Etiological and immunological aspects of equine hoof canker

Abstract

Chronic equine proliferative pododermatitis (canker) is a hypertrophic hoof disease characterized by excessive proliferation of horn-producing tissues resulting from dyskeratosis of keratinocytes. Since its etiology remains unknown, this study investigated potential etiological agents and evaluated biochemical and immunological profiles. Ten horses with hoof canker and ten healthy controls of the Norik of Muran breed were included. Dermatophytes and bovine papillomavirus 1 and 2 (BPV 1/2) were not detected. Candida glabrata was the most frequently identified species in both affected and healthy groups. In horses with canker, Aspergillus fumigatus, Mucor spp., and Trichosporon spp. were also detected. Biochemical analysis showed significantly higher creatine kinase myoglobin binding activity in affected horses compared to controls (12.58 ± 5.86 vs. 4.11 ± 1.57; p < 0.01), with a slight increase in aspartate aminotransferase activity. Principal component analysis revealed substantial overlap between healthy and affected groups. Immunologically, monocyte phagocytic activity was significantly reduced in affected horses (83.78% ± 3.40 vs. 90.26% ± 2.88; p < 0.001). Affected horses had increased CD4⁺ (Th) lymphocyte counts and a higher CD4⁺:CD8⁺ ratio (p < 0.01), along with higher IgM⁺ B lymphocyte levels (27.20% ± 8.23 vs. 18.51% ± 4.53; p < 0.05). Serum IgG levels were also significantly elevated (11.15 ± 3.42 vs. 5.88 ± 1.91 mg/mL; p < 0.001). These results indicate stimulation of adaptive cellular and humoral immune responses in canker-affected horses. However, the lack of consistent pathogen detection suggests that the precise etiology of hoof canker remains unresolved and requires further investigation.

Introduction

Chronic equine proliferative pododermatitis, also known as equine hoof canker, is a chronic infectious and proliferative disease characterized by hypertrophy of the horn-producing epidermal tissues. It occurs in horses of all breeds, regardless of age or sex, although a higher prevalence has been observed in cold-blooded breeds. This disease poses a serious threat to the usability and welfare of affected horses (Dietz 2006; Reinthaler et al. 2025). Clinically, hoof canker is a debilitating disorder that may affect one or more hooves. It most frequently originates in the frog and can extend to the adjacent sole, collateral sulci, hoof wall, and coronary band. Hoof canker is generally confined to the epidermal tissue, with proliferation originating there. In advanced stages, it can invade the underlying dermal structures (O’Grady, 2018). Typical clinical manifestations include cauliflower-like or filamentous formations in the frog area and collateral sulci, often accompanied by a characteristic foul odor and bleeding (Baxter and Stashak 2011). Therapeutic approaches are based on a multimodal strategy that combines intravenous administration of antibiotics and corticosteroids, local application of antiseptics, radical excision of altered tissues, and a pressure bandage (O’Grady and Madison, 2004).

The etiology of hoof canker remains poorly understood and is considered multifactorial. Several infectious agents have been detected in affected tissue, including anaerobic bacteria such as Fusobacterium necrophorum and Bacteroides spp., as well as spirochetes of the genus Treponema (Redding and O’Grady 2012; Sykora and Brandt 2015; Marčeková et al. 2021). Viral involvement, particularly bovine papillomavirus types 1 and 2 (BPV-1/2), has also been investigated (Apprich et al. 2020). Environmental and management factors such as wet or unhygienic conditions, poor hoof care, and prolonged exposure to moisture have been identified as predisposing factors. However, canker is also frequently observed in horses kept under optimal conditions with regularly trimmed hooves (Dietz 2006; Redding and O’Grady, 2012). Some authors have suggested that an (auto) immune mechanism or genetic predisposition may play a role in the pathogenesis of the disease (Jongbloets et al. 2005; Tregnago Azzolini et al. 2019). Furthermore, hoof canker has been reported in association with coronary band dystrophy, a condition classified as an autoimmune disease (Twitchell et al. 2014).

Given that the etiology of hoof canker is complex and multifactorial, with limited information available on both infectious agents and the immunological responses of affected horses, the aim of this study was to investigate potential etiological agents and analyze selected immunological parameters in the blood of healthy horses and those diagnosed with hoof canker.

Materials and methods

Animal subjects

The study population consisted of Norik of Muran horses from a stud farm in eastern Slovakia, all maintained under the same management conditions. A total of 20 horses aged 5–17 years, including 10 healthy mares (5.50 ± 0.53 years) and 10 mares diagnosed with hoof canker (13.00 ± 3.27 years), were included in the study (Table 1). Before sample collection, all horses underwent a comprehensive clinical examination, including a thorough inspection of all four hooves. The healthy control mares showed no clinical signs of hoof canker or other hoof abnormalities and had no previous history of the disease. All animals were otherwise clinically healthy and exhibited no signs of other diseases. The horses were routinely dewormed and vaccinated according to standard protocols, and no systemic or topical medication was applied before sample collection.

Table 1.

General information about the horses involved in the study

Horse	Age (years)	Diagnosis	Horse	Age (years)	Diagnosis	Affected leg
1.	5	Healthy	11.	17	Canker	RF frog
2.	6	Healthy	12.	13	Canker	RF frog and hoof wall
3.	6	Healthy	13.	12	Canker	LF frog
4.	5	Healthy	14.	9	Canker	RF frog
5.	6	Healthy	15.	17	Canker	LH frog and lateral heel
6.	5	Healthy	16.	13	Canker	LF hoof wall
7.	6	Healthy	17.	12	Canker	RF frog, LH frog and lateral heel
8.	5	Healthy	18.	7	Canker	LH frog
9.	5	Healthy	19.	16	Canker	RF frog
10.	6	Healthy	20.	14	Canker	LH frog and lateral heel

RF right foreleg, LF left foreleg, LH left hindleg

Hoof biopsies

Hoof biopsies (two to four samples per hoof) were collected from affected horses (Fig. 1) during debridement for histopathological examination. Biopsies were obtained from the frog and adjacent collateral sulci, primarily at the margins of active lesions. Tissue samples were collected as excisional biopsies using a sterile scalpel, with an approximate size of 5–10 mm, including both epidermal and superficial dermal layers. All samples were collected prior to the initiation of any antimicrobial or anti-inflammatory treatment. Procedures were performed with the horse standing under regional anesthesia (abaxial sesamoid nerve block) using mepivacaine (Mepivastesin 3% 30 mg/mL, 3 M ESPE, Neuss, Germany). A tourniquet was applied to a lower limb to prevent excessive bleeding. Immediately after collection, biopsy samples intended for histopathological examination were fixed in 10% neutral buffered formalin and submitted for analysis. In addition, two biopsy swabs were taken from each hoof: one for general mycological examination and one for dermatophyte detection. As controls, hoof samples and swabs were collected from healthy horses during regular trimming. One biopsy sample and two swab samples (for mycological examination and dermatophyte detection) were collected from each control hoof from anatomically comparable regions using the same technique as in affected horses. Histopathological examination of hoof biopsies was performed by IDEXX Laboratories (Leipzig, Germany).

Fig. 1.

Chronic proliferative pododermatitis of the equine hoof; the scalpel indicates one of the sampling sites

Detection of pathogens

Swabs for mycological and dermatophyte examination were taken from the hoof biopsies and placed in Amies agar gel without charcoal (Copan Italia, Brescia, Italy). Samples were transported to the laboratory at 4 °C and processed within 24 h. Mycological examination and cultivation for dermatophytes on Sabouraud dextrose agar were carried out by the State Veterinary and Food Institute (Košice, Slovak Republic).

To investigate the presence of BPV-1/2, hoof tissue and hair with roots were collected from each horse from both affected and healthy areas. The samples were sent to Laboklin s.r.o. (Bratislava, Slovak Republic) for analysis.

Therapy

In horses with hoof canker, thorough debridement (surgical removal of affected tissue) was performed under regional anesthesia (abaxial sesamoid nerve block) while the horse was standing. All procedures were performed by the same clinician. This was followed by topical application of metronidazole powder (Metrozol, Metronidazolum 200 mg/g, PharmaGal s.r.o., Nitra, Slovak Republic) and a hoof bandage, which was changed daily for the first 10 days, then every other day, and continued until the disease resolved (defined as the absence of proliferative lesions and restoration of normal horn tissue). In eight horses, the treatment lasted approximately three to six months, with no recurrence observed. In the remaining two horses, the treatment was unsuccessful, and complete resolution of the disease was not achieved.

Blood collection

Blood samples were collected from two groups of horses: clinically healthy individuals (n = 10) and affected horses one month after starting treatment (n = 10). All mares in the study were housed in a loose housing system. The horses were routinely fed twice daily with oats and had free access to water and hay (ad libitum). Sampling was performed in the morning between 7:00 and 9:00 a.m., before feeding. Blood samples were taken from the jugular vein into different types of collection tubes according to the intended analysis: for biochemical analysis, blood was collected in serum tubes (without anticoagulant); for hematological analysis, in tubes containing K₃EDTA (Greiner Bio-One Vacuette, Kremsmünster, Austria); and for immunological analysis, in Eppendorf tubes containing heparin (8–10 U/mL).

Biochemical and hematological analyses

During transport, samples were stored at 8–10 °C and analyzed within six hours after collection due to the long transport distance. After clotting, samples intended for biochemical analysis were centrifuged for 10 min at 3500 rpm. The mineral profile (calcium, phosphorus, magnesium, copper, and zinc) was determined using an Ellipse analyzer (AMS, Rome, Italy) and Dialab commercial kits (Dialab GmbH, Wiener Neudorf, Austria). Hematological analysis was performed using a BC-2008 VET automatic analyzer (Mindray, Shenzhen, China). Results are expressed in appropriate units as mean ± standard deviation (SD).

Immunological analyses

Immunological analyses were performed on the horses’ heparinized blood samples. As part of these examinations, phagocytic activity (PA) was monitored, specifically the activity of neutrophils, monocytes, and the total PA, using a commercial kit (Phagotest, Celonic, Germany) designed for flow cytometry analysis. In addition, the proportions of selected lymphocyte subpopulations were evaluated. Immune cells were identified using fluorescently labeled monoclonal antibodies and subsequently analyzed on a BD FACSCanto flow cytometer (Becton Dickinson Biosciences, USA). The following subpopulations were selected to assess the cellular immune response: Th (helper) lymphocytes characterized by CD4⁺CD8⁻; Tc (cytotoxic) lymphocytes characterized by CD4⁻CD8⁺; double-positive T lymphocytes characterized by CD4⁺CD8⁺; total T lymphocytes characterized by CD2⁺; and subpopulation of B lymphocytes characterized by IgM⁺. The ratio of CD4⁺:CD8⁺ lymphocytes was used to assess the level of immunostimulation. For the identification of CD4 + and CD8 + lymphocytes was used direct immunostaining protocol, where 25 µL of heparinized blood was mixed with 2 µL of anti-CD4 monoclonal antibody (MoAb) conjugated with FITC (clone: DWT5; CD Creative Diagnostics, NY, USA) and 1 µL of anti-CD8a MoAb conjugated with R-PE (clone: DWT9; CD Creative Diagnostics, NY, USA). The mixture was incubated for 20 min at 25 °C in the dark. Subsequently, erythrocytes were lysed by adding 1 mL of BD FACS Lysing Solution (BD Biosciences, USA) and incubating for an additional 20 min under the same conditions. Samples were then centrifuged at 300 × g for 5 min, and the resulting cell pellets were washed twice with 1 mL phosphate-buffered saline (PBS; MP Biomedicals, France), with centrifugation at 300 × g for 5 min after each wash. Following the final washing step, cells were resuspended in 100 µL PBS and analyzed by flow cytometry.

Indirect immunofluorescence staining was used for the identification of CD2⁺ and IgM⁺ lymphocytes. For CD2⁺ lymphocyte detection, 50 µL of heparinized blood was incubated with 10 µL of anti-CD2 MoAb (clone: HB88A; Monoclonal Antibody Center, Washington State University, USA) under the same conditions as described for direct staining. Subsequently, erythrocytes were lysed using 1 mL of ammonium chloride-based lysis buffer (8.29 g NH₄Cl, 1.0 g KHCO₃, and 0.393 g Na₂EDTA per liter) during a 20 min incubation at 25 °C. Samples were then washed with 1 mL PBS and centrifuged at 300 × g for 5 min. For the identification of IgM⁺ lymphocytes, erythrocytes were first removed using the same ammonium chloride-based lysis procedure. Following erythrocyte lysis, leukocytes were incubated with the primary anti-IgM MoAb (clone: 1.9/3.2; Monoclonal Antibody Center, Washington State University, USA) as described above. Subsequently, 50 µL of secondary antibody conjugated with FITC (clone: M1-14D12; eBioscience, San Diego, USA) diluted 1:100 in PBS was added. After two washing steps with PBS, the samples were diluted in 200 µL PBS and analyzed by flow cytometry. The proportions of lymphocyte populations are expressed as percentages ± SD. Serum IgG concentrations were determined using the Horse IgG ELISA Kit (ab190531, Abcam, UK).

Statistical analysis

The data were analyzed using GraphPad Prism version 11.0.0 (GraphPad Software, San Diego, CA, USA). Differences between groups were assessed with an unpaired t-test with Welch’s correction, considering p < 0.05 as significant. Principal component analysis (PCA) was performed on the biochemical parameters after data standardization (z-score normalization) to explore data patterns and assess potential clustering of samples by disease status. Correlation analysis of hematological and immunological parameters was conducted using Pearson’s correlation coefficient.

Results

Detection of pathogens

In affected horses, mycological analysis showed that Candida glabrata was the most frequently detected species (6/10), followed by Trichosporon spp. (3/10). Aspergillus fumigatus and Mucor spp. were each identified in 2 of 10 animals. The remaining detected species – Aspergillus terreus, Candida parapsilosis, Penicillium spp., and Rhodotorula rubra – were each found in a single horse (1/10). In healthy hooves, Candida glabrata was also the most frequently detected species (6/10), followed by Candida albicans and Penicillium spp. (4/10 each), while Candida krusei was detected in only one horse (1/10). Dermatophyte cultures were negative in all samples. All samples tested for BPV-1/2 were also negative.

Hematological analyses

Statistical analysis of hematological parameters showed an increased red blood cell count and mean corpuscular hemoglobin concentration (MCHC), and a decreased mean corpuscular volume in affected horses compared to healthy horses (Table 2). However, values in both groups remained within reference values, except for MCHC, which was above the reference values in both groups.

Table 2.

Hematological profiles of horses affected by hoof canker and healthy controls

Parameter	WBC	LYM	MONO	GRAN	RBC	HGB
Units	G/L	G/L	G/L	G/L	T/L	g/L
Healthy	7.48 ± 1.03	1.14 ± 0.42	0.44 ± 0.08	5.91 ± 1.18	6.47 ± 0.57	113.90 ± 8.05
Affected	7.62 ± 0.97	2.19 ± 1.50	0.54 ± 0.24	4.88 ± 1.05	7.26 ± 0.87	130.95 ± 26.31
T-test	ns	ns	ns	ns	p < 0.05	ns
Values*	3.0–12.2	1.5–5.0	0–0.62	2.6–7.2	5.0–9.5	93–160
Parameter	HCT	MCV	MCH	MCHC	PLT
Units	L/L	fL	pg	g/L	G/L
Healthy	0.32 ± 0.02	50.26 ± 2.25	17.68 ± 0.74	352.70 ± 4.60	139.85 ± 34.63
Affected	0.32 ± 0.05	44.03 ± 7.22	17.14 ± 1.51	394.58 ± 31.32	140.19 ± 33.70
T-test	ns	p < 0.05	ns	p < 0.01	ns
Values*	0.24–0.44	36.6–67.6	12.2–22.5	276–342	100–250

WBC white blood cell count, LYM lymphocyte count, MONO monocyte count, GRAN granulocyte count, RBC red blood cell count, HGB hemoglobin, HCT hematocrit, MCV mean corpuscular volume, MCH mean corpuscular hemoglobin, MCHC mean corpuscular hemoglobin concentration, PLT platelet count. Data are presented as mean ± standard deviation. Unpaired t-test with Welch’s correction was used for statistical analysis: ns not significant; statistically significant differences were considered at p < 0.05. Values* – reference values for Norik of Muran horses or cold-blooded horses (Pađen et al., 2014; Prvanović Babić et al., 2019; Witkowska-Piłaszewicz et al., 2021; Czech et al., 2025)

Biochemical analyses

Analysis of biochemical parameters revealed several changes in the evaluated parameters. AST activity (µkat/L) was slightly above the physiological reference range in horses with canker (p < 0.01), while CK-MB activity (µkat/L) was markedly elevated in the canker-affected group (p < 0.01). Cholesterol concentration (mmol/L) was lower in both groups of horses. In contrast, TG (mmol/L), LDL (mmol/L), and Mg (mmol/L) concentrations were above the physiological reference range in both groups. Although HDL (mmol/L) and zinc (µmol/L) differed significantly between groups (p < 0.001 and p < 0.01, respectively), their values remained within the physiological reference range (Table 3).

Table 3.

Biochemical profiles of horses affected by hoof canker and healthy controls

Parameter	TP	Urea	Albumin	Creatinine	Glucose	TG
Units	g/L	mmol/L	g/L	µmol/L	mmol/L	mmol/L
Healthy	67.46 ± 11.31	5.16 ± 1.14	29.58 ± 4.00	153.40 ± 32.48	5.25 ± 0.60	0.82 ± 0.14
Affected	75.46 ± 11.36	3.45 ± 2.27	30.43 ± 5.60	152.40 ± 23.34	4.99 ± 0.87	0.80 ± 0.50
T-test	ns	ns	ns	ns	ns	ns
Values*	55–75	3.5–8.0	25–38	71–156	4.2–6.4	< 0.65
Parameter	Cholesterol	HDL	LDL	AST	ALT	GGT
Units	mmol/L	mmol/L	mmol/L	µkat/L	µkat/L	µkat/L
Healthy	1.64 ± 0.46	0.95 ± 0.25	0.75 ± 0.20	4.35 ± 1.18	0.07 ± 0.05	0.29 ± 0.08
Affected	1.75 ± 0.39	1.41 ± 0.15	0.88 ± 0.28	5.72 ± 0.80	0.10 ± 0.06	0.36 ± 0.27
T-test	ns	p < 0.001	ns	p < 0.01	ns	ns
Values*	1.9–2.7	0.8–1.6	0.27–0.42	2.5–5.5	0.06–0.35	0.10–0.35
Parameter	ALP	Bilirubin	CK-MB	LDH	GLDH	α-amylase
Units	µkat/L	µmol/L	µkat/L	µkat/L	µkat/L	µkat/L
Healthy	2.04 ± 0.58	13.78 ± 4.05	4.11 ± 1.57	5.48 ± 2.90	0.07 ± 0.03	0.17 ± 0.06
Affected	2.41 ± 0.82	17.10 ± 6.30	12.58 ± 5.86	6.23 ± 3.23	0.09 ± 0.03	0.21 ± 0.14
T-test	ns	ns	p < 0.01	ns	ns	ns
Values*	0.4–4.0	1.7–39.0	1.0–5.0	1.3–6.8	< 0.136	0.2–0.3
Parameter	Ca	P	Mg	Cu	Zn	Cl
Units	mmol/L	mmol/L	mmol/L	µmol/L	µmol/L	mmol/L
Healthy	2.22 ± 0.29	1.42 ± 0.32	1.28 ± 0.30	18.24 ± 2.34	17.01 ± 3.70	106.40 ± 9.70
Affected	2.34 ± 0.20	1.09 ± 0.42	1.05 ± 0.23	19.72 ± 3.61	11.71 ± 2.30	101.51 ± 13.74
T-test	ns	ns	ns	ns	p < 0.01	ns
Values*	2.2–3.3	0.7–1.5	0.5–0.9	18.8–21.2	9.2–19.9	95–109

TP total protein, TG triacylglycerols, HDL high-density lipoprotein cholesterol, LDL low-density lipoprotein cholesterol, AST aspartate aminotransferase, ALT alanine aminotransferase, GGT gamma-glutamyl transferase, ALP alkaline phosphatase, CK-MB creatine kinase myoglobin binding, LDH lactate dehydrogenase, GLDH glutamate dehydrogenase, Ca calcium, P phosphorus, Mg magnesium, Cu copper, Zn zinc, Cl chloride. Data are presented as mean ± standard deviation. Unpaired t-test with Welch’s correction was used for statistical analysis: ns not significant; statistically significant differences were considered at p < 0.05. Values* – reference values (Pađen et al., 2014; Burlikowska et al., 2015; Maksymovych et al., 2015; Pošivák et al., 2015)

PCA of biochemical parameters showed only partial separation between healthy and canker-affected groups, primarily along the first principal component (PC1). The pronounced overlap between clusters, together with the relatively low variance explained by the first two components (45.59%), indicates that group differences are present but not fully captured in the two-dimensional projection (Fig. 2).

Fig. 2.

Principal component analysis of biochemical parameters. PC1 principal component 1, PC2 principal component 2. Points represent horses with hoof canker (red) and healthy controls (blue)

Immunological analyses

To evaluate the cellular immune response in horses affected by hoof canker, PA and lymphocyte subpopulations in peripheral blood were analyzed and compared with healthy controls. When comparing PA, no significant differences were observed in total PA or in the PA of neutrophils. However, the PA of monocytes was significantly lower in horses affected by hoof canker than in healthy horses (Table 4).

Table 4.

Phagocytic activity in the peripheral blood of horses affected by hoof canker and healthy controls

Parameter	PA total	PA Neu	PA Mo
Healthy	94.00 ± 1.58	95.06 ± 1.24	90.26 ± 2.88
Affected	92.19 ± 2.54	93.83 ± 2.64	83.78 ± 3.40
T-test	ns	ns	p < 0.001

PA total total phagocytic activity, PA Neu phagocytic activity of neutrophils, PA Mo phagocytic activity of monocytes. Data are presented as mean ± standard deviation. Unpaired t-test with Welch’s correction was used for statistical analysis: ns not significant; statistically significant differences were considered at p < 0.05

Analysis of lymphocyte subpopulations revealed that affected horses had a significantly higher proportion of CD4⁺ (Th) lymphocytes, resulting in a higher CD4⁺:CD8⁺ lymphocyte ratio, which indicates immune stimulation in affected horses. Similarly, the proportion of IgM⁺ (B) lymphocytes was higher in affected horses (Table 5). The serum IgG level was significantly higher in horses with hoof canker (11.15 ± 3.42 mg/mL) than in healthy control horses (5.88 ± 1.91 mg/mL; p < 0.001), indicating the presence of long-term inflammation.

Table 5.

Lymphocyte subpopulations and CD4+:CD8 + ratio in the peripheral blood of horses affected by hoof canker and healthy controls

Parameter	CD4+	CD8+	CD4+CD8+	CD2+	IgM+	CD4:CD8
Healthy	32.37 ± 4.21	11.75 ± 1.57	0.52 ± 0.15	45.55 ± 2.43	18.51 ± 4.53	2.79 ± 0.50
Affected	43.18 ± 7.56	13.95 ± 3.07	0.82 ± 0.46	48.99 ± 7.97	27.20 ± 8.23	3.47 ± 0.38
T-test	p < 0.01	ns	ns	ns	p < 0.05	p < 0.01

CD4⁺ Th lymphocytes, CD8⁺ Tc lymphocytes, CD4⁺CD8⁺ double-positive T lymphocytes, CD2⁺ total T lymphocytes, IgM⁺ subpopulation of B lymphocytes, CD4:CD8 ratio of CD4⁺ to CD8⁺ lymphocytes. Data are presented as mean ± standard deviation. Unpaired t-test with Welch’s correction was used for statistical analysis: ns not significant; statistically significant differences were considered at p < 0.05

Correlation analysis

Correlation analysis showed that neutrophil PA was significantly positively correlated with total PA. In contrast, monocyte PA was negatively correlated with lymphocyte and monocyte counts, as well as with CD4⁺ and CD8⁺ cell counts and IgG levels, and positively correlated with granulocyte counts. CD4⁺ lymphocytes were positively correlated with lymphocyte counts and CD2⁺ cells. CD8⁺ cells were positively correlated with CD4⁺CD8⁺ double-positive lymphocytes, which were, in turn, positively associated with IgM⁺ cells. Additionally, IgM⁺ lymphocytes were positively correlated with serum IgG levels (Fig. 3).

Fig. 3.

Correlation matrix of hematological and immunological parameters. WBC white blood cell count, LYM lymphocyte count, MONO monocyte count, GRAN granulocyte count, RBC red blood cell count, HGB hemoglobin, HCT hematocrit, PLT platelet count, PA total total phagocytic activity, PA Neu phagocytic activity of neutrophils, PA Mo phagocytic activity of monocytes, CD4⁺ Th lymphocytes, CD8⁺ Tc lymphocytes, CD4⁺CD8⁺ double-positive T lymphocytes, CD2⁺ total T lymphocytes, IgM⁺ subpopulation of B lymphocytes, IgG immunoglobulin G. Strong positive correlations are shown in teal, while negative correlations are represented in brown. The analysis was conducted using Pearson’s correlation coefficient and statistical significance was set at p < 0.05

Discussion

Given that the etiology of equine hoof canker remains unknown and no standardized therapeutic protocol has yet been established, treatment is often time-consuming and frustrating (Reinthaler et al. 2025). Various systemic and topical treatments have been attempted, with inconsistent results. Surgical debridement combined with appropriate hoof care is considered essential (Oosterlinck et al. 2011). Topical cisplatin chemotherapy has also been used (Apprich and Licka 2013), although recurrences were common. Treatment typically involves debridement, application of a hoof bandage to maintain pressure on the healing solar wound and keep the area clean, and the use of topical antiseptic solutions or antimicrobial agents such as metronidazole (Dabareiner et al. 2011). In the present study, thorough debridement, topical application of metronidazole, and hoof bandaging were performed. Another therapeutic approach combines extensive surgical debridement with cryotherapy, followed by topical application of 10% benzoyl peroxide in acetone and crushed metronidazole tablets, which has yielded consistent and predictable results in all treated horses (56 cases). Emphasis should be placed on keeping the surgical wound clean and dry until the defect begins to cornify (O’Grady and Madison, 2004). Hoof canker predominantly affects older horses, in which a higher incidence of autoimmune disorders has been reported. Some authors (Jongbloets et al. 2005; Oosterlinck 2018; Azzolini et al. 2019) suggest that these disorders may contribute to the development of the disease. Additional oral prednisolone treatment has been shown to significantly reduce hospitalization duration and is now considered part of the therapeutic protocol. In contrast, horses receiving inappropriate preceding treatments were significantly more likely to relapse, highlighting the need for prompt diagnosis and appropriate subsequent treatment (Oosterlinck 2018).

Metronidazole is an antimicrobial agent effective against anaerobic bacteria, including species such as F. necrophorum and spirochetes of the genus Treponema, which are commonly associated with necrotic and poorly oxygenated tissues. However, in our study, F. necrophorum was not detected in hoof canker biopsies or in healthy hooves (unpublished results). On the other hand, our previous research demonstrated that T. pedis was found in horses with hoof canker as well as in one clinically unaffected horse, without causing clinical signs. Additionally, 70% of 10 horses affected with canker had hoof biopsies positive for T. pedis, 20% for T. denticola and 20% for T. brennaborense (Marčeková et al. 2021).

In addition to anaerobic bacteria, other microorganisms such as fungi and viruses can also contribute to the development of hoof canker (Apprich et al. 2017). In the present study, all samples of hoof tissue and hair with roots collected from both affected and healthy areas were negative for BPV-1/2, which is consistent with our previous investigations in other stud farms (Marčeková et al. 2021). In contrast, Brandt et al. (2011a) detected BPV-1/2 DNA by PCR in all 24 canker-affected tissues and in 12 of 13 clinically normal skin samples, suggesting a possible association with the disease process. Apprich et al. (2020) detected BPV-1/2 in 5 of 18 horses affected by canker and in 2 of 6 control horses, but found no association between the presence of BPV-1/2 and the degree of koilocyte-like keratinocytes or other morphological changes in hoof canker tissue. This suggests that virus-induced pathogenesis is only one of several possible causal pathways for the disease.

Keller et al. (2000) isolated several species of molds and dermatophytes from equine hooves, belonging to the genera Microsporum and Trichophyton. However, no dermatophytes were detected in the present study, which is consistent with our previous findings (Marčeková et al. 2021). The results of the mycological analysis are again in agreement with our previous study, in which Candida glabrata and Trichosporon spp. were among the most frequently isolated yeasts. In the present study, Candida glabrata was also the most frequently detected yeast species in clinically healthy hooves. Candida spp., including less common species such as Candida glabrata, are considered opportunistic yeasts that may colonize equine mucosal surfaces and can cause infection under conditions of dysbiosis or immunosuppression (Neira-Egea et al. 2025).

PCA revealed moderate group differentiation; however, substantial overlap between healthy and canker-affected horses indicates that separation based on biochemical parameters is not well defined. Biochemical analysis in horses with hoof canker showed mildly increased AST activity and markedly elevated CK-MB levels. Muscle enzyme activity in horses, particularly creatine kinase (CK) and AST, is commonly used to diagnose muscle damage (Giers et al. 2023; Connysson and Jansson 2025), while CK-MB is traditionally considered a marker of myocardial injury. However, in equine medicine, the diagnostic specificity of CK-MB is limited. Unlike in humans, CK-MB in horses is not exclusively of cardiac origin (Argiroudis et al. 1982; Hohnadel 2017) and may also reflect skeletal muscle metabolism (Kurapati and Zubair 2026). Based on these considerations, we propose that the elevated CK-MB and AST levels are likely associated with altered biomechanics of movement, lameness, and compensatory muscle activity induced by hoof canker.

To the authors’ knowledge, no studies have systematically evaluated immunological parameters in horses with hoof canker. Previous research has focused primarily on histopathological and microbiological characteristics of the disease (Moe et al. 2010; Brandt et al. 2011b; Apprich et al. 2017). Our findings suggest the presence of persistent inflammation in the bodies of horses with hoof canker, characterized by elevated IgG levels and an increased proportion of both B lymphocytes and T helper lymphocytes. Age-related changes in immune cell populations should also be considered when interpreting these findings. In horses older than 20 years, total lymphocyte, CD4⁺, CD8⁺, T-regulatory, and B cell counts have been reported to decrease, whereas the CD4⁺:CD8⁺ ratio increases with age (DeNotta and McFarlane 2023). In contrast, serum concentrations of the immunoglobulin subtypes IgG, IgG(T), IgM, and IgA did not differ with age (McFarlane et al. 2001). In the present study, the affected horses were older than the healthy controls but all were younger than 20 years, suggesting that the increased CD4⁺:CD8⁺ ratio observed in affected horses is unlikely to be attributable to advanced age and may instead reflect immune activation associated with hoof canker. As all affected horses were receiving treatment, the observed immunostimulation may indicate that the immune system remained actively engaged in the response to the disease.

Given the chronic nature of hoof canker, rather than an acute inflammatory response, neutrophil PA is no longer reduced, whereas monocyte PA is decreased. Reduced monocyte PA is commonly reported in conditions such as sepsis, trauma, chronic disease, and advanced cancer (Liu et al. 1999; Arsenijevic et al. 2001; Vollrath et al. 2021; Gaire et al. 2024). In horses, decreased monocyte PA may also be associated with aging. Previous studies have shown that while healthy aged horses do not exhibit differences in absolute monocyte counts, geriatric horses demonstrate reduced monocyte phagocytic capacity (Herbst et al. 2022; DeNotta and McFarlane 2023). As the horses with hoof canker in this study were older than the healthy controls, it cannot be determined whether the observed decrease in monocyte PA is attributable to the disease itself, age-related immune changes, or a combination of both.

Correlation analysis, along with differences in immunological parameters between healthy and affected horses, suggests a shift in immune balance associated with the disease. Negative correlations between monocyte PA and immune cell counts (including lymphocytes and monocytes), as well as IgG levels, may reflect a compensatory mechanism between innate and adaptive immunity. Similar interactions between impaired innate immune function and compensatory lymphocyte activation have been described in horses and other species during inflammatory or infectious conditions (Zhao et al. 2009; Holmes and Wagner 2024; Wang et al. 2024).

The observed increase in CD4⁺ lymphocytes and the CD4:CD8 ratio in affected horses, along with positive correlations among CD4⁺, CD8⁺, and CD2⁺ cells, may indicate a shift toward T-cell–mediated immune responses. Expansion and activation of both CD4⁺ and CD8⁺ T-cell populations have been documented in horses during viral infections (Holmes and Wagner 2024). Proliferation of antigen-specific CD8⁺ lymphocytes has also been demonstrated in horses following immunization (Pretorius et al. 2012). Additionally, lesions in horses with insect bite hypersensitivity contain significantly higher numbers of CD4⁺ cells compared to the skin of healthy controls (Heimann et al. 2011). Together, these findings support the central role of T lymphocytes in adaptive immune responses in horses.

The association of CD8⁺ cells with double-positive CD4⁺CD8⁺ lymphocytes and their relationship to IgM⁺ cells further indicates ongoing immune activation and possible lymphocyte differentiation. CD4⁺CD8⁺ double-positive cells have attracted interest because of their involvement in immune disorders, inflammation, and cancer. Their higher prevalence in advanced cancers, along with strong lytic activity and a distinct cytokine profile, suggests these cells may play a key role in modulating immune responses to cancer (Alam et al. 2025). There is also considerable evidence for an increased frequency of peripheral CD4⁺CD8⁺ T cells during viral infections (Nascimbeni et al. 2004). Increased levels of IgM⁺ lymphocytes, along with their positive correlation with serum IgG levels, suggest B-cell activation and stimulation of the humoral immune response. Since the production of IgG antibodies by B cells is a crucial component of adaptive immunity, this process provides specific and long-lasting protection following infections or vaccinations (Rival et al. 2024).

Conclusion

The lack of consistent detection of specific pathogens across samples, along with the presence of opportunistic microorganisms in both affected and healthy hooves, supports the idea that equine hoof canker is a multifactorial disease rather than a condition driven by a single etiological agent. Principal component analysis based on biochemical parameters showed substantial overlap between healthy and canker-affected horses, indicating that differentiation based solely on these parameters remains unclear. These biochemical changes likely reflect secondary effects of lameness and altered biomechanics rather than primary disease drivers. In contrast, immunological findings indicate a shift in immune balance, characterized by activation of adaptive immunity and a reduction in innate immune function. Increased proportions of CD4⁺ lymphocytes, a higher CD4⁺:CD8⁺ ratio, elevated counts of IgM⁺ cells, and increased IgG levels, together with observed associations among lymphocyte subpopulations, further support ongoing immune stimulation.

Overall, these findings highlight the importance of both local and systemic factors in the pathogenesis of hoof canker. Future research, especially longitudinal studies focusing on immune dynamics and host–microbiome interactions, will be essential to further clarify disease mechanisms and optimize treatment approaches.

Author contributions

E.S.: Writing – original draft, review & editing, Conceptualization, Methodology, Data curation, Funding acquisition. D.M.: Methodology, Data curation, Writing – review & editing. S.G.: Methodology, Writing – review & editing. Z.Ž.: Methodology, Writing – review & editing. J.K.: Writing – original draft, Visualization, Software, Data curation. I.V.: Methodology, Writing – review & editing.

Funding

This publication was supported by the Scientific Grant Agency VEGA, grant number 1/0108/26.

Data availability

The data supporting the findings of this study are available from the corresponding author on reasonable request.

Declarations

Ethics approval

This study was approved by the Ethics Committee of the University of Veterinary Medicine and Pharmacy in Košice, Slovak Republic (permit number EKVP/2022-14). The study was conducted in accordance with local legislation and institutional requirements.

Competing interests

The authors declare no competing interests.