerobic digester foam, and anaerobic digester
sludge consisted of Gordona rRNA; rRNAs from G. amarae
SE6 and SE102 were minor contributors to the Gordona
rRNA. Even though G. amarae SE6 and SE102 were present,
the hybridization results suggest that the foaming problem in
the Urbana wastewater treatment system may be linked to
Gordona strains other than G. amarae SE6 or SE102. SSU
rDNA sequencing of other Gordona strains and development
of strain-specific probes should resolve such questions.
The observation that Gordona species were predominant
among the mycolic acid-containing actinomycetes (Mycobacte-
rium complex) was made possible through a “probe nesting”
approach and illustrates the advantage of using probes for
groups representing different phylogenetic levels. We also em-
phasize that relative rRNA concentrations do not necessarily
correspond to filament numbers, but should be used to evalu-
ate target group activity relative to total microbial activity (43).
Significance for control of filamentous foaming. Control of
foaming problems in activated sludge plants can only be fur-
thered through a thorough understanding of the mechanisms
that cause foaming. The role of microbiological factors, such as
the growth of filamentous bacteria, must be understood. Iden-
tification and quantification of important organisms are thus
integral components of treatment plant operation and labora-
tory-scale studies (52). The filamentous organism identification
keys of Eikelboom (13) and later modifications by Jenkins et
al. (17) have been critical in addressing this need. However,
these techniques, which are based on staining characteristics
and other morphological criteria, may not be able to identify
and quantify causative microorganisms in all cases. The use of
hybridization probes allows for the in situ detection of foam-
causing bacteria regardless of morphological characteristics.
This advantage may be important in laboratory- and full-scale
studies in which microbial populations must be monitored over
periods of time across treatment processes or for the detection
of morphologically diverse nocardioform actinomycetes (fuga-
cious mycelia may break up into rod-shaped or coccoid ele-
ments [20]). In some cases, total counts of filaments may not be
adequate to represent the abundance of Gordona spp. For
example, Jenkins and coworkers compared the Nocardia (Gor-
dona) filament counts from two treatment plants employing
different methods for dealing with foam and recycle streams
(11, 17, 29). They hypothesized that seeding of the influent
with Nocardia (Gordona) organisms from different process re-
cycle streams would explain the differences in Nocardia (Gor-
dona) filament counts. However, attempts to confirm this by
detecting Nocardia (Gordona) filaments in the activated sludge
influent were not successful (17, 29). A method that does not
rely on morphology may have been beneficial in this study.
In conclusion, we developed and characterized a “nested”
set of oligonucleotide probes for filamentous bacteria which
are thought to be important in activated sludge foam as an
alternative to conventional identification methods. The use of
hybridization probes allowed for the identification and quan-
tification of filamentous microbial populations in activated
sludge and anaerobic digester foam. Further probe develop-
ment is necessary to fully assess the microbial community
structure in foam. At the same time, current limitations of
FISH and membrane hybridization methods must be ad-
dressed. For example, we demonstrated the importance of
optimizing cell fixation methods, as well as the applicability of
a universal base analog in probe design. The development of
methods that allow for microbial characterization of foams and
mixed liquor is an essential step in understanding the biolog-
ical basis of foaming. When combined with chemical analyses
and carefully designed reactor studies, the use of hybridization
probes may provide insights into the causes and control of
foaming.
ACKNOWLEDGMENTS
We are thankful to Mark Hernandez for providing G. amarae
strains, Elizabeth Wheeler Alm for providing other reference cultures,
Lala de los Reyes for help with constructing the phylogenetic tree,
Volodya Gelfand and Steve Rogers for assistance with fluorescence
microscopy, and Rudi Amann and Michi Wagner for helpful sugges-
tions on FISH. The support of Tim Bachman and the Urbana-Cham-
paign Sanitary District is also gratefully acknowledged.
This research was supported by the U.S. National Science Founda-
tion (grant BES 9410476).
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