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Exploiting Molecular Methods to Explore Endodontic
Infections: Part 1—Current Molecular Technologies for
Microbiological Diagnosis
J. F. Siqueira, Jr, DDS, MSc, PhD, and I. N. Rôças, DDS, MSc, PhD
Abstract
Endodontic infections have been traditionally studied
by culture-dependent methods. However, as with other
areas of clinical microbiology, culture-based investigations are plagued by significant problems, including the
probable involvement of viable but uncultivable microorganisms with disease causation and inaccurate microbial identification. Innumerous molecular technologies have been used for microbiological diagnosis in
clinical microbiology, but only recently some of these
techniques have been applied in endodontic microbiology research. This paper intended to review the main
molecular methods that have been used or have the
potential to be used in the study of endodontic infections. Moreover, advantages and limitations of current
molecular techniques when compared to conventional
methods for microbial identification are also discussed.
Key Words
Apical periodontitis, endodontic microbiology, molecular technology, microbiological diagnosis
From the Department of Endodontics, Estácio de Sá University, Rio de Janeiro, Rio de Janeiro, Brazil.
Address requests for reprint to José F. Siqueira, Jr, R.
Herotides de Oliveira 61 / 601, Icaraı́, Niterói, RJ, Brazil 24230230. E-mail address: [email protected].
Copyright © 2005 by the American Association of
Endodontists
I
n essence, endodontic infection is the infection of the root canal of the tooth and is the
primary etiologic agent of the different forms of periradicular inflammatory diseases
(1, 2). The pathological process is started when the dental pulp becomes necrotic,
usually as a sequel to caries, and then infected by micro-organisms that are usually
normal inhabitants of the oral cavity. The root canal containing a necrotic pulp affords
micro-organisms a moist, warm, nutritious, and anaerobic environment, which is, by
and large, protected from host defenses. Such conditions are highly conducive to
microbial colonization and multiplication. After the endodontic infection is established,
micro-organisms enter in close contact with the periradicular tissues via the apical
foramen and occasional foraminas, inflict damage to these tissues, and give rise to
inflammatory changes. Periradicular inflammatory diseases are arguably among the
most common diseases that affect human beings (3, 4).
Traditionally, endodontic bacteria have been studied by means of cultivationbased techniques, which rely on isolation, growth, and laboratory identification by
morphology and biochemical tests. However, cultivation and other traditional identification methods have been demonstrated to have several limitations when it comes to
microbiological diagnosis (5). The past decade has brought many advances in microbial molecular diagnostics, the most prolific being in DNA-DNA hybridization as well as
in polymerase chain reaction (PCR) technology and its derivatives. Indeed, findings
from cultivation-based methods with regard to the microbiota living in diverse ecosystems have been supplemented and significantly expanded with molecular biology techniques, and the impact of these methods on the knowledge about the oral microbiota in
healthy and diseased conditions is astonishing and breathtaking.
The first part of this paper describes the state of the art of molecular methodologies
already used or with potential to be applied to the study of the microbiota associated
with endodontic infections. The significant contribution brought about by molecular
technology and its impact on the redefinition of endodontic infections are discussed in
the second part of this paper.
Traditional Identification Methods
Culture
For more than a century, cultivation using artificial growth media has been the
standard diagnostic test in infectious diseases. The microbiota associated with different
sites in the human body has been extensively and frequently scrutinized by studies using
cultivation approaches. The success in cultivation of important pathogenic bacteria
probably led microbiologists to feel satisfied with and optimistic about their results and
to recognize that there is no dearth of known pathogens (6, 7). But should we be so
complacent with what we know about human pathogens?
Making micro-organisms grow under laboratory conditions presupposes some
knowledge of their growth requirements. Nevertheless, very little is known about the
specific growth factors that are utilized by innumerous micro-organisms to survive in
virtually all habitats, including within the human body (8). A huge proportion of the
microbial species in nature are difficult to be tamed in the laboratory. Certain bacteria
are fastidious or even impossible to cultivate. Some well-known human pathogens, such
as Mycobacterium leprae and Treponema pallidum continue to defy scientists regarding their cultivation under laboratory conditions (9).
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TABLE 1. Reasons for bacterial unculturability
a)
b)
c)
d)
e)
f)
Lack of essential nutrients or growth factors in the artificial culture medium.
Toxicity of the culture medium itself, which can inhibit bacterial growth.
Production of substances inhibitory to the target microorganism by other species present in a mixed consortium.
Metabolic dependence on other species for growth.
Disruption of bacterial intercommunication systems induced by separation of bacteria on solid culture media.
Bacterial dormancy, which is a state of low metabolic activity that some bacteria develop under certain stressful environmental
conditions, such as starvation. Dormant bacterial cells can be unable to divide or to form colonies on agar plates without a
preceding resuscitation phase.
Data according to references 7, 19, 20.
Of 36 bacterial divisions noted by Hugenholtz et al. (10), 13 divisions are exclusively comprised of as-yet uncultivated bacteria. Updated
analyses have indicated that presently 52 phyla can be discerned, of
which 26 are candidate phyla, that is, they are uncultivable and known
only by gene sequences (11). In fact, there has been a pronounced bias
towards the study of representatives of four bacterial phyla, namely
Proteobacteria, Firmicutes, Actinobacteria, and Bacteroidetes, out
of 52 bacterial phyla (12). This bias is arguably related to the fact that
representatives of these phyla are easier to cultivate with the methods
available nowadays. It is glaringly obvious that there is an urgent need
for this bias to be rectified.
Infectious disease professionals have long been aware of the fact
that when cultivation is applied to diagnose the causative agents of
diseases of suspected microbial etiology, including pneumoniae, encephalitis, meningitis, pericarditis, acute diarrhea, and sepsis, a large
number of cases can not be explained microbiologically (13, 14).
Moreover, the etiology of a number of chronic inflammatory diseases
with features suggestive of infection, such as rheumatoid arthritis, systemic lupus erythematosis, atherosclerosis, and diabetes mellitus, remain obscure (15). Taking into consideration that known bacterial
pathogens fall within 7 out of the 52 candidate bacterial divisions and
that cultivation-independent approaches have shown that 40 to 75% of
the human microbiota in different sites are composed of as-yet uncultivated bacteria (16 –18), it is fair to realize that there can be many
pathogens which remain to be identified. Therefore, it is of concern that
clinical microbiology continues to rely on cultivation-based identification procedures (6).
Advantages and Limitations
The main advantages of cultivation approaches are related to their
broad-range nature, which makes it possible to identify a great variety of
microbial species in a sample, including those that are not being sought
after. Still, cultivation makes it possible to determine antimicrobial susceptibilities of the isolates and to study their physiology and pathogenicity.
However, cultivation-based identification approaches have several
limitations: they are costly; they can take several days to weeks to identify
some fastidious anaerobic bacteria (that can delay antimicrobial treatment); they have a very low sensitivity (particularly for fastidious anaerobic bacteria); their specificity may be also low and is dependent on the
experience of the microbiologist; they have strict dependence on the
mode of sample transport; they are time-consuming and laborious.
Finally, the impossibility of cultivating a large number of bacterial species as well as the difficulties in identifying many cultivable species
represent the major drawbacks of cultivation-based approaches.
Difficulties in Cultivation
There are many possible reasons for bacterial unculturability (7,
19, 20) (Table 1). Obviously, if micro-organisms can not be cultivated,
they can not be identified by phenotype-based methods. While we stay
relatively ignorant on the requirements of many bacteria to grow, iden412
Siqueira and Rôças
tification methods that are not based on bacterial culturability are required. This would avoid that many pathogens pass unnoticed when one
is microbiologically surveying clinical samples.
It is worth pointing out that the fact that a given species is uncultivable does not necessarily imply that the same species will remain
indefinitely impossible to cultivate. For instance, a myriad of strict anaerobic bacteria were uncultivable 100 yr ago, but further developments in cultivation techniques have to a large extent helped solve this
problem. There is a growing trend to develop approaches and culture
media that allow cultivation of previously uncultivated bacteria. Strategies may rely on application of cultivation procedures that better mimic
conditions existing in the natural habitat from which the samples were
obtained. Recent efforts to accomplish this have met with some success
by including the following: the use of agar media with little or no added
nutrients; relatively lengthy periods of incubation (more than 30 days);
and inclusion of substances that are typical of the natural environment
in the artificial growth media (21, 22). However, it is not unreasonable
to surmise that a huge number of species will remain uncultivable for
years to come.
Difficulties in Identification
Accurate identification of microbial isolates is paramount in clinical microbiology. For a given microbial species to be identified by
means of their phenotypic features, this species has to be cultivated.
However, one should be mindful that in some circumstances even the
successful cultivation of a given microorganism does not necessarily
mean that this micro-organism can be successfully identified.
For slow-growing and fastidious bacteria, traditional phenotypic
identification is difficult and time-consuming. In addition, interpretation of phenotypic test results can involve a substantial amount of subjective judgment and personnel’s expertise. Still, one major difficulty
associated with microbial identification based on phenotypic features is
that of divergence and convergence. Divergence occurs for strains of the
same species, which are genetically similar, but have evolved to be
different phenotypically. Convergence occurs for strains of different
species, which are genetically different, but have evolved to have similar
phenotypic behavior (23). In both situations, phenotypically based diagnostic tests would result in misidentification.
16S rRNA gene (rDNA) sequencing approach was first proposed to
identify uncultivable bacteria (see section on Broad-Range PCR in this
article), but it has also been recently used for identification of cultivable
bacteria that shows atypical phenotypic behavior and cannot be accurately identified by culture. Unlike phenotypic identification, which can
be modified by the variability of expression of characters, 16S rDNA
sequencing provides unambiguous data even for rare isolates. Bosshard
et al. (24) reported that 14% of all aerobic gram-positive rods isolated
in their clinical microbiology laboratory required 16S rDNA sequencebased identification. Drancourt et al. (25) reported that about 0.5 to 1%
of the bacterial isolates recovered in their laboratory were not possible
to be identified by phenotypic criteria. Using 16S rDNA sequence analysis, they reported that several atypical isolates corresponded to 27 new
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TABLE 2. Advantages of molecular genetic methods over other methods for microbial identification
Advantages of Molecular Genetic Methods
a) Detection of not only cultivable species but also of uncultivable microbial species or strains.
b) Higher specificity and accurate identification of microbial strains with ambiguous phenotypic behavior, including divergent or
convergent strains.
c) Detection of microbial species directly in clinical samples, without the need for cultivation.
d) Higher sensitivity.
e) Faster and less time-consuming.
f) They offer a rapid diagnosis, which is particularly advantageous in cases of life-threatening diseases or diseases caused by slowgrowing micro-organisms.
g) They do not require carefully controlled anaerobic conditions during sampling and transportation, which is advantageous since
fastidious anaerobic bacteria and other fragile micro-organisms can lose viability during transit.
h) They can be used during antimicrobial treatment.
i) When a large number of samples are to be surveyed in epidemiological studies, samples can be stored and analyzed all at once.
bacterial species associated with human diseases, of which 11 isolates
were previously uncharacterized species and 16 isolates had never been
reported in humans, being previously regarded as environmental. Drancourt et al. (26) evaluated the utility of 16S rDNA sequencing as a means
to identify a collection of 177 isolates obtained from environmental,
veterinary, and clinical sources. Conventional identification failed to
produce accurate results because of inappropriate biochemical profile
determination in 76 isolates (58.7%), Gram staining in 16 isolates
(11.6%), oxidase and catalase activity determination in five isolates
(3.6%) and growth requirement determination in two isolates (1.5%).
They reported that the overall performance of 16S rDNA sequence analysis was excellent, because it succeeded in identifying 90% of phenotypically unidentifiable bacterial isolates. Song et al. (27) evaluated the
utility of 16S rDNA sequencing as a means of identifying clinically important gram-positive anaerobic cocci. Based on the sequences of the
13 type strains obtained in their study, 84% (131 of 156) of the clinical
isolates were accurately identified to species level, with the remaining
25 clinical strains revealing nine unique sequences that could represent
eight novel species. This finding was in contrast to the phenotypic identification results, by which only 56% of isolates were correctly identified
to species level. These studies clearly shown that sequence analysis of
the 16S rDNA can be used to overcome difficulties in identifying cultivable species that are notoriously refractory to identification by phenotypic means.
Microscopy
Microscopy may suggest an etiologic agent, but it rarely provides
definitive evidence of infection by a particular species. Microscopic
findings regarding bacterial morphology may be misleading, because
many species can be pleomorphic and conclusions can be influenced by
subjective interpretation of the investigator. In addition, microscopy has
limited sensitivity and specificity to detect micro-organisms in clinical
samples. Limited sensitivity is because a relatively large number of microbial cells are required before they are seen under microscopy (e.g.
104 bacterial cells/ml of fluid) (9). Some micro-organisms can even
require appropriate stains and/or approaches to become visible. Limited specificity is because our inability to speciate micro-organisms
based on their morphology and staining patterns.
Immunological Methods
Immunological methods are based on the specificity of antigenantibody reaction. It can detect micro-organisms directly or indirectly,
the latter by detecting host immunoglobulins specific to the target micro-organism. The enzyme-linked immunosorbent assay (ELISA) and
the direct or indirect immunofluorescence tests are the most commonly
used immunological methods for microbial identification. Advantages
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clude: (a) they take no more than a few hours to identify a microbial
species; (b) they can detect dead micro-organisms; (c) they can be
easily standardized; and (d) they have low cost (28). However, they
have also important limitations as they can detect only target species,
they have low sensitivity (about 104 cells), their specificity is variable
and depends on types of antibodies used, and they can detect dead
micro-organisms (28, 29).
Molecular Genetic Methods
Investigations of many aquatic and terrestrial environments using
cultivation-independent methods have revealed a great deal of previously unsuspected microbial diversity (30). In fact, the cultivable members of these living systems represent less than 1% of the total extant
population (31, 32). Novel culture-independent methods for microbial
identification that involve DNA amplification of 16S rDNA followed by
cloning and sequencing have recently been used to determine the bacterial diversity within diverse environments, such as deep-sea sediments
(33), hot springs (34), and human diseased and healthy sites (16 –18,
35–39). Perhaps not surprisingly, the number of recognized bacterial
phyla has exploded from the original estimate of 11 in 1987 to 36 in
1998 (10). The latest tally of bacterial phyla is now probably near 52, of
which one-half has only uncultivable representatives (11).
A significant contribution of molecular methods to medical microbiology relates to the identification of previously unknown human
pathogens (40 – 44). In addition, molecular studies have revealed that
about 40 to 50% of the bacterial clones in the oral cavity (17) and about
70% of clones in the gut and colon (16, 35) represent unknown and
as-yet uncultivable species. As a consequence, it is fair to realize that
there can exist a number of uncharacterized pathogens in this uncultivable proportion of the human microbiota.
Molecular diagnostic methods have several advantages over other
methods with regard to microbial identification (Table 2). Limitations
of molecular approaches will be discussed further ahead in this paper.
There are a plethora of molecular methods for the study of microorganisms and the choice of a particular approach depends on the
questions being addressed (Fig. 1). Broad-range polymerase chain reaction (PCR) followed by sequencing can be used to investigate the
microbial diversity in a given environment. Microbial community structure can be analyzed via fingerprinting techniques, such as denaturing
gradient gel electrophoresis (DGGE) and terminal restriction fragment
length polymorphism (T-RFLP). Fluorescence in situ hybridization
(FISH) can measure abundance of particular species and provide information on their spatial distribution in tissues. Among other applications, DNA-DNA hybridization macroarrays and microarrays, speciesspecific PCR, nested PCR, multiplex PCR, and real-time PCR can be used
to survey a large number of clinical samples for the presence of target
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Figure 1. Examples of molecular techniques used or with potential to be used in the study of endodontic infections. The choice for a particular method will depend
on the question being addressed.
species. Variations in PCR technology can also be used to type microbial
strains. This review will restrict discussion to the most commonly used
approaches applied to the research of the endodontic microbiota and
some with potential to be used with this intent.
Gene Targets for Microbial Identification
Molecular approaches for microbial identification rely on the fact
that certain genes contain revealing information about the microbial
identity. Ideally, a gene to be used as target for microbial identification
should contain regions that are unique to each species. Following the
pioneer studies by Woese (45), the genes encoding rRNA molecules,
which are present in all cellular forms of life, namely, the domains
Bacteria, Archaea, and Eucarya, have been widely used for comprehensive identification of virtually all living organisms and inference of
their natural relationships.
Ribosomes are intracellular particles composed of rRNA and proteins. In Escherichia coli, about two-thirds of the ribosome is rRNA and
the remainder is protein (46). The sizes of ribosomes are given in
Svedburg (S) units, which represent a measure of how rapidly particles
or molecules sediment in an ultracentrifuge. Bacterial and archaeal
cells have 70S ribosomes composed of 30S and 50S subunits. The 30S
subunit contains a 16S rRNA molecule, having approximately 1540 nucleotides. The 50S subunit contains a 23S rRNA molecule, having ap414
Siqueira and Rôças
proximately 2900 nucleotides, and a small 5S rRNA molecule having
only about 120 nucleotides. Fungi have 80S ribosomes composed of 40S
and 60S subunits. The 40S subunit contains 18S rRNA and the larger 60S
subunit has 25S rRNA and 5.8S rRNA (47).
Large subunit genes (23S and 25S rDNA) and small subunit genes
(16S and 18S rDNA) have been widely used for microbial identification,
characterization and classification. The small subunit rDNA is among
the most evolutionary conserved macromolecules in all living systems
(45). The advantages of using small subunit rDNA is that it is found in all
organisms, is long enough to be highly informative and short enough to
be easily sequenced, particularly with the advent of automated DNA
sequencers (45). The small subunit rDNA contains some regions that
are virtually identical in all representative of a given domain (conserved
regions) and other regions that vary in sequence from one species to
another (variable regions) (Fig. 2). Variable regions contain the most information about the genus and species of the bacterium, with unique signatures that allow specific identification. The 16S rRNA of bacteria and archaea
and the 18S rDNA of fungi and other eukaryotes have been extensively
examined and sequenced and have been used to determine phylogenetic
relationships among living organisms. In addition, data from rDNA sequences can also be used for accurate and rapid identification of known
bacterial species, using techniques that do not require microbial cultivation.
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Figure 2. Schematic drawing of the 16S rRNA gene (rDNA). Orange areas correspond to variable regions, which contain information about the genus and the
species. Red areas correspond to conserved regions of the gene.
There are now more than 90,000 different bacterial 16S rDNA
sequences in public databases, while the number of 23S rDNA sequences is still considerably smaller (⬎1400), though it keeps increasing (48). Consequently, 16S rDNA is the most useful target for bacterial
identification by molecular approaches, and 23S rDNA is becoming a
suitable alternative. Recently, the intergenic, noncoding spacer sequences have also been used, because they are more likely to be variable than the rDNAs. The ones that have been used include the 16S-23S
intergenic spacer for bacteria (49) and the internal transcribed spacer
for fungi (50). These regions are useful for differentiating between
closely related species because of heterogeneity of its length and sequence but are not adequate to establish phylogenetic relationships
(51).
PCR
The PCR process was conceived by Kary Mullis in 1983 and ever
since has revolutionized the field of molecular biology by being able to
amplify as few as one copy of a gene into millions to billions of copies of
that gene (52). The impact of PCR on biological and medical research
has been remarkable, dramatically speeding the rate of progress of the
study of genes and genomes. Nowadays, it is possible to isolate essentially any gene from any organism using PCR, which made this technique
a cornerstone of genome sequencing projects (53). Since its introduction, PCR has spawned an increasing number of associated technologies
for diverse applications. Perhaps the most widespread advance in clinical diagnostic technology has come from the application of PCR for
detection of microbial pathogens (54 –56).
The PCR method is based on the in vitro replication of DNA through
repetitive cycles of denaturation, primer annealing and extension steps
(Fig. 3). The target DNA serving as template melts at temperatures high
enough to break the hydrogen bonds holding the strands together, thus
liberating single strands of DNA. Two short oligonucleotides (primers)
are annealed to complementary sequences on opposite strands of the
target DNA. Primers are selected to encompass the desired genetic
material, defining the two ends of the amplified stretch of DNA. In
sequence, a complementary second strand of new DNA is synthesized
through the extension of each annealed primer by a thermostable DNA
polymerase in the presence of excess deoxyribonucleoside triphosphates. All previously synthesized products act as templates for new
primer-extension reactions in each ensuing cycle. The result is the
exponential amplification of new DNA products, which confers extraordinary sensitivity in detecting the target DNA. In fact, PCR has unrivaled
sensitivity—it is at least 10 to 100 times more sensitive than the other
more sensitive identification method (29, 57).
There are several methods to check if the intended PCR product
was generated. The most commonly used method for detecting PCR
products is electrophoresis in an agarose gel. Aliquots of the PCR reac-
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tion are loaded into the gel and an electrical gradient is applied through
a buffer solution. The products migrate through the gel according to
size, with larger products running a shorter distance in the gel because
they experience more resistance in the gel matrix. DNA ladder digests
represent DNA fragments of known size and are run in the same gel to
serve as molecular size standard. This allows the size of the PCR products to be estimated. The gel is usually visualized using ethidium bromide staining and ultraviolet transillumination. Designed primers are
expected to generate a PCR product of a given size and the observation
of a PCR amplicon of the predicted size in the electrophoretic gel is
consistent with a positive PCR result. Identity of PCR products can still be
confirmed by sequencing of the PCR product, hybridization of a specific
oligonucleotide probe to a region of the PCR product that is internal to
the priming sites, or restriction enzyme cleavage of the PCR product
using an enzyme that is known to cut a specific sequence within the
product [restriction fragment length polymorphism (RFLP)].
Numerous derivatives in PCR technology have been developed
since its inception. The most used PCR-derived assays are described in
the following sections.
Nested PCR
Nested PCR uses the product of a primary PCR amplification as
template in a second PCR reaction and was devised mainly to have
increased sensitivity (58). The first PCR products are subjected to a
second round of PCR amplification with a separate primer set, which
anneals internally to the first products. This approach shows increased
sensitivity allowing the detection of the target DNA several folds lower
than conventional PCR. Increased sensitivity is because of the large total
number of cycles. In addition, target DNA is amplified in the first round
of amplification, with subsequent dilution of other DNA and inhibitors
present in the sample. The set of primers used in the second round of
PCR results in additional specificity. The second reaction is performed
with reduced background of eukaryotic DNA and other regions of the
bacterial DNA (57). Even if nonspecific DNA amplification occurs in the
first round of amplification, the nonspecific PCR product does not serve
as template in the second reaction, since it is highly unlikely to possess
regions of DNA complementary to the second set of specific primers
(59).
The major drawback of nested-PCR protocol is the high probability
of contamination during transfer of the first-round amplification products to a second reaction tube and special precautions should be taken
to avoid this.
Reverse Transcriptase PCR (RT-PCR)
RT-PCR was developed to amplify RNA targets and exploits the use
of the enzyme reverse transcriptase, which can synthesize a strand of
complementary DNA (cDNA) from an RNA template. Most RT-PCR assays employ a two-step approach. In the first step, reverse transcriptase
converts RNA into single-stranded cDNA. In the second step, PCR primers, DNA polymerase, and nucleotides are added to create the second
strand of cDNA. Once the double-stranded DNA template is formed, it
can be used as template for amplification as in conventional PCR (60).
The RT-PCR process may be modified into a one-step approach by using
it directly with RNA as the template. In this approach, an enzyme with
both reverse transcriptase and DNA polymerase activities is used, such
as that from the bacteria Thermus thermophilus.
Multiplex PCR
Most PCR assays have concentrated on the detection of a single
microbial species by means of individual reactions. In multiplex PCR,
two or more sets of primers specific for different targets are introduced
in the same reaction tube. Since more than one unique target sequence
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Figure 3. Scheme for PCR.
in a clinical specimen can be amplified at the same time, multiplex PCR
assays permit the simultaneous detection of different microbial species.
Multiplex PCR assays have been used to minimize the time and expenditure needed for detection approaches. Primers used in multiplex assays must be designed carefully to have similar annealing temperatures
and to lack complementarity (61, 62).
PCR-Based Microbial Typing
PCR technology can also be used for clonal analysis of microorganisms. An example of the PCR techniques used for this purpose
includes the arbitrarily primed PCR (AP-PCR), also referred to as random amplified polymorphic DNA (RAPD) (63). AP-PCR is a relatively
rapid PCR-based genomic fingerprinting technique that can be used as
a tool to determine whether two isolates of the same species are epidemiologically related. The method utilizes a single 10- to 20-base random sequence primer that anneals to unspecified DNA target sites under
conditions that allow for mismatched base-pairing. The use of a random-sequence primer at low stringency allows for priming at sites with
imperfect matches. Genetic variations between two DNA templates result in discriminative DNA fingerprints because of the differences in the
priming sites. The amplicons generated form a pattern in the electrophoretic gel that may be strain specific. The advantage of AP-PCR is its
ability to furnish highly specific DNA profiles with no prerequisite for
knowing the DNA sequences. Primers may also be designed to target
known genetic elements, such as enterobacterial repetitive intergenic
consensus sequences (ERIC-PCR) (64, 65) and repetitive extragenic
palindromic sequences (REP-PCR) (66). Clonal analysis may help elucidate whether certain strains of a given species are more associated
with signs or symptoms of a given disease. Clonal analysis also may help
to track the origin of micro-organisms infecting a given site. For instance, by comparing bacterial strains isolated from the root canal and
other oral sites, one can have information as to where micro-organisms
present in the root canal system came from. Clonal analysis can also
track the origin of the micro-organisms present in a suspected focal
disease by comparing the isolates found in the infected site with others
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present in the suspected focus of infection, including the oral cavity
(and infected root canals).
Real-Time PCR
Conventional PCR assays are qualitative or can be adjusted to be
semi-quantitative. One exception is the real-time PCR, which is characterized by the continuous measurement of amplification products
throughout the reaction (67).
There are several different real-time PCR approaches. The three
general real-time PCR chemistries for amplifying and detecting DNA
targets are SYBR-Green, TaqMan, and molecular beacon (68, 69).
SYBR Green is the simplest and most affordable method, and consists of
a fluorescent dye that binds to double-stranded DNA. During extension,
increasing amounts of dye bind to the increasing amount of newly
formed double-stranded DNA. Fluorescence is measured at the end of
the extension step of every PCR cycle to monitor the increasing amount
of amplified DNA. Dye that remains unbound exhibits little fluorescence
in solution. The SYBR-Green assay is very sensitive but has diminished
specificity, as the dye binds to all double-stranded DNA present, and
primer dimers can result in a false reading (70).
The TaqMan method can be more specific than the SYBR-Green
assay. Increased specificity of the TaqMan assay results from the utilization of a specific labeled oligonucleotide probe along with the primers
(71, 72). The TaqMan probe is a 20- to 30-base-long oligonucleotide
sequence that specifically anneals to a sequence flanked by the two
primers. The TaqMan probe contains a reporter fluorescent dye at the
5⬘ end and a quencher dye at the 3⬘ end that quenches the emission
spectrum of the reporter dye (Fig. 4). As long as the probe remains
unbound, it is intact and no signal is generated. During the extension
step of real-time PCR, the Taq DNA polymerase enzyme cleaves the
TaqMan probe, resulting in separation of the reporter from the
quencher (Fig. 4). This results in increased fluorescence emission.
Another real-time PCR assay uses molecular beacons, which are
single-stranded nucleic acid molecules with a stem-and-loop structure
(69). The loop portion is complementary to a sequence in the target
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Figure 4. Probes for real-time PCR. (A) TaqMan probes. The probe in solution folds and bring the reporter fluorescent dye close enough to the quencher dye to
quench the fluorescence. The probe bound to the PCR product is digested by the 5⬘ nuclease activity of Taq polymerase during the extension step of the cycle. (B)
Molecular beacons. Hybridization to its target induces a conformational change in the molecular beacon probe that forces the arm sequences apart and causes the
fluorophore to move away from the quencher.
DNA. The stem is formed by the annealing of complementary arm sequences located on either side of the probe. A fluorescent marker is
attached to the end of one arm, and a quencher is attached to the end of
the other arm. Free molecular beacons acquire a hairpin structure
when in solution, and the stem keeps the arm sequences in close proximity. This results in efficient quenching of the fluorescent dye (Fig. 4).
When hybridizing to their complementary target, molecular beacons are
forced to undergo a conformational change, with consequent formation
of probe that hybridizes to the template. The conformational change
forces the fluorescent dye and the quencher apart, generating fluorescence (73) (Fig. 4).
By monitoring the release of fluorescence with each PCR cycle, the
progress of the reaction is recorded in real time and the amount of DNA
in the sample is quantified. Thus, accumulation of PCR products is
measured automatically during each cycle in a closed tube format using
a thermocycler combined with a fluorimeter. Direct measurement of the
accumulated PCR product allows the phases of the reaction to be monitored. The initial amount of target DNA in the reaction can be related to
a cycle threshold defined as the cycle number at which there is a statistically significant increase in fluorescence. Target DNA can then be
quantified by construction of a calibration curve that relates cycle
threshold to known amounts of template DNA (74).
Real-Time PCR technology is commercially available as the TaqMan (Applied Biosystems, Perkin-Elmer Corp.) and LightCycler (Roche
Diagnostics Corp.) systems. Real-time PCR assays allow the quantification of individual target species as well as total bacteria in clinical
samples. The advantages of real-time PCR are the rapidity of the assay
(30 – 40 min), the ability to quantify and identify PCR products directly
without the use of agarose gels, and the fact that contamination of the
nucleic acids can be limited because of avoidance of postamplification
manipulation (75).
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Broad-Range PCR
PCR technology can also be used to investigate the whole microbial
diversity in a given environment. In broad-range PCR, primers are designed that are complementary to conserved regions of a particular
gene that are shared by a group of micro-organisms. For instance,
primers that are complementary to conserved regions of the 16S rDNA
have been used with the intention of exploiting the variable internal
regions of the amplified sequence for sequencing and further identification (76). The strength of broad-range PCR lies in the relative absence
of selectivity, so that (in principle) any kind of bacteria present in a
sample can be detected and identified. This aspect is in analogy to
cultivation and in contrast to species-specific molecular approaches
(48). Thus, broad-range PCR can detect the unexpected and in this
regard it is far more effective and accurate than culture. Broad-range
PCR has allowed the identification of several novel, fastidious or uncultivable bacterial pathogens directly from diverse human sites (5, 7, 77).
Initially, bulk nucleic acids are extracted directly from samples.
Afterwards, the 16S rDNA is isolated from the bulk DNA via PCR with
oligonucleotide primers specific for conserved regions of the gene (universal or broad-range primers). Longer distances between primer pairs
generally result in less sensitivity, but this provides more variable sequence for more accurate identification and may reduce the risks of
amplifying DNA from contaminants in PCR reagents, which appear to be
fragmented into smaller sizes (48). Amplification with universal primers results in a mixture of all of the 16S rDNA amplified from the bacteria
in the sample. In mixed infections, direct sequencing of the PCR products cannot be performed because there are mixed products from the
different species composing the consortium. PCR products are then
cloned into a plasmid vector, which is used to transform Escherichia
coli cells, establishing a library of 16S rDNA from the sample. Cloned
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genes are then sequenced individually and submitted for identification
to databases, usually via the World Wide Web. Preliminary identification
can be done by using similarity searches in public databases. A ⬎99%
identity in 16S rDNA sequence has been the most accepted criterion
used to identify a bacterium to the species level (25, 26). A 97 to 99%
identity in 16S rDNA sequence has been the criterion used to identify a
bacterium at the genus level, and ⬍97% identity in 16S rDNA gene
sequence has been the criterion used to define a potentially new bacterial species (25, 26). In addition to the similarity search, phylogenetic
analysis should be accomplished since it provides a much more accurate assessment (48, 51).
The analytical sensitivity of most broad-range PCR assays is in
practice above 10, if not 100, gene copies per PCR, which is significantly
lower when compared to most species-specific PCR assays (48). Because broad-range primers are used, there is a high risk for DNA from
microbial contaminants to be amplified. A wide range of precautions is
necessary to avoid contamination, including separate room for pre- and
post-PCR work, UV decontamination of surface areas, use of high-quality reagents, and adequate sampling techniques and vials for clinical
specimens (48, 59, 78).
Denaturing Gradient Gel Electrophoresis
Techniques for genetic fingerprinting of microbial communities
can be used to determine the diversity of different micro-organisms
living in diverse ecosystems and to monitor microbial community behavior over time. A commonly used strategy for genetic fingerprinting of
complex microbial communities encompasses the extraction of DNA,
the amplification of the 16S rDNA using broad-range primers, and then
the analysis of PCR products by denaturing gradient gel electrophoresis
(DGGE).
In DGGE, DNA fragments of the same length but with different
base-pair sequences can be separated (79, 80). The DGGE technique is
based on electrophoresis of PCR-amplified 16S rDNA (or other genes)
fragments in polyacrylamide gels containing a linearly increasing gradient of DNA denaturants (a mixture of urea and formamide). As the
PCR product migrates in the gel, it encounters increasing concentrations of denaturants and, at some position in the gel, it will become
partially or fully denatured. Partial denaturation causes a significant
decrease in the electrophoretic mobility of the DNA molecule. Molecules with different sequences may have a different melting behavior
and will therefore stop migrating at different positions in the gel. The
position in the gel at which the DNA melts is determined by its nucleotide
sequence and composition (81). A GC-rich sequence (GC-clamp) is
added to the 5⬘-end of one of the PCR primers, which ensures that at
least part of the DNA remains double stranded (82). DNA bands in
DGGE can be visualized using ethidium bromide, SYBR Green I, or silver
staining.
In DGGE, multiple samples can be analyzed concurrently, making
it possible to compare the structure of the microbial community of
different samples and to follow changes in microbial populations over
time, including after antimicrobial treatment. Specific bands can also be
excised from the gels, re-amplified and sequenced to allow microbial
identification. The DGGE method and its application in endodontic microbiology research have been recently reviewed (83). Temperature
gradient gel electrophoresis (TGGE) uses the same principle as DGGE,
except for the fact that the gradient is temperature rather than chemical
denaturants.
Terminal-RFLP
T-RFLP is a recent molecular approach that can assess subtle genetic
differences between microbial strains as well as provide insight into the
structure and function of microbial communities (84). T-RFLP analysis
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Siqueira and Rôças
measures the size polymorphism of terminal restriction fragments from a
PCR amplified marker. T-RFLP is a modification of the conventional RFLP
approach. In T-RFLP, rDNA from different species in a community is PCRamplified using one of the PCR primers labeled with a fluorescent dye, such
as 4,7,2⬘,7⬘-tetrachloro-6-carboxyfluorescein (TET) or phosphoramidite
fluorochrome 5-carboxyfluorescein (6-FAM) (85). PCR products are then
digested with restriction enzymes, generating different fragment lengths.
Digestion of PCR products with judiciously selected restriction endonucleases produces terminal fragments appropriate for sizing on high resolution
sequencing gels. The latter step is performed on automated systems
such as the ABI gel or capillary electrophoresis systems that provide
digital output (85).
The use of a fluorescently labeled primer limits the analysis to only
the terminal fragments of the enzymatic digestion. This simplifies the
banding pattern, thus allowing the analysis of complex communities as
well as providing information on diversity as each visible band represents a single species. All terminal fragment sizes generated from digestion of a PCR product pool can be compared with the terminal fragments
derived from sequence databases to derive phylogenetic inference.
Through application of automated DNA sequencer technology, T-RFLP
has considerably greater resolution than gel-based community profiling
techniques, such as DGGE/TGGE (84, 85).
DNA-DNA Hybridization
DNA-DNA hybridization methodology employs DNA probes, which
entail segments of single-stranded DNA, labeled with an enzyme, radioactive isotope or a chemiluminescence reporter, that can locate and
bind to their complementary nucleic acid sequences. DNA-DNA hybridization arrays on macroscopic matrices, such as nylon membranes,
have been often referred to as “macroarrays.” DNA probe may target
whole genomic DNA or individual genes. Whole genomic probes are
more likely to cross-react with nontarget micro-organisms because of
the presence of homologous sequences between different microbial
species. Oligonucleotide probes based on signature sequences of specific genes may display limited or no cross-reactivity with nontarget
micro-organisms when under optimized conditions. In addition, oligonucleotide probes can differentiate between closely related species or
even subspecies and can be designed to detect uncultivable bacteria.
Socransky et al. (86) introduced a method for hybridizing large
numbers of DNA samples against large numbers of digoxigenin-labeled
whole genomic DNA or 16S rDNA-based oligonucleotide probes on a
single support membrane—the checkerboard DNA-DNA hybridization
method. Denatured DNA from clinical samples is placed in lanes on a
nylon membrane using a Minislot device. After fixation of the samples to
the membrane, the membrane is placed in a Miniblotter 45 with the
lanes of samples at 90° to the lanes of the device. Digoxigenin-labeled
whole genomic DNA probes are then loaded in individual lanes of
the Miniblotter. After hybridization, the membranes are washed at
high stringency and the DNA probes detected using antibody to
digoxigenin conjugated with alkaline phosphatase and chemiluminescence detection.
The checkerboard method permits the simultaneous determination of the presence of a multitude of bacterial species in single or
multiple clinical samples. Thus, it is particularly applicable in largescale epidemiological research. In addition to the reported advantages
of molecular methods, DNA-DNA hybridization technology has the additional feature that microbial contaminants are not cultivated, nor is
their DNA amplified (87). Because the numbers of contaminating micro-organisms are not increased, one may assume that, if present, they
would be in numbers below detection limits of the checkerboard DNADNA hybridization method, which have been reported to be by the order
of 103 to 104 cells (86, 88).
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Review
A modification of the checkerboard method was proposed by Paster et al. (89) and consists of a PCR based, reverse-capture checkerboard hybridization methodology. The procedure circumvents the need
for in vitro bacterial cultivation, necessary for preparation of whole
genomic probes. Up to 30 reverse-capture oligonucleotide probes that
target regions of the 16S rDNA are deposited on a nylon membrane in
separate horizontal lanes using a Minislot apparatus. Each probe is
synthesized with a polythymidine tail, which is cross-linked to the membrane support via UV irradiation, leaving the probe available for hybridization. The 16S rDNA from clinical samples are PCR amplified using a
digoxigenin labeled primer. Hybridizations are performed in vertical
channels in a Miniblotter apparatus with digoxigenin-labeled PCR amplicons from up to 45 samples. Hybridization signals are detected using
chemifluorescence procedures.
DNA Microarrays
Developing a high-density macroarray is a difficult task, as large
membranes represent a practical limitation when it comes to the volume of probes that must be used, and particularly when small amounts
of samples are available. In addition, because of the slow hybridization
kinetics involved, they usually require longer periods for analysis. Miniaturization of these assays saves time while cutting costs in diagnostic
applications and research (90). Low volumes reduce reagent consumption and increase sample concentration, thus improving reaction kinetics. The use of nonporous supports allows faster hybridization and
easier washing steps. As a consequence, hundreds to thousands of results can be obtained in the time needed for a single macroarray experiment.
DNA microarray methods were first described in 1995 (91) and
essentially consist of many probes that are discretely located on a nonporous solid support, such as a glass slide (90, 92). Printed arrays and
high-density oligonucleotide arrays are the most commonly used types
of microarrays. A printed array comprises probes that are spotted onto
specific locations on the solid support platform and range in size from
500 to 2,000 base pairs in length. Probes can be DNA fragments such as
library clones or PCR products. By means of a robotic arrayer and
capillary printing tips, thousands of elements can be printed on a single
microscope slide. A high-density oligonucleotide microarray has a large
number of probes attached to a solid support substrate. In this method,
arrays are constructed by synthesizing single-stranded oligonucleotides
in situ by use of photolithographic techniques. These probes are typically shorter gene sequences comprised of 15 to 30 nucleotides (93).
Advantages of the former method include relatively low cost and substantial flexibility. Moreover, sequence information is not needed to
print a probe. Advantages of the latter method include higher density
and elimination of the need to obtain and store cloned DNA or PCR
products (94).
Once microarrays have been printed, targets are prepared for
hybridization. Targets may be PCR products, genomic DNA, total RNA,
cDNA and so on, and usually incorporate either a fluorescent label or
some other moiety, such as biotin, that permits subsequent detection
with a secondary label. Targets are applied to the array and those that
hybridize to complementary probes are detected using some type of
reporter molecule. Following hybridization, arrays are imaged using a
high-resolution scanner. Spatial resolution of these systems can be as
low as of 3 to 5 ␮m, although 10 ␮m of resolution is sufficient. Sophisticated computer software programs can then be used to analyze the
results.
DNA microarrays can be used to enhance PCR product detection
and identification. When PCR is used to amplify microbial DNA from
clinical specimens, microarrays can then be used to identify the PCR
products by hybridization to an array that is composed of species-
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specific probes. Using broad-range primers, such as those that amplify
the 16S rDNA, a single PCR can be used to detect hundreds of bacterial
species simultaneously. When coupled to PCR, microarrays have detection sensitivity similar to conventional molecular methods with the
added ability to discriminate several species at a time.
The advent of DNA microarrays has already given a marked contribution to many fields, including cellular physiology (95, 96), cancer
biology (97–99), and pharmacology (100). In clinical microbiology, in
addition to being applied to microbial detection and identification using
rDNA information, microarrays have been used to analyze the genetic
polymorphisms of specific loci associated with resistance to antimicrobial agents, to explore the distribution of genes among isolates from the
same and similar species, to analyze quorum-sensing systems, to understand the evolutionary relationship between closely related species
and to study host-pathogen interactions, particularly by identifying
genes from pathogens that may be involved in pathogenicity and by
surveying the host response to infection (101–107).
Fluorescence In Situ Hybridization
Fluorescence in situ hybridization (FISH) with rRNA-targeted oligonucleotide probes has been developed for in situ identification of
individual microbial cells (108). This technique detects nucleic acid
sequences by a fluorescently labeled probe that hybridizes specifically
to its complementary target sequence within the intact cell. In addition
to provide identification, FISH gives information about presence, morphology, number, organization and spatial distribution of micro-organisms (109). FISH not only allows the detection of cultivable microbial
species, but also of as-yet uncultivable micro-organisms (110).
rRNAs are the main target molecules for FISH. This is because they
can be found in all living organisms; they are relatively stable and occur
in high copy numbers (usually several thousands per cell); and they
include both variable and highly conserved sequence domains (45).
Signature sequences unique to a given group of micro-organisms, ranging from whole phyla to individual species, can be identified. In the vast
majority of applications for identification of bacteria or archaea, FISH
probes target 16S rRNA (109). The oligonucleotide probes used in FISH
are generally between 15 and 30 nucleotides long and covalently linked
at the 5⬘-end to a single fluorescent dye molecule. Common fluorophors
include fluorescein, tetramethylrhodamine, Texas red and carbocyanine dyes like Cy3 or Cy5 (108, 109).
A typical FISH protocol includes four steps: the fixation and permeabilization of the sample; hybridization with the respective probes for
detecting the respective target sequences; washing steps to remove unbound probe; and the detection of labeled cells by microscopy or flow
cytometry (111).
Limitations of Molecular Methods
Molecular techniques have been used to overcome the limitations
of cultivation procedures. Nonetheless, as with all methods, they are not
without their own limitations. The following discussion verses on the
major shortcomings associated with the most used molecular techniques.
DNA-DNA Hybridization
As with most molecular assays, DNA-DNA hybridization assays,
such as the checkerboard technique, have been used only to detect
target microbial species and consequently fail to detect the unexpected.
For whole genomic probe preparation, bacteria must be initially cultured. Consequently, DNA-DNA hybridization assays using whole genome probes can only detect cultivable species. In addition, whole
genomic probes used in the DNA-DNA hybridization assays may show
cross-reactivity with nontarget micro-organisms because of the presMolecular Methods in Endodontic Microbiology: Part 1
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TABLE 3. Limitations of PCR-derived technologies
a)
b)
c)
d)
e)
f)
g)
Most PCR assays used for identification purposes qualitatively detect the target microorganism but not its levels in the sample.
Quantitative results can however be obtained in real-time PCR assays.
Most PCR assays only detect one species or a few different species (multiplex PCR) at a time. However, broad-range PCR
analysis can provide information about the identity of virtually all species in a community.
Like DNA-DNA hybridization, most PCR assays only detect target species and consequently fail to detect unexpected species.
This can be overcome by broad-range PCR assays.
In addition to being laborious and costly, broad-range PCR analyses can be affected by some factors, such as biases in
homogenization procedures (76), preferential DNA amplification (112, 113) and differential DNA extraction (114, 115).
Microorganisms with thick cell walls, such as fungi, may be difficult to break open and may require additional steps for lysis
and consequent DNA release to occur.
False positive results have the potential to occur because of PCR amplification of contaminant DNA. The most important
means of contamination is through carryover of amplification product and special precautions should be taken to avoid this.
False negatives may occur because of enzyme inhibitors or nucleases present in clinical samples, which may abort the
amplification reaction and degrade the DNA template, respectively. Analysis of small sample volumes may also lead to false
negative results, particularly if the target species is present in low numbers.
ence of homologous sequences between different bacterial species. This
can give rise to false positive results. However, Socransky et al. (86, 88)
reported that most probes tested in the checkerboard assay did not
exhibit cross-reactions under the experimental conditions. Some
probes, as those to Tannerella forsythia, Porphyromonas gingivalis,
and Treponema denticola, did not exhibit cross-reactions with any
heterologous species tested. When cross-reactions were observed they
were always within genera and were quite limited (88).
Reverse-capture checkerboard assay using oligonucleotide
probes may show higher specificity than the conventional checkerboard
technique and probes can be devised to target and detect uncultivable
species (89).
PCR-Derived Techniques
Table 3 lists the main limitations of PCR technology. Many of them
can be gotten around by variations in technology. The issues related to
the ability of PCR to detect either an extremely low number of cells or
dead cells are of special interest when one interprets the results of PCR
identification procedures in endodontic microbiology research. Therefore, these issues are worth a separate detailed discussion.
The “Too-High Sensitivity” Issue
The high detection rate of PCR may be a reason of concern, specifically when nonquantitative assays are employed. It has been commonly claimed that because PCR can detect a very low number of cells
of a given microbial species, the results obtained by this method may
have no significance with regard to disease causation. Nevertheless,
some other factors should be also addressed in this discussion, which
represent great advantages of such highly sensitive identification procedure in endodontic research.
When taking samples from endodontic infections, difficulties
posed by the physical constraints of the root canal system and by the
limitations of the sampling techniques can make it difficult the attainment of a representative sample from the main canal (116). If cells of a
given species are sampled in a number below of the detection rate of the
diagnostic test, species prevalence will be underestimated.
It is also important to take into consideration the analytical sensitivity needed for the specific clinical sample. For example, a sensitivity of
about 104 microbial cells per ml is required for urine, while a sensitivity
of one cell may be of extreme relevance for samples from blood or
cerebrospinal fluid (117). There is no clear evidence as to the microbial load necessary for a periradicular lesion to be induced. Endodontic
infections are characterized by a mixed infection and individual species
may play different roles in the consortium or dominate various stages of
the infection. At least theoretically, all bacterial species established in
the infected root canal have the potential to be considered endodontic
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pathogens (118). Based on this, it would be glaringly prudent to use the
method with the highest sensitivity to detect all species colonizing the
root canal.
In fact, detection of very low numbers of cells in clinical samples
may not be as common as anticipated. There are numerous factors that
can influence PCR reactions, sometimes dramatically reducing sensitivity for direct microbial detection in clinical samples. Thus, the analytical
sensitivity of the method does not always correspond to its “clinical”
sensitivity. It is well known that the effects of inhibitors are magnified in
samples with low number of target DNA and therefore can significantly
decrease the sensitivity of the method (62). Another impediment refers
to the aliquots of the whole sample used in PCR reactions. Most of the
PCR assays that have been used so far in endodontic research have an
analytical sensitivity of about 10 to 100 cells. If one takes into account
the dilution factor dictated by the use of small aliquots (5–10%) of the
whole sample in each amplification reaction, the actual sensitivity of the
assay is 200 to 400 to 2000 to 4000 cells in the whole sample, without
discounting the effects of inhibitors (57). These numbers are still lower
than the detection limits of other methods, but can represent more
significance with regard to etiology.
Therefore, the use of highly sensitive techniques is welcome in the
study of endodontic infections, decreasing the risks for potentially important species to pass unnoticed during sample analysis. Although
qualitative results do not lack significance, the use of quantitative molecular assays, like the real-time PCR, can allow inference of the role of
a given species in the infectious process while maintaining high sensitivity and the ability to detect fastidious or uncultivable microbial species.
The “Dead-Cell” Issue
Detection of dead cells by a given identification method can be
regarded as a double-edged sword. This can be at the same time an
advantage and a limitation of the method. On the one hand, this ability
can allow detection of uncultivable bacteria or fastidious bacteria that
can die during sampling, transportation or isolation procedures (57,
119, 120). On the other hand, if the bacteria were already dead in the
infected site, they may also be detected and this might give rise to a false
assumption about their role in the infectious process (121, 122).
Several studies have addressed this issue. In a mouse model of
Lyme disease, it was revealed by PCR that Borrelia burgdorferi DNA
disappeared from tissues immediately after a 5-day antibiotic treatment
course, and that the PCR results correlated well with culture results
(123). Similarly, investigators using a chinchilla model of otitis media
found a strong correlation between PCR results and culture results when
animals were injected with viable Haemophilus influenza (124).
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When animals were injected with purified bacterial DNA or with killed
bacteria, the amplifiable DNA rapidly disappeared. The authors concluded that DNA, and DNA from intact but nonviable bacteria, does not
persist in an amplifiable form for more than a day in the presence of an
effusion. In a rabbit model of syphilis in which live or heat-killed Treponema pallidum was injected into the skin and testes, heat-killed treponemes were no longer detectable by PCR after 15 to 30 days, whereas
viable micro-organisms were detected by PCR for months (125). A
study of pulmonary tuberculosis treatment showed that sputum smears
and cultures for Mycobacterium tuberculosis convert to negative before PCR results, but that PCR results do correlate with clinical response
to antibiotics and also can predict relapse (126). It was not clear if the
persistent PCR signal seen in some of these patients was a result of low
levels of viable micro-organisms, or because of amplifiable DNA from
nonviable micro-organisms. All these studies show that bacterial DNA
can be cleared from the host sites after bacterial death and that DNA
from different species may differ as to the elimination kinetics at different body sites. It remains to be clarified for how long bacterial DNA from
dead cells can remain detectable in the infected root canal system.
Detection of microbial DNA sequences in clinical specimens does
not indicate viability of the micro-organism. However, this issue should
be addressed with common sense and without any sort of biases. The
fact that some micro-organisms die during the course of an infectious
process does not necessarily implicate that in a determined moment
these micro-organisms did not participate in the pathogenesis of the
disease. In addition, the fate of DNA from micro-organisms that have
entered and not survived in root canals is unknown. DNA from dead
cells might be adsorbed by dentine because of affinity of hydroxyapatite
to this molecule (127). However, it remains to be shown if DNA from
microbial dead cells can really be adsorbed in dentinal walls and, if
even, it can be retrieved during sampling with paper points. In fact, it is
highly unlikely that free microbial DNA can remain intact in an environment colonized by living micro-organisms. Dnases released by some
living species as well as at cell death can degrade free DNA in the
environment. It has been reported the presence of Dnase activity on
whole bacterial cells and vesicles thereof that can degrade linear DNA
(128). These bacteria include common putative endodontic pathogens,
such as Porphyromonas endodontalis, P. gingivalis, T. forsythia, Fusobacterium species, Prevotella intermedia, and Prevotella nigrescens. Indeed, Dnases are of concern during sample storage, because
they can be carried along with the sample and induce DNA degradation,
with consequent false negative result after PCR amplification.
Therefore, although there is a possibility of detecting DNA from
dead cells in endodontic infections, this possibility is conceivably low. In
the event DNA from dead cells is detected, the results by no means lack
significance with regard to participation in disease causation. However,
the ability to detect DNA from dead cells poses a major problem when
one is investigating the immediate effectiveness of antimicrobial treatment, because DNA from cells that have recently died can be detected.
To circumvent or at least minimize this problem, one can use some
adjustments in the PCR assay or take advantage of PCR technology derivatives, such as RT-PCR. McCarty and Atlas (129) investigated the
effect of amplicon size on the PCR detection of Legionella pneumophila
after chlorine inactivation. Two amplicons specific to the L. pneumophila mip gene were used for the PCR analyses. A 168-bp amplicon was
detectable longer after chlorine treatment than a 650-bp amplicon. On
the basis of their data, they concluded that larger amplicons appear to
correlate better with bacterial viability. Findings from their study
showed that smaller fragments of DNA may persist for a longer time after
cell death than larger sequences. Therefore, designing primers to generate large amplicons may reduce the risks of positive results because of
DNA from dead cells. Moreover, because RNAs are more labile and have
JOE — Volume 31, Number 6, June 2005
a shorter half-life when compared to DNA, they can be rapidly degraded
after cell death (122). Thus, assays directed towards the detection of
rRNA or mRNA through RT-PCR can be more reliable for detection of
living cells.
Unraveling the Oral Microbiota
The oral cavity harbors one of the highest accumulations of microorganisms in the body. Even though viruses, yeasts and protozoa can be
found as constituents of the oral microbiota, bacteria are by far the most
dominant inhabitants of the oral cavity. There are an estimated 1010
bacteria in the mouth (130). A high diversity of bacterial species has
been revealed in the oral cavity by cultivation and application of molecular methods to the analysis of the bacterial diversity has revealed a still
more broad and diverse spectrum of extant oral bacteria. Overall, bacteria detected from the oral cavity fall into 11 phyla that comprised over
700 different species or phylotypes (17, 131, 132). About 40 to 50% of
these bacteria are novel uncultivated species or phylotypes, which are
known only by 16S rDNA sequences (17). This raises the interesting
possibility that uncultivated and as-yet-uncharacterized species that
have remained invisible to studies using traditional identification methods may make up a large fraction of the living oral microbiota, and may
participate in the etiology of oral diseases.
In fact, the introduction of molecular approaches in oral microbiology research has brought about a significant body of new knowledge
with regard to the oral microbiota in health and disease. The development of molecular bacterial identification methods has made it possible
to study the role that uncultivable bacteria or even other micro-organisms such as archaea play in oral diseases (133–137). Consequently, a
significant revolution in the knowledge of the oral microbiota in health
and disease has taken place after the advent of molecular techniques for
microbial identification. In this context, endodontic infections are far
from being an exception. The second part of this review deals with the
significant impact of molecular genetic methods for microbial identification on the knowledge of endodontic infections.
Acknowledgment
This study was supported in part by grants from CNPq, a Brazilian Governmental Institution.
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