Hydrotherapy is an integral component of equine rehabilitation, with 82% of respondents from an “International Survey Regarding the Use of Rehabilitation Modalities in Horses” choosing cold water hydrotherapy as part of their top 10 treatment recommendations for horses.1 Thermal hydrotherapy, both hot and cold, is commonly utilized to assist with athletic recovery2–5 and for the treatment of equine distal limb injuries in clinical practice.6,7 Previous equine publications7–10 highlight the utility of this therapy for conditions such as distal limb cellulitis and lymphangitis, tendinitis, desmitis, arthritis, and inflammatory conditions that cannot be treated with systemic medications when horses are in active competition or suffering from a preexisting condition such as renal compromise. The Equine Hydrotherapy Working Group currently cites skin lesions and recent IA injection (within 4 days) as contraindications to underwater treadmill exercise11; however, no such guidelines have been established for saltwater hydrotherapy systems.
While saltwater can have antimicrobial properties depending on its salinity,12–14 human saltwater spas and other recreational saltwater supplies have demonstrated pathogenic microbial growth even with the addition of chlorine filtration.15,16 The ability of bacteria to grow in saltwater from equine hydrotherapy units has yet to be evaluated. In a clinical setting, small abrasions, lacerations, or cutaneous bacterial infections may accompany the common equine conditions listed above that benefit from immersive thermal hydrotherapy. Previous literature17–19 has documented Escherichia coli, Pseudomonas aeruginosa, Staphylococcus aureus, and Streptococcus equi subspecies zooepidemicus as common isolates of equine skin lesions, with deeper and more chronic wounds often being polymicrobial.20 Therefore, the purpose of this study was to understand the ability of these common bacterial isolates to survive in saltwater obtained from a commercially available equine hydrotherapy unit when maintained at either full salinity (20 ppt) or half salinity (10 ppt) and at temperatures of either 2 °C used for cryotherapy, 44 °C used for heat therapy, and 22 °C for room temperature at which hydrotherapy holding tanks may be kept over 96 hours. We hypothesized that a salinity of 20 ppt would effectively eliminate common equine pathogenic bacteria at all temperatures examined.
Methods
A schematic of the study design is shown (Figure 1). Briefly, sterilized saltwater samples at full and half salinity were inoculated with the 4 bacterial isolates being investigated. Salinity was measured every 24 hours to ensure consistency. Inoculated samples were maintained at the 3 temperatures being evaluated and plated on blood agar plates every 24 hours up to and including 96 hours for bacterial quantification. The entire study was performed over a 2-week period (January 22, 2024 to February 2, 2024).
Saltwater samples were obtained over a 2-week period (January 22, 2024 to February 2, 2024) from a clinical equine hydrotherapy unit. Sterilized samples of both full and half salinity were inoculated in duplicate with single colonies of each bacterial strain prepared to 0.5 McFarland Standards (McF) and maintained at each respective temperature. Bacterial quantification dilutions were plated and counted in triplicate at time 0 hours and then every 24 hours up to and including 96 hours for all samples.
Citation: American Journal of Veterinary Research 85, 11; 10.2460/ajvr.24.07.0184
Bacterial isolates
Individual commercially available bacterial strains of E coli (ATCC 25922), P aeruginosa (ATCC 27853), S aureus (ATCC 976), and S zooepidemicus (ATCC 35246) were inoculated onto blood agar plates containing 5% sheep RBC and incubated at 37 °C for 18 to 24 hours. Single, pure colonies were selected and utilized to prepare McFarland Standards (McF).
Saltwater sample preparation and inoculation
Saltwater prepared according to the manufacturer's directions to reach a maximum salinity of 20 ppt (specific gravity [SG], 1.014) was collected from the holding tank of a commercially available equine hydrotherapy unit (Nautilus Equine Saltwater Spa; Equine Spa Therapy LLC) used in our hospital for the treatment of clinical patients. This saltwater is created by the addition of 25 kg of Fine Bokek Dead Sea Salt sodium chloride (Bokek) and 22.7 kg of Ultra Epsom magnesium sulfate (Ultra Epsom; Saltworks) to 1,582 L of tap water. Salinity was measured using a refractometer (AgTec LLC) and then characterized and maintained as either full salinity (20 ppt; SG, 1.014) or diluted with distilled water and characterized and maintained at half salinity (10 ppt; SG, 1.007) for further testing. All water samples were sterilized using a steam autoclave to eliminate any potential existing bacterial contamination before bacterial inoculation for the purpose of this study.
Samples of full salinity (20 ppt) and half salinity (10 ppt) were then aliquoted in duplicate for inoculation of 1 of 4 bacterial strains (E coli, P aeruginosa, S aureus, and S zooepidemicus) and incubation at each temperature condition (2, 22, and 44 °C). Each bacterial inoculum was prepared to a 0.5 McF (approx 108 CFU/mL) using a calibrated standard colorimeter (DensiCHEK Plus; bioMerieux), and 100 μL of this preparation was added to each saltwater sample to reach a concentration of 1.5 X 106 CFU/mL. After inoculation, each respective water sample was maintained at each respective temperature condition. The 2 °C samples were kept in a laboratory refrigerator set to 2 °C, the 22 °C samples were kept on the laboratory benchtop in a room with the thermostat set to 22 °C, and the 44 °C samples were kept in a water bath set at 44 °C. Salinity measurements were repeated every 24 hours to ensure the maintenance of salinity concentrations.
Bacterial concentration
To quantify bacteria, dilutions of each duplicate sample were prepared using 1:10 dilutions in sterile PBS within 96-well plates. Dilutions were made up to 10−4, and aliquots of each dilution were plated on blood agar plates. Dilutions were plated in triplicate with 20 μL of sample each. Plates were incubated at 37 °C for 18 to 24 hours. Counts were performed for each dilution sample plated on the concentration with between 10 and 100 visible colonies; the average count of all 3 was the reported concentration. The water samples were maintained at each respective temperature throughout the duration of the experiment. The lower limit of detection was set at 1 CFU/mL (1.22 log10) per triplicate plated in the event no isolates were obtained.
Statistical analyses
Bacterial concentrations were log10 transformed for analysis. Data analysis was performed using SPSS Statistics (version 29.0; IBM). A γ-regression mixed effects model was chosen to account for the larger variability of log counts. Effects of salinity, temperature, time, and interactions up to and including all 3 were assessed. Post hoc analyses were performed to determine differences between temperatures and how bacterial concentrations changed over time within each temperature. For temperatures with bacterial concentration difference significance, pairwise comparisons were performed and adjusted for familywise error rates using a sequential Bonferroni adjustment. Significance was set at P < .05. All graphs were generated with GraphPad Prism (version 10.1.1; Graphpad Software Inc), and composite figures were created with BioRender.21
Results
For all bacterial strains examined, there was a significant interaction of temperature and time on bacterial concentrations (P < .001). In contrast, there were no significant effects involving salinity for any bacterial strain examined indicating that salinity did not have any relationship with bacterial concentration. For this reason, the effect of salinity was excluded from further analyses and only temperature and time are discussed below. Additionally, duplicate wells for each salinity concentration at each temperature for each bacterial strain were then grouped by temperature only for each bacterial strain to yield a sample size of n = 4 per group.
To determine how bacterial concentrations changed at different temperatures within each time point, joint post hoc tests of temperature by time point were performed and identified significant differences for all time points after 0 hours for all bacterial strains. When performing pairwise comparisons of temperatures within 24- to 72-hour time points, significant differences in bacterial concentrations were identified between the 44 °C temperature and both the 2 and 22 °C temperatures for all time points (P < .001) for E coli, P aeruginosa, and S aureus, with the lower temperatures having higher bacterial concentrations (Figure 2). No differences in bacterial concentrations were identified when comparing between the 2 and 22 °C at any time point for E coli, P aeruginosa, and S aureus. However, S zooepidemicus was unique in that when comparing between temperatures, significant differences in bacterial concentrations were identified between all 3 temperatures for all time points from 24 to 72 hours (P ≤ .013), with the lower temperatures having higher bacterial concentrations. At 96 hours, significant differences in bacterial concentrations of S zooepidemicus were identified between the 2 °C temperature and both the 22 and 44 °C temperatures (P < .001), with the 2 °C having higher bacterial concentrations and with no differences in bacterial concentrations between the 22 and 44 °C temperatures.
Line graphs of bacterial concentrations in CFU/mL (log10) over time in hours with pairwise comparisons of temperatures within each time point for E coli (A), P aeruginosa (B), S aureus (C), and S zooepidemicus (D). The different temperatures are denoted by color and shape, the blue circles represent 2 °C, the green diamonds 22 °C, and the red squares 44 °C. Significant differences between temperatures (P < .05) at each time point: *between the 44 °C temperature and both the 2 and 22 °C temperatures; #between all 3 temperatures; and ^between the 2 °C temperature and both the 22 and 44 °C temperatures; n = 4/group. LLOD = Lower limit of detection.
Citation: American Journal of Veterinary Research 85, 11; 10.2460/ajvr.24.07.0184
To determine how bacterial concentrations changed over time within each temperature, joint post hoc tests of temperature by time point were performed. Significant differences in bacterial concentrations over time at 2 °C were found for E coli and S zooepidemicus, and significant differences in bacterial concentrations over time at 22 °C were found for S zooepidemicus only. Significant differences in bacterial concentrations over time at 44 °C were found for all bacterial strains. Pairwise comparisons of time points by temperature are displayed for each bacterial strain at 2 (Figure 3), 22 (Figure 4), and 44 °C (Figure 5). Of note is the fact that reduction in the bacterial concentrations of E coli and S zooepidemicus at 2 °C was very modest at less than or equal to 1 log10 reduction, while at 22 °C, S zooepidemicus concentrations steadily decreased to the lower limit of detection of the quantification method by 96 hours. At 44 °C, E coli, S aureus, and S zooepidemicus all had substantial reductions in bacterial concentrations to the lower limit of detection of the quantification method by 24 to 48 hours. P aeruginosa concentrations also decreased over the first 24 hours at 44 °C, but with a modest 2 log10 reduction, and then stabilized with no further decrease.
Bar graphs of bacterial concentrations in CFU/mL (log10) over time at 2 °C with pairwise comparisons of time points and P values for all significant differences (P < .05) E coli (A), P aeruginosa (B), S aureus (C), and S zooepidemicus (D); n = 4/group. Each bar represents a 24-hour time increment between 0 and 96 hours. LLOD = Lower limit of detection.
Citation: American Journal of Veterinary Research 85, 11; 10.2460/ajvr.24.07.0184
Bar graphs of bacterial concentrations in CFU/mL (log10) over time at 22 °C with pairwise comparisons of time points and P values for all significant differences (P < .05) for E coli (A), P aeruginosa (B), S aureus (C), and S zooepidemicus (D); n = 4/group. Each bar represents a 24-hour time increment between 0 and 96 hours. LLOD = Lower limit of detection.
Citation: American Journal of Veterinary Research 85, 11; 10.2460/ajvr.24.07.0184
Bar graphs of bacterial concentrations in CFU/mL (log10) over time at 44 °C with pairwise comparisons of time points and P values for all significant differences (P < .05) for E coli (A), P aeruginosa (B), S aureus (C), and S zooepidemicus (D); n = 4/group. Each bar represents a 24-hour time increment between 0 and 96 hours. LLOD = Lower limit of detection.
Citation: American Journal of Veterinary Research 85, 11; 10.2460/ajvr.24.07.0184
Discussion
Equine saltwater hydrotherapy units offer a hypertonic environment with the capability to provide cryotherapy or heat to the equine limbs depending on the condition being treated. All limbs are submerged at once, and aeration may be initiated to increase circumferential hydrostatic pressure.7,22 Anti-inflammatory and analgesic effects are appreciated for pain-causing conditions of equine limbs.7,23–25 The versatility of this modality and the multiple benefits achieved at once make this a practical treatment option. Between treatments, the water is meant to be recycled back into holding tanks, one maintained as cold as 2 °C and one maintained as hot as 44 °C. Depending on the time between treatments as well as what temperature the unit is set to and/or any disruptions in power to the chiller or heater elements, it is also possible that the holding tanks may reach room temperature of approximately 22 °C between treatments. The plumbing system from the hydrotherapy unit does provide an option to discard the treatment water rather than have it return to a holding tank. This is often time and cost prohibitive as the tank it came from must be completely refilled and all sodium chloride and magnesium sulfate replaced. The goal of this study, therefore, was to take a first step towards identifying the potential risk of bacterial contamination of saltwater within these tanks when horses with limb wounds or skin infections are treated. Contrary to our hypothesis, salinity had no effect on the survival ability of 4 bacterial strains commonly found in equine limb wounds, while higher temperatures did for certain strains.
Bacterial survival is often negatively correlated with salinity through 2 main methods, osmotic and ionic stress.13,26 The concentration of salt to produce these effects is dependent on the bacterial type.12 Although the full salinity evaluated in this study of 20 ppt is less than saltwater found in the ocean, with an average salinity of 35 ppt,14 it has been established that the nutrient content of ocean saltwater is more impactful to bacterial activity than the water salinity itself, as demonstrated in an estuary environment where bacterial activity was increased in the saltwater compared to freshwater.13 Future investigation is required to determine if a higher salinity of saltwater used for an equine hydrotherapy unit could cause detrimental osmotic and ionic effects on the common equine limb bacteria assessed in this study or if higher concentrations of salinity would remain ineffective against these bacterial strains.
All 4 strains examined in this study, the gram-negative E coli and P aeruginosa and the gram-positive S aureus and S zooepidemicus, had minimal to no reduction in bacterial concentrations at 2 °C over the duration of the 96-hour study. Even the minimal reductions in E coli and S zooepidemicus that were statistically significant at this temperature are not clinically relevant at less than a 1 log10 CFU/mL decrease in bacterial concentration. Similarly, at 22 °C, there was no reduction in E coli, P aeruginosa, or S aureus concentrations. Interestingly, survival of S zooepidemicus was markedly impacted at 22 °C, with concentrations steadily decreasing to the lower limit of detection of the quantification method by 96 hours. At 44 °C, survival of all strains except P aeruginosa was markedly impacted, with concentrations of E coli, S aureus, and S zooepidemicus quickly decreasing to the lower limit of detection by 24 to 48 hours and remaining there for the duration of the study. Although the concentrations of P aeruginosa initially decreased over the first 24 hours at 44 °C, they stabilized and remained well above the lower limit of detection at all time points of the study. The clinical impact of these results is not understood at this time, and further attention must be given to hydrotherapy unit protocols such as case inclusion/exclusion criteria and disinfection procedures.
Each bacterium assessed in this study has a unique nutritional requirement, likely accounting for the variability in survival observed. Escherichia coli and P aeruginosa are both gram-negative bacteria; however, they grew to different concentrations under the conditions examined.27,28 Pseudomonas aeruginosa prefers moist environments and has simple growth requirements with excellent nutritional versatility, making Pseudomonas species a particular concern for hydrotherapy environments where it is likely to survive.19,28 Staphylococcus aureus and S zooepidemicus are both gram-positive bacteria, and they also showed variability in survival likely because of their nutritional requirements.17,28 Streptococcus zooepidemicus has a more complex nutritional requirement,28 which may be why it was more easily eliminated at both 22 and 44 °C compared to the other common equine limb bacteria examined.
Several limitations of this study exist, most notably this is an in vitro isolated study based on the salinity and temperature recommendations of one commercially available equine hydrotherapy unit only. Different salinity preparation instructions and recommended holding tank and treatment temperatures may be in place for other units. While our hydrotherapy unit specifically uses a sand filtration system (Triton II Fiberglass Sand Filter; Pentair Pool) between the treatment unit and the holding tanks capable of eliminating larger 20- to 30-μm–sized particles, we did not examine the ability of this filtration system to potentially eliminate or reduce bacterial contamination. It is also possible that other hydrotherapy units may contain a chlorine filtration step, even though our unit does not, and the effect of such a chlorine filtration step was not examined. In addition, there was no investigation in this study into the potential for biofilm formation within the hydrotherapy unit, filtration piping, or holding tanks, a concern that is well documented in recreational water systems and piping.29 Finally, we have only assessed 4 specific bacteria, which although common, are still a small percentage of potential pathogens inhabiting equine limb wounds, cutaneous tissue, and a recycled water environment.
In conclusion, this is the first study to examine the ability of bacteria common to equine limb wounds to survive in saltwater at different salinity concentrations and temperatures relevant to equine hydrotherapy in a clinical setting. Although only an initial step, the results are concerning and warrant further investigation into specific hydrotherapy unit disinfection protocols and the need for wastage of treatment water following patient use, particularly at cooler temperatures and/or when Pseudomonas species are suspected or cultured from a patient. The bacterial contamination burden (concentration) from the cutaneous surface or limb wound into the saltwater with or without aeration is still unknown as is how this could impact bacterial survival within a hydrotherapy unit or holding tank. Similarly, it is unknown how much bacteria would need to survive in the saltwater to potentially infect a subsequent patient. With the information obtained from this investigation, future studies can be designed to help answer these questions and determine the safest protocols for the utilization of this form of immersive hydrotherapy in a treatment and rehabilitation plan.
Acknowledgments
The authors thank Rebecca Melendez and John Shamoun for their input on study design and data interpretation. The authors also thank the North Carolina State University Clinical Microbiology and Molecular Diagnostics laboratory and Melissa West for bacterial strain acquisition.
Disclosures
The authors have nothing to disclose. No AI-assisted technologies were used in the generation of this manuscript.
Funding
This project was generously funded by the Carolina Equine Sports Medicine Education Foundation.
ORCID
Lauren V. Schnabel https://orcid.org/0000-0002-1993-8141
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