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Department of Oral Biology, School of Dental Medicine1 and Department of Otolaryngology, School of Medicine and Biomedical Sciences2, State University of New York at Buffalo, Buffalo, NY 14214, USA
Correspondence Frank A. Scannapieco fas1{at}acsu.buffalo.edu
Received September 3, 2002
Accepted February 24, 2003
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Abstract |
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 TOP Abstract INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPAbstractINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Reports of bacterial growth in chronic maxillary sinusitis range from 36 % of mucosal specimens taken by translabial antroscopy (Jiang et al., 1998) to 92 % of sinus aspirates (Brook et al., 1997). However, only a limited number of bacterial genera have been detected using conventional culture techniques (Wilson et al., 1997). Novel culture-independent methods that involve PCR amplification of bacterial small-subunit rDNA followed by cloning and sequencing have recently been used to determine the diversity of bacterial flora within specific environments, such as deep-sea sediments (Kato et al., 1997) and hot springs (Hugenholtz et al., 1998). In humans, studies using these methods suggest that about 40 % of clones in the oral cavity (Paster et al., 2001) and about 70 % of clones in the gut and colon (Suau et al., 1999; Wilson & Blitchington, 1996) do not correspond to known organisms, and are therefore novel (and probably non-cultivable) species. A previous study found PCR to be more efficient than standard culture for the detection of aerobic bacteria in chronic rhinosinusitis; such bacteria were detected in 62 % of samples tested by PCR, compared to 50 % using culture techniques (Keech et al., 2000). However, definitive studies that describe the total bacterial flora of the chronically inflamed maxillary sinus using molecular methods are not yet available.
Oral infections and inflammatory diseases such as periodontitis may influence systemic health by contributing to underlying mechanisms responsible for chronic coronary heart disease (Beck et al., 1999), pregnancy complications (Offenbacher et al., 1998), diabetes (Grossi & Genco, 1998) and respiratory infections (Scannapieco, 1999). The oral cavity may also serve as a reservoir for infection of contiguous structures such as the sinuses or middle ear. Oral bacteria may ascend from the mouth through the middle meatus to the maxillary sinus, or through the Eustachian tube to the middle ear, causing infections. Dental conditions such as periodontitis or periapical granuloma have been associated with 40 % of chronic maxillary sinusitis cases (Melen et al., 1986). Typically oral anaerobic bacteria such as Porphyromonas, Prevotella and Fusobacterium species can be detected by culture methods in infections of the middle ear (Brook et al., 2000), tonsils (Rajasuo et al., 1996) and sinuses (Le Moal et al., 1999). Aerobic species that normally reside in the oral cavity, such as viridans streptococci, have also been found in chronic sinusitis (Biel et al., 1998). Additionally, elevated serum levels of antibodies against F. nucleatum and Prevotella intermedia have been observed in patients with chronic maxillary sinusitis (Brook & Yocum, 1999).
The objective of this study was to determine whether bacteria were present in clinical samples from chronic maxillary sinusitis. We also attempted to determine if bacteria detected within the chronically inflamed maxillary sinus were simultaneously present in the oral cavity of the same patient. Using culture-independent methods, we also aimed to detect novel as-yet-uncultivable bacterial species in the chronically inflamed maxillary sinus.
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METHODS |
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TOPAbstractINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Specimen collection.
Washes and/or tissue samples from maxillary sinuses were obtained during standard surgical treatment as follows: after sterilization of the nasal vestibule and inferior meatus with ethanol, a sterile 18 G trocar needle was introduced into the maxillary sinus. Tissue specimens were aspirated directly from the sinus, or the sinus was irrigated with sterile saline and the fluid was removed with a sterile 10 cm3 syringe attached to the trocar. Oral swabs from the buccal marginal gingiva in the maxillary premolar regions of both sides of the mouth were also collected at the time of surgery. A portion of each maxillary sinus sample was sent to a clinical laboratory for microbiological culture, and the remaining samples were stored at -70 °C until they were processed for PCR analysis.
Microbiological culture.
Samples were transported to the clinical laboratory in brain heart infusion solution for routine cultivation of bacteria and fungi. Samples were cultured in thioglycollate broth and on blood, chocolate, MacConkey, Columbia CNA, Sabouraud and Mycosel (Becton Dickinson) agars. All media were maintained aerobically at 37 °C in 5 % CO2. Any growth in thioglycollate broth was subcultured to blood agar and incubated both aerobically and anaerobically. The lower limit of culture detection was approximately 100 c.f.u. (ml sample)-1.
Specimen digestion and DNA extraction.
Tissue samples and oral swabs were suspended in sterile dH2O. Maxillary washes (at least 4 ml) were centrifuged at 10 000 g for 30 min and the resulting pellet was suspended in sterile dH2O. An equal volume of lysis buffer containing 400 µg proteinase K ml-1, 2 % SDS (w/v), 100 mM Tris/HCl (pH 8.5) and 2 mM EDTA (Kroes et al., 1999) was added to all samples, which were then incubated at 55 °C overnight. DNA was extracted by phenol/chloroform extraction and ethanol precipitation. Isolation of DNA from each sample was confirmed by agarose gel electrophoresis and ethidium bromide staining; the DNA was stored at -70 °C until use.
PCR amplification of 16S rDNA.
PCR was performed in a reaction volume of 50 µl, consisting of 0.2 mM each dNTP (Invitrogen), 0.4 µM each primer, 5 µl 10x PCR buffer (Promega), 1.5 mM MgCl2 and 1 U Taq DNA polymerase (Promega). The three universal primer pairs used to amplify bacterial 16S rDNA were 27f (5'-AGAGTTTGATCCTGGCT CAG-3') and 1492r (5'-GGTTACCTTGTTACGACTT-3') (Lane et al., 1985), 63f (5'-CAGGCCTAACACATGCAAGTC-3') and 1387r [5'-GGGCGG(A/T)GTGTACAAGGC-3'] (Marchesi et al., 1998), and SDBact0338aA18 (Bact-338) (5'-ACTCCTACGGGAGGCAGC-3') and SDBact1525aS17 (Bact-1525) (5'-AAGGAGGTGATCCAGCC-3') (Kroes et al., 1999). Negative controls, consisting of a reagent control (sterile water used as a template in the PCR reagent mixture) and a control for sample preparation (sterile water prepared in the same way as the samples), were included in each set of PCR amplifications. Chromosomal DNA from Escherichia coli was used as a positive control. PCR amplification was performed with 30 cycles as follows: denaturation at 94 °C for 1 min, annealing at 55 °C for 1 min and extension at 72 °C for 2 min. Amplification products were separated by 1.0 % agarose gel electrophoresis and visualized under UV light after staining with ethidium bromide. To exclude false negative results caused by inhibitors in the sample, several dilutions of the original sample were used as a template, followed by a second PCR using the amplified product as a template. Each negative result was confirmed at least four times. In the literature, the lower detection limit for PCR has been estimated to be from a few bacterial cells to more than 100 c.f.u. per PCR (Ahmet et al., 1999; Gram et al., 1996; Rantakokko-Jalava et al., 2000); this variation is most likely to depend on the type of clinical sample studied (fluid versus tissue, or mixed versus single bacterial species).
Cloning of amplified 16S rDNA.
The 16S rDNA products were purified using the Wizard PCR Preps DNA purification system (Promega). The purified amplicon was ligated into the vector pGEM-T (Promega) and transformed into E. coli DH5
competent cells (Invitrogen), according to the manufacturers’ instructions. From each sample, 100–200 ampicillin-resistant transformants, identified as white colonies on LB agar containing IPTG (Invitrogen) and X-Gal (Fisher Scientific) after overnight incubation at 37 °C, were selected for further study. Cells from representative colonies were suspended in 50 µl dH2O, boiled, and the supernatant was used as a template in PCR with primers M13 forward (5'-GTAAAACGACGGCCAGT-3') and M13 reverse (5'-GGAAACAGCTATGACCATG-3') to determine if the clone had a correctly sized insert. PCR products with inserts of appropriate size were further used as templates in a second PCR with the bacterial-specific 16S rDNA primer pair used in the original PCR amplification. To reduce the overrepresentation of any species when selecting clones for sequencing, PCR products were digested with the restriction endonucleases HaeIII, BamHI and Sau3AI for 1–2 h, enzymes were inactivated by heating, and RFLP profiles were run on 1 % agarose gel (Kroes et al., 1999; Dymock et al., 1996). At least 20 positive inserts from each patient were analysed by RFLP, and additional clones were examined if more than one RFLP profile was distinguished. Up to five representative clones of each unique restriction pattern were selected for sequencing.
Sequencing and phylogenetic analysis.
Plasmids for sequencing were prepared by using the Wizard Plus SV Minipreps DNA purification system (Promega). DNA sequencing was performed using primer T7, complementary to the vector pGEM-T, by the Biopolymer Facility at the Roswell Park Cancer Institute, Buffalo, NY, USA. Sequences of inserts were compared to the 16S rRNA gene sequences in GenBank using BLAST software. DNA sequences sharing 
98 % identity with known sequences were assigned to that phylotype.
Detection of bacterial species identified from maxillary sinuses in oral samples from the same patient.
For each patient, an attempt was made to detect the species obtained from the sinus within the oral sample obtained at the time of sinus surgery. Primers specific to Streptococcus oralis/Streptococcus mitis (5'-GTCGAAGGTGATGATAT GAC-3' and 5'-CTGCATTCTGACGCATGACAG-3') (Garnier et al., 1997), Streptococcus gordonii (5'-TGATGAAGCTACTGATGC-3' and 5'-TAACAACGCTGCAGAAGACAA-3') (Brown et al., 1999), Staphylococcus aureus (5'-GTTATTAGGGAAGAACATATGTG-3' and 5'-CCACCTTCCTCCGGTTTGTCACC-3') (Jaffe et al., 2000), Pseudomonas aeruginosa (5'-AGGGCAGTAAGTTAATACCTTGCTG-3' and 5'-CCACCTCTACCGTACTCTAGCTCAG-3') (Wilson et al., 1999) and Serratia marcescens (5'-GGGAGCTTGCTCACTGGGTG-3' and 5'-GCGAGTAACGTCAGTTGATGAGCGTATTA-3') (Wilson et al., 1999) were used in PCR amplifications as described in this study and the original publications. Respective sinus clones were used as positive controls in PCR amplifications.
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RESULTS |
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TOPAbstractINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Possible novel species
The majority of the bacterial sequences identified in this study demonstrated 98 % identity with previously known bacterial species. No previously unrecognized species were recovered, but one clone from patient 4 demonstrated 94 % sequence identity with a Prevotella sp. oral clone, and one clone from patient 3 demonstrated 96 % sequence identity with an Enterococcus avium strain when > 1000 bp were compared. As < 97 % similarity is considered to differentiate species within genera (Stackebrandt & Goebel, 1994) and < 90 % similarity to represent a species from an unknown genus, these clones may represent novel species related to Prevotella or Enterococcus.
Relationship between bacteria detected in the maxillary sinus and the oral cavity by PCR
In patient 3, nine different bacterial clones were identified from the maxillary sinus; the majority of these clones were typically oral species such as Streptococcus sp., nutritionally variant streptococci, Eubacterium sp. or Neisseria sp. In the same patient, an attempt was made to detect the same species in the oral cavity. Indeed, a positive result was obtained with Streptococcus mitis/Streptococcus oralis primers, but a negative result was obtained with Streptococcus gordonii primers. In patient 4, a Prevotella clone with a possible oral origin was detected in the maxillary sinus. Also, in patient 4, attempts to find the same species by molecular methods from the oral cavity gave positive results with Pseudomonas aeruginosa primers and negative results with Staphylococcus aureus and Serratia marcescens primers. In patients 6 and 11, Staphylococcus aureus, which was present in the maxillary sample, was not detected in the oral sample by molecular methods. For the other species identified in the maxillary sinuses, specific primers were not available or the primers designed as described in previous studies did not work in our laboratory with the matched species DNA.
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DISCUSSION |
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TOPAbstractINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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We detected bacteria and/or bacterial 16S rDNA in five of 11 patients (45 %). Our results corroborate those of some previous studies, where bacteria have been detected in mucosal specimens from approximately 50 % of chronic maxillary sinusitis cases (Jiang et al., 1998; Hwang et al., 1999). However, other studies have reported that up to 92 % of chronic sinusitis samples contain bacteria (Brook et al., 1997). The type of specimen and the methods of detection may contribute to the different results observed. Our results are based upon analysis of mucosal tissues or maxillary sinus washes, because no exudates were obtained from these patients. The results must also be interpreted with caution, as the present study was limited to 11 subjects. Studies of larger populations using culture-independent methods are necessary.
Of the three primer pairs used to amplify bacterial DNA in our study, primers 27f and 1492r are broadly conserved and have previously been widely used to detect bacteria in human biopsies and body fluids (Dymock et al., 1996; Rantakokko-Jalava et al., 2000). To enhance detection of some specific oral species such as Prevotella, two additional bacteria-specific primer pairs that were designed to detect oral species were used (Kroes et al., 1999; Marchesi & Weightman, 2000). It is acknowledged that the absence of bacterial DNA in chronic maxillary sinusitis samples may simply be due to the fact that the number of bacteria in the sample was below the detection limit of the PCR method used.
Three maxillary sinus samples were positive by both PCR and culture methods. One sample was positive only by PCR, and one was positive only by culture. PCR has previously been found to be more sensitive than standard culture methods when mucosal specimens from chronic sinusitis were analysed for the detection of aerobic bacteria (Keech et al., 2000). However, in three cases where Staphylococcus aureus was detected by culture, it was PCR-negative or faintly positive, and therefore was not suitable for further analysis using universal PCR primers. Whilst every effort was made to collect sterile specimens, some of these culture-positive results could be the result of contamination of the sample during collection or processing. This may be particularly true in the case of patient 11, who received moxifloxacin, a fluoroquinolone that is effective against Staphylococcus aureus, 2 weeks before sample collection. This sample was PCR-negative but culture-positive for Staphylococcus aureus. Alternatively, the Staphylococcus-specific primers used here may not bind to the strains found in this study.
The species identified in the present study are not commonly associated with acute bacterial rhinosinusitis by culture methods (Gwaltney, 1996). The bacteria detected in maxillary sinuses in this study included several typically oral species, suggesting that the oral flora may be a source for bacterial colonization of the chronically inflamed maxillary sinus. Of these species, viridans streptococci have previously been identified in chronic maxillary sinusitis by culture methods (Biel et al., 1998), and elevated antibody levels to Prevotella intermedia have been observed in patients with chronic maxillary sinusitis (Brook & Yocum, 1999). However, the presence of specific antibodies against an organism does not necessarily define it as a pathogen. Interestingly, Abiotrophia defectiva and Granulicatella elegans, previously classified as nutritionally variant streptococci and members of the normal flora of the oral cavity, throat, urogenital and intestinal tracts, were identified by molecular methods from the same maxillary sinus. These species are rarely cultured from clinical samples because of their fastidious nature, which may have contributed to the fact that they were detected and identified here only by molecular methods. Recent results from a multicentre study corroborate our findings by showing that Granulicatella species are associated with chronic maxillary sinusitis and may contribute to antibiotic treatment failure (Finegold et al., 2002).
Possible contiguous spread of bacterial flora from the oral cavity to the maxillary sinus was investigated by attempting to find the species detected in the sinus also in the oral cavity, using published species-specific primers. There were no primers available for some species, or the primers that were available did not work in our laboratory as described in the original publications, so our efforts were limited to about half of the species detected in the sinus. Two of five bacterial species found in the maxillary sinus were also found in the oral cavity by PCR using species-specific primers. The same Streptococcus mitis and/or Streptococcus oralis clone(s) were found in the oral cavity and the maxillary sinus in one patient. In another patient, Pseudomonas aeruginosa, identified in the maxillary sinus, was also found in the oral cavity. Previous studies suggest that oral colonization by potential respiratory pathogens may occur, and that dental plaque may serve as a reservoir for these organisms (Scannapieco et al., 1992). Unexpectedly, Streptococcus gordonii, a common colonizer of the oral cavity, was not found in the mouth of the patient who carried it in the maxillary sinus. Also, Staphylococcus aureus and Serratia marcescens were not detected in the oral samples. However, this does not rule out the possibility that these species may have been present in very low numbers in our samples and thus below the detection limit for the method.
As PCR does not distinguish viable from dead bacteria, we cannot say that the present cases of chronic maxillary sinusitis were definitely caused by the bacterial species isolated. It is possible that there were only dead bacteria or released DNA left in the sinuses from a past infection by these organisms. Bacterial DNA has been demonstrated to persist in human synovial fluid and tissue samples as long as 10–22 days after the initiation of antimicrobial treatment, whereas all samples became culture-negative by 2–3 days following the start of therapy (van der Heijden et al., 1999). In contrast, studies on chinchillas have shown that bacterial DNA can be found only for 1 day in the presence of middle ear effusion (Aul et al., 1998). The elimination of bacterial DNA may therefore depend on the body site and fluid. The role of bacteria in chronic sinusitis and related disorders has been questioned, and fungal pathogens have been proposed to be responsible for the inflammatory reaction and mucosal response in the sinus. However, in the present study, no fungi were detected in chronic maxillary sinusitis by culture and we did not attempt to detect fungi by molecular methods.
In conclusion, the present results suggest that both aerobic and anaerobic bacteria can be detected in nearly half of chronic maxillary sinusitis cases. The species identified in the present study are rarely associated with acute bacterial rhinosinusitis and include several oral species, suggesting that these are present in the inflamed maxillary sinus. The simultaneous detection of some bacterial species in the maxillary sinus and in the oral cavity shows that the direct connection between these two sites may allow oral bacteria to contribute to non-oral inflammatory conditions.
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Acknowledgments |
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