Original Paper Caries Res 2014;48:451–460 DOI: 10.1159/000357920
Received: October 8, 2013 Accepted after revision: December 8, 2013 Published online: May 15, 2014
Molecular Studies of the Structural Ecology of Natural Occlusal Caries Irene Dige Lene Grønkjær Bente Nyvad Section for Dental Pathology, Operative Dentistry and Endodontics, Department of Dentistry, Aarhus University, Aarhus, Denmark
Abstract Microbiological studies of occlusal dental biofilms have hitherto been hampered by inaccessibility to the sampling site and demolition of the original biofilm architecture. This study shows for the first time the spatial distribution of bacterial taxa in vivo at various stages of occlusal caries, applying a molecular methodology involving preparation of embedded hard dental tissue slices for fluorescence in situ hybridization (FISH) and confocal microscopy. Eleven freshly extracted teeth were classified according to their occlusal caries status. The teeth were fixed, embedded, sectioned and decalcified before FISH was performed using oligonucleotide probes for selected abundant species/genera associated with occlusal caries including Streptococcus, Actinomyces, Veillonella, Fusobacterium, Lactobacillus and Bifidobacterium. The sites showed distinct differences in the bacterial composition between different ecological niches in occlusal caries. Biofilm observed along the entrance of fissures showed an inner layer of microorganisms organized in palisades often identified as Actinomyces, covered by a more loosely structured bacterial layer consisting of diverse genera, similar to supragingival biofilm. Biofilm within the fissure proper seemed less metabolically active, as judged by
© 2014 S. Karger AG, Basel 0008–6568/14/0485–0451$39.50/0 E-Mail [email protected] www.karger.com/cre
low fluorescence signal intensity and presence of material of non-bacterial origin. Bacterial invasion (often Lactobacillus and Bifidobacterium spp.) into the dentinal tubules was seen only at advanced stages of caries with manifest cavity formation. It is concluded that the molecular methodology represents a valuable supplement to previous methods for the study of microbial ecology in caries by allowing analysis of the structural composition of the undisturbed biofilm in caries lesions in vivo. © 2014 S. Karger AG, Basel
The etiological role of biofilms in the development of caries is indisputable. Much effort has been made to study and analyze the bacterial composition and structure of biofilms in caries lesions, especially in relation with occlusal caries. Occlusal surfaces continue to carry a major burden of caries in contemporary populations [Schwarz et al., 1994; Brown and Selwitz, 1995]. Therefore, studies exploring the ecology of occlusal caries are highly warranted. Previous studies of the microbiology of occlusal caries have typically used needles or dental explorers for sampling of bacteria in natural fissures [Meiers et al., 1982; Loesche et al., 1984; Meiers and Schachtele, 1984], in artificial fissure models [Loe et al., 1973; Igarashi et al., 1990], or in in situ models [Theilade et al., 1976, 1978]. Evaluation of the composition of the biofilm was done by classical cultivation of the bacteria. These studIrene Dige Section for Dental Pathology, Operative Dentistry and Endodontics Department of Dentistry, Aarhus University Vennelyst Boulevard 9, DK–8000 Aarhus C (Denmark) E-Mail irene.dige @ odontologi.au.dk
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Key Words Biofilms · Confocal laser scanning microscopy · Fluorescence in situ hybridization · Occlusal caries
ies have contributed with important information about the predominant composition of the biofilm in sound and carious occlusal surfaces but do not include yet uncultured organisms [Mikkelsen et al., 2000; Thurnheer et al., 2001]. These organisms may now be identified in studies using molecular techniques, most of which are based on 16S rRNA sequencing data such as PCR-based, reverse capture checkerboard hybridization methods. However, most molecular studies of the microbial composition of dental biofilms comparing the microbiota in patients with and without caries have primarily been performed on pooled plaque samples [Becker et al., 2002; Aas et al., 2008; Peterson et al., 2013]. The lack of individual site-specific sampling is an obvious disadvantage when trying to relate bacterial compositions with the stage of caries lesion progression and the ecology of caries [Nyvad et al., 2013]. Molecular studies dealing with the microbiota in occlusal caries [Arif et al., 2008; Mantzourani et al., 2009; Lima et al., 2011] have performed site-specific sampling but mainly focused on severe dentin caries, leaving out information about non-cavitated stages. Sampling of occlusal fissures is hampered by inaccessibility to the defined site and may remove only a limited fraction of the microbiota in fissures [Meiers and Schachtele, 1984]. It is also important to appreciate that sampling always damages the natural structure of the biofilm and therefore does not include information about the architecture of the biofilms in different locations. Early studies described the structural composition of the microbiota in occlusal fissures by electron microscopy [Theilade et al., 1976; Ekstrand and Bjørndal, 1997]. These studies offered detailed information about bacterial structure (cell wall, intracellular material), differentiation between Gram-positive and Gram-negative bacteria as well as between different morphologies of bacteria. However, differentiation between species or genera was not possible in vivo until fluorescence in situ hybridization (FISH) was introduced [Amann et al.,
1990]. This method has successfully been used to explore the structural composition of in situ dental biofilms [Diaz et al., 2006; Al-Ahmad et al., 2009; Dige et al., 2009] and may help expand our understanding of the spatial distributions and functions of intact microbial communities on teeth. Recently, FISH methodologies were refined for the application on in vivo biofilms using embedded teeth for visualization of colonization patterns in periapical lesions [Sunde et al., 2003] and biofilm architecture of supra- and subgingival biofilms [Zijnge et al., 2010]. So far no studies have studied in vivo biofilms of carious dental tissues such as occlusal caries by the use of FISH and fluorescence microscopy. Therefore, the aim of the present study was to explore the spatial distribution of commonly encountered bacterial taxa in vivo at various stages of occlusal caries, applying a new methodology involving preparation of embedded hard dental tissue slices and FISH and confocal microscopy.
Fig. 1. Images of in vivo biofilms on dental occlusal surfaces. a– c Toluidine blue-stained sections showing an overview of occlusal surfaces with shallow-fissure-like morphology (a), groovelike morphology (b) and cavitated caries lesion (c). Arrows refer
(purple/magenta in e, g–i) nor Fusobacterium spp. (purple/magenta in f). Note that the biofilm could be divided into an inner compact layer of palisade-like bacteria (d–h) often with a columnar pattern (g, h) on top of which a looser-structured layer (d, e, f, h, i) with non-stained voids (d, i) was seen. The outermost part of the decalcified enamel showed a thin auto-fluorescent layer without bacteria (blue or green in d, g), and invaginations of developmental origin were often filled with bacteria (d, g, arrows). All images are oriented with the biofilm surface upwards. Scale bars: 500 μm (a–c) and 25 μm (d–i). (For figure see next page.)
to the areas illustrated in figure 2d, 2b, 2i, 2j, 1g and 2l, respectively. d–i Confocal laser scanning microscopy images of microbial colonization pattern from above the entrance of shallow fissures and groove-like occlusal surfaces. In all confocal laser scanning microscopy images, red represents all bacteria that are neither Streptococcus spp. (yellow-green in d–i) nor Actinomyces spp.
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Caries Res 2014;48:451–460 DOI: 10.1159/000357920
Sample Collection The teeth were extracted as part of the patients’ general treatment in a private practice in Denmark. Eleven freshly extracted teeth obtained from eleven patients were included in the study. Collection of the teeth was performed anonymously and approved by the Central Denmark Region Committees on Biomedical Research Ethics (case no. 1-10-72-50-13). Patients gave oral informed consent that the teeth could be used for research purposes. Clinical Characterization of Lesions The teeth were photographed using a digital camera (Zeiss AxioCam MRc5, Carl Zeiss Micro Imaging GmbH, Göttingen, Germany) mounted on a Wild Macroskop M420 (Wild, Heerbrugg, Switzerland) and classified according to the occlusal caries status (active/inactive/sound; cavitated/non-cavitated) using the visual criteria of Nyvad [Nyvad et al., 1999]. Twelve occlusal sites were examined, of which five were classified as active and noncavitated, three as active and cavitated, two as inactive and noncavitated, and two as clinically sound.
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Materials and Methods
Structural Ecology of Natural Occlusal Caries in vivo
Caries Res 2014;48:451–460 DOI: 10.1159/000357920
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1
Table 1. Target bacteria and sequences, labels and hybridization conditions of used oligonucleotide probes Probe
Target bacteria
Target sequence (5′–3′)
Reference Formamide 5′ modification concentration1
STR405
most streptococci
TAG CCG TCC CTT TCT GGT
Alexa488
25–30%
Paster et al., 1998
ACT476
Actinomyces spp.
ATC CAG CTA CCG TCA ACC
Atto550
30%
Gmür and Lüthi-Schaller, 2007; Dige et al., 2009
FUS664
most Fusobacterium
CTT GTA GTT CCG CYT ACC TC (Y=C/T) Cy3
25%
Thurnheer et al., 2004; Al-Ahmad et al., 2009; Zijnge et al., 2010
E79
Veillonella spp.
AAT CCC CTC CTT CAG TGA
Atto633
25%
Al-Ahmad et al., 2007; Al-Ahmad et al., 2009
MIT447
mitis group of streptococci
CAC YCG TTC TTC TCT TAC A (Y=C/T)
Atto633
25%
Thurnheer et al., 2001; Quevedo et al., 2011
LGC358a
most lactobacilli
CCA TTG TGG AAG ATT CCC T
Cy3
25%
Quevedo et al., 2011
25%
Quevedo et al., 2011
LGC358b-comp competitor probe for LGC358a
CCA TTG CGG AAG ATT CCC T
MUT590
Streptococcus mutans
ACT CCA GAC TTT CCT GAC
Cy3
25%
Thurnheer et al., 2001; Quevedo et al., 2011
BIF164-mod
Bifidobacterium genus
CAT CCG GYA TTA CCA CCC
Cy3
0%
Sim et al., 2012
EUB338
most bacteria
GCT GCC TCC CGT AGG AGT
Atto633/Cy3 0–50%
formamide concentration in hybridization buffer.
Sample Preparation Immediately after extraction, the teeth were rinsed briefly in running tap water and stored in individual vials at –20 ° C until they could be fixed in 3% paraformaldehyde in PBS for 16 h at 4 ° C. The specimens were subsequently washed with PBS and stored in a mixture of 60% ethanol and 40% PBS at –20 ° C until embedding in Technovit 8100 (Heraeus Kultzer GmbH, Wehrheim, Germany) at 4 ° C according to the method described by Zijnge et al. [2010]. The embedded teeth were cut into 1 mm plano-parallel slices perpendicularly to the occlusal surface (Leitz Saw Microtome 1600, Ernst Leitz Wetzlar GmbH, Wetzlar, Germany). Cut slices were photographed (Zeiss AxioCam MRc5 mounted on a Wild Macroskop M420) to obtain an overview of the lesion. Subsequently, the embedded slices were decalcified (17% ethylenediamine-tetraacetic acid) for 14–25 days. Complete decalcification was verified by digital X-ray analysis. The decalcified slices were photographed, re-embedded in Technovit 8100 and stored at 4 ° C. For technical reasons slices were subdivided into smaller areas of interest before re-embedding, e.g. carious enamel and dentin were often analyzed separately. Then 1–2-μm-thin sections were cut perpendicular to the fissure or groove of the occlusal surface as illustrated in figure 1a–c using a glass knife in an ultramicrotome (Reichert Jung Ultracut E, Reichert Jung Optische Werke AG, Vienna, Austria). The sections were stretched on water and mounted on polysine precoated glass slides (Menzel-Gläser, Thermo Scientific, Braunschweig, Germany) for FISH analysis. One random section of each lesion/site was stained by toluidine blue.
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FISH The 16S rRNA targeted oligonucleotide probes used in the study are summarized in table 1. The STR405 were purchased from MWG Biotech (Ebersberg, Germany) and the remaining from IBA (Göttingen, Germany). The probe LGC358a was used in in conjunction with a competitor probe LGC358b-comp. FISH was carried out according to the procedure described by Dige et al. [2009], modified from Manz et al. [1992], using formamide concentrations as recorded in table 1, except for BIF164-mod, which was hybridized at 0% formamide at 50 ° C for 16 h [Sim et al., 2012]. Prior to the FISH procedure all specimens were permeabilized with 25 μl lysozyme (Sigma) (70 U/μl in 100 mM Tris/HCl pH 7.5 [Sigma], 5 mM EDTA [Merck]) for 9 min at 37 ° C in a humid chamber except for the probe LGC358a where lysozyme, achromopeptidase (1 mg/ ml; Sigma-Aldrich A-3547) and lipase (Sigma-Aldrich L-1754) were used according to Quevedo et al. [2011]. The following combination of probes was used on all lesion sites: ACT476-Atto 550/STR405-Alexa488/EUB338-Atto633, FUS664-Cy3/STR405Alexa488/EUB338-Atto633, E79-Atto633/STR 405-Alexa488/EUB 338-Cy3, MIT447-Atto633/STR405-Alexa488/EUB338-Cy3, LGC 358a-Cy3/LGC358b-comp/STR405-Alexa488/EUB338-Atto633, MUT590-Cy3/STR405-Alexa488/EUB338- Atto633, and BIF164mod-Cy3/EUB338-Atto633. All 16S rRNA targeted oligonucleotide probes used in the study have been used and published before (table 1). The sequence specificities of the probes were tested against pure cultures of strains with zero number of mismatches to the probe sequence using the following strains: Fusobacterium nucleatum CCUG 9126 (=ATCC10953), F. nucleatum NCO 10562
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1 Optimum
Amann et al., 1990
Microscopy Toluidine blue sections were examined under a light microscope (Olympus BX51, Olympus Corporation, Tokyo, Japan) to get an overview of the biofilm in the fissures in vivo. The biofilms were examined on an inverted Zeiss LSM 510 META (Carl Zeiss, Jena, Germany) confocal laser scanning microscope. Laser lines 488, 543 and 633 nm were used for excitation of Alexa448, Atto550/ Cy3 and Atto633, respectively. Fluorescence emission from Alexa488 was detected between 500 and 550 nm with a bandpass filter (BP500–550), while fluorescence emission from Atto550/ Cy3 was detected above 560 nm with a long pass filter (LP560). Atto633 fluorescence was detected with the META detector set to 651–704 nm. Images of each section were obtained using a PlanApochromat 63×/1.40 oil immersion objective at fixed settings. Color micrographs were processed using ImageJ and Adobe Photoshop CS4 (Adobe) without any qualitative changes of the raw images.
Results
All of the occlusal surfaces of the extracted teeth were colonized by biofilms, and the architecture of the biofilms was easily observed by FISH and confocal laser scanning microscopy. The occlusal surfaces had different morphologies of the groove-fossa system. Three sites were classified as having a fissure-like morphology and six sites as having a groove-like morphology (as defined by Ekstrand and Bjørndal [1997]), whereas three sites showed cavitation in the enamel (for examples see fig. 1a– c, respectively). Distinct differences were noted in the structural composition between the different sites and stages of occlusal caries.
layer of palisade-like bacteria, mostly short rods and often Actinomyces spp. (fig. 1g). A columnar pattern between different species/genera perpendicular to the enamel surface was observed (fig. 1g, h). On top of the condensed inner layer a looser biofilm layer was seen with a random arrangement of bacteria (fig. 1d–f). The bacteria were often arranged around voids of non-stained material (fig. 1i). The bacterial composition was diverse (based on positive signal after hybridizations with species/genera-specific oligonucleotide probes and different morphologies of EUB338-labeled cells). At all sound and non-cavitated carious sites Streptococcus spp., including S. mitis, Veillonella spp. and Fusobacterium spp. were observed. Veillonella spp. appeared most often in areas with Streptococcus spp. (fig. 2a), whereas Fusobacterium spp. were scattered in the inner part of the outer layer (fig. 1f, 2b). Streptococcus spp., including S. mitis, appeared both as distinct colonies and scattered in the outer layer of the biofilm (fig. 2c). S. mutans was observed on sites with both active and inactive caries, but not on clinically sound enamel, whereas Bifidobacterium spp. were only detected in sites with active caries. Lactobacillus spp. did neither appear on clinically sound nor on non-cavitated sites. Apart from that, there was no obvious difference in the structural composition and amount of microbial biofilm between clinically sound, active and inactive carious lesion sites. The outermost part of the enamel was lined by a thin auto-fluorescent layer, likely to represent proteinaceous tissue of developmental origin (for examples see fig. 1d, g). In some cases bacteria were found to colonize irregular projections of the auto-fluorescent material extending into the enamel space, such as Tomes processes pits, openings of the striae of Retzius, and focal holes [Nyvad et al., 1988] (fig. 1d, g, arrows).
Groove-Like Occlusal Surfaces and above the Entrance of Shallow Fissures The pattern and degree of microbial coverage as well as the thickness of the biofilm covering the enamel along the occlusal surface including the entrance of the fissures differed between sections. In general, the biofilm could be divided into two layers (fig. 1d–f) of which the inner layer adhering to the enamel surface consisted of a compact
Inside the Shallow Fissures All shallow fissures contained biofilms. However, the biofilm deep in the fissures seemed less metabolically active judged by the fluorescence intensities of EUB338-labeled cells (fig. 2d). The fissures also comprised auto-fluorescent material of non-bacterial origin. The fluorescing bacteria were either surrounding or intermingled with this material (fig. 2e). Irrespectively of the caries activity status of the fissures, no principal differences in the architecture and amount of biofilm were observed. The composition of bacteria deep in the fissures seemed less diverse than the biofilm at the entrance of the fissures. Actinomyces spp. and a few colonies of streptococci as well as scattered streptococci were observed. Fu-
Structural Ecology of Natural Occlusal Caries in vivo
Caries Res 2014;48:451–460 DOI: 10.1159/000357920
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(NCTC; =ATCC10953) for the probe FUS664, Veillonella parvula CCUG 5123T (=ATCC 10790) for the probe E79, Streptococcus mitis SK24 for the probe MIT447, Lactobacillus orisT DSM 4864 for the probe LGC358a, Streptococcus mutans DSM 20523 for the probe MUT590, and Bifidobacterium dentiumT DSM 20436 for the probe BIF164-mod. Testing of STR405 and ACT476 has previously been reported [Dige et al., 2009]. F. nucleatum, V. parvula and S. mitis were kindly provided by M. Kilian, Department of Biomedicine, Aarhus University, Denmark. Before microscopy a cover glass was mounted with Citifluor AF1 (Citifluor Ltd., Leicester, UK). A total of 105 sections were analyzed.
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Dige/Grønkjær/Nyvad
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(For legend see next page.)
This study used a new methodology [Zijnge et al., 2010] based on FISH analysis of embedded dental tissue slices to study the structural composition of the undisturbed biofilm at varying stages of caries lesion formation in occlusal surfaces in vivo. Different ecological niches in occlusal caries showed distinct differences in the bacterial composition of the most abundant species/genera of the microbiota and in metabolic activity, as judged by fluorescence intensity. For the first time, the spatial arrangement of aciduric bacteria such as Lactobacillus spp. and
Bifidobacterium spp. in manifest dentin caries lesions was visualized. The new methodology used in the present study overcomes the latent problem of bacterial dispersion of microbial samples for studies of the ecology of caries in vivo by allowing a three-dimensional reconstruction of the spatial arrangement of bacteria within caries lesions. However, there are limitations to this method as well. Studies of population dynamics cannot be carried out because the caries-associated biofilm can only be evaluated at the time of tooth extraction. Moreover, only labeled bacteria may be visualized and FISH studies are limited by the small amount of available oligonucleotide probes that currently allow discrimination of the most commonly encountered bacterial taxa in dental biofilms. Another obstacle with fluorescence microcopy is the limited number of available fluorophores with non-overlapping emission signals, leaving only few microbial taxa to be differentiated in one image. This problem may, however, be surmounted by analyzing many adjacent sections using differently labeled oligonucleotide probes, as done in this study. The use of combinatorial labeling and spectral imaging FISH (CLASI-FISH), which may allow simultaneous visualization of 15 different taxa in one image, may also potentially solve the problem with overlapping emission signals in the future [Valm et al., 2011]. However, so far this method presents some complications with inaccurate image segmentation when different taxa overlap within the plane of focus or touching cells are not identified as separate objects. In the future, when more fluorophores and more fluorophore combinations can be mixed successfully, CLASI-FISH might be very useful for studies of thick multispecies biofilm such as in caries lesions. It should also be appreciated that the new methodology is technically difficult and time-consuming. Nevertheless, the technical difficulties are amply returned by the new information gained about the distribution and co-localization of bacterial species and genera in intact carious tissue.
Fig. 2. Images of microbial colonization patterns of in vivo biofilm on occlusal surfaces. Red represents all bacteria except for Streptococcus spp. (yellow-green in a–e, j), Veillonella spp. (purple/magenta in a), Fusobacterium spp. (purple/magenta in b), S. mitis (turquoise in c), Actinomyces spp. (purple/magenta in d), Lactobacillus spp. (purple/magenta in f, j, l), Bifidobacterium spp. (purple/ magenta in g, i, k) and S. mutans (yellow-green in h). Veillonella spp. (a) and S. mitis (c) appeared most often in the outer layers together with streptococci. Fusobacterium spp. formed an intermediate layer between the bottom and the superficial layer (b). In
the deeper part of shallow fissures (d, e) the biofilm showed less fluorescence signaling of the oligonucleotide probe-labelled bacteria. The fissures often contained auto-fluorescent material of nonbacterial origin (green in e) intermingled with bacteria. In cavitated caries lesions with penetration into the dentin (f–l), Lactobacillus spp. (f, j, l) and Bifidobacterium spp. (g, i, k) were abundant both inside the cavities (f, g) and in the surface layers of the biofilm at the entrance of the cavity (i, j). Bifidobacterium spp. (k) and Lactobacillus spp. (l) invaded the dentinal tubules. S. mutans were also present in all cavitated lesions (h). Scale bars: 25 μm.
Structural Ecology of Natural Occlusal Caries in vivo
Caries Res 2014;48:451–460 DOI: 10.1159/000357920
Cavitated Enamel and Dentin The principal structural composition of the biofilm in active cavitated enamel lesions was rather similar to that observed inside the shallow fissures. However, rods and filamentous bacteria often identified as Lactobacillus spp. (fig. 2f) or Bifidobacterium spp. (fig. 2g) were prominent. S. mutans was observed in all sections with cavitation (fig. 2h). When cavitation was observed, Bifidobacterium spp. and Lactobacillus spp. were also seen in the outer layers of the biofilm at the entrance to the cavity (fig. 2i, j). Bacterial invasion with penetration into the dentinal tubules occurred only at advanced stages of the caries process with manifest cavity formation. The bacteria were identified by the use of the EUB338 probe and had a filamentous morphology, some of them being Bifidobacterium spp. (fig. 2k) or Lactobacillus spp. (fig. 2l). Autofluorescent material was detected within the enamel cavities and at the dentin-enamel junction, probably representing tuft protein [Robinson and Hudson, 2011].
Discussion
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sobacterium spp., Veillonella spp., Lactobacillus spp. and Bifidobacterium spp. were not frequent residents inside the shallow fissures. Abundance of S. mutans was detected in one active lesion site with a fissure-like morphology.
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ence of both inter- and intracellular mineralization in shallow fissures [Galil and Gwinnett, 1975; Theilade et al., 1976; Ekstrand and Bjørndal, 1997]. Galil and Gwinnett [1975] reported that 70% of 574 fissures showed progressive mineralization, especially at the bottom part of fissures, and concluded that the mineralization process begins there. This corresponds well with experimental in situ studies of artificial fissures constructed with ion-sensitive field-effect transistor electrodes that showed a decreased acidogenic response at the base of mature fissure biofilms [Igarashi et al., 1990]. At cavitation we observed a marked enrichment of aciduric bacteria including S. mutans, Bifidobacterium spp. and Lactobacillus spp., all of which have previously been associated with advanced caries with cavitation [Edwardsson, 1974; Munson et al., 2004; Aas et al., 2008; Mantzourani et al., 2009; Lima et al., 2011]. This supports the idea that it is the biochemical processes such as environmental acidification that are the main driving force of the phenotypic and genotypic changes of the microbiota in caries [Takahashi and Nyvad, 2008, 2011]. In agreement with previous studies [Thylstrup and Qvist, 1987; Ricketts et al., 2002], bacterial invasion into dentin was noted only in stages of caries with manifest cavity formation. Furthermore, only filamentous bacteria, many of which belonged to Bifidobacterium and Lactobacillus, were found to invade the dentinal tubules. It is interesting that we did not localize S. mutans in the demineralized dentin and therefore this species may not seem to have a propensity to penetrate into dentinal tubules. Because of the small number of samples examined, it is not possible to say whether such differences in the colonization pattern between S. mutans and Bifidobacterium/Lactobacillus spp. reflect different ecological preferences. The finding of mutans streptococci in other studies of deep carious dentin may be explained by contamination during sampling, as suggested by Edwardsson [1974]. Careful sampling of dentin caries from well-defined sites of extracted teeth [Edwardsson, 1974; Hoshino, 1985] has shown a similar pattern of distribution of the above-mentioned bacteria as in our study. In conclusion, this descriptive study using a new hard tissue preparation technique in combination with FISH has shown that there is a specific pattern of bacterial invasion into carious dentin. This might indicate that conventional sampling techniques for microbiological analysis are often too crude for making conclusions on caries ecology. Irrespective of the precision of the sampling technique, the architecture of the biofilm and the spatial distribution of the most abundant species/genera associated Dige/Grønkjær/Nyvad
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The structural features and the bacterial composition of biofilm in grooves and above the entrance of fissures had a striking resemblance with supragingival in vivo biofilms [Zijnge et al., 2010] and confirmed observations from transmission electron microscopic studies [Theilade et al., 1976; Ekstrand and Bjørndal, 1997]. For example in this study, a palisade arrangement of the bacteria in the basal layer of the biofilm at the entrance of occlusal grooves was often identified as Actinomyces spp. and thus verifies the preferential localization of Actinomyces spp. in the inner part of biofilms [Dige et al., 2009]. By contrast, the outer part of the biofilm was more loosely structured and the bacteria were arranged between non-stained voids, possibly representing extracellular material [Newman, 1979; Nyvad and Fejerskov, 1989]. Interestingly, Fusobacterium spp. preferred a niche between the inner and outer layer, an observation that has also recently been reported in supragingival biofilm [Zijnge et al., 2010], supporting the theory that Fusobacterium spp. act as a bridging organism by binding to both early and later colonizers [Kolenbrander et al., 2006]. Streptococci did not show a clear preferential distribution in the biofilm. However, it is a new observation that the streptococci in the outer layer predominantly belonged to the S. mitis group, which may imply that these bacteria were new colonizers [Nyvad and Kilian, 1987; Li et al., 2004]. Likewise, Veillonella spp. were located in the outer layer, often in close association with streptococci. A metabolic partnership between streptococci and veillonellae has been suggested to dampen the caries challenge because of veillonellae converting lactic acid produced by streptococci into weaker acids [Mikx et al., 1972; van der Hoeven et al., 1978; Periasamy and Kolenbrander, 2010]. The biofilm within the fissure proper was less diverse than that found above the entrance to the fissures. In agreement with Ekstrand and Bjørndal [1997], we observed that the bacteria in the fissures were less metabolically active. In the latter study low metabolic activity of the microorganisms was judged by morphological criteria such as disintegrating microorganisms, non-dividing microorganism or empty ghost-like cells, whereas in our FISH study viability relied on the occurrence of ribosomal RNA in the bacteria. While both methods may be inadequate for evaluating cell viability the findings are, nevertheless, indicative of a low metabolic activity deep in the fissures. In addition, we observed auto-fluorescent material possibly consisting of developmental protein [Ekstrand et al., 1991] and/or organic matrix of calculus [Embery, 1989]. Several researchers have shown the pres-
with occlusal caries can only be studied reliably on whole tissue sections. No single method will be able to answer all relevant questions of caries ecology; however, we believe that the current methodology represents a valuable supplement to previous methods for the study of microbial communities in caries. Together with other molecular techniques dealing with composition and function, it may contribute to a better understanding of the role of biofilms in the caries process.
fully acknowledge the financial support from the Danish Dental Association (FORSKU). The funders had no role in study design, data collection and analysis, decision to publish or preparation of the manuscript.
Author Contributions I. Dige, B. Nyvad and L. Grønkjær conceived and designed the experiments; I. Dige and L. Grønkjær performed the experiments; I. Dige and B. Nyvad analyzed the data; I. Dige, B. Nyvad and L. Grønkjær wrote and approved the paper.
Acknowledgments We thank the dentists in the dental practice ‘Tandlægerne i Møldrup’ for providing freshly extracted teeth, and Dr. Merete Raarup for help with the confocal microscope set-up. We grate-
Disclosure Statement The authors declare that there are no conflicts of interest.
References
Structural Ecology of Natural Occlusal Caries in vivo
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Takahashi N, Nyvad B: Caries ecology revisited: microbial dynamics and the caries process. Caries Res 2008;42:409–418. Takahashi N, Nyvad B: The role of bacteria in the caries process: ecological perspectives. J Dent Res 2011;90:294–303. Theilade E, Fejerskov O, Karring T, Theilade J: A microbiological study of old plaque in occlusal fissures of human teeth. Caries Res 1978; 12:313–319. Theilade J, Fejerskov O, Horsted M: A transmission electron microscopic study of 7-day old bacterial plaque in human tooth fissures. Arch Oral Biol 1976;21:587–598. Thurnheer T, Gmür R, Giertsen E, Guggenheim B: Automated fluorescent in situ hybridization for the specific detection and quantification of oral streptococci in dental plaque. J Microbiol Methods 2001;44:39–47. Thurnheer T, Gmür R, Guggenheim B: Multiplex FISH analysis of a six-species bacterial biofilm. J Microbiol Methods 2004;56:37–47. Thylstrup A, Qvist V: Principal enamel and dentine reactions during caries progressions; in Thylstrup A, Leach SA, Qvist V (eds): Dentine and Dentine Reactions in the Oral Cavity. Oxford, IRL Press, 1987, pp 3–16. Valm AM, Mark Welch JL, Rieken CW, Hasegawa Y, Sogin ML, Oldenbourg R, Dewhirst FE, Borisy GG: Systems-level analysis of microbial community organization through combinatorial labeling and spectral imaging. Proc Natl Acad Sci USA 2011;108:4152–4157. van der Hoeven JS, Toorop AI, Mikx RH: Symbiotic relationship of Veillonella alcalescens and Streptococcus mutans in dental plaque in gnotobiotic rats. Caries Res 1978;12:142–147. Zijnge V, van Leeuwen MB, Degener JE, Abbas F, Thurnheer T, Gmür R, Harmsen HJ: Oral biofilm architecture on natural teeth. PLoS One 2010;5:e9321.
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Mikx FH, van der Hoeven JS, König KG, Plasschaert AJ, Guggenheim B: Establishment of defined microbial ecosystems in germ-free rats. I. The effect of the interactions of Streptococcus mutans or Streptococcus sanguis with Veillonella alcalescens on plaque formation and caries activity. Caries Res 1972;6:211–223. Munson MA, Banerjee A, Watson TF, Wade WG: Molecular analysis of the microflora associated with dental caries. J Clin Microbiol 2004; 42:3023–3029. Newman HN: The host-organism interface in natural human dental plaque. J Dent 1979; 7: 235–245. Nyvad B, Crielaard W, Mira A, Takahashi N, Beighton D: Dental caries from a molecular microbiological perspective. Caries Res 2013; 47:89–102. Nyvad B, Fejerskov O: Structure of dental plaque and the plaque-enamel interface in human experimental caries. Caries Res 1989;23:151– 158. Nyvad B, Fejerskov O, Josephsen K: Organic structures of developmental origin in human surface enamel. Scand J Dent Res 1988; 96: 288–292. Nyvad B, Kilian M: Microbiology of the early colonization of human enamel and root surfaces in vivo. Scand J Dent Res 1987;95:369–380. Nyvad B, Machiulskiene V, Baelum V: Reliability of a new caries diagnostic system differentiating between active and inactive caries lesions. Caries Res 1999;33:252–260. Paster BJ, Bartoszyk IM, Dewhirst FE: Identification of oral streptococci using PCR-based, reverse-capture, checkerboard hybridization. Methods Cell Sci 1998;20:223–231. Periasamy S, Kolenbrander PE: Central role of the early colonizer Veillonella sp. in establishing multispecies biofilm communities with initial, middle, and late colonizers of enamel. J Bacteriol 2010;192:2965–2972.
Received: October 8, 2013 Accepted after revision: December 8, 2013 Published online: May 15, 2014
Molecular Studies of the Structural Ecology of Natural Occlusal Caries Irene Dige Lene Grønkjær Bente Nyvad Section for Dental Pathology, Operative Dentistry and Endodontics, Department of Dentistry, Aarhus University, Aarhus, Denmark
Abstract Microbiological studies of occlusal dental biofilms have hitherto been hampered by inaccessibility to the sampling site and demolition of the original biofilm architecture. This study shows for the first time the spatial distribution of bacterial taxa in vivo at various stages of occlusal caries, applying a molecular methodology involving preparation of embedded hard dental tissue slices for fluorescence in situ hybridization (FISH) and confocal microscopy. Eleven freshly extracted teeth were classified according to their occlusal caries status. The teeth were fixed, embedded, sectioned and decalcified before FISH was performed using oligonucleotide probes for selected abundant species/genera associated with occlusal caries including Streptococcus, Actinomyces, Veillonella, Fusobacterium, Lactobacillus and Bifidobacterium. The sites showed distinct differences in the bacterial composition between different ecological niches in occlusal caries. Biofilm observed along the entrance of fissures showed an inner layer of microorganisms organized in palisades often identified as Actinomyces, covered by a more loosely structured bacterial layer consisting of diverse genera, similar to supragingival biofilm. Biofilm within the fissure proper seemed less metabolically active, as judged by
© 2014 S. Karger AG, Basel 0008–6568/14/0485–0451$39.50/0 E-Mail [email protected] www.karger.com/cre
low fluorescence signal intensity and presence of material of non-bacterial origin. Bacterial invasion (often Lactobacillus and Bifidobacterium spp.) into the dentinal tubules was seen only at advanced stages of caries with manifest cavity formation. It is concluded that the molecular methodology represents a valuable supplement to previous methods for the study of microbial ecology in caries by allowing analysis of the structural composition of the undisturbed biofilm in caries lesions in vivo. © 2014 S. Karger AG, Basel
The etiological role of biofilms in the development of caries is indisputable. Much effort has been made to study and analyze the bacterial composition and structure of biofilms in caries lesions, especially in relation with occlusal caries. Occlusal surfaces continue to carry a major burden of caries in contemporary populations [Schwarz et al., 1994; Brown and Selwitz, 1995]. Therefore, studies exploring the ecology of occlusal caries are highly warranted. Previous studies of the microbiology of occlusal caries have typically used needles or dental explorers for sampling of bacteria in natural fissures [Meiers et al., 1982; Loesche et al., 1984; Meiers and Schachtele, 1984], in artificial fissure models [Loe et al., 1973; Igarashi et al., 1990], or in in situ models [Theilade et al., 1976, 1978]. Evaluation of the composition of the biofilm was done by classical cultivation of the bacteria. These studIrene Dige Section for Dental Pathology, Operative Dentistry and Endodontics Department of Dentistry, Aarhus University Vennelyst Boulevard 9, DK–8000 Aarhus C (Denmark) E-Mail irene.dige @ odontologi.au.dk
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Key Words Biofilms · Confocal laser scanning microscopy · Fluorescence in situ hybridization · Occlusal caries
ies have contributed with important information about the predominant composition of the biofilm in sound and carious occlusal surfaces but do not include yet uncultured organisms [Mikkelsen et al., 2000; Thurnheer et al., 2001]. These organisms may now be identified in studies using molecular techniques, most of which are based on 16S rRNA sequencing data such as PCR-based, reverse capture checkerboard hybridization methods. However, most molecular studies of the microbial composition of dental biofilms comparing the microbiota in patients with and without caries have primarily been performed on pooled plaque samples [Becker et al., 2002; Aas et al., 2008; Peterson et al., 2013]. The lack of individual site-specific sampling is an obvious disadvantage when trying to relate bacterial compositions with the stage of caries lesion progression and the ecology of caries [Nyvad et al., 2013]. Molecular studies dealing with the microbiota in occlusal caries [Arif et al., 2008; Mantzourani et al., 2009; Lima et al., 2011] have performed site-specific sampling but mainly focused on severe dentin caries, leaving out information about non-cavitated stages. Sampling of occlusal fissures is hampered by inaccessibility to the defined site and may remove only a limited fraction of the microbiota in fissures [Meiers and Schachtele, 1984]. It is also important to appreciate that sampling always damages the natural structure of the biofilm and therefore does not include information about the architecture of the biofilms in different locations. Early studies described the structural composition of the microbiota in occlusal fissures by electron microscopy [Theilade et al., 1976; Ekstrand and Bjørndal, 1997]. These studies offered detailed information about bacterial structure (cell wall, intracellular material), differentiation between Gram-positive and Gram-negative bacteria as well as between different morphologies of bacteria. However, differentiation between species or genera was not possible in vivo until fluorescence in situ hybridization (FISH) was introduced [Amann et al.,
1990]. This method has successfully been used to explore the structural composition of in situ dental biofilms [Diaz et al., 2006; Al-Ahmad et al., 2009; Dige et al., 2009] and may help expand our understanding of the spatial distributions and functions of intact microbial communities on teeth. Recently, FISH methodologies were refined for the application on in vivo biofilms using embedded teeth for visualization of colonization patterns in periapical lesions [Sunde et al., 2003] and biofilm architecture of supra- and subgingival biofilms [Zijnge et al., 2010]. So far no studies have studied in vivo biofilms of carious dental tissues such as occlusal caries by the use of FISH and fluorescence microscopy. Therefore, the aim of the present study was to explore the spatial distribution of commonly encountered bacterial taxa in vivo at various stages of occlusal caries, applying a new methodology involving preparation of embedded hard dental tissue slices and FISH and confocal microscopy.
Fig. 1. Images of in vivo biofilms on dental occlusal surfaces. a– c Toluidine blue-stained sections showing an overview of occlusal surfaces with shallow-fissure-like morphology (a), groovelike morphology (b) and cavitated caries lesion (c). Arrows refer
(purple/magenta in e, g–i) nor Fusobacterium spp. (purple/magenta in f). Note that the biofilm could be divided into an inner compact layer of palisade-like bacteria (d–h) often with a columnar pattern (g, h) on top of which a looser-structured layer (d, e, f, h, i) with non-stained voids (d, i) was seen. The outermost part of the decalcified enamel showed a thin auto-fluorescent layer without bacteria (blue or green in d, g), and invaginations of developmental origin were often filled with bacteria (d, g, arrows). All images are oriented with the biofilm surface upwards. Scale bars: 500 μm (a–c) and 25 μm (d–i). (For figure see next page.)
to the areas illustrated in figure 2d, 2b, 2i, 2j, 1g and 2l, respectively. d–i Confocal laser scanning microscopy images of microbial colonization pattern from above the entrance of shallow fissures and groove-like occlusal surfaces. In all confocal laser scanning microscopy images, red represents all bacteria that are neither Streptococcus spp. (yellow-green in d–i) nor Actinomyces spp.
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Sample Collection The teeth were extracted as part of the patients’ general treatment in a private practice in Denmark. Eleven freshly extracted teeth obtained from eleven patients were included in the study. Collection of the teeth was performed anonymously and approved by the Central Denmark Region Committees on Biomedical Research Ethics (case no. 1-10-72-50-13). Patients gave oral informed consent that the teeth could be used for research purposes. Clinical Characterization of Lesions The teeth were photographed using a digital camera (Zeiss AxioCam MRc5, Carl Zeiss Micro Imaging GmbH, Göttingen, Germany) mounted on a Wild Macroskop M420 (Wild, Heerbrugg, Switzerland) and classified according to the occlusal caries status (active/inactive/sound; cavitated/non-cavitated) using the visual criteria of Nyvad [Nyvad et al., 1999]. Twelve occlusal sites were examined, of which five were classified as active and noncavitated, three as active and cavitated, two as inactive and noncavitated, and two as clinically sound.
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Materials and Methods
Structural Ecology of Natural Occlusal Caries in vivo
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1
Table 1. Target bacteria and sequences, labels and hybridization conditions of used oligonucleotide probes Probe
Target bacteria
Target sequence (5′–3′)
Reference Formamide 5′ modification concentration1
STR405
most streptococci
TAG CCG TCC CTT TCT GGT
Alexa488
25–30%
Paster et al., 1998
ACT476
Actinomyces spp.
ATC CAG CTA CCG TCA ACC
Atto550
30%
Gmür and Lüthi-Schaller, 2007; Dige et al., 2009
FUS664
most Fusobacterium
CTT GTA GTT CCG CYT ACC TC (Y=C/T) Cy3
25%
Thurnheer et al., 2004; Al-Ahmad et al., 2009; Zijnge et al., 2010
E79
Veillonella spp.
AAT CCC CTC CTT CAG TGA
Atto633
25%
Al-Ahmad et al., 2007; Al-Ahmad et al., 2009
MIT447
mitis group of streptococci
CAC YCG TTC TTC TCT TAC A (Y=C/T)
Atto633
25%
Thurnheer et al., 2001; Quevedo et al., 2011
LGC358a
most lactobacilli
CCA TTG TGG AAG ATT CCC T
Cy3
25%
Quevedo et al., 2011
25%
Quevedo et al., 2011
LGC358b-comp competitor probe for LGC358a
CCA TTG CGG AAG ATT CCC T
MUT590
Streptococcus mutans
ACT CCA GAC TTT CCT GAC
Cy3
25%
Thurnheer et al., 2001; Quevedo et al., 2011
BIF164-mod
Bifidobacterium genus
CAT CCG GYA TTA CCA CCC
Cy3
0%
Sim et al., 2012
EUB338
most bacteria
GCT GCC TCC CGT AGG AGT
Atto633/Cy3 0–50%
formamide concentration in hybridization buffer.
Sample Preparation Immediately after extraction, the teeth were rinsed briefly in running tap water and stored in individual vials at –20 ° C until they could be fixed in 3% paraformaldehyde in PBS for 16 h at 4 ° C. The specimens were subsequently washed with PBS and stored in a mixture of 60% ethanol and 40% PBS at –20 ° C until embedding in Technovit 8100 (Heraeus Kultzer GmbH, Wehrheim, Germany) at 4 ° C according to the method described by Zijnge et al. [2010]. The embedded teeth were cut into 1 mm plano-parallel slices perpendicularly to the occlusal surface (Leitz Saw Microtome 1600, Ernst Leitz Wetzlar GmbH, Wetzlar, Germany). Cut slices were photographed (Zeiss AxioCam MRc5 mounted on a Wild Macroskop M420) to obtain an overview of the lesion. Subsequently, the embedded slices were decalcified (17% ethylenediamine-tetraacetic acid) for 14–25 days. Complete decalcification was verified by digital X-ray analysis. The decalcified slices were photographed, re-embedded in Technovit 8100 and stored at 4 ° C. For technical reasons slices were subdivided into smaller areas of interest before re-embedding, e.g. carious enamel and dentin were often analyzed separately. Then 1–2-μm-thin sections were cut perpendicular to the fissure or groove of the occlusal surface as illustrated in figure 1a–c using a glass knife in an ultramicrotome (Reichert Jung Ultracut E, Reichert Jung Optische Werke AG, Vienna, Austria). The sections were stretched on water and mounted on polysine precoated glass slides (Menzel-Gläser, Thermo Scientific, Braunschweig, Germany) for FISH analysis. One random section of each lesion/site was stained by toluidine blue.
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FISH The 16S rRNA targeted oligonucleotide probes used in the study are summarized in table 1. The STR405 were purchased from MWG Biotech (Ebersberg, Germany) and the remaining from IBA (Göttingen, Germany). The probe LGC358a was used in in conjunction with a competitor probe LGC358b-comp. FISH was carried out according to the procedure described by Dige et al. [2009], modified from Manz et al. [1992], using formamide concentrations as recorded in table 1, except for BIF164-mod, which was hybridized at 0% formamide at 50 ° C for 16 h [Sim et al., 2012]. Prior to the FISH procedure all specimens were permeabilized with 25 μl lysozyme (Sigma) (70 U/μl in 100 mM Tris/HCl pH 7.5 [Sigma], 5 mM EDTA [Merck]) for 9 min at 37 ° C in a humid chamber except for the probe LGC358a where lysozyme, achromopeptidase (1 mg/ ml; Sigma-Aldrich A-3547) and lipase (Sigma-Aldrich L-1754) were used according to Quevedo et al. [2011]. The following combination of probes was used on all lesion sites: ACT476-Atto 550/STR405-Alexa488/EUB338-Atto633, FUS664-Cy3/STR405Alexa488/EUB338-Atto633, E79-Atto633/STR 405-Alexa488/EUB 338-Cy3, MIT447-Atto633/STR405-Alexa488/EUB338-Cy3, LGC 358a-Cy3/LGC358b-comp/STR405-Alexa488/EUB338-Atto633, MUT590-Cy3/STR405-Alexa488/EUB338- Atto633, and BIF164mod-Cy3/EUB338-Atto633. All 16S rRNA targeted oligonucleotide probes used in the study have been used and published before (table 1). The sequence specificities of the probes were tested against pure cultures of strains with zero number of mismatches to the probe sequence using the following strains: Fusobacterium nucleatum CCUG 9126 (=ATCC10953), F. nucleatum NCO 10562
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1 Optimum
Amann et al., 1990
Microscopy Toluidine blue sections were examined under a light microscope (Olympus BX51, Olympus Corporation, Tokyo, Japan) to get an overview of the biofilm in the fissures in vivo. The biofilms were examined on an inverted Zeiss LSM 510 META (Carl Zeiss, Jena, Germany) confocal laser scanning microscope. Laser lines 488, 543 and 633 nm were used for excitation of Alexa448, Atto550/ Cy3 and Atto633, respectively. Fluorescence emission from Alexa488 was detected between 500 and 550 nm with a bandpass filter (BP500–550), while fluorescence emission from Atto550/ Cy3 was detected above 560 nm with a long pass filter (LP560). Atto633 fluorescence was detected with the META detector set to 651–704 nm. Images of each section were obtained using a PlanApochromat 63×/1.40 oil immersion objective at fixed settings. Color micrographs were processed using ImageJ and Adobe Photoshop CS4 (Adobe) without any qualitative changes of the raw images.
Results
All of the occlusal surfaces of the extracted teeth were colonized by biofilms, and the architecture of the biofilms was easily observed by FISH and confocal laser scanning microscopy. The occlusal surfaces had different morphologies of the groove-fossa system. Three sites were classified as having a fissure-like morphology and six sites as having a groove-like morphology (as defined by Ekstrand and Bjørndal [1997]), whereas three sites showed cavitation in the enamel (for examples see fig. 1a– c, respectively). Distinct differences were noted in the structural composition between the different sites and stages of occlusal caries.
layer of palisade-like bacteria, mostly short rods and often Actinomyces spp. (fig. 1g). A columnar pattern between different species/genera perpendicular to the enamel surface was observed (fig. 1g, h). On top of the condensed inner layer a looser biofilm layer was seen with a random arrangement of bacteria (fig. 1d–f). The bacteria were often arranged around voids of non-stained material (fig. 1i). The bacterial composition was diverse (based on positive signal after hybridizations with species/genera-specific oligonucleotide probes and different morphologies of EUB338-labeled cells). At all sound and non-cavitated carious sites Streptococcus spp., including S. mitis, Veillonella spp. and Fusobacterium spp. were observed. Veillonella spp. appeared most often in areas with Streptococcus spp. (fig. 2a), whereas Fusobacterium spp. were scattered in the inner part of the outer layer (fig. 1f, 2b). Streptococcus spp., including S. mitis, appeared both as distinct colonies and scattered in the outer layer of the biofilm (fig. 2c). S. mutans was observed on sites with both active and inactive caries, but not on clinically sound enamel, whereas Bifidobacterium spp. were only detected in sites with active caries. Lactobacillus spp. did neither appear on clinically sound nor on non-cavitated sites. Apart from that, there was no obvious difference in the structural composition and amount of microbial biofilm between clinically sound, active and inactive carious lesion sites. The outermost part of the enamel was lined by a thin auto-fluorescent layer, likely to represent proteinaceous tissue of developmental origin (for examples see fig. 1d, g). In some cases bacteria were found to colonize irregular projections of the auto-fluorescent material extending into the enamel space, such as Tomes processes pits, openings of the striae of Retzius, and focal holes [Nyvad et al., 1988] (fig. 1d, g, arrows).
Groove-Like Occlusal Surfaces and above the Entrance of Shallow Fissures The pattern and degree of microbial coverage as well as the thickness of the biofilm covering the enamel along the occlusal surface including the entrance of the fissures differed between sections. In general, the biofilm could be divided into two layers (fig. 1d–f) of which the inner layer adhering to the enamel surface consisted of a compact
Inside the Shallow Fissures All shallow fissures contained biofilms. However, the biofilm deep in the fissures seemed less metabolically active judged by the fluorescence intensities of EUB338-labeled cells (fig. 2d). The fissures also comprised auto-fluorescent material of non-bacterial origin. The fluorescing bacteria were either surrounding or intermingled with this material (fig. 2e). Irrespectively of the caries activity status of the fissures, no principal differences in the architecture and amount of biofilm were observed. The composition of bacteria deep in the fissures seemed less diverse than the biofilm at the entrance of the fissures. Actinomyces spp. and a few colonies of streptococci as well as scattered streptococci were observed. Fu-
Structural Ecology of Natural Occlusal Caries in vivo
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(NCTC; =ATCC10953) for the probe FUS664, Veillonella parvula CCUG 5123T (=ATCC 10790) for the probe E79, Streptococcus mitis SK24 for the probe MIT447, Lactobacillus orisT DSM 4864 for the probe LGC358a, Streptococcus mutans DSM 20523 for the probe MUT590, and Bifidobacterium dentiumT DSM 20436 for the probe BIF164-mod. Testing of STR405 and ACT476 has previously been reported [Dige et al., 2009]. F. nucleatum, V. parvula and S. mitis were kindly provided by M. Kilian, Department of Biomedicine, Aarhus University, Denmark. Before microscopy a cover glass was mounted with Citifluor AF1 (Citifluor Ltd., Leicester, UK). A total of 105 sections were analyzed.
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(For legend see next page.)
This study used a new methodology [Zijnge et al., 2010] based on FISH analysis of embedded dental tissue slices to study the structural composition of the undisturbed biofilm at varying stages of caries lesion formation in occlusal surfaces in vivo. Different ecological niches in occlusal caries showed distinct differences in the bacterial composition of the most abundant species/genera of the microbiota and in metabolic activity, as judged by fluorescence intensity. For the first time, the spatial arrangement of aciduric bacteria such as Lactobacillus spp. and
Bifidobacterium spp. in manifest dentin caries lesions was visualized. The new methodology used in the present study overcomes the latent problem of bacterial dispersion of microbial samples for studies of the ecology of caries in vivo by allowing a three-dimensional reconstruction of the spatial arrangement of bacteria within caries lesions. However, there are limitations to this method as well. Studies of population dynamics cannot be carried out because the caries-associated biofilm can only be evaluated at the time of tooth extraction. Moreover, only labeled bacteria may be visualized and FISH studies are limited by the small amount of available oligonucleotide probes that currently allow discrimination of the most commonly encountered bacterial taxa in dental biofilms. Another obstacle with fluorescence microcopy is the limited number of available fluorophores with non-overlapping emission signals, leaving only few microbial taxa to be differentiated in one image. This problem may, however, be surmounted by analyzing many adjacent sections using differently labeled oligonucleotide probes, as done in this study. The use of combinatorial labeling and spectral imaging FISH (CLASI-FISH), which may allow simultaneous visualization of 15 different taxa in one image, may also potentially solve the problem with overlapping emission signals in the future [Valm et al., 2011]. However, so far this method presents some complications with inaccurate image segmentation when different taxa overlap within the plane of focus or touching cells are not identified as separate objects. In the future, when more fluorophores and more fluorophore combinations can be mixed successfully, CLASI-FISH might be very useful for studies of thick multispecies biofilm such as in caries lesions. It should also be appreciated that the new methodology is technically difficult and time-consuming. Nevertheless, the technical difficulties are amply returned by the new information gained about the distribution and co-localization of bacterial species and genera in intact carious tissue.
Fig. 2. Images of microbial colonization patterns of in vivo biofilm on occlusal surfaces. Red represents all bacteria except for Streptococcus spp. (yellow-green in a–e, j), Veillonella spp. (purple/magenta in a), Fusobacterium spp. (purple/magenta in b), S. mitis (turquoise in c), Actinomyces spp. (purple/magenta in d), Lactobacillus spp. (purple/magenta in f, j, l), Bifidobacterium spp. (purple/ magenta in g, i, k) and S. mutans (yellow-green in h). Veillonella spp. (a) and S. mitis (c) appeared most often in the outer layers together with streptococci. Fusobacterium spp. formed an intermediate layer between the bottom and the superficial layer (b). In
the deeper part of shallow fissures (d, e) the biofilm showed less fluorescence signaling of the oligonucleotide probe-labelled bacteria. The fissures often contained auto-fluorescent material of nonbacterial origin (green in e) intermingled with bacteria. In cavitated caries lesions with penetration into the dentin (f–l), Lactobacillus spp. (f, j, l) and Bifidobacterium spp. (g, i, k) were abundant both inside the cavities (f, g) and in the surface layers of the biofilm at the entrance of the cavity (i, j). Bifidobacterium spp. (k) and Lactobacillus spp. (l) invaded the dentinal tubules. S. mutans were also present in all cavitated lesions (h). Scale bars: 25 μm.
Structural Ecology of Natural Occlusal Caries in vivo
Caries Res 2014;48:451–460 DOI: 10.1159/000357920
Cavitated Enamel and Dentin The principal structural composition of the biofilm in active cavitated enamel lesions was rather similar to that observed inside the shallow fissures. However, rods and filamentous bacteria often identified as Lactobacillus spp. (fig. 2f) or Bifidobacterium spp. (fig. 2g) were prominent. S. mutans was observed in all sections with cavitation (fig. 2h). When cavitation was observed, Bifidobacterium spp. and Lactobacillus spp. were also seen in the outer layers of the biofilm at the entrance to the cavity (fig. 2i, j). Bacterial invasion with penetration into the dentinal tubules occurred only at advanced stages of the caries process with manifest cavity formation. The bacteria were identified by the use of the EUB338 probe and had a filamentous morphology, some of them being Bifidobacterium spp. (fig. 2k) or Lactobacillus spp. (fig. 2l). Autofluorescent material was detected within the enamel cavities and at the dentin-enamel junction, probably representing tuft protein [Robinson and Hudson, 2011].
Discussion
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sobacterium spp., Veillonella spp., Lactobacillus spp. and Bifidobacterium spp. were not frequent residents inside the shallow fissures. Abundance of S. mutans was detected in one active lesion site with a fissure-like morphology.
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Caries Res 2014;48:451–460 DOI: 10.1159/000357920
ence of both inter- and intracellular mineralization in shallow fissures [Galil and Gwinnett, 1975; Theilade et al., 1976; Ekstrand and Bjørndal, 1997]. Galil and Gwinnett [1975] reported that 70% of 574 fissures showed progressive mineralization, especially at the bottom part of fissures, and concluded that the mineralization process begins there. This corresponds well with experimental in situ studies of artificial fissures constructed with ion-sensitive field-effect transistor electrodes that showed a decreased acidogenic response at the base of mature fissure biofilms [Igarashi et al., 1990]. At cavitation we observed a marked enrichment of aciduric bacteria including S. mutans, Bifidobacterium spp. and Lactobacillus spp., all of which have previously been associated with advanced caries with cavitation [Edwardsson, 1974; Munson et al., 2004; Aas et al., 2008; Mantzourani et al., 2009; Lima et al., 2011]. This supports the idea that it is the biochemical processes such as environmental acidification that are the main driving force of the phenotypic and genotypic changes of the microbiota in caries [Takahashi and Nyvad, 2008, 2011]. In agreement with previous studies [Thylstrup and Qvist, 1987; Ricketts et al., 2002], bacterial invasion into dentin was noted only in stages of caries with manifest cavity formation. Furthermore, only filamentous bacteria, many of which belonged to Bifidobacterium and Lactobacillus, were found to invade the dentinal tubules. It is interesting that we did not localize S. mutans in the demineralized dentin and therefore this species may not seem to have a propensity to penetrate into dentinal tubules. Because of the small number of samples examined, it is not possible to say whether such differences in the colonization pattern between S. mutans and Bifidobacterium/Lactobacillus spp. reflect different ecological preferences. The finding of mutans streptococci in other studies of deep carious dentin may be explained by contamination during sampling, as suggested by Edwardsson [1974]. Careful sampling of dentin caries from well-defined sites of extracted teeth [Edwardsson, 1974; Hoshino, 1985] has shown a similar pattern of distribution of the above-mentioned bacteria as in our study. In conclusion, this descriptive study using a new hard tissue preparation technique in combination with FISH has shown that there is a specific pattern of bacterial invasion into carious dentin. This might indicate that conventional sampling techniques for microbiological analysis are often too crude for making conclusions on caries ecology. Irrespective of the precision of the sampling technique, the architecture of the biofilm and the spatial distribution of the most abundant species/genera associated Dige/Grønkjær/Nyvad
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The structural features and the bacterial composition of biofilm in grooves and above the entrance of fissures had a striking resemblance with supragingival in vivo biofilms [Zijnge et al., 2010] and confirmed observations from transmission electron microscopic studies [Theilade et al., 1976; Ekstrand and Bjørndal, 1997]. For example in this study, a palisade arrangement of the bacteria in the basal layer of the biofilm at the entrance of occlusal grooves was often identified as Actinomyces spp. and thus verifies the preferential localization of Actinomyces spp. in the inner part of biofilms [Dige et al., 2009]. By contrast, the outer part of the biofilm was more loosely structured and the bacteria were arranged between non-stained voids, possibly representing extracellular material [Newman, 1979; Nyvad and Fejerskov, 1989]. Interestingly, Fusobacterium spp. preferred a niche between the inner and outer layer, an observation that has also recently been reported in supragingival biofilm [Zijnge et al., 2010], supporting the theory that Fusobacterium spp. act as a bridging organism by binding to both early and later colonizers [Kolenbrander et al., 2006]. Streptococci did not show a clear preferential distribution in the biofilm. However, it is a new observation that the streptococci in the outer layer predominantly belonged to the S. mitis group, which may imply that these bacteria were new colonizers [Nyvad and Kilian, 1987; Li et al., 2004]. Likewise, Veillonella spp. were located in the outer layer, often in close association with streptococci. A metabolic partnership between streptococci and veillonellae has been suggested to dampen the caries challenge because of veillonellae converting lactic acid produced by streptococci into weaker acids [Mikx et al., 1972; van der Hoeven et al., 1978; Periasamy and Kolenbrander, 2010]. The biofilm within the fissure proper was less diverse than that found above the entrance to the fissures. In agreement with Ekstrand and Bjørndal [1997], we observed that the bacteria in the fissures were less metabolically active. In the latter study low metabolic activity of the microorganisms was judged by morphological criteria such as disintegrating microorganisms, non-dividing microorganism or empty ghost-like cells, whereas in our FISH study viability relied on the occurrence of ribosomal RNA in the bacteria. While both methods may be inadequate for evaluating cell viability the findings are, nevertheless, indicative of a low metabolic activity deep in the fissures. In addition, we observed auto-fluorescent material possibly consisting of developmental protein [Ekstrand et al., 1991] and/or organic matrix of calculus [Embery, 1989]. Several researchers have shown the pres-
with occlusal caries can only be studied reliably on whole tissue sections. No single method will be able to answer all relevant questions of caries ecology; however, we believe that the current methodology represents a valuable supplement to previous methods for the study of microbial communities in caries. Together with other molecular techniques dealing with composition and function, it may contribute to a better understanding of the role of biofilms in the caries process.
fully acknowledge the financial support from the Danish Dental Association (FORSKU). The funders had no role in study design, data collection and analysis, decision to publish or preparation of the manuscript.
Author Contributions I. Dige, B. Nyvad and L. Grønkjær conceived and designed the experiments; I. Dige and L. Grønkjær performed the experiments; I. Dige and B. Nyvad analyzed the data; I. Dige, B. Nyvad and L. Grønkjær wrote and approved the paper.
Acknowledgments We thank the dentists in the dental practice ‘Tandlægerne i Møldrup’ for providing freshly extracted teeth, and Dr. Merete Raarup for help with the confocal microscope set-up. We grate-
Disclosure Statement The authors declare that there are no conflicts of interest.
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