SHORT COMMUNICATION
The Microbial Generation of Nitric Oxide in the
Human Oral Cavity
A. J. Smith1, N. Benjamin2, D. A. Weetman1, D. Mackenzie1 and T. W. MacFarlane1
From the 1Infection Research Group, Level 9, Glasgow Dental School, Glasgow and the 2Department of
Clinical Pharmacology, St. Bartholomew’s Hospital Medical College, University of London, West Smithfield,
London, UK
Correspondence to: Dr A. J. Smith, Infection Research Group, Level 9, Glasgow Dental School, 378 Sauchiehall
Street, Glasgow, UK. Fax: +44 141 353 1593; E-mail: [email protected]
Microbial Ecology in Health and Disease 1999; 11: 23–27
Nitric oxide a potent antimicrobial agent and vasodilator, may be synthesized in the human mouth due to the bacterial reduction of
salivary nitrate. The purpose of this study was to determine whether intra-oral pH changes produced by the oral flora may augment the
production of nitric oxide. To test this hypothesis a sucrose mouthrinse was administered to human volunteers who were free from caries
and periodontal disease and production of oral nitric oxide was related to intra-oral pH changes. Baseline measurements consisted of
pooled supragingival plaque and anterior and posterior tongue scrapings for microbiological analysis. Additional samples of saliva were
collected for nitrate and nitrite analysis. Intra-oral pH measurements were recorded with a microelectrode at selected sites and NO
intra-oral levels were measured by chemiluminescence. In addition selected members of the oral flora were tested for their ability to reduce
nitrate to nitrite. A decrease in pH produced by the sucrose rinse was significantly associated with an increase in the intra-oral generation
of NO after 5 min (p =0.006). The most frequently recovered nitrate reducing organisms were Veillonella spp. and Actinomyces spp.
whilst the acidogenic Streptococcus spp. were negative nitrate reducers. The oral generation of NO appears to be influenced by pH levels
within the oral cavity. Key words: nitric oxide, nitrate, nitrite, saliva, nitrate reductase, intra-oral pH.
INTRODUCTION
Nitric oxide (NO) a labile and highly reactive gas can be
generated endogenously by mammalian cells such as activated macrophages by nitric oxide synthase and L-arginine
(1). The alternative source of nitric oxide is via exogenous
sources usually by the action of microbial nitrate reductase
enzymes on nitrate compounds to produce nitrites. Under
acidic conditions nitrites dissociate to give gaseous nitrogen oxides including NO which can readily diffuse through
cell membranes to exert its effects (2).
The precise role of generated NO is unknown but it has
been suggested that this agent may act in the regulation of
microbial communities by its antimicrobial activities. A
number of microorganisms are amongst a growing list of
sensitive organisms (3, 4). The mechanism(s) by which the
gaseous free-radical NO, exerts its antimicrobial activity
has not been identified clearly but interference by highly
reactive NO intermediates on iron-dependent enzymes and
with bacterial DNA appear likely (5).
In man, nitrate derived from the diet is concentrated in
saliva and salivary nitrate levels can be correlated to
dietary intake of nitrate compounds (6). Increased
amounts of dietary nitrate can lead to increased generation
of oral nitric oxide (7) but in man the mechanisms behind
this are unclear although it seems likely that nitrate reduc© Scandinavian University Press 1999. ISSN 0891-060X
tase produced by the oral flora may reduce salivary nitrate
to nitrite which under acidic conditions may dissociate to
form nitric oxide (8). The pH levels necessary for NO
generation in the oral cavity are more likely to occur when
the pH is lowered following carbohydrate ingestion.
The aim of this study was to investigate microbial
nitrate metabolism and NO generation within the oral
cavity of man. A cross-sectional study in healthy volunteers was performed to investigate the relationships between selected members of the oral flora, intra-oral pH
and oral NO levels.
MATERIALS AND METHODS
Subject recruitment
Ethical approval for the study was obtained from Glasgow
Dental School Ethics Committee. Volunteers were medically fit, not receiving any drug therapy, were non-smokers, dentate and free from active caries and periodontal
disease. Subjects (n= 9) refrained from normal oral hygiene procedures the night before and on the day of testing
and abstained from food and drink 2 h prior to testing.
Samples
Sali6a. Pooled unstimulated saliva was collected for 30 s
from each volunteer at baseline and after 5, 10, 15, 30 and
Microbial Ecology in Health and Disease
24
A. J. Smith et al.
60 min. Saliva samples were stored in sterile universals at
4 °C until they could be transported to the laboratory and
stored at − 10 °C. The samples were centrifuged (Microspin 12S, Sorvall, Stevenage, UK) at 17000 g for 15
min at 4 °C and the supernatants assayed for nitrite and
nitrate using high pressure liquid chromatography. Separations were performed on a Wescan Anion/R column
(250×4.1 mm, Alltech, Carnforth, UK) fitted with an
Anion/R guard column using 4 mmol/l 4-hydroxybenzoic
acid, containing 2% (v/v) methanol adjusted to pH 8.5 by
the addition of lithium hydroxide, as eluent. The flow rate
through the column was 1.5 ml/min and detection was by
suppressed conductivity (Model 430, Waters, Watford,
UK).
Intra-oral pH measurements. The measurement of pH
was performed using a micro-electrode (Beetrode®,
MEPH1,World Precision Instruments, Stevenage, UK)
with a micro reference electrode (Microelectrodes Inc,
Bedford, NH, USA) connected to a meter (Model
220,Corning, Halsted, UK) set to read mV. The meter was
calibrated using pH 7 and pH 4 buffers, subsequent readings were converted from mV to pH using a calibration
curve. When the pH measurements took place subjects
were asked to place a finger into a container of 4 M
potassium chloride together with the reference electrode.
The posterior dorsum and anterior portion of the tongue
were also measured for pH. In addition the pH of the
interdental plaque distal to the upper first molars and
central incisors and distal to the lower first molars and
incisors was measured at baseline and after 5, 10, 15, 30
and 60 min.
Microbiological assessment
Supragingival plaque was collected using a sterile dental
excavator and pooled from the mesial surface of one upper
and lower molar tooth and the mesial surface of one upper
and lower incisor tooth. The tongue was sampled using a
sterile round ended periodontal curette. The posterior dorsum of the tongue was sampled approximately 6 cm from
the anterior tip of the tongue by scraping for 2 cm across
the midline. The anterior surface of the tongue was sampled 1 cm from the tip by scraping 2 cm across the
midline. Each sample taken at baseline and at 30 min was
placed immediately into 0.5 ml of pre-reduced Fastidious
Anaerobe broth (FAB) (Lab M, Bury, UK) and processed
within 15 min of being collected. Samples were vortexmixed for 30 s and serial dilutions made in FAB from neat
to 10 − 4. All dilutions were inoculated to Columbia agar
(Pro-lab diagnostics, Cheshire, UK) plates containing 7.5%
v/v sterile defibrinated horse blood and 1% v/v vitamin
K/haemin solution (Life Technologies, Paisley, UK) using
a spiral plater (Don-Whitely, Shipley, UK). MacConkey
Agar, Sabouraud Dextrose Agar (Pro-lab diagnostics,
Cheshire, UK), Rogosa agar and Mitis Salivarius agar
(Difco, Surrey, UK) containing 20 units/ml bacitracin
(Sigma) were inoculated with 50 ml of neat sample and this
inoculum was spread evenly over the surface of the agar
with a sterile L spreader (Bioprobe, Hughes Whitlock Ltd,
Gwent, UK). Plates were incubated as follows: MacConkey and Sabouraud agar aerobically at 37 °C for 48 h,
Mitis Salivarius Bacitracin agar in a CO2 incubator (5%
CO2 and 95% air) for 48 h and Rogosa and Columbia
Blood Agar in an anaerobic incubator in an atmosphere of
nitrogen, hydrogen and carbon dioxide (80:10:10 by volume) for 72 h.
Nitrate reductase acti6ity
The two predominant colony types from each anaerobic
plate were selected for purity plating and identification. In
addition type and laboratory cultures of common oral
species were selected. Both wild and type cultures were
tested for nitrate reductase activity (9). Briefly, nitrate
disks were prepared by the addition of 20 ml of a 3 M
potassium nitrate, 4 mM sodium molybdenate sterile solution to 4 mm diameter filter paper disks (Mast Laboratory,
Liverpool, UK) which were allowed to dry at room temperature. Reduction of nitrate was tested by removing the
nitrate disk from the surface of a blood agar plate on
which there was growth of the test organism and placed in
a sterile petri dish. Twenty microlitres of a solution containing 0.8% v/v sulphanilic acid in 5 N glacial acetic acid
were added to the disk followed by 20 ml of 0.6% N-Ndimethyl-1-napthylamine in 5 N glacial acetic acid. Reduction of nitrate to nitrite is indicated by a pink to red
colour. If no colour was seen within 3 min a small amount
of zinc dust was added and left for 5 min. Development of
a red colour indicates that nitrate was not reduced; if the
disk remained colourless, nitrate was reduced beyond nitrate to gaseous compounds (positive test). Each organism
was tested in duplicate on two separate occasions with
known positive (Escherichia coli NCTC 10418) and negative (Staphylococcus aureus NCTC 7447) controls.
Nitric oxide analysis
A sample of air was introduced into the closed and empty
mouth via a 50 ml plastic syringe and the individual asked
to close their lips around the tip of the syringe. The air was
withdrawn with the syringe after 1 min and analysed for
NO concentration (parts per billion) with a chemiluminescence meter (Thermo-Electron Instruments, UK) at baseline and after 5, 10, 15, 30 and 60 min.
Effect of sucrose mouthrinse
Following collection of baseline data each subject was
asked to rinse for 30 s with a standard sterile sucrose (10%
w/v) containing solution. The samples for the analysis of
salivary nitrate and nitrite ions, intra-oral pH and nitric
oxide generation were then collected after 5, 10, 15, 30 and
60 min. Microbiological parameters were measured only
after 30 min.
The microbial generation of nitric oxide
25
Fig. 1. Intra oral pH changes and nitric oxide levels following a sucrose
rinse. =Nitric oxide levels (parts per
billion); = pH changes; --- = pH
changes at gingival margin; — =pH
changes at anterior dorsum of tongue
and – – =pH changes at posterior
dorsum of tongue. Error bars represent
SD.
RESULTS AND DISCUSSION
Following the sucrose rinse the interdental supragingival
plaque pH fell from a baseline value of 6.8 to a minimum
of 5.8 after 5 min and had returned to baseline by 30 min
(Fig. 1). The plaque pH response to the sucrose rinse
followed that of the classical Stephan curve (10). Similar
pH changes were mirrored by plaque from the posterior
but not the anterior dorsum of the tongue.
There was a large intersubject variation in the nitrite
and nitrate concentrations in saliva which probably reflects
the variation in dietary nitrate intake (Table I). There was
a significant decrease in salivary nitrate from baseline at 5
min (p=0.039, paired Wilcoxan signed rank test). However, there was no significant relationship between the rate
of nitric oxide production or peak nitric oxide levels and
salivary nitrate/nitrite concentrations.
Analysis of the production of NO during the study
period (Fig. 1) demonstrated a significant increase over
baseline (paired Wilcoxan signed rank test) at 5 min
(p =0.006), 10 min (p=0.06) and 15 min (p =0.048).
Analysis of the microflora in dental plaque and tongue
samples from the nine healthy volunteers demonstrated no
significant difference in microbiological parameters over
the time period of the study. No significant numbers of
black pigmenting anaerobes, yeasts, coliforms, lactobacilli
or mutans streptococci were isolated from any of the
subjects. The predominant bacteria isolated from the anterior and posterior dorsum of the tongue were Streptococcus species, none of which reduced nitrate. Analysis of type
strain cultures and selected isolates from the volunteers for
nitrate reductase activity (Tables IIa and IIb) demonstrated a relatively small range of genera capable of reducing nitrate. The most common nitrate reducing organism
isolated in the volunteers from dental plaque and tongue
dorsum were Actinomyces spp. and Veillonella spp.
This study has observed a significant increase in intraoral levels of NO following a sucrose rinse in the oral
cavity of volunteers clinically free from active caries and
periodontal disease. The generation of nitric oxide from
within the oral cavity was affected significantly by changes
in the intra-oral pH produced by a sucrose rinse.
Although there were differences in salivary nitrate and
nitrite levels between individuals this did not seem to
influence the rate or amount of NO production. Dietary
nitrate appears to be excreted via saliva into the oral cavity
where it can be converted into nitrite. It has been estimated that 25% of ingested nitrate is recirculated in the
saliva (11). While parotid duct saliva is free of nitrite
whole saliva is positive (12) and the level can increase
some 30-fold following ingestion of food or drink rich in
nitrate containing compounds, mostly derived from leafy
vegetables (6, 7). Large interindividual variations in nitrate
and nitrite concentrations similar to the present results
have been observed previously (6). The acid catalyzed
reactions of nitrite are largely dependent on pH and the
presence or absence of inhibitors — such as urea, ammonia
and vitamin C — or catalysts — such as thiocyanate, chloride and some phenolics (13) — may account for the intersubject variation in NO production.
Of particular interest was the pH drop on the dorsum of
the tongue. The tongue dorsum has been suggested as the
Table I
Sali6ary nitrate/nitrite le6els following a sucrose rinse
Time
Saliva nitrate mmol/l
mean (SD)
Saliva nitrite mmol/l
mean (SD)
Baseline
5 min
10 min
15 min
30 min
60 min
531.7
496.4
505.8
510.2
508.7
524.4
263.5
260.4
262.5
269.5
253.6
260.0
(114.7)
(77.8)*
(82.4)
(98.9)
(136.7)
(105.6)
(34.7)
(39.4)
(30.0)
(31.2)
(42.8)
(31.5)
* p = 0.039 paired Wilcoxan signed rank test
26
A. J. Smith et al.
Table IIa
Reduction of nitrate by some common oral commensals
Isolate
Nitrate
reduction
Type strains
Haemophilus paraphrophilus NCTC 29242
H. aphrophilus NCTC 5886
Streptococcus oralis NCTC 11427
S. sanguis NCTC 7863
S. intermedius NCTC 11324
S. constellatus NCTC 11325
S. mutans NCTC 10449
S. mitis NCTC 10712
Bacteroides forsythus ATCC 43037
Escherichia coli NCTC 10418
Peptostreptococcus micros NCTC 9821
Peptostreptococcus magnus NCTC 9815
Peptostreptococcus asacharolyticus NCTC 9820
Pre6otella nigrescens ATCC 33563
Pre6otella intermedia ATCC 25611
Porphyromonas gingi6alis NCTC 11834
Actinobacillus actinomycetemcomitans NCTC 9710
Fusobacteria nucleatum NCTC 10596
Candida albicans NCPF 3153
+
+
−
−
−
−
−
−
+
+
−
−
−
−
−
−
+
+
−
Table IIb
Clinical isolates
S. mitis
S. mutans
A. naeslundii
A. odontolyticus
F. nucleatum
Veillonella spp.
Lactobacillus spp.
Peptostreptococcus spp.
Capnocytophaga spp.
P. acnes
B. corrodens
H. aphrophilus
Number of
strains tested
Nitrate
reduction
7
1
2
2
2
4
2
1
2
2
1
3
−
−
++
++
+
++
−
−
++
−
++
+
++ =strong positive reaction; + =positive reaction; −= no
reaction.
major site of NO production on the basis of animal work
(8) and its relatively large surface area which is due to its
highly fissured surface particularly in the posterior dorsum
would favour the growth of nitrate reducing anaerobes, for
example Veillonella spp. Although the tongue flora has
been relatively poorly studied to date, the data collected in
this small study of healthy individuals lend support to the
observation of others (14–16) that the posterior dorsum
has higher anaerobic counts than the anterior portion of
the tongue. The major bacterial species collected from the
tongue of healthy volunteers that were capable of reducing
nitrate were Veillonella spp. and Actinomyces spp. organisms with poor acidogenic potential. The tongue flora also
contains high numbers of oral Streptococci species capable
of producing the observed change in pH.
Concentrations of NO lower than 1 ppm have been
shown to inhibit the growth of several bacteria and fungi,
such as E. coli and C. albicans (8, 17 – 19). This raises the
interesting possibility that the anaerobic flora in the posterior dorsum of the tongue and within the depths of the
periodontal pocket generate sufficient NO to inhibit colonization by other microbial species of this particular niche.
In addition to its antimicrobial properties, it has been
suggested (20) that NO generation may influence the inflammatory response (vasodilation) in the oral cavity particularly at the gingival margins in gingivitis and periodontitis.
In conclusion, intra-oral production of NO may depend
on many factors, such as nitrate intake and salivary flow
rates, specific nitrate reductase activities of the microbial
flora, their carriage rate and spatial distribution in relation
to the salivary nitrate flow. However, we have demonstrated that a significant increase in NO — a potent antimicrobial agent and vasodilator — is associated with a drop
in intra-oral pH. The full range of the physiological interactions of NO in the maintenance of the oral flora and
normal mucosa remain to be elucidated.
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