A Postbiotic Positively Shifts the Canine Oral Microbiome

Abstract

The oral microbiome is an important aspect of overall oral health in dogs. To investigate the ability of a postbiotic, canine oral health postbiotic (COHP), to support oral health, a double-blind, placebo-controlled study was conducted with a dirty tooth model to assess its ability to reduce volatile sulfur compound (VSC) producing microbes that cause halitosis and modulate canine oral microbiome composition. Ten client-owned dogs were randomly split into 2 groups. The groups received either the COHP or a placebo as a powder topper on food for 7 days. Oral microbiome samples were collected on day 0 and day 7 along with the buccal gingival margin. Hydrogen sulfide (H₂S)-producing colonies were quantified by plating the oral microbiome samples and counting pigment-producing colonies. Additionally, oral microbiome samples were sequenced, and taxa abundance was quantified. A trend toward a reduction in H₂S-producing colonies was observed in the COHP group (P = .06), but not in the placebo group (P = .9). Canine oral health postbiotic reduced the abundance of 9 times as many taxa as the placebo, including taxa that form biofilms and produce VSCs. The placebo reduced the abundance of only one taxon, which is not associated with biofilms or VSCs. The findings provide evidence for COHP's ability to promote a positive shift in the canine oral microbiome, and, together with previous results, provides evidence that it may broadly help to maintain canine oral health.

Keywords

canine halitosis postbiotic microbiome

Introduction

The oral microbiome plays a critical role in overall canine dental health. The oral microbiome is the community of microorganisms and their collective genetic material that inhabit the mouth. Within the canine oral cavity, dental plaque is composed of complex, polymicrobial biofilms,^1–4 and its formation ultimately leads to further bacterial proliferation, which shifts the oral microbiome.^5–10 In the first step of dental plaque formation, biofilm-forming facultative anaerobes adhere to the surface of the teeth.^1,2,10 The inflammatory and oxygen-scavenging activity of these early colonizers creates a microenvironment more favorable to obligate anaerobes such as Fusobacterium and Porphyromonas,^1,6 allowing them to colonize the subgingival crevice.^11,12 Early plaque colonizers can indeed elicit a localized gingival inflammatory response via host immune activation, which may be accompanied by increased inflammatory exudation into the gingival sulcus.^1,6,12 Subgingival plaque is characterized by a higher abundance of black-pigmented gram-negative obligate anaerobes, such as Porphyromonas, Prevotella, and Treponema.^6–9

Plaque and calculus are closely linked to halitosis, in both humans and dogs,^13–15 and in dogs, many of the genera associated with canine periodontal disease also produce volatile sulfur compounds (VSCs). Volatile sulfur compounds are small, sulfur-containing gases produced when oral bacteria break down proteins and other substrates, and they include molecules such as hydrogen sulfide (H₂S), methyl mercaptan, and dimethyl sulfide. They are offensive to humans because these compounds have very low odor detection thresholds.¹⁶ Porphyromonas gulae and Porphyromonas gingivalis, 2 of the main periodontitis-associated pathogens in dogs,^1,11,17,18 have been found to produce high levels of VSCs in studies of canine-associated isolates.^17,19,20 Measuring changes in VSC levels in the oral cavity is a common way to assess the effectiveness of canine oral care interventions in vivo.^17,20,21 Volatile sulfur compounds, particularly H₂S, may exacerbate periodontal disease progression and severity^16,22–24; however, much of the mechanistic evidence supporting this concept comes from human oral cell studies and noncanine experimental models. Hydrogen sulfide has been shown to increase cell death in human gingival fibroblasts²⁵ and epithelial cells,²⁶ stimulate inflammatory responses through pro-inflammatory cytokine release in human-based systems,^27–29 and enhance oral mucosal permeability in experimental tissue models.³⁰ These mechanisms collectively contribute to halitosis and its impact on oral health.³¹ In dogs, halitosis is not just a clinical sign but it can negatively affect the emotional bond between pets and their owners.³² Beyond species-level associations, periodontitis research has also proposed that certain organisms such as P. gingivalis can play a disproportionate role in shaping dysbiotic communities.³³

As halitosis is driven by microbes, there is a need for convenient solutions that effectively modulate the canine oral microbiome.³⁴ Chemical ingredients such as sodium hexametaphosphate target oral microbes indiscriminately, potentially disrupting beneficial bacteria.³⁵ The antimicrobial peptide nisin has also been shown to be potentially disruptive to oral commensals,³ and gallic acid failed to show a robust effect on canine oral taxa abundance.³⁶ While dental chews have recently been shown to shift the oral microbiome,^37,38 they work through mechanical action, which is not applicable to all pet products.

Microbial-based ingredients, such as probiotics and postbiotics, offer a promising alternative for supporting oral health by modulating the oral microbiome.³⁹ Probiotics are live microorganisms that, when administered in adequate amounts, confer a health benefit to the host, whereas postbiotics are preparations of inanimate microorganisms and or their structural components and metabolites that also confer a health benefit to the host.⁴⁰ While one study found that probiotics failed to shift the canine oral microbiome,⁴¹ another showed that a Lactobacillus acidophilus probiotic decreased pathogenic taxa in dogs.⁴² However, probiotics may not be successfully integrated into pet products that require high heat and pressure in their manufacturing. Postbiotics are defined as “a preparation of inanimate microorganisms and/or their components that confer a health benefit to the host.”⁴³ This class of ingredients contains microbial metabolites that may support a healthy oral microbiome effectively while offering stability across various product formulations. In humans, postbiotics have been shown to positively shift the oral microbiome⁴⁴ and decrease microbes associated with VSCs.⁴⁵ Nevertheless, clinical evidence demonstrating that postbiotics will support a healthy canine microbiome is limited.^46,47

Previously, a novel canine oral health postbiotic (COHP) was found to effectively reduce canine halitosis, as assessed by VSC levels in the oral cavity.⁴⁶ The postbiotic group showed a significant 22% decrease in VSC levels after 7 days, and prevented an increase of VSC levels at Day 14, whereas there was a 35% increase in VSC in the placebo group.⁴⁶ To further investigate COHP's ability to support oral health, a double-blind, placebo-controlled study was conducted to assess the ability of COHP to modulate the canine oral microbiome composition and reduce the abundance of H₂S-producing bacteria.

Materials and Methods

Animals

The protocol was reviewed and approved by the Institutional Animal Care and Use Committee. All dogs were classified as USDA Category C for the full duration of the study. This classification includes animal use activities that involve no more than momentary or slight pain or distress for which there is no need for the use of pain-relieving drugs.

The general inclusion criteria in the study were as follows: (1) dogs were not taking any prescribed medications, (2) dogs were a healthy weight for their breed (a dog's weight is outside the healthy range if a veterinarian has previously stated that the dog was overweight), (3) dogs were not pregnant and had not been pregnant in the last 6 months, (4) the dogs allowed a veterinary technician to perform the required sampling, (5) the pet owner was willing to withhold water from their dog for the duration of one meal per day (∼15 min) on each day of the study, and (6) the dog was between 1 and 7 years old. The oral hygiene inclusion criteria in the study were as follows: (1) participants did not receive any oral care by a veterinary dentist in the 3 months preceding the study, and (2) all participants had not had their teeth brushed and did not consume or use any oral health product, including dental chews, treats, and food supplements in the week preceding the study.

All dogs enrolled in the study were privately owned animals of various breeds between 1 and 5 years old (Table 1). The dogs were housed at home for the duration of the study. Prior to enrollment, owners of the eligible dogs consented to the use of their dog's deidentified information for publication purposes, as well as all study guidelines and parameters. Dogs did not receive a physical examination or laboratory evaluation for this study.

Table 1.

Participant Signalment and Group Allocation.

Participant ID	Spayed/Neutered	Age (years)	Sex	Weight (kg)	Breed	Group
COHP111C01	Yes	5	Male	20.4	Samoyed	Placebo
COHP111C02	Yes	4	Male	8.6	Shihtzu Poodle Mix	COHP
COHP111C03	No	1	Male	3.2	Shihtzu Poodle Mix	COHP
COHP111C04	No	2	Male	5	Toy Poodle	Placebo
COHP111C06	Yes	5	Male	11.4	Cocker Spaniel Chihuahua Mix	COHP
COHP111C07	Yes	2	Male	12.3	Chihuahua German Shepherd Mix	Placebo
COHP111C09	Yes	3	Female	5.9	Mini Golden Doodle	COHP
COHP111C14	No	2	Female	18.2	Australian Shepherd	Placebo
COHP111C16	Yes	1	Female	2.7	Chihuahua Mix	COHP
COHP111C17	No	2.5	Female	10	American Eskimo	Placebo

Study Design

The study was double-blind, placebo-controlled, and randomized, and 7 days in duration. The intervention was administered for 7 days (days 1-7). Dogs were randomly split into 2 groups of 5 with balanced numbers of male and female in each (Table 1).

The study used a dirty tooth model (no dental cleaning occurred as part of the study). No dental chews, treats, or supplements were provided throughout the study outside of the test materials. In addition, ad libitum access to routine dental enrichment, such as chew toys and bones that may disrupt plaque via mechanical action, was interrupted for the duration of the study.

There were 3 termination criteria for the study: (1) Abnormal changes in the dog’s health, such as unusually loose stools, are observed, and the owner decides to terminate participation in the study based on the recommendation from their veterinarian. (2) Health issues surpassing minor adverse effects as judged by the owner are observed, and emergency veterinary care is sought out by the pet parent. The health issue need not be related to the study. (3) An animal refuses to eat more than 2 consecutive meals containing the ingredient, or an owner is observing a significant impact on the eating behavior of their dog.

Intervention

Canine oral health postbiotic^a is a commercially available ingredient composed of a tapioca maltodextrin carrier, dried Pediococcus pentosaceus fermentation product, and dried Bacillus subtilis fermentation product. Both fermentation products are heat-treated to inactivate live cells, then spray or freeze-dried. The placebo was tapioca maltodextrin^b, the same as the carrier utilized in the test ingredient.

Participants were fed at home for the duration of the study. Dog owners received blinded, prepackaged powder toppers (either COHP or placebo) for each meal and clear instructions on how to incorporate the powder topper into the dog's meal.

On days 1 to 7, each dog received 250 mg of either COHP or the placebo added to their only meal of the day as a powder topper. Half of the daily food portion was placed into a bowl. If the dog was receiving dry kibble, it was moistened with a small amount of water to promote adhesion of the powder topper to the food. The dog's water source was removed; the dog was served the food containing the powder topper and was allowed to eat. Fifteen minutes after the dog ate the food containing the powder topper, the dog was served the second half of their daily food portion without the powder topper. The dog's water source was replaced upon completion of the meal.

Oral Microbiome Sampling

Oral sampling was performed on day 0 (the day prior to the first intervention) and on day 7 (after the last intervention). Each dog was swabbed once in each of the 4 quadrants of the mouth along the gingival margin: maxillary left, maxillary right, mandibular left, mandibular right. A trained veterinary technician performed the oral microbiome sampling, and dog owners assisted in controlling the dog during sampling. Wearing clean gloves, the technician swabbed along with the buccal gingival margin in the appropriate location for 5 to 10 s, turning the swab to expose all parts of the swab to the gumline. Once swabbing was completed, the swab was immediately placed in the collection tube containing 1 mL of modified Liquid Amies medium^c. A unique swab was used for each quadrant. Oral sampling in each quadrant was repeated twice on each sampling day: once for H₂S colony counting and once for oral microbiome sequencing. For oral microbiome sequencing, the collection tubes were stored at room temperature for no more than 8 h prior to processing. For H₂S colony counting, the collection tubes were processed within 2 h of collection.

Hydrogen Sulfide Colony Counting

The oral microbiome samples were placed in a Coy anaerobic chamber^d (5% H₂, 10% CO₂, 85% N₂) within 30 min of being collected. All methods involved the use of an anaerobic chamber. The 4 samples from each dog were pooled. Each pooled sample was serially diluted 10-fold in previously deoxygenated 1× PBS 4 times to a final dilution of 1E4; 50 µL from the 1E2, 1E3, and 1E4 dilutions was spread on Oral Hydrogen Sulfide Organism Carbohydrate Agar^e petri dishes to obtain final dilutions of 2E3, 2E4, and 2E5. Petri dishes were incubated in anaerobic container systems^f for 96 h at 37°C. Petri dishes were imaged using a color imaging camera^g. Colonies were counted manually and rated as H₂S-positive if they were black or dark brown or had a black or dark brown ring around them. All other colonies were rated as H₂S-negative. Plates with very dense colonies were divided into quadrants. Colonies from one quadrant were graded, and the result was multiplied by 4 to obtain representative counts.

The petri dishes with the most total colonies were used for statistical analysis (17/18 plates included were from the same dilution) to make fair comparisons across samples. Hydrogen sulfide-producing colonies in each plate were normalized to the total colony number. The relative change in colonies between day 0 and day 7 was calculated. Nonparametric tests (Wilcoxon rank tests) were used to assess within and between-group differences, due to small sample size (n = 4-5).

Oral Microbiome Sequencing

Oral microbiome samples for all dogs at both time points were processed in parallel in a single multi-well plate; samples were not processed in batches. Samples were processed in duplicate from the raw oral microbiome samples. The V4 hypervariable region of the 16S rRNA gene was amplified with barcoded sequencing primers^h and equal volumes of the resulting libraries were pooled and sequenced on the Illumina iSeqⁱ. Counts of the zero-radius operational taxonomic units (ZOTUs)⁴⁸ were determined for each library using USEARCH,⁴⁹ and taxonomic assignments for the ZOTUs were made using the RDP classifier.⁵⁰ For fold change calculations, taxa with less than 20 reads in at least 20% of samples were removed, and read counts were standardized to the median sequencing depth. Read counts were analyzed in R using phyloseq⁵¹ and DESeq2⁵² packages.

The log fold change in number of reads was calculated for each taxon for each group of dogs by comparing the day 0 and day 7 data for each dog. Taxa are considered significantly changed if the log fold change from day 0 to day 7 has a False Discovery Rate (FDR)-adjusted P < .01.

Results

One dog in the placebo group was removed from the study during the intervention period due to voluntary withdrawal. No adverse events were observed in participants receiving COHP.

Canine oral health postbiotic administration led to a trend toward a reduction in H₂S-positive colonies at day 7 (mean relative change of −45 ± 32%, P = .06, Wilcoxon signed rank test) (Figure 1). No effect on the H₂S-positive colonies was observed in the placebo group (+82 ± 220%, P = .9, Wilcoxon signed rank test). The overall abundance of microorganisms (eg, total plate count) did not change across groups and timepoints (placebo P = .38, COHP P > .99, Wilcoxon ranked test) (Figure 2).

Figure 1.

Relative changes in H₂S-producing colonies. On day 7, the mean relative change in H₂S-producing colonies was +82% in the placebo group and −45% in the COHP group. Bars indicate the mean and error bars indicate the standard deviation. H₂S-producing colonies were normalized to the total colonies on the plate. COHP, canine oral health postbiotic; H₂S, hydrogen sulfide.

Figure 2.

Total plate counts remained consistent across groups and timepoints. The bars indicate the mean and the error bars represent the standard deviation. The day 0 values did not differ between groups (P = .45; Mann-Whitney test) and did not change over the course of the study in either group (Placebo P = .38; COHP P > .99; Wilcoxon ranked test). COHP, canine oral health postbiotic.

Within the microbiome composition, COHP reduced the abundance of more taxa compared to the placebo. There was no significant change in the abundance of most taxa, and over 130 operational taxonomic units (OTUs) remained unchanged in both groups. The number of OTUs that increased in abundance was similar for both groups, with 17 OTUs increasing for the control group and 18 increasing for the intervention group (alpha = .01, FDR correction, Wald test) (Figure 3, Tables 2 and 3). Only one OTU decreased for the placebo group, while 9 OTUs exhibited reduced abundance in the COHP group, demonstrating that the administration of COHP had a greater effect on the microbiome composition compared to placebo.

Figure 3.

Taxa with significant log fold changes. Taxa with significant log fold change increase and decrease between day 0 and day 7 for (A) placebo and (B) postbiotic intervention groups (FDR-corrected P < .01). COHP, canine oral health postbiotic; FDR, False Discovery Rate.

Table 2.

Operational Taxonomic Units With Significant Log2 Fold Changes in the Placebo Group.

Phylum	Genus	Log2 fold change	FDR adjusted P value	Pathogenic?
Pseudomonadota	Lysobacter	−1.03	.0078	No
Bacillota	Serpentinicella	1.19	.0063	No
Bacteroidota	Mediterranea	1.19	.00042	No
Pseudomonadota	Enterobacillus	1.26	.0038	Yes¹
Bacillota	Velocimicrobium	1.38	.0078	No
Bacillota	Tepidibacter	1.52	.0038	No
Spirochaetota	Treponema	1.55	.0038	Yes^1,2
Bacillota	Aminipila	1.61	.0075	No
Cyanobacteriota	Bangiophyceae	1.98	.0073	No
Bacteroidota	Paramuribaculum	2.14	.00042	No
Bacillota	Lentihominibacter	2.14	.00013	No
Bacillota	Aminipila	2.21	.00042	No
Bacillota	Catonella	2.30	.00031	Yes³
Bacillota	Pseudoclostridium	2.34	.00065	No
Bacillota	Catonella	2.42	.00013	Yes³
Bacillota	Anaerovibrio	2.52	.00042	No
Bacillota	Tepidibacter	2.66	.00042	No
Pseudomonadota	Desulfovulcanus	4.21	.0075	No

Table 3.

Operational Taxonomic Units With Significant Log2 Fold Changes in the Canine Oral Health Postbiotic (COHP) Group.

Phylum	Genus	Log2 fold change	FDR adjusted P value	Pathogenic?
Campylobacterota	Arcobacter	−2.11	.0011	Yes⁴
Bacteroidota	Ulvibacter	−1.97	.0011	No
Cyanobacteriota	Bangiophyceae	−1.71	.0012	No
Bacteroidota	Bergeyella	−1.50	.0002	Yes⁵
Actinomycetota	Corynebacterium	−1.43	.0046	Yes^6,7
Bacillota	Abiotrophia	−1.21	.0052	Yes⁸
Pseudomonadota	Neisseria	−1.15	.0046	Yes^6,9
Bacillota	Trichococcus	−1.12	.0012	Yes¹⁰
Pseudomonadota	Frederiksenia	−0.94	.0002	Yes¹¹
Bacteroidota	Mediterranea	0.89	.0099	No
Bacillota	Serpentinicella	1.02	.0011	No
Clostridia	Tepidibacter	1.20	.0011	No
Bacteroidota	Porphyromonas	1.22	.0011	Yes¹²
Clostridia	Pseudoclostridium	1.23	.0008	No
Bacillota	Tepidibacter	1.32	.0011	No
Bacillota	Lacrimispora	1.35	.0011	No
Bacillota	Floccifex	1.35	.0011	No
Spirochaetota	Brucepastera	1.40	.0040	No
Bacteroidia	Falsiporphyromonas	1.46	.0016	No
Bacillota	Peptoanaerobacter	1.49	.0040	No
Pseudomonadota	Desulfovibrio	1.60	.00037	Yes¹³
Bacteroidota	Porphyromonas	1.62	.0011	Yes¹²
Bacillota	Fusibacter	1.70	.000078	No
Clostridia	Aminipila	1.77	.0011	No
Bacillota	Aminipila	1.88	.0009	No
Pseudomonadota	Desulfovibrio	2.30	.0046	Yes¹³
Bacillota	Serpentinicella	2.55	.00019	No

To assess the nature of microbiome changes, the log fold change in number of reads was calculated for each taxon for each group of dogs, and taxa are considered significantly changed if the log fold change from day 0 to day 7 has a FDR-adjusted P < .01. Overall, COHP showed a decrease in pathogen-associated OTUs, while the placebo did not. In the COHP and placebo groups, a similar number of taxa increased in abundance throughout the study (18 and 17, respectively). Pathogen-associated OTUs in the control group represent 23% (4/17) of the total number of OTUs that increased in abundance from day 0 to day 7 (Table 2). Similarly, 22% (4/18) of the OTUs that increased in the COHP group are associated with pathogenicity (Table 3). In contrast, COHP reduced the abundance of 9 times as many taxa as the placebo at day 7 (9 and 1, respectively). The single OTU that decreased in the control group, Lysobacter, is not associated with pathogenicity, while 78% of the 9 OTUs (7/9) that decreased in abundance in the intervention group are pathogen-associated.^53–60 These results indicate that COHP may support a healthy oral microbiome by decreasing the abundance of bacteria associated with pathogenicity while maintaining microbes that are normally found in a healthy oral microbiome.³⁴ In comparison, the placebo only decreased the abundance of Lysobacter, which is not associated with oral biofilms.

Canine oral health postbiotic was found to decrease the abundance of microbes associated with biofilm formation (Table 4), which is an important process to control in the context of oral health to limit the formation of plaque and tartar.^3,6,66 Overall, 6 out of 9 genera that were reduced in abundance in the COHP group are associated with oral biofilms: Bergeyella, Corynebacterium, Neisseria, Arcobacter, Frederiksenia, and Abiotrophia. Bergeyella, Corynebacterium, and Neisseria are all facultative anaerobes that are known to form biofilms in vivo and are associated with early colonization of the tooth surface in canines.^1,6

Table 4.

Taxa Reduced in the Canine Oral Health Postbiotic (COHP) Group That Are Associated With Oral Biofilm Formation or Volatile Sulfur Compound (VSC) Production.

Phylum	Genus	Log2 fold change	FDR adjusted P value	Oral biofilm formation	Oral volatile sulfur compound production
Campylobacterota	Arcobacter	−2.11	.0011	Yes^37,38	No
Bacteroidota	Bergeyella	−1.50	.0002	Yes^1,6,55,61	No
Actinomycetota	Corynebacterium	−1.43	.0046	Yes^1,6	No
Bacillota	Abiotrophia	−1.21	.0052	Yes^4,57	Yes⁵⁷
Pseudomonadota	Neisseria	−1.15	.0046	Yes^37,55,62,63	Yes⁶⁴
Pseudomonadota	Frederiksenia	−0.94	.0002	Yes⁶⁵	No

Discussion

The majority of the microbes of the oral microbiome were stable over the course of the study in both groups, and overall microbial abundance, as assessed by plate count, did not change. This stability is consistent with the short duration of the study, as well as published longitudinal studies showing that the oral microbiome for an individual is largely unchanged over time.^67,68 Importantly, this provides evidence that the postbiotic does not work by eliciting broad antimicrobial activity as antimicrobial substances would be expected to substantially impact microbiome composition and/or microbial abundance.^10,34

The predominant genera found in the oral cavities of healthy adult dogs include Porphyromonas, Prevotella, Treponema, Fusobacterium, Enhydrobacter, Moraxella, Corynebacterium, and Fusibacter,^5,69 which all contain some pathogen-associated species. Thus, it is not unexpected that some of the taxa that have increased abundance in both the control and intervention groups may be pathogen-associated.^{19,53–60,70–73} However, COHP was distinct from the control group in its ability to decrease the abundance of taxa associated with oral biofilms. Bergeyella and Neisseria are associated with supragingival plaque in dogs and in humans,^62,63 and Corynebacterium and Neisseria have also been found to be highly abundant in subgingival plaque of dogs.^37,55 Additionally, Corynebacterium and Neisseria are associated with human gingivitis and periodontitis.^55,61 Arcobacter is another taxon found to be highly abundant in canine supragingival dental plaque, and its relative abundance is positively correlated with periodontal disease, plaque, calculus, gingivitis, and pocket scores in dogs.^37,38 Frederiksenia contains species shown to exhibit synergistic biofilm growth with the canine pathogen P gulae in vitro,⁶⁵ while Abiotrophia is associated with early dental plaque both in vitro and in humans.^4,57

In addition to biofilm formation, the taxa that were reduced in abundance in the COHP group are associated with VSC production (Table 4). Volatile sulfur compounds are the causative driver of bad breath and can have further undesirable oral health effects.^25–31 Neisseria is known to be associated with high H₂S concentration in the human oral cavity,⁶⁴ and Abiotrophia is known to produce H₂S in humans.⁵⁷ In contrast, COHP reduced levels of Bergeyella, which is negatively correlated with VSCs in humans.⁷⁴ The reduced abundance of VSC-associated taxa in the COHP group aligns with the observed trend toward a reduction of H₂S-positive colonies observed at day 7, while no effect on the H₂S-positive colonies was observed in the placebo group. Again, Lysobacter, which was reduced in abundance in the placebo group, is not a known oral VSC-producer.^75,76

The results from the H₂S Colony Counting assay, albeit not statistically significant, are consistent with the results of the oral microbiome sequencing analysis as they show a potential differential impact between COHP and the placebo interventions on the oral microbiome composition. The variability of the H₂S colony counting assay was relatively large, which is typical for this type of assay. Nonetheless, it provides corroborating evidence for the primary findings of the oral microbiome sequencing analysis through an independent analytical methodology. A similar approach was previously used to support VSC-related findings.⁷⁷

The primary objective of this investigation was to determine whether COHP suppresses VSC-associated microorganisms and whether it shifts the composition of the canine oral microbiome. Consistent with that aim, the authors selected a dirty tooth model in a diverse group of client-owned dogs to reflect the conditions under which many dogs are maintained, namely, the absence of routine professional dental prophylaxis. Therefore, baseline plaque, gingivitis, and calculus assessments were not made. Canine oral health postbiotic drove a shift in the canine oral microbiome, decreasing the abundance of 9 times as many taxa as the control group. The taxa that were reduced are generally known for undesirable properties, including association with pathogens, biofilm formation, and VSC production. Overall, this indicates that after 7 days, COHP helped to promote a healthier microbiome, suggesting that COHP may affect microbiome diversity and help support a balanced oral microbiome, a factor which has been linked to long-term oral health in another longer study.^78–81 Additionally, COHP was previously shown to reduce VSCs in the canine oral cavity at both day 7 and day 14, suggesting that COHP's effects on the oral microbiome may extend for at least 14 days.⁴⁶

While data on the performance of oral health postbiotics in canines is limited, a recent study demonstrated that a Lactiplantibacillus plantarum postbiotic only increased the abundance of one taxon compared to the placebo group at Day 57 and did not decrease the abundance of any taxa.⁴⁷ This taxon was one of many anticorrelated with gingivitis, plaque, and calculus in their study, and the change only occurred in the group receiving the lower of 2 doses tested. Canine oral health postbiotic affected ∼20× more taxa in only one week.

This pilot study had methodological limitations that should be considered when interpreting the findings. First, the sample size was small (5 dogs per group), and although the authors did observe statistically significant differences in this cohort, evaluation in a larger cohort would strengthen confidence in the robustness and generalizability of the results. Second, the intervention period was short (7 days). While this duration was well-suited to assess a rapid shift in the oral microbiome following COHP administration, longer studies would be valuable to determine whether observed microbiome changes persist over time and would better characterize temporal dynamics beyond the initial response. Also, while groups were matched according to sex, stratification by age would have been preferred, as age is closely associated with halitosis in dogs. More importantly, stratification based on the initial state of the oral microbiome would have been preferred as a stratification method based on the primary analytical variable. Finally, a longer washout period might have established a more uniform oral microbiome baseline; however, the authors do not expect the selection of a 7-day duration for the washout period to have a major impact on the conclusion of this study, as the oral microbiome of individual dogs can vary greatly, and each dog served as its own reference in the metagenomic analysis.

Conclusion

This clinical study demonstrated COHP's ability to promote a positive shift in the canine oral microbiome. Canine oral health postbiotic reduced the abundance of 9 times as many taxa as the placebo at day 7, 78% of which are pathogen-associated, and a substantial portion of which are also oral biofilm formers and VSC-producers. Aligned with these results, COHP showed a trend in reduction of H₂S-positive colonies on day 7. These results indicate that COHP modulates the oral microbiome, suggesting that COHP may be a good alternative to chemical ingredients that target oral microbes indiscriminately, and thus potentially disrupt beneficial bacteria.³⁵ Overall, the findings suggest that this novel postbiotic is able to support a healthy oral microbiome.

Materials

Superculture Pet Oral^®, Kingdom, Brooklyn, NY, USA

Tapioca maltodextrin, Mike's Mix, Mazomanie, WI, USA

ESwab Collection Kit (95037-744), Becton, Dickinson and Company, Franklin Lakes, NJ, USA

Anaerobic chamber, Coy Laboratory Products, Inc., Grass Lake, MI, USA

Oral H2S Organisms Agar with Lead Acetate (OHO-C; AS-6430), Anaerobe Systems, Morgan Hill, CA, USA

GasPak EZ Anaerobe Container System (catalog no. 260001), BD Diagnostic Systems, Sparks, MD, USA

Color camera (PL-D7715CU-S-BL-AF7.5), Pixelink Incorporated, Ottawa, ON, Canada

Custom oligonucleotides for 16S rRNA V4 amplification, Integrated DNA Technologies, Coralville, IA, USA

iSeq 100 Sequencing System, Illumina, Inc., San Diego, CA, USA

Footnotes

Acknowledgments

The authors would like to thank all Kingdom employees for their contributions to this project. In particular, the authors thank Antonio Diaz for his contributions to the project administration, partner management, and help with manuscript preparation, and Emily Daley for her review of the manuscript.

ORCID iD

Raphaël Turcotte

Institutional Review Board Statement

The study was conducted at a registered research facility that complied with all local regulations governing the care and use of laboratory animals and was conducted in accordance with OMAFRA, the CCAC Guide to the Care and Use of Experimental Animals. To ensure compliance, the protocol was reviewed and approved by the facility's Institutional Care and Use Committee (IACUC). Ethical Approval Code: IACUC# COHP111C.

Author Contributions

Conceptualization, R.U.S., R.T., and A.S.; methodology, A.S., R.T., and J.H.; formal analysis, A.S. and J.H.; data curation, A.S. and J.H.; writing—original draft preparation, J.H. and A.S.; writing—review and editing, J.H., A.S., R.T., and R.U.S.; visualization, A.S. and J.H.; supervision, R.T.; validation, A.S., J.H., and R.T.; project administration, A.S. All authors have read and agreed to the published version of the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding. Kingdom provided support in the form of salaries for the authors, and funding and resources for study execution.

Declaration of Conflicting Interests

The authors declared the following potential conflicts of interest with respect to the research, authorship, and/or publication of this article: All authors are employees of Kingdom and hold stocks and/or stock options in the company. Kingdom funded this research and is the supplier of the commercial material assessed in this study. The authors are committed to maintaining scientific integrity and adhering to ethical research practices. The paper reflects the view of the scientists and not the company.

Data Availability Statement

Datasets are available on reasonable request from the authors.

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