• View in gallery

    Photograph of a hoof wall specimen obtained from the dorsum of a hoof (A) and NMR microscopic images of the same specimen acquired before (B) and after (C) water absorption. Notice the color scale for the signal intensity.

  • View in gallery

    A series of NMR microscopic images of the hoof wall specimen in Figure 1 obtained from the dorsum of the hoof and allowed to absorb water for up to 24 hours. Representative images are provided, with intervals of 1.25 mm in the direction of water permeation (water was placed in a tube in contact with the bottom of each specimen and was absorbed toward the upper region of each specimen). Signal intensity is indicated by a color scale. See Figure 1 for remainder of key.

  • View in gallery

    Number of high local maxima, in relation to the distance from the water surface in contact with each specimen and the duration of water absorption, for specimens obtained from the dorsum (A and B), lateral quarter (C), and lateral heel (D) of a healthy equine hoof. Maxima were determined before (time 0 [white square and dashed line]) and 0.3 (white triangle and dashed line), 0.6 (white circle and dashed line), 0.9 (black square and solid line), 2 (black triangle and solid line), 4 (black circle and solid line), 6 (white square and solid line), 8 (white triangle and solid line), and 24 (white circle and solid line) hours after initiation of water absorption. On the x-axis, 1 U of height is approximately 0.31 mm; height = 0 is the bottom of each specimen (ie, the location nearest to the water surface in contact with the specimen). AU = Arbitrary units.

  • View in gallery

    Relationships between local maximal and minimal signal intensities in each segment of equine hoof at various times after initiation of water absorption in the same specimen as in Figure 1. The range of the axis for maximal and minimal signals in the plots is 0 to 100,000 and 0 to 80,000, respectively. AU = Arbitrary units.

  • 1. Bertram JE, Gosline JM. Functional design of horse hoof keratin: the modulation of mechanical properties through hydration effects. J Exp Biol 1987; 130: 121136.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 2. Hinterhofer C, Stanek C, Binder K. Elastic modulus of equine hoof horn, tested in wall samples, sole samples and frog samples at varying levels of moisture. Berl Munch Tierarztl Wochenschr 1998; 111: 217221.

    • Search Google Scholar
    • Export Citation
  • 3. Wagner IP, Hood DM, Hogan HA. Comparison of bending modulus and yield strength between outer stratum medium and stratum medium zona alba in equine hooves. Am J Vet Res 2001; 62: 745751.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 4. Borderas TF, Pawluczuk B & de Passille AM et alClaw hardness of dairy cows: relationship to water content and claw lesions. J Dairy Sci 2004; 87: 20852093.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 5. Kasapi MA, Gosline JM. Exploring the possible functions of equine hoof wall tubules. Equine Vet J Suppl 1998;(26):1014.

  • 6. Higuchi H, Kurumado H & Mori M et alEffects of ammonia and hydrogen sulfide on physical and biochemical properties of the claw horn of Holstein cows. Can J Vet Res 2009; 73: 1520.

    • Search Google Scholar
    • Export Citation
  • 7. Eccles CD, Callaghan PT. High-resolution imaging. The NMR microscope. J Magn Reson 1986; 68: 393398.

  • 8. Neeman M, Sillerud LO. NMR microscopy. In: Gillies RJ, ed. NMR in physiology and biomedicine. San Diego: Academic Press Inc, 1994;101118.

    • Search Google Scholar
    • Export Citation
  • 9. Murray RC, Dyson SJ & Schramme MC et alMagnetic resonance imaging of the equine digit with chronic laminitis. Vet Radiol Ultrasound 2003; 44: 609617.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 10. Keller MD, Galloway GJ, Pollitt CC. Magnetic resonance microscopy of the equine hoof wall: a study of resolution and potential. Equine Vet J 2006; 38: 461466.

    • Search Google Scholar
    • Export Citation
  • 11. Pollitt CC. Anatomy and physiology of the inner hoof wall. Clin Tech Equine Pract 2004; 3: 321.

  • 12. Butler D. The principles of horseshoeing. 2nd ed. Maryville, Mo: D. Butler Publisher, 1985.

  • 13. Kempson SA, Campbell EH. A permeability barrier in the dorsal wall of the equine hoof capsule. Equine Vet J Suppl 1998;(26):1521.

  • 14. Rasband WS. ImageJ. Bethesda, Md: US National Institutes of Health, 2009.

  • 15. Wertz PW, Downing DT. Cholesteryl sulfate: the major polar lipid of horse hoof. J Lipid Res 1984; 25: 13201323.

  • 16. Wertz PW, Kremer M, Squier CA. Comparison of lipids from epidermal and palatal stratum-corneum. J Invest Dermatol 1992; 98: 375378.

  • 17. Douglas JE, Mittal C & Thomason JJ et alThe modulus of elasticity of equine hoof wall: implications for the mechanical function of the hoof. J Exp Biol 1996; 199: 18291836.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 18. O'Grady SE. White line disease-an update. Equine Vet Educ 2002; 14: 5155.

  • 19. Wagner IP, Hood DM. Effect of prolonged water immersion on equine hoof epidermis in vitro. Am J Vet Res 2002; 63: 11401144.

  • 20. Squier CA, Hall BK. The permeability of skin and oral mucosa to water and horseradish peroxidase as related to the thickness of the permeability barrier. J Invest Dermatol 1985; 84: 176179.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 21. Goodman AM, Haggis L. Regional variation in the flexural properties of the equine hoof wall. Comp Exerc Physiol 2008; 5: 161168.

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Investigation of hydration processes of the equine hoof via nuclear magnetic resonance microscopy

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  • 1 Laboratory of Animal Physiology and Functional Anatomy, Graduate School of Agriculture, Kyoto University, Kitashirakawa-Oiwake-cho, Sakyo-ku, Kyoto, Kyoto 606-8502, Japan.
  • | 2 Clinical Science and Pathobiology Division, Equine Research Institute, Japan Racing Association, 321-4, Tokami-cho, Utsunomiya, Tochigi 320-0856, Japan.
  • | 3 Laboratory of Animal Physiology and Functional Anatomy, Graduate School of Agriculture, Kyoto University, Kitashirakawa-Oiwake-cho, Sakyo-ku, Kyoto, Kyoto 606-8502, Japan.
  • | 4 Laboratory of Animal Physiology and Functional Anatomy, Graduate School of Agriculture, Kyoto University, Kitashirakawa-Oiwake-cho, Sakyo-ku, Kyoto, Kyoto 606-8502, Japan.
  • | 5 Clinical Science and Pathobiology Division, Equine Research Institute, Japan Racing Association, 321-4, Tokami-cho, Utsunomiya, Tochigi 320-0856, Japan.
  • | 6 Clinical Science and Pathobiology Division, Equine Research Institute, Japan Racing Association, 321-4, Tokami-cho, Utsunomiya, Tochigi 320-0856, Japan.

Abstract

Objective—To examine the distribution of water in hoof wall specimens of horses via nuclear magnetic resonance (NMR) microscopy and determine changes in water distribution during hydration.

Sample—4 hoof wall specimens (2 obtained from the dorsum and 1 each obtained from the lateral quarter and lateral heel regions) of the stratum medium of healthy hooves of 1 horse.

Procedures—Equine hoof wall specimens were examined via NMR microscopy. Proton density–weighted 3-D images were acquired. Changes during water absorption were assessed on sequential images.

Results—The inner zone of the stratum medium had higher signals than did the outer zone. Areas of high signal intensity were evident in transverse images; these corresponded to the distribution of horn tubules. During water absorption, the increase in signal intensity started at the bottom of a specimen and extended to the upper region; it maintained the localization pattern observed before hydration. The relationship between the local maximal signals in areas corresponding to the horn tubules and minimal signal intensities in areas corresponding to the intertubular horn was similar and maintained approximately a linear distribution.

Conclusions and Clinical Relevance—Based on the premise that signal intensity reflects water content, hydration in the equine hoof wall during water absorption occurred concurrently in the tubules and intertubular horn, and there was maintenance of the original water gradients. This technique can be applied for the assessment of pathophysiologic changes in the hoof wall on the basis of its hydration properties.

Abstract

Objective—To examine the distribution of water in hoof wall specimens of horses via nuclear magnetic resonance (NMR) microscopy and determine changes in water distribution during hydration.

Sample—4 hoof wall specimens (2 obtained from the dorsum and 1 each obtained from the lateral quarter and lateral heel regions) of the stratum medium of healthy hooves of 1 horse.

Procedures—Equine hoof wall specimens were examined via NMR microscopy. Proton density–weighted 3-D images were acquired. Changes during water absorption were assessed on sequential images.

Results—The inner zone of the stratum medium had higher signals than did the outer zone. Areas of high signal intensity were evident in transverse images; these corresponded to the distribution of horn tubules. During water absorption, the increase in signal intensity started at the bottom of a specimen and extended to the upper region; it maintained the localization pattern observed before hydration. The relationship between the local maximal signals in areas corresponding to the horn tubules and minimal signal intensities in areas corresponding to the intertubular horn was similar and maintained approximately a linear distribution.

Conclusions and Clinical Relevance—Based on the premise that signal intensity reflects water content, hydration in the equine hoof wall during water absorption occurred concurrently in the tubules and intertubular horn, and there was maintenance of the original water gradients. This technique can be applied for the assessment of pathophysiologic changes in the hoof wall on the basis of its hydration properties.

The stratum medium of the equine hoof wall is a keratinized tissue with material characteristics and material properties that facilitate the accommodation of forces associated with static and dynamic weight bearing. The mechanical properties of keratinized structures are modulated by their hydration state.1–3 Water content affects the stiffness and fracture resistance of the hoof wall.1 Although excessive dehydration causes fractures in the hoof, swelling attributable to excess water absorption into the hoof wall is linked to a high incidences of hoof damage.4 In other studies,1,5,6 the water content of the hoof wall has been examined by gravimetric changes via various drying techniques because limited methods exist for determination of water distribution in the tissue. It is difficult to detect localization patterns for water in specimens with these techniques. Therefore, we attempted to use NMR microscopy, which can reveal water localization through visual results.

Nuclear magnetic resonance microscopy is a method that provides enhanced spatial resolution because of the use of a strong static magnetic field and high magnetic field gradients. In NMR microscopy, spatial resolution is provided to submillimeter dimensions,7,8 whereas that in traditional MRI is on the order of 1 mm. Magnetic resonance imaging, including NMR microscopy, is suitable for visual localization of water molecules in tissue specimens.

Magnetic resonance imaging or NMR microscopic investigation of equine hoof specimens has been reported.9,10 In these studies,9,10 the lamellae of the inner hoof wall (stratum internum) and the coronary papillae of the coronet were clearly visible; however, structures in the stratum externum and stratum medium of the hoof wall were less clearly visible. The stratum medium has a characteristic structural organization of individual horn tubules surrounded and separated by intertubular horns. The horn tubules are elongated cylinders approximately 0.1 to 0.2 mm in diameter that morphologically consist of 2 components: the outer circular layer, known as the cortex, and the central area (marrow). These tubules are believed to be involved in regulating the mechanical properties of the hoof wall as well as the gradient of water content11 and water transfer12 in the hoof wall on the basis of their morphology and distribution pattern. These beliefs raise the possibility that water distribution in the hoof wall may be associated with these tubules; however, the relationship of horn tubules to water distribution or permeation in the hoof wall has not been determined, but has been evaluated via an indirect observation with horseradish peroxidase as a water-soluble tracer.13

The objective of the study reported here was to determine via sequential NMR microscopic observations the distribution of water in equine hoof wall specimens and changes in this distribution during hydration. The process of water absorption from the surface of the horn specimen contacting the water and the relationship between the absorption pattern of the horn tubules and intertubular horns in this process were analyzed to assess the involvement of horn tubules in water transfer.

Materials and Methods

Sample—Hoof wall specimens were obtained from a healthy equine hoof of a horse that was euthanized. The horse received a sedative (5 μg of medetomidine hydrochloride/kg, IV) and was subsequently injected with an anesthetic (2.5 g of thiopental sodium, IV). After the horse was unconscious, a muscle relaxant (2 g of suxamethonium chloride, IV) was administered to cause respiratory arrest. In addition, exsanguination from the carotid artery was used to cause immediate cardiac arrest. Specimens of approximately 5×5×10 mm, with the long axis oriented parallel to the horn tubules, were cut with an electric band sawa from the dorsum, lateral quarter, and lateral heel regions of the stratum medium at the midpoint of the height of the hoof wall. All euthanasia and sample-procurement procedures were approved by the Animal Welfare and Ethics Committee of the Equine Research Institute of the Japan Racing Association.

Preparation for NMR microscopic evaluation—The stratum externum and internum were removed, and only the stratum medium was prepared for evaluation. Each stratum medium was wrapped with plastic sealing filmb to prevent dehydration and was stored refrigerated at 4° to 10°C.

The sealing film around each specimen was removed, and the 4 sides of each specimen parallel to the tubules were covered with nail varnish. The specimen was set in a flat-bottomed NMR sample tube, with the axis of the tubules oriented in the vertical direction. The region predicted to be out of the field of view (> 8 mm from the bottom) was covered with aluminum foil to prevent aliasing artifacts. Each sample tube contained a flat polytetrafluoroethylene ring at the bottom as a spacer and a polyethylene tube for injection of water into the bottom of the tube. A plug consisting of a piece of silicone sponge was placed in each sample tube just above the hoof wall specimen.

NMR microscopy—Examinations were performed with an NMR microscope,c which consisted of a 400-MHz wide-bore spectrometer equipped with a microimaging attachment for acquisition of the proton signal. The sample tube was set in the detection coil (diameter, 10 mm) of the imaging probe. The imaging probe was inserted in a magnet with a static magnetic field of 9.4 T equipped with coils that generated magnetic field gradients. Nuclear magnetic resonance microimages were acquired with 3-D Fourier transformation variable flip-angle (flip-back) spin-echo sequences to reduce the effect of the T1 relaxation (spin-lattice relaxation) time and allow acquisition of proton density–weighted images with a short repetition time. Variables were set as follows: field of view, 10 × 10 × 10 mm; repetition time, 500 milliseconds; echo time, 5 milliseconds; flip angle, 165°; data matrix, 128 × 128 × 16 (16 encoding along the direction parallel to the tubules); and number of excitations, 1. Receiver gain was fixed. Imaging time was approximately 20 minutes.

For each specimen, an initial image was acquired (time 0). The sample tube then was removed from the magnet, and 0.2 mL of ultrapure water was placed at the bottom of the sample tube to initiate water absorption. The specimen was replaced in the magnet as quickly as possible after the addition of the water, and 3 images were acquired. The specimen remained in the magnet throughout the remainder of the experiment, and images were acquired 2, 4, 6, 8, and 24 hours after initiating water absorption.

Image reconstruction and analysis—Image reconstruction and analysis were performed by Fourier transformation. The image matrix was zero-filled to 256 × 256 × 32. Image analysis was performed with computer software.14,d To estimate signal changes in horn tubules, local signal maxima were detected in transverse images of the hoof wall. Local maxima that had signal intensities higher than the upper 2.5% of the local maxima in the image prior to water absorption were counted as high local maxima. Segmentation and analysis of images were also performed to enable us to examine the relationship between the signal intensities of the horn tubules and intertubular horn areas, which were represented by maximal and minimal signals in each segment, respectively.

Results

Localized patterns and increases in signal intensities were observed in NMR microscopic images during water absorption (Figure 1). In specimens before hydration, the inner zone of the stratum medium had higher signal intensities than did the outer zone. Areas of high signal intensity were evident in the inner area of the stratum medium in transverse images; these areas corresponded to the distribution of horn tubules. Occasionally, low signal intensity was observed in the center of these areas of high signal intensity; these low-intensity areas corresponded to the horn tubule marrow.

Figure 1—
Figure 1—

Photograph of a hoof wall specimen obtained from the dorsum of a hoof (A) and NMR microscopic images of the same specimen acquired before (B) and after (C) water absorption. Notice the color scale for the signal intensity.

Citation: American Journal of Veterinary Research 73, 11; 10.2460/ajvr.73.11.1775

Similar structures were less clear in the outer area of the stratum medium before hydration but became apparent after water absorption. The differences in signal intensities between the inner and outer zones remained consistent after water absorption. Immediately after the initiation of water absorption, signal increases were observed at the bottom of each specimen (ie, near the surface in contact with the water). Increases in signal intensity were evident in the upper region of each specimen (approx 6 mm above the surface in contact with the water) at 6 hours after initiation of water absorption (Figure 2). There were no marked differences in these processes among the specimens, except that the proportion of areas with high signal intensity was larger in the specimens obtained from the lateral heel regions than in the specimens obtained from the dorsum and lateral quarter regions.

Figure 2—
Figure 2—

A series of NMR microscopic images of the hoof wall specimen in Figure 1 obtained from the dorsum of the hoof and allowed to absorb water for up to 24 hours. Representative images are provided, with intervals of 1.25 mm in the direction of water permeation (water was placed in a tube in contact with the bottom of each specimen and was absorbed toward the upper region of each specimen). Signal intensity is indicated by a color scale. See Figure 1 for remainder of key.

Citation: American Journal of Veterinary Research 73, 11; 10.2460/ajvr.73.11.1775

Image analysis revealed that the number of local maximal points detected in transverse images of the hoof wall typically increased after water absorption. The observed numbers of high local maxima were plotted in relation to their respective distances from the water surface in contact with the bottom of each specimen and the duration of water absorption (Figure 3). The number of high local maxima increased near the water surface immediately after the initiation of water absorption. Increases were evident in the upper region of each specimen at 6 hours after initiation of water absorption. Gradients in the number of high local maxima were preserved during water absorption for at least 8 hours. Analysis of the relationships between the local maximal and minimal signal intensities in each area revealed that they were similar and maintained approximately a linear distribution (r > 0.8) during water absorption (Figure 4).

Figure 3—
Figure 3—

Number of high local maxima, in relation to the distance from the water surface in contact with each specimen and the duration of water absorption, for specimens obtained from the dorsum (A and B), lateral quarter (C), and lateral heel (D) of a healthy equine hoof. Maxima were determined before (time 0 [white square and dashed line]) and 0.3 (white triangle and dashed line), 0.6 (white circle and dashed line), 0.9 (black square and solid line), 2 (black triangle and solid line), 4 (black circle and solid line), 6 (white square and solid line), 8 (white triangle and solid line), and 24 (white circle and solid line) hours after initiation of water absorption. On the x-axis, 1 U of height is approximately 0.31 mm; height = 0 is the bottom of each specimen (ie, the location nearest to the water surface in contact with the specimen). AU = Arbitrary units.

Citation: American Journal of Veterinary Research 73, 11; 10.2460/ajvr.73.11.1775

Figure 4—
Figure 4—

Relationships between local maximal and minimal signal intensities in each segment of equine hoof at various times after initiation of water absorption in the same specimen as in Figure 1. The range of the axis for maximal and minimal signals in the plots is 0 to 100,000 and 0 to 80,000, respectively. AU = Arbitrary units.

Citation: American Journal of Veterinary Research 73, 11; 10.2460/ajvr.73.11.1775

Discussion

Magnetic resonance imaging enables the nondestructive examination of the localization and status of molecules in materials. In proton MRI, major signals arise from water and lipids during examination of biological materials. Signal intensities are affected by the density of the nuclei and by their T1 relaxation and T2 relaxation (spin-spin relaxation) properties. Although lipids generate high magnetic resonance signals because of their short T1 relaxation time, the lipid content of the hoof wall has been reported to be 1.5% on a dry-weight basis15; therefore, the overall contribution of lipids to the magnetic resonance signals in the study reported here was likely to be small. Some ceramides are likely to be covalently bound to the cellular envelope, as has been reported in the epidermis16; however, these are considered to generate only low-intensity signals because highly restricted molecules have extremely short T2 relaxation times. Water in the tissue interacts with macromolecules such as proteins and carbohydrates, and bound water has shorter T1 and T2 relaxation times than does free water. Therefore, we expected that local variations in water content in the hoof wall and changes during hydration would result in differences in relaxation times. In the study reported here, our objective was to examine the localization of water molecules, regardless of whether they were bound or in a free state; however, in a preliminary study (data not shown), we observed local variations of T1 in the hydrated hoof wall specimens, which ranged from < 0.1 to 1.2 seconds, with longer T1 times in tubular areas. In addition, a shorter duration for imaging time (ie, a short repetition time sequence) was also needed to acquire transient images during the hydration process in the present study. These requirements were met by use of a variable flip-angle pulse, albeit with incomplete suppression of the T1 variation effect. Therefore, we believe that the signal intensities in the NMR microimages reflected the water content in the corresponding sites in the specimens.

The water content in the hoof wall is higher in the inner zone than in the outer zone.1,3,17 Similar differences were detected with NMR microscopy in the study reported here. Areas of high signal intensity in transverse images of the horn wall have also been described as horn tubules.10 Points of low signal intensity within these areas correspond macroscopically to dark spots at the centers of the horn tubules. The horn tubules originate from coronary papillae that contain blood vessels to nourish the germinative layer of the stratum medium. After the coronet has received a strong impact, these papillae often hemorrhage slightly, and the horn tissue can take up the hemorrhagic debris. Thus, deformed erythrocytes, which are probably derived from this hemorrhage, are occasionally observed in the marrow of the horn tubules. The dark spots observed in the NMR images were assumed to be hemosiderin, a hemoglobin derivative. Hemosiderin shortens apparent spin-spin relaxation time as well as T2 and may thereby lower MR signals. Gradients of signal intensities between the inner and outer zones of the stratum medium were evident in both the tubular and intertubular areas. These results suggested that the horn tubules had higher moisture content than did the intertubular areas and that even though the moisture content in these compartments varied, depending on the location in the hoof wall, the moisture content of each horn tubule typically was higher than that in the surrounding intertubular region.

Water content of the equine hoof wall specimen is 18.2% at 75% relative humidity, and it increases to 40% at 100% relative humidity.1 Fully hydrated hoof wall material has a 50% decrease in fracture resistance, compared with the maximal fracture resistance observed at 75% relative humidity; hydration also decreases the stiffness of the hoof wall.1 These changes increase the incidence of hoof wall damage4 and may be a risk factor for white line disease.18 Another study19 revealed that hoof wall specimens immersed in water had a significant increase in mass after 24 hours; however, local differences in the hydrating hoof wall were not reported. In 1 study,13 researchers investigated the permeability barrier in the dorsal wall of the equine hoof with horseradish peroxidase as a water-soluble tracer and concluded that there was a permeable barrier in the normal hoof wall. In the study reported here, we found that absorption of water (estimated from changes in signal intensities) started immediately after contact with the water; however, diffusion of water into the hoof wall was slow because the moisture gradient, represented by the number of hydrated horn tubules, was maintained during at least 8 hours of hydration.

To confirm the hydration of hoof wall specimens for the conditions of the present study, we also examined changes in the hydration status of the specimens in a similar setting by measuring changes in weight. Mean ± SD moisture content of the specimens increased from 29.77 ± 1.08% to 37.85 ± 2.98% during 8 hours of hydration and subsequently reached 39.01 ± 0.08% after 24 hours of hydration; these values are comparable to those reported for the physiologic or hydrated state.1,2,17 The permeation of water observed in our experiments differed from that reported in another study,13 presumably because of differences in the molecular sizes of water and horseradish peroxidase. This difference is plausible on the basis of analogous observations in skin and oral mucosa.20 In heel specimens, the area of high signal intensity in the inner zone of the stratum medium was broader than that in the corresponding areas in the dorsum and lateral specimens; however, the signal intensity of each area and changes detected during the hydration process were similar among all the specimens. The moisture content is higher in the heel than in other parts of the hoof wall.21 Differences in the ratio of the areas for the inner and outer zones of the stratum medium are likely to contribute to variations in moisture content between the heel and other hoof sites, although this may also be attributable to differences in the hygroscopic properties of each zone among the hoof sites.

We also observed that hydration progressed concurrently in the horn tubules and intertubular regions. It is thought that the horn tubules facilitate water transfer in the hoof wall.12 If water first penetrates into the tubules and then diffuses into the intertubular region, it would be expected that signal intensities at the local maxima of NMR microimages would increase first and those at the corresponding minimal points would remain low during the early stages of water permeation. Our image analyses revealed that the signal intensities at the local maxima and the surrounding regions of minimal signal remained highly associated during the hydration process. In another study,5 investigators determined that, on the basis of weight changes in hoof wall specimens exposed to water vapor in a hydration chamber, horn tubules do not facilitate hydration. The observations and conclusions of that study5 are in agreement with results of the study reported here. We believe that horn tubules have no specific effects on the transfer of water.

In the present study, we evaluated zonal and structural differences in water localization in the equine hoof wall. Analysis of the results suggested that hydration in the hoof wall during water absorption occurs concurrently in the tubules and intertubular horn, thereby maintaining water gradients. Results of the present study indicated that NMR microimaging techniques can be used in the assessment of pathophysiologic changes of the hoof wall on the basis of its hydration properties.

ABBREVIATION

NMR

Nuclear magnetic resonance

a.

2114C, Makita Co Ltd, Aichi, Japan.

b.

Parafilm, Pechiney Plastic Packaging Inc, Chicago, Ill.

c.

NM-AIM imaging equipment, JEOL, Tokyo, Japan.

d.

ImageJ, version 1.45, National Institutes of Health, Bethesda, Md. Available at: rsbweb.nih.gov/ij/index.html. Accessed Mar 2, 2011.

References

  • 1. Bertram JE, Gosline JM. Functional design of horse hoof keratin: the modulation of mechanical properties through hydration effects. J Exp Biol 1987; 130: 121136.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 2. Hinterhofer C, Stanek C, Binder K. Elastic modulus of equine hoof horn, tested in wall samples, sole samples and frog samples at varying levels of moisture. Berl Munch Tierarztl Wochenschr 1998; 111: 217221.

    • Search Google Scholar
    • Export Citation
  • 3. Wagner IP, Hood DM, Hogan HA. Comparison of bending modulus and yield strength between outer stratum medium and stratum medium zona alba in equine hooves. Am J Vet Res 2001; 62: 745751.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 4. Borderas TF, Pawluczuk B & de Passille AM et alClaw hardness of dairy cows: relationship to water content and claw lesions. J Dairy Sci 2004; 87: 20852093.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 5. Kasapi MA, Gosline JM. Exploring the possible functions of equine hoof wall tubules. Equine Vet J Suppl 1998;(26):1014.

  • 6. Higuchi H, Kurumado H & Mori M et alEffects of ammonia and hydrogen sulfide on physical and biochemical properties of the claw horn of Holstein cows. Can J Vet Res 2009; 73: 1520.

    • Search Google Scholar
    • Export Citation
  • 7. Eccles CD, Callaghan PT. High-resolution imaging. The NMR microscope. J Magn Reson 1986; 68: 393398.

  • 8. Neeman M, Sillerud LO. NMR microscopy. In: Gillies RJ, ed. NMR in physiology and biomedicine. San Diego: Academic Press Inc, 1994;101118.

    • Search Google Scholar
    • Export Citation
  • 9. Murray RC, Dyson SJ & Schramme MC et alMagnetic resonance imaging of the equine digit with chronic laminitis. Vet Radiol Ultrasound 2003; 44: 609617.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 10. Keller MD, Galloway GJ, Pollitt CC. Magnetic resonance microscopy of the equine hoof wall: a study of resolution and potential. Equine Vet J 2006; 38: 461466.

    • Search Google Scholar
    • Export Citation
  • 11. Pollitt CC. Anatomy and physiology of the inner hoof wall. Clin Tech Equine Pract 2004; 3: 321.

  • 12. Butler D. The principles of horseshoeing. 2nd ed. Maryville, Mo: D. Butler Publisher, 1985.

  • 13. Kempson SA, Campbell EH. A permeability barrier in the dorsal wall of the equine hoof capsule. Equine Vet J Suppl 1998;(26):1521.

  • 14. Rasband WS. ImageJ. Bethesda, Md: US National Institutes of Health, 2009.

  • 15. Wertz PW, Downing DT. Cholesteryl sulfate: the major polar lipid of horse hoof. J Lipid Res 1984; 25: 13201323.

  • 16. Wertz PW, Kremer M, Squier CA. Comparison of lipids from epidermal and palatal stratum-corneum. J Invest Dermatol 1992; 98: 375378.

  • 17. Douglas JE, Mittal C & Thomason JJ et alThe modulus of elasticity of equine hoof wall: implications for the mechanical function of the hoof. J Exp Biol 1996; 199: 18291836.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 18. O'Grady SE. White line disease-an update. Equine Vet Educ 2002; 14: 5155.

  • 19. Wagner IP, Hood DM. Effect of prolonged water immersion on equine hoof epidermis in vitro. Am J Vet Res 2002; 63: 11401144.

  • 20. Squier CA, Hall BK. The permeability of skin and oral mucosa to water and horseradish peroxidase as related to the thickness of the permeability barrier. J Invest Dermatol 1985; 84: 176179.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 21. Goodman AM, Haggis L. Regional variation in the flexural properties of the equine hoof wall. Comp Exerc Physiol 2008; 5: 161168.

Contributor Notes

Dr. Yoshihara's present address is Racehorse Hospital, Ritto Training Center, Japan Racing Association, 1028, Misono, Rittoshi, Shiga, 520-3005, Japan.

Dr. Wada's present address is Equine Department of Veterinary Section, Japan Racing Association, 11-1, Roppongi-6-chome, Minato-ku, Tokyo, 106-8401, Japan.

Supported in part by a grant from the Japan Racing Association.

Presented in abstract form at the Annual Meeting of the Japanese Society of Equine Science, Tokyo, November 2009.

Address correspondence to Dr. Sugimoto (sugimoto@kais.kyoto-u.ac.jp).