Author Names: Coralie Wright (BSc (Hons) Bioveterinary Science) and Aisling Carroll
The number of pet dogs (Canis lupus familiaris) in the common household is continually rising. The increasingly close contact between humans and cohabitant pets is leading to concerns regarding bacterial transmission of zoonoses. The dog water bowl has been identified as the third most contaminated item within the household, suggesting that it is able to act as a fomite for bacterial transmission, particularly where young or immunocompromised individuals are present. Studies in livestock have identified that water trough construction material influences bacterial count; however no similar research has been conducted for dog water bowls. The objectives of the current study were to identify which dog bowl material – plastic, ceramic or stainless steel – harbours the most bacteria over a 14 day period and whether the species identified varies between bowl materials. The study took place over 6 weeks. A sample of 6 medium sized (10-25kg) dogs aged 2-7 (mean= 3.8 ± 1.95) was used. All dogs were clinically healthy, housed individually and located within a rural environment. All bowls were purchased brand new and sterilised prior to a two week sampling period. On day 0, day 7 and day 14 swabs were taken from each bowl and 10-fold serial dilutions were conducted on blood agar. The cultured bacteria were subjected to biochemical testing and the most prominent bacteria from day 14 were further identified using PCR. A significant difference was identified for all bowl materials when comparing total CFU/ml between day 0 and day 7 and day 0 and day 14 (p<0.05). No significant difference was identified between total CFU/ml and bowl material (P>0.05), however descriptive statistics suggest that the plastic bowl material maintains the highest bacterial count after 14 days. Several medically important bacteria were identified from the bowls, including MRSA and Salmonella, with the majority of species being identified from the ceramic bowl. This could suggest that harmful bacteria may be able to develop biofilms more successfully on ceramic materials. Further research is required to identify the most suitable or alternative materials for dog water bowls.
The domestic dog (Canis lupus familiaris) has lived alongside humans for at least 15,000 years (Frantz et al., 2016; Wang et al., 2016; Shannon et al., 2015). Now one of humankind’s closest companions, the number of dogs kept within households is increasing, with an estimated population of 8.5 million in the UK (PFMA, 2018; Damborg et al., 2016; Wang et al., 2016; da Costa, Loureiro and Matos, 2013; Buma et al., 2006). Increasingly close contact between humans and their dogs allows favourable conditions for bacterial transmission both directly and indirectly (Damborg et al., 2016; Chomel and Sun, 2011; Deplazes et al., 2011; Bowman and Lucio-Forster, 2010; Fèvre et al., 2006; Guardabassi, Schwartz and Lloyd, 2004). Dogs spend a considerable amount of time outdoors, where they come into contact with a plethora of bacterial pathogens through contaminated soil, water, faeces and other animals (Lambertini et al., 2016). Additionally, the canine skin and oral microbiome is naturally colonised by a diverse range of bacteria (Hoffmann et al., 2014; Dewhirst et al., 2012; Elliot et al., 2005). Sixty percent of emerging pathogens are considered zoonotic (Dula and Pal, 2017; Cutler, Fooks and van der Poel, 2010), emphasising the importance of minimising the risk of bacterial transmission between humans, particularly the young and immunocompromised, and their cohabitant pets (Damborg et al., 2016; Moyaert et al., 2006). Lambertini et al. (2016) state that contact between owners and their dog occurs multiple times a day and can also be mediated by surfaces. The dog water bowl has been identified by Donofrio et al. (2012) as the third most contaminated item in the household, suggesting it is able to act as a fomite in bacterial transmission. There is currently no research exploring the most suitable material for dog water bowls with regards to bacterial load and species.
1.1 Development of Biofilms
Water is essential for life and almost every canine bodily function (Kenssington, 2014; Jéquier and Constant, 2010; Aspinall, 2012); however, the visual purity of water can disguise the proliferation of microorganisms (Folorunso, Kayode and Onibon, 2014). Bacterial biofilms can develop on many abiotic surfaces, particularly those in contact with non-sterile water (Flemming, 2011; Morita et al., 2011; Fuster-Valls et al., 2008). Livestock water troughs have been previously identified as reservoirs for bacterial growth and biofilm formation (Folorunso, Kayode and Onibon, 2014; Watson et al., 2012; Cook, Britt and Bolster, 2010; Avery et al., 2008; Sargeant et al., 2003; McGee et al., 2002; LeJeune et al., 2001a; LeJeune et al., 2001b); however research on dog water quality is incredibly limited. Biofilm formation in dog water bowls could serve as temporary or long term habitats for hygienically important bacteria (Wingender and Flemming, 2011). In addition to continuous re-infection of bacteria to dogs via ingestion of contaminated water, it is suggested that biofilms are responsible for 65% of all bacterial infections in humans (Rowson and Townsend, 2016). When washing or refilling a water bowl, humans may come into contact with bacterial biofilms and subsequently transfer pathogens from their hands to household surfaces (Lambertini et al., 2016). This is cause for concern as cohabitant pets often reside within the kitchen, which may ultimately lead to ingestion of contaminant bacteria (Lambertini et al., 2016; Behravesh et al., 2010).
Bacteria in biofilms exhibit increased resistance and tolerance to antimicrobials and environmental stressors (Ciofu et al., 2017; Wall and Mah, 2017; Rowson and Townsend, 2016; Steenackers et al., 2012), suggesting that they may be able to persist for longer within dog water bowls. This is concerning where zoonotic transfer is possible because of the shared antimicrobials in both human and veterinary medicine (Rendle and Page, 2018; Pomba et al., 2017; da Costa, Loureiro and Matos, 2013; Moyaert et al., 2006; Guardabassi, Loeber and Jacobson, 2004). This highlights the importance of minimising the risk of zoonotic spread from water bowls by identifying the most suitable material.
1.2 Influence of Materials on Bacterial Adhesion
The strength of bacterial adhesion depends on the organism’s surface properties and the material being colonised (Ribeiro, Monteiro and Ferraz, 2012; Van Houdt and Michiels, 2010; Faille et al., 2002). Material properties, including surface roughness and hydrophobicity influence the ability of bacterial cells to adhere to the surface and therefore determine the hygienic status of the material (Gharechahi, Moosavi and Forghani, 2012; Van Houdt and Michiels, 2010). A study by LeJeune et al. (2001a) into the hygiene of cattle water troughs identified a significant difference in bacterial coliform count between troughs constructed of different materials, with metal having lower counts than concrete and plastic. This study collected samples of water from each trough, however it is thought that 95% of the bacteria present in water systems are located at the surfaces and only 5% can be identified in the water phase (Flemming, Percival and Walker, 2002). Therefore, these findings may not accurately represent material influence on water quality, despite the large sample used by LeJeune et al. (2001a). Cook, Britt and Bolster (2010) examined decay rates of biofilms on different materials, artificially inoculated with Mycobacterium avium subsp. Paratuberculosis (Map), within glass water troughs. They identified that biofilm decay rates were fastest on stainless steel materials, followed closely by plastic. No significant difference was identified between stainless steel and plastic, however these findings are in agreement with the results of LeJeune et al. (2001a). Despite this, they only examined decay rates for Map and these results are therefore not representative for all bacterial species and their specific adherence to these materials as described by Van Houdt and Michiels (2010) and Faille et al. (2002).
Donofrio et al. (2012) identified high levels of bacteria within dog water bowls; however no specific links between bowl material and heterotrophic plate counts (HPC) were made. Samples of 26 items were tested, in 22 households, and the data revealed that sponge materials had the highest HPC, followed by porcelain, stainless steel and plastic. Dog bowls are commonly constructed of stainless steel, plastic and ceramic. These materials may permit heavy growth of bacterial pathogens as they lack antimicrobial effects (Fatoba et al., 2014). The results of this study suggest that these materials can harbour bacteria in high volumes and thus research specifically applied to bacterial load in dog water bowls constructed with these materials and the species present in them would be beneficial. Furthermore, identification of the most hygienic dog bowl material could allow for evidence-based pet ownership guidelines to be developed and thus reduce zoonotic transmission (Damborg et al., 2016; Leonard, 2014; Moyaert et al., 2006).
1.3 Dogs as a Reservoir of Zoonotic Transmission
There is substantial research which provides evidence of dogs acting as a reservoir of both zoonotic and resistant bacteria (Schwartz, Loeffler and Kadlec, 2017; Damborg et al., 2016; Chomel and Sun, 2013; Martins et al., 2013; Buma et al., 2006; Guardabassi, Loeber and Jacobson, 2004; Guardabassi, Schwartz and Lloyd, 2004). Several hygienically important bacteria have been isolated from dog mouths including Pasteurella spp., Staphylococcus spp., Bacillus spp. and many more (Zambori et al., 2013; Dewhirst et al., 2012; Elliot et al., 2005); therefore these species may potentially be present in dog water bowls.
A pathogen of major importance, Methicillin-resistant Staphylococcus aureus (MRSA), is of growing concern due to increased reports of zoonotic transmission (Han, Yang and Park, 2016; Loeffler et al., 2011; Abbott et al., 2010; Aklilu et al., 2010; Rutland et al., 2009). MRSA can potentially lead to mild or life-threatening skin infections in both humans and animals and it is highly resistant (Pantosti, 2012; Catry et al., 2010; Weese, 2010). Balen et al. (2013) conducted a year-long MRSA surveillance study in a veterinary hospital and discovered that some water bowls were contaminated for 2 consecutive months with the same pulsotype of MRSA, which was also detected on doors and floor surfaces. This study was conducted within a veterinary hospital with a large number of animals present and thus these findings cannot be extrapolated to household environments (Balen et al., 2013). However, the results demonstrate how bacteria, such as MRSA, can persist within a water bowl and can be spread throughout the environment – acting as a possible reservoir of bacterial transmission. Additionally, many studies have identified MRSA strain sharing amongst humans and their cohabitant pets (Faires, Tater and Weese, 2009; Sing, Tuschak and Hörmansdorfer, 2008; Boost, O’donoghue and Siu, 2007; Weese, 2006). However, cross-sectional studies of MRSA colonisation cannot identify direction of transmission and they do not eliminate common source infection (Weese, 2010).
Another microorganism with high zoonotic potential is Salmonella spp., which causes salmonellosis, a severe, common enteric disease in both humans and animals (Lowdon et al., 2015; Verma, Sinha and Singh, 2011). Dogs are thought to be asymptomatic carriers of salmonella and have been shown to act as a vector for bacterial contamination throughout the household (Lambertini et al., 2016; Hoelzer, Switt and Wiedmann, 2011; Bagcigil et al., 2007; Cherry et al., 2004; Sato et al., 2000). Additionally, recent studies have also identified human salmonella outbreaks caused by feeding dogs both dry and raw diets (CDC, 2012; Behravesh et al., 2010; Lenz et al., 2009; CDC, 2008). Weese and Rousseau (2006) state that food bowls can act as a source of infection for humans and animals if not sufficiently disinfected. They analysed the ability of salmonella to persist within both plastic and stainless steel food bowls, however no significant difference was determined when comparing overall persistence of salmonella and bowl material. A small sample size of 10 bowls was used, which significantly limits the statistical power of this study (Kylie et al., 2017). The presence of bacteria in food bowls may lead to cross contamination to water bowls, which are typically kept within the same area (Lambertini et al., 2016).
Current research suggests that animal drinking systems have the potential to act as a reservoir for multiple bacteria and that material does influence bacterial quantities (Folorunso, Kayode and Onibon, 2014; Martins et al., 2013; Donofrio et al., 2012; Watson et al., 2012; Cook, Britt and Bolster, 2010; Avery et al., 2008; LeJeune et al., 2001a). Weese, Rousseau and Arroyo (2005) state that pathogen survival in dog water bowls has not been adequately evaluated. Furthermore, to the author’s knowledge, there is no current research evaluating the influence of water bowl material on bacterial count and species present.
1.4 Aims and Objectives
The aim of the current study was to contribute to the limited existing literature by identifying whether the material of a dog’s water bowl – ceramic, plastic or stainless steel – and the length of use influences the quantity of bacteria present. Additionally, it was investigated whether the species identified from the water bowls varied between the three materials.
- To determine whether there was a difference in bacterial count with different bowl materials.
Null hypothesis: There will be no significant difference between bacterial count with different bowl materials.
Alternative hypothesis: There will be a significant difference between bacterial count with different bowl materials.
- To identify whether the length of use of the water bowl influences the bacterial count.
Null hypothesis: There will be no significant effect of the length of use of the water bowl on the bacterial count.
Alternative hypothesis: There will be a significant effect of the length of use of the water bowl on the bacterial count.
- To qualitatively determine, by observation, whether there is a difference in the species identified between bowl materials.
2.0 Materials and Methods
2.1 Pilot Study
A two week pilot study was conducted in order to eliminate any problems that may be encountered during the six week data collection period. It was acknowledged that the use of nutrient broth to transport the samples encouraged growth and increased contamination risk, and was thus rendering the bacterial counts inaccurate. To rectify this, phosphate buffer saline (PBS) solution was used as the transport medium, similar to that of Donofrio et al. (2012), as this does not encourage bacterial growth.
2.2 Bowl Sterilisation
The three construction materials tested were plastic, stainless steel and ceramic. All bowls were purchased brand new and sterilised in the same way. Gloves were worn throughout the process to prevent contamination. Following adapted methods of Jensen et al. (2013), a sterile basin was filled with boiling water, Dettol and washing up liquid. Each bowl was submerged, scrubbed gently, rinsed thoroughly and shaken dry. Bowls were immediately placed into a sterile autoclave bag and sealed until required.
2.3 Agar Preparation
Prior to each two week sampling period, Blood Agar Base (Oxoid, UK, CM0055) was prepared following the manufacturer’s instructions. This was autoclaved at 121oC, 103 kPa for 15 minutes in order to eliminate microbial contamination (Choi, Rodrigez and Sobsey, 2014). Once the agar had cooled to approximately 50-60oC, defibrinated horse blood was added aseptically at a 5% concentration and approximately 20ml of the solution was poured into petri dishes – in an aseptic environment – and allowed to set. Blood agar was used to culture the bacteria due to it being non-selective, rich in nutrients and able to grow a variety of bacteria (Dortet et al., 2014).
2.4 Broth Preparation
Prior to each two week sampling period MRVP broth (Merck, UK, V964212) and Nutrient Broth (Oxoid, UK, CM0001) was produced according to the manufacturer’s guidelines. Once the MRVP broth had dissolved, 10ml was syringed into individual test tubes and sealed with a lid. The test tubes and nutrient broth were autoclaved at 121oC, 103 kPa for 15 minutes and kept in the fridge until required.
2.5 Sample Collection
A sample of 6 dogs (5 female and 1 male), classified as medium weight (10-25kg) were used in this study (Vicente and Hammond, 2017). The age of the subjects ranged from 2-7 years (mean= 3.8 ± 1.95), and subjects were of various breeds (3 Cockapoos, 1 Border terrier, 1 Springer spaniel and 1 Golden retriever). All dogs were clinically healthy, fed a dry diet and had similar lifestyles. For example, all subjects were walked twice daily, housed individually within a rural location and no other pets were present in the household.
Bowls were placed in the kitchen by the back door in all households and the same position was used for all three construction materials. All owners were informed to thoroughly wash their hands before and after filling the dog bowl, and asked not to rinse, wash or disinfect the bowl. Samples were always collected at the same time in the morning. Gloves were worn throughout sample collections, sterile cotton swabs were used and all swabs were conducted by the same person (Donofrio et al., 2012), in the same manner – rubbed three times around the inner circumference of the bowl and once over the inner base, before being placed into 3ml sterile PBS. Samples were kept on ice and transported to the laboratory within two hours.
For the controls, on day 0 the bowl was filled with tap water, emptied and immediately swabbed, before being re-filled and placed in the selected position. Samples were collected again at day 7 and day 14.
2.6 Enumeration of Bacteria
2.6.1 Serial Dilution and Bacterial Count
Using modified methods of Folorunso, Kayode and Onibon (2014), Donofrio et al. (2010), LeJeune et al. (2001a) and LeJeune et al. (2001b), spread plate 10 fold serial dilutions were performed for each sample from 10-1 to 10-8. All serial dilutions were conducted within a biosafety cabinet to minimise contamination risk. The original sample was vortexed for 15 seconds using a Mini Vortex Mixer (Fisher Scientific, UK); 100μl aliquots were aseptically withdrawn using a micropipette and transferred into 900μl of sterile PBS (10-1). The solution was pipetted in and out to ensure thorough mixing. A 100μl aliquot was then aseptically withdrawn from the 10-1 solution and transferred into 10-2. This process was repeated until 10-8 was reached. A 100μl aliquot of each dilution from 10-3 to 10-8 was inoculated onto blood agar and spread using an L-shaped spreader. All plates were incubated at 37oC for 18-24 hours (Folorunso, Kayode and Onibon 2014).
Following incubation, total colony forming units (CFU) were counted and recorded. This was performed manually which is considered the gold standard approach (Davey, 2011; Clark et al., 2010). An average total CFU was calculated from the dilutions, for each sample, for each bowl type. Similar to LeJuene et al. (2001a) Plates with less than 30 colonies, or more than 300 colonies were excluded from the results as they are considered not statistically viable (Sutton, 2011; Rawling et al., 2009).
2.7 Isolation of Pure Bacteria
In a biosafety cabinet, the serial dilution plates were observed for individual, varying colonies. All colonies, from each sample, with a morphological difference were individually collected using a sterile, plastic inoculation loop and then streaked, using the four quadrant streaking method, onto blood agar plates to produce pure colonies. Plates were incubated at 37oC for 24 hours (Folorunso, Kayode and Onibon 2014).
2.8 Bacterial Identification
Pure bacterial samples were identified by their morphological characteristics, gram stain, and biochemical test results, similar to the methods of Folorunso, Kayode and Onibon (2014). The catalase, oxidase, Voges-Proskauer and methyl red test were conducted and interpreted following methods presented in literature (Sawain et al., 2018; Cobos-Trigueros et al., 2017; den Bakker et al., 2014; Hemraj, Diksha and Aveneet, 2013).
2.9 Freezing of Bacterial Isolates
The pure bacterial isolates from day 14, which were in the late log phase, were frozen at -80oC until required for polymerase chain reaction (PCR) (Kataoka et al, 2013).
2.10 Identification by Polymerase Chain Reaction
Following biochemical test results, further identification for the presence of potential medically important bacteria, isolated from day 14, was carried out using PCR.
2.10.1 DNA Extraction
The frozen samples were thawed at room temperature. Following 15 seconds of vortexing, using a Mini Vortex Mixer (Fisher Scientific, UK), a 100μl aliquot of each sample was added to an individual, labelled blood agar plate and spread using an inoculation ‘L-shaped’ spreader. The plates were incubated at 37oC for 48 hours. After incubation, DNA was extracted following the guidelines of Dashti et al. (2009).
2.10.2 PCR and Primers
The specific primers (Thermo Fisher Scientific, UK) selected for PCR and expected amplicon product size (bp) are displayed in table 1.
All primers were centrifuged for 30 seconds, hydrated using sterile distilled water according to manufacturer’s guidelines and stored at -20oC until required.
Table 1: A table to show the forward and reverse primer sequences used and the size of the amplicon product (bp)
|Organism/ gene||Primer Sequence
|Size of amplicon product (bp)||Reference|
|E.coli O157:H7/ eaeA (Forward)||AAG CGA CTG AGG TCA CT||450||(Holland et al., 2000; Louie et al., 1994)|
|E.coli O157:H7/ eaeA
|ACG CTG CTC ACT AGA TGT|
|MRSA/ mecA (Forward)||AAA ATC GAT GGT AAA GGT TGG C||533||(Bühlmann et al., 2008)|
|MRSA/ mecA (Reverse)||AGT TCT GGA GTA CCG GAT TTG C|
|Pasteurella multocida/ kmt
|TGC CAC TTG AAA TGG GAA ATG||168||(Król et al., 2011)|
|Pasteurella multocida/ kmt
|AAT AAC GTC CAA TCA GTT GCG|
|Pasteurella canis/ sod A (Forward)||GTA AAT AAT GCA AAT GCG G||186|
|Pasteurella canis/ sod A
|GCC TTG CAA AGT AGT AC|
|Staphylococcus species/ TstaG422
|GGC CGT GTT GAA CGT GGT CAA ATC A||370||(Martineau et al., 2001)|
|Staphylococcus species/ Tstag765
|TIA CCA TTT CAG TAC CTT CTG GTA A|
|Bacillus species/ p8FPL
|AGT TTG GAT CCT GGC TCA G||78||(Fernández-No et al., 2011)|
|Bacillus species/ p806R
|GGA CTA CCA GGG TAT CTA AT|
|Salmonella species/ ttr
|CTC ACC AGG AGA TTA CAA CAT GG||94||(Gwida and Al-Ashmawy, 2014)|
|Salmonella species/ ttr
|AGC TCA GAC CAA AAG TGA CCA TC|
A QuantiTect® Probe PCR Kit was used to prepare samples for PCR. A total volume of 20μl was used for amplification. Following the protocol, 10μl of QuantiTect® Probe PCR 2x Master Mix, 0.1μl of specific forward primer, 0.1μl of specific reverse primer and 4.8μl of RNase-free water were added aseptically to each sample tube. A 5μl aliquot of the DNA template was added to each tube. A blank was created using 5μl of RNase-free water instead of template DNA. Aseptic techniques were used throughout to eliminate contamination. All PCR was run using a Techne TC-3000 thermocycler (VWR, UK).
2.10.3 Gel Electrophoresis
A 50x stock solution of Tris acetate EDTA (TAE) was prepared and diluted to a 1x concentration by adding 980ml of distilled water to 20ml of 50x TAE. Each amplified DNA fragment required a different percentage of agarose gel for analysis. The appropriate weight of Electran® Agarose DNA grade (VWR,Belgium) was added to 150ml of TAE in a conical flask and stirred until dissolved. The solution was heated for 2 minutes and then heated gently until the solution was clear. After slight cooling, the gel was poured into a horizontal gel tank and combs were added. Once the gel had set, the combs were removed gently to produce the wells. TAE x1 was poured over the gel until it was covered.
Using a micropipette, 5μl of 100bp DNA marker, containing loading dye, was added to 2μl of 10x GelRed and 6μl of this solution was added to the first and last well. A 5μl aliquot of DNA sample was mixed with 2μl of 10x GelRed and 1μl of 6x loading dye (bromophenol blue) and then 6μl was added to the remaining wells. All runs of gel electrophoresis were examined by UV light using the UVP gel Doc-it reading system. Observations were made for clear bands at the specific amplicon product size (bp), displayed in table 1.
2.11 Statistical analysis
All statistical analysis was carried out using SPSS software (IBM SPSS, version 24, USA). Tests were conducted on the average total CFU/ml for each sample, for each bowl type. Descriptive statistics including mean and standard deviation were calculated. The Kolmogorov Smirnoff test was conducted to determine normality. A Kruskal Wallis was used to determine whether there is a significant difference between the average total CFU/ml for each bowl construction material. A Mann Whitney or independent T test, depending on normality, was used to determine if there is a significant difference in average total CFU/ml between day 0 and day 7, day 7 and day 14, and day 0 and day 14.
2.12 Ethical considerations
Recent studies have found evidence of dogs having the ability to remember the physical properties of objects (Kundey et al., 2010; Pattison et al., 2010). Additionally, exposure to novel surroundings has been shown to cause dogs stress (Rooney, Gaines and Bradshaw, 2007). Changing the material of a dog’s bowl could induce stress. If any signs of stress or adversity to drinking were displayed, owners were advised to remove their dog from the study.
Owners provided informed consent for each subject via the completion of a participant consent form. All data provided was held in accordance of The Data Protection Act (1998). The researcher and owners had the right to withdraw a subject at any time, for any reason, until the point of data analysis.
3.1 Bacterial Enumeration
For each bowl material on each day, an average total CFU/ml was calculated using all samples. The average total CFU/ml for each bowl material over the two week period is presented in table 2 and figure 1. No bacterial growth was observed from the initial control samples taken on day 0. Bacteria were cultured from all three bowl materials on day 7 and day 14. However, for the plastic bowl material, two of the six samples showed no growth on both day 7 and day 14. On day 7, the stainless steel bowl had the highest bacterial count of 250.33 ± 236.47 CFU/ml x 106. However, although all counts were lower after 14 days, the plastic bowl had the highest remaining bacterial count with 43.26 ± 62.40 CFU/ml x 106. The ceramic bowl had the lowest bacterial count throughout all test days of the study.
Table 2: A table to show the average total CFU/ml ± standard deviation (SD) (1.0 x 106) for each bowl material over time.
|Bowl material||Average total ± SD CFU/ml (1.0 x 106)|
|Day 0||Day 7||Day 14|
|Ceramic||0.00 ± 0.00||18.56 ± 40.50||2.20 ± 2.65|
|Plastic||0.00 ± 0.00||92.28 ± 191.21||43.26 ± 62.40|
|Stainless steel||0.00 ± 0.00||250.33 ± 236.40||10.95 ± 16.96|
The Kolmogorov Smirnoff test revealed the data to be mostly non-parametric (P<0.05). The Kruskal Wallis identified no significant difference when comparing bowl material and total CFU/ml (P>0.05); therefore the null hypothesis that there is no significant difference between bacterial count and bowl material must be retained. When testing for difference between day and average total CFU/ml, the Mann Whitney U test revealed a significant difference between day 0 and day 7 and between day 0 and day 14 for all bowl materials (P<0.05); thus the null hypothesis that there is no significant difference between the length of use of the water bowl and the bacterial count can be rejected. However, no significant difference in total CFU/ml was identified for all bowl materials between day 7 and day 14 (P>0.05).
Figure 1: A bar chart presenting the average total CFU/ml (1.0 x 106) over time.
3.2 Bacterial Identification
3.2.1 Biochemical test results
If bacterial characteristics identified on day 7 were identified again on day 14, the suspected genus was used for the selection of specific PCR primers, which are displayed in table 1. The colony morphology and biochemical test results for these isolates are presented in table 3. The results for the Voges-Proskauer tests are considered anomalies as no colour change was observed for this test for all isolates on day 7 and day 14.
Table 3: A table to show the colony morphology, biochemical test results and possible genus for all isolates that appeared in samples at both day 7 and day 14.
|Colony morphology||Biochemical test||Possible genus|
3.2.2 Polymerase Chain Reaction Results
Following the biochemical test results, all day 14 samples were subjected to PCR for identification of seven hygienically important bacteria, presented in table 1. The results for PCR are presented in table 4. The E.coli O157:H7 primer identified one isolate from the ceramic bowl material, displayed by the faint amplification at 450 base pairs (bp). The MRSA primer detected seven samples with positive amplification at 533bp. From the seven positive isolates, three originated from the stainless steel bowl and four from the ceramic bowl. The primer for the genus of staphylococcus only displayed 3 positive amplifications at 370bp, two from the stainless steel bowl and one from ceramic. No clear bands were present at 168bp for identification of Pasteurella multocida. Two samples, one from both the stainless steel and plastic bowl were identified as positive for Pasteurella canis, displayed by the bands at 186bp. Three ceramic and one plastic bowl displayed amplification at 78bp for detection of Bacillus species. Five Salmonella species were identified by the presence of faint bands at 94bp. One of the positive isolates originated from the plastic bowl and the remaining four from the ceramic.
Table 4: A table to show the results from PCR. S1 – S6 represents the stainless steel bowl samples, C1 – C6 the ceramic bowl samples and P1-P5 the plastic bowl samples. ‘+’ represents the presence of the specific amplicon product.
|– Control (blank)||–||–||–||–||–||–||–|
At present, dogs are the most popular pet in the UK and concerns of zoonotic disease transmission are rising due to their increasingly close contact with owners (PMFA, 2018; Damborg et al., 2016; Wang et al., 2016; da Costa, Loureiro and Matos, 2013; Buma et al., 2006). Dog water bowls have the potential to act as a reservoir of zoonotic bacteria, similar to the findings in livestock water troughs (Folorunso, Kayode and Onibon, 2014; Watson et al., 2012; Cook, Britt and Bolster, 2010; Avery et al., 2008; Sargeant et al., 2003; McGee et al., 2002; LeJeune et al., 2001a; LeJeune et al., 2001b). The current study aimed to explore the effects of material, and length of use, on the quantity and species of bacteria present in dog water bowls. The results indicate that bowl material may influence bacterial count and the species isolated.
4.1 Bacterial Enumeration
4.1.1 Influence of bowl material on bacterial count
On day 7, there was no significant difference identified between bowl material and average total CFU/ml (P>0.05). Although not significantly different, the descriptive statistics suggest that the stainless steel bowl had the highest bacterial count, followed by plastic and then ceramic. Stainless steel is considered a hydrophilic material with a high surface energy and negative charge (Hamadi et al., 2014; Hočevar et al., 2014; Di Bonaventura et al., 2008; Sinde and Carballo, 2000; An and Friedman, 1998). Bacteria frequently attach in higher numbers to hydrophobic materials, such as plastic (Hočevar et al., 2014; Di Bonaventura et al., 2008; Sinde and Carballo, 2000); however Gebhardt et al. (2012) state that cell growth on hydrophobic surfaces is slower than on hydrophilic surfaces. Additionally, Flint, Brooks and Bremer (2000) examined the bacterial adhesion of two streptococcus species to stainless steel, glass and other metal coupons and concluded that negatively charged surfaces attracted more bacteria than positively charged surfaces. This study examined two bacterial species and therefore is not representative of all bacterial species adhesion mechanisms (Greene et al., 2016). Despite this, these findings may suggest why stainless steel had a higher bacterial count than plastic on day 7.
Similarly, on day 14, no significant difference was identified between bowl material and average total CFU/ml (P>0.05). In contrast to day 7, the plastic bowl had the highest bacterial count. Plastic materials are known to degrade rapidly and surface scratches and wear increase bacterial adhesion (Bohinc et al., 2014; Crawford et al., 2012; Verran et al., 2008). An investigation over a longer time period may be beneficial. Cook, Britt and Bolster (2010) examined bacterial decay rate of Map over one year and identified that plastic disks have slower bacterial decay rates than stainless steel disks, although not significantly different. This study was conducted in controlled laboratory conditions, and may not be applicable in vivo (Sorrentino et al., 2018; Cook, Britt and Bolster, 2010). Despite this, LeJeune et al. (2001a) observed metal water troughs to have significantly lower bacterial counts than plastic water troughs over 8 months. A large sample of 467 troughs was studied, suggesting this data to be reliable with high statistical power (Halsey et al., 2015). The lack of statistical significance found in the current study is likely to be due to the small sample used and larger samples should be considered in future research (Halsey et al., 2015).
The ceramic bowl obtained the lowest bacterial count on both day 7 and day 14, although it was not significantly different from the other materials. There have been several reports that ceramic materials exhibit lower bacterial adhesion than other materials due to its higher hydrophilicity (Hofs et al., 2011; Eick et al., 2004). Sorrentino et al. (2018) studied the bacterial adhesion of S.aureus and S.epidermis to ceramic, metal and plastic disks and identified that ceramic materials had a significantly lower bacterial adhesion, slower biofilm development and thinner biofilm formation than the other materials (P<0.05). Additionally, the plastic material showed the highest bacterial adhesion, in line with the findings of this study. Similarly, Fatoba et al. (2014) placed various materials into individual liquid cultures of Bacillus, E.coli and S.aureus and conducted bacterial counts after various time periods. Their findings suggest that the ceramic material obtained lower bacterial counts, followed by plastic and then stainless steel when immersed in E.coli. However, all three materials had very similar counts for both Bacillus and S.aureus. Although there is a body of evidence that suggests ceramic materials obtain lower bacterial counts – similar to the findings of the current study – some studies demonstrate that adhesion is bacteria dependent (Fatoba et al., 2014; Ribeiro, Monteiro and Ferraz, 2012; Hofs et al., 2011; Van Houdt and Michiels, 2010; Eick et al., 2004; Faille et al., 2002). This highlights the need for further research in this area.
Although there was no significant difference identified between bowl material and average total CFU/ml in the current study, descriptive statistics and the above studies support the findings that the ceramic bowl has lower bacterial adhesion. However, further research is required to identify if there is a more hygienic bowl material. Additionally, Crawford et al. (2012) state that it is imperative to have a comprehensive characterisation of the specific surface properties when studying bacterial adhesion to materials and therefore thorough examination of the materials should be implemented in future research.
4.1.2 Influence of length of use on bacterial count
A significant difference was identified for all bowl materials when comparing average total CFU/ml between both day 0 and day 7, and day 0 and day 14 (P<0.05). Bacterial count increased significantly between days 0 and 7. This demonstrates that when bowls are refilled without cleaning, they produce a favourable environment for bacterial proliferation (Folorunso, Kayode and Obinon, 2014). This is in agreement with the findings of Folorunso, Kayode and Onibon (2014) who studied the bacterial count in poultry water troughs at three farms and found that the bacterial load progressively increases over 7 days.
Interestingly, the bacterial counts reduced for all bowl materials on day 14; however they were not significantly different from day 7. This suggests that high quantities of bacteria can persist within dog bowl biofilms. Many environmental factors, such as competition and nutrient availability, influence bacterial survival and adhesion – which may suggest why the bacterial counts slightly reduce (Crawford et al., 2012; Di Bonaventura et al., 2008; Katsikogianni and Missirlis, 2004). Many cells aggregated as part of a biofilm inhibit their motility, preventing them from searching for optimal environments when nutrient depletion occurs, ultimately causing cell death (Tuson and Weibel, 2013). Furthermore, the bacteria may have been removed by the dog or upon refilling. Additionally, the swab taken from the bowl surfaces on day 7 is likely to have removed and disrupted bacteria present in the biofilm, reducing the count on day 14. This represents a major limitation of the current study which may be overcome in future by conducting swabs on day 7 and then beginning a new 14 day sampling period with no day 7 swab. Despite this, the lack of significant difference between day 7 and day 14 highlights the importance of daily cleaning regimes as high levels of bacteria in dog drinking water could significantly impact their health and welfare and increase transmission to human cohabitants – particularly immunocompromised individuals (Damborg et al., 2016; Folorunso, Kayode and Obinon, 2014; Weese et al., 2010). Furthermore, this study demonstrates that the water bowl acts as a fomite; therefore it may be a vector of diseases with high veterinary importance.
4.2 Bacterial Identification
4.2.1 Biochemical tests
Bacteria with the same biochemical characteristics isolated from both day 7 and day 14 were identified to the genus level using biochemical tests and morphological characteristics (Vaseekaran, Balakumar and Arasaratnam, 2010). The isolates were identified as E.coli, Salmonella, Staphylococcus, Pasteurella and Bacillus, according to biochemical results presented in literature (Folorunso, Kayode and Onibon, 2014; Patnaik, Prasad and Ganguly, 2014; Verma et al., 2013; Vaseekaran, Balakumar and Arasaratnam, 2010). The limited number of biochemical tests used in this study meant that accurate identification was difficult (Janda and Abbott, 2002). Additionally, the Voges-Proskauer test results were identified as anomalies and excluded from the results. Biochemical tests have low specificity and interpretation of can be difficult if untrained (vanVeen, Claas and Kuijper, 2010). Furthermore, many diverse species can share the same biochemical characteristics, causing unreliability (Srinivasan et al., 2015; Heikens et al., 2005). Despite this, they allowed for selection of appropriate primers for PCR identification. Accurate identification of microorganisms is fundamental in order to implement suitable infection control and improve general hygiene (Nomura, 2015; Pahlow et al., 2015; Srinivasan et al., 2015).
4.2.2 Polymerase Chain Reaction
PCR provides a highly sensitive and specific method for bacterial species identification and avoids the limitations of biochemical tests (Pahlow et al., 2015, Cherkaoui et al., 2010; Vasoo, Stevens and Singh, 2009). Interestingly, the bacteria tested for by PCR were most frequently isolated from the ceramic bowl, followed by stainless steel and then plastic, suggesting an influence of bowl material on species.
E.coli O157:H7 was identified in one sample for the ceramic bowl material. The genus E.coli was identified in many bowls from the biochemical tests; however many species of E.coli are commensal and are therefore typically harmless (Chandran et al., 2017; Odonkor and Ampofo, 2013). E.coli O157:H7 is a highly pathogenic serotype of E.coli often found on the intestinal tract of many animals; therefore dogs are likely to come into contact with contaminated faecal matter when on walks (Lambertini et al., 2016; Mendonça et al., 2012). The presence of E.coli O157:H7 in one ceramic water bowl is concerning as infection can cause bloody diarrhoea, hemorrhagic colitis and fatal haemolytic uremic syndrome and it is consequently a major threat to public health (Albanese et al., 2018; Gould et al., 2016; Li et al., 2014).
Staphylococcus spp. was confirmed by PCR to be present in two stainless steel bowls and one ceramic bowl. The low presence of Staphylococcus is surprising as S.aureus has been previously isolated from dog water bowls (Donofrio et al., 2012). Furthermore, a study by Bean and Wigmore (2013) examined 117 healthy dogs for the presence of S.aureus and S.pseudointermedius and concluded that the mouth and perineum were the most common sites for detection of these bacteria. This suggests that these species are likely to be present within water bowls. However, this study was conducted in Australia and thus cannot be generalised to the worldwide dog population (Jaeger et al., 2010). Despite this, studies in Denmark have also suggested the mouth and perineum as the area’s most highly colonised with S.pseudointermedius (Paul et al., 2012). Additionally, studies assessing the microbiology of canine bite wounds have identified Staphylococcus spp. as one of the most commonly isolated bacteria (Abrahamian and Goldstein, 2011; Thomas and Brook, 2011; Talan et al., 1999). These studies suggest that Staphylococcus spp. are highly prevalent in dog mouths. The low presence of these species in this study may be as a result of an unsuitable primer, as it was designed for qPCR, which was not performed in this study (Martineau et al., 2001); however the positive control amplified with a very clear band.
Interestingly, seven isolates of MRSA were identified, three in the stainless steel bowl and four in the ceramic. This may suggest that false negative or false positive results were obtained due to the discrepancies between the Staphylococcus and MRSA findings. The clearest amplicon band for MRSA was seen in sample C5, which was negative in the Staphylococcus PCR. This suggests the error is in the Staphylococcus primer where minor modifications were made to the cycling conditions and this may have impacted the efficacy. Despite this, the identification of MRSA in the water bowl is concerning as it can be transferred to humans and cause life threatening illness (Faires, Tater and Weese, 2009; Sing, Tuschak and Hörmansdorfer, 2008; Boost, O’donoghue and Siu, 2007; Weese, 2006). Mafu et al. (2013) state that there is limited knowledge about the adhesion of MRSA to surfaces and the current study suggests it preferentially adheres to hydrophilic materials. Alternatively, the other bowls may not have been contaminated with the bacterium. Balen et al. (2013) showed that MRSA can survive within a dog water bowl for 2 months; suggesting the need for a suitable cleaning regime that is effective against this highly resistant bacteria.
Salmonella species were confirmed in four ceramic bowls and one plastic bowl. Salmonella has hydrophobic cell properties and has been previously reported to preferentially adhere to hydrophobic surfaces, such as plastic (Veluz, Pitchiah and Alvarado, 2012; Chia et al., 2009; Chia et al., 2008). This does not agree with the findings of the current study; however, Oliveira et al. (2006) suggest that adherence is entirely strain dependent. Furthermore, the bacterium may not have been introduced into all bowls. Although only 9% of human infections are attributable to animal contact, Salmonella results in severe gastrointestinal disorders (Lowdon et al., 2015; Verma, Sinha and Singh, 2011; Majowicz et al., 2010). It can also lead to abdominal pain, weakness and fever in dogs (Marks et al., 2011). Identifying water bowls as a vector for Salmonella transmission may lead to a reduction of infections if owners’ education is increased.
PCR identified Bacillus in three ceramic bowls and one plastic bowl. The primer identified B.cereus, B.licheniformis and B.subtilis and although it is not clear which exact species were isolated from these bowls, all of these species can be pathogenic (Fernández-No et al., 2011). Bacillus species are often found within soil, suggesting that dogs are likely to come into contact with it on walks (Lambertini et al., 2016; Earl, Losick and Kolter, 2008). Identification of Bacillus species is concerning as they develop spores; this increases their resistance to cleaning procedures and Faille et al. (2002) found that removal was harder on hydrophobic materials.
Pasteurella canis was identified in one stainless steel bowl and one plastic bowl. Pasteruella multocida was not identified. Pasteurella spp. are responsible for several soft tissue infections, occasional respiratory tract infections and in rare cases, meningitis (Faceira et al., 2017; Zambori et al., 2013).The low presence of these species was not expected as Pasteurella species were identified frequently from the biochemical tests and are also frequently isolated from dog mouths and bites (Faceira et al., 2017; Oehler et al., 2009; Talan et al., 1999). This may suggest PCR errors were encountered, for example, the heat lysis method of DNA extraction may not have been successful and thus there was no DNA present to be amplified. Additionally, the ladder did not migrate clearly for P.multocida which may suggest that the gel did not run correctly.
The limited sample used within this experiment does not allow for findings to be extrapolated to all dogs. In future, a larger sample, including a larger variety of breeds should be used. Additionally, the findings of this study are only applicable to dogs in a rural location of Devon and geographical studies should be considered to identify the difference between urban and rural housed dogs.
Although great care was taken to ensure all components of the PCR were managed correctly, the high number of samples meant that maintaining the reagents at 4oC was difficult. Prolonged exposure to higher temperatures can denature the Taq polymerase enzyme, reducing the sensitivity of PCR (Peters et al., 2004). In addition, non-specific activation of Taq polymerase can cause non-specific amplification (Ashrafi, Yee and Paul, 2009). Furthermore, the lack of positive controls throughout limited the ability to compare the amplifications to a known positive amplicon product.
4.4 Future Research
Although not tested in this study, Fatabo et al. (2014) demonstrated that copper and brass materials had antimicrobial and bactericidal effects against bacteria over time. Additionally, Mehtar et al. (2008) discovered that copper surfaces decreased bacterial loads from 107 CFU/ ml to below detectable limits within 180 minutes. However, these studies are conducted in laboratory settings and may not be representative of the antimicrobial properties of copper in heterogeneous bacterial environments (Gould et al., 2009). Despite this, the antimicrobial properties in these materials should be studied in vivo for the development of antimicrobial dog water bowls, reducing bacterial count and viability. This would subsequently improve dog welfare and health and reduce the risk of zoonotic transmission.
Additionally, due to the high level of antimicrobial resistance in biofilm bacteria, antimicrobial susceptibility testing should be carried out in order to identify effective disinfectants against the bacteria isolated from dog water bowls (Ciofu et al., 2017; Wall and Mah, 2017; Rowson and Townsend, 2016; Steenackers et al., 2012; Reller et al., 2009). This would allow for evidence-based cleaning routines to be developed (Leonard, 2014), improving both owner and dog health and welfare.
The current study contributes to the limited research on dog water bowl microbiology and begins to determine the impact of bowl material on the bacteria present. The findings demonstrate how ceramic, plastic and stainless steel water bowls allow for proliferation of bacteria – a health risk for owners and their dogs. No significant difference was identified between the bowl material and the bacterial count over two weeks, although descriptive statistics indicate that differences might be present, suggesting that further research involving a larger sample size is required. However, ceramic bowls appear to reduce bacterial proliferation more than stainless steel and plastic bowls. Additionally, other materials, such as copper, should be studied due to their antimicrobial properties. This would allow for identification of a more hygienic bowl that could reduce the risk of zoonotic transmission.
The length of use influences the bacterial count in all three bowl types, which demonstrates the need for daily cleaning regimes and increased owner awareness on the impacts of bowl hygiene. A study over a longer time period would be beneficial in order to identify the bowl material that minimises bacterial load. This would lead to improvements in both human and canine health.
PCR accurately identified multiple bacterial species, many of which have pathogenic potential. This presents the dog water bowl as a possible vector in disease transmission, both to humans and other animals. The frequent observation of MRSA and Salmonella isolated from the bowls is concerning due to their infectious nature for both humans and animals. However, this identifies an area where good hygiene implementation could reduce the number of infections from these bacteria and ultimately improve dog welfare. The ceramic bowl isolated the highest number of harmful bacteria, despite having the lowest bacterial count. This suggests that these bacteria may be able to adhere more easily to ceramic materials, demonstrating how bowl material may impact the species present in the bowl. Ultimately, more research is required before suitable conclusions can be drawn.
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