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. 2018 Jun 6;15(4):623–632. doi: 10.1111/iwj.12906

Influence of digestive enzymes on development of incontinence‐associated dermatitis: Inner tissue damage and skin barrier impairment caused by lipidolytic enzymes and proteases in rat macerated skin

Yuko Mugita 1, Takeo Minematsu 2,3, Gojiro Nakagami 1,3, Hiromi Sanada 1,3,✉
PMCID: PMC7949763  PMID: 29877066

Abstract

One of the most common complications in patients with incontinence is incontinence‐associated dermatitis. This study aimed to examine the influences of lipidolytic enzymes and/or proteases on skin barrier and tissue structure on the development of incontinence‐associated dermatitis. Two animal experiments, ex vivo and in vivo, were performed using rats to examine the influences of 3 factors (maceration, proteases, and lipidolytic enzymes) alone or in various combinations on the barrier function and histology of the skin. As a result, skin treatments, including both of the skin maceration and proteases application, caused erythrocyte leakage from the blood vessels in the dermis. The erythrocyte leakage was observed in a larger area in the skin treated with proteases and lipidolytic enzymes with maceration than in the skin treated with proteases with maceration, that is, the addition of lipidolytic enzymes to skin maceration with proteases enhanced erythrocyte leakage. Lipidolytic enzymes in macerated skin are factors that accelerate tissue damage via skin barrier impairment, and proteases are the factors that trigger the development of incontinence‐associated dermatitis via tissue damage. Advanced nursing care of perineal skin in patients with faecal incontinence is required because of the deleterious influence of lipidolytic enzymes and proteases.

Keywords: digestive enzymes, incontinence‐associated dermatitis, inner tissue damage, skin barrier function, skin maceration

1. INTRODUCTION

One of the most common complications in patients with incontinence is skin disorders caused by exposure to urine or faeces. Such a skin disorder is globally identified as incontinence‐associated dermatitis (IAD).1, 2 IAD is defined as the inflammation of the skin that occurs when urine or stool comes into contact with the perineal or perigenital skin1 and is macroscopically identified as rash, erythema, blister, or erosion.3, 4, 5

The elderly population frequently has incontinence.6 It has been reported that the prevalence of incontinence among nursing home residents was 59.8%, including 7.7% of residents with urinary incontinence only, 12.4% with faecal incontinence only, and 39.7% with both urinary and faecal incontinence.7 The prevalence of IAD was reported to be 36.0% in elderly patients with urinary and/or faecal incontinence in Japanese long‐term medical facilities,8 and the incidence of IAD among community‐living adults with faecal incontinence was 41%.3 The condition of the elderly population is also complicated by structural changes in the skin that occur with the aging of the skin, which increase skin fragility, decrease the ability of the skin to heal, and promote the development of various cutaneous disorders.9 Therefore, elderly patients with incontinence are at a particularly high risk of the development of IAD. Once IAD is developed, it is accompanied by discomfort, such as tingling, itching, burning, and pain,1 which negatively affect the patients’ quality of life.10 Clinical nurses recognise that patients can develop IAD‐related discomfort. Therefore, the development of effective care for IAD is a pressing nursing issue. Conventional care for established IAD is mainly performed by the removal of urine or faeces from the area by cleansing or the application of a skin barrier product to prevent repeated exposures. Such care does not directly contribute to IAD healing. Effective care for the treatment of IAD should be provided as early as possible because prolonged IAD poses a risk of bacterial infection caused by faecal exposure11; risk of ulceration, including pressure ulcer12; and leads to the eventual decline in the patients’ quality of life.10 The provision of effective therapeutic care for IAD requires the recognition of the pathophysiology of IAD. However, no concept of IAD treatment has existed yet because the pathophysiology of IAD development and healing is as yet unknown. Recognition of the pathophysiology of IAD is required for the development of an effective therapeutic approach for IAD.

Skin maceration, which is caused by exposure to excessive water, has been focused on as one of the risk factors of IAD. A previous clinical investigation revealed that the adjacent area of IAD shows a higher skin hydration, indicating skin maceration, than the perineal skin of patients without IAD.8 Skin maceration is defined as a functional disorder of the skin barrier.13 In macerated skin, structural alterations in the intercellular lipid layers14, 15, 16, 17 and junctions between the keratinocytes in the epidermis17 contribute to skin barrier impairments. Skin barrier impairments in macerated skin are clinically recognised by increased transepidermal water loss (TEWL)17, 18 and enhancement of the transdermal penetration of soluble molecules larger than 500 Da into the deeper layer of the dermis16, 17; TEWL is a parameter of skin barrier function against water penetration from the inside to the outside of the body. Skin barrier impairment caused by skin maceration is considered to be a basic factor of IAD development.

We previously demonstrated the histological and functional investigations of macerated skin with excessive normal saline in rats. We did not observe any skin lesions or inflammation, although the skin barrier function was remarkably disrupted in macerated skin.17 These results suggest that skin maceration is not enough to contribute to IAD development. IAD is supposed to develop via the transdermal penetration of irritants, which is caused by an impairment of the skin barrier function related to skin maceration.

Bliss et al.7 have reported that nursing home residents with faecal incontinence only or with double incontinence were significantly correlated with perineal dermatitis; conversely, urinary incontinence alone did not have a correlation, which indicates that contact with faeces is a crucial factor for IAD development. Faeces contain several digestive enzymes, including trypsin, α‐chymotrypsin, lipase, and phospholipase.19, 20 Trypsin and lipase have been reported as risk factors for erythema and skin barrier impairment.20 In particular, trypsin is an enhancer factor for the transdermal penetration of macromolecules.21, 22 It is suggested that the barrier impairment in the perineal skin exposed to urine and faeces is accelerated because of the influence of proteases as well as skin maceration. Moreover, faeces also contain intestinal bacteria.23, 24 In patients with faecal incontinence, the perineal skin is frequently exposed to high concentrations of bacterial flora. Therefore, the perineal skin of patients with faecal incontinence is exposed to high‐density bacterial flora.

In our previous study on the examination of IAD pathophysiology,25 we considered proteases and bacteria in the faeces to be important factors for IAD development. At first, the influence of protease‐associated skin maceration, termed “proteolytic maceration”, was examined in dorsal rat skin. A comparison of proteolytic skin maceration and no treatment skin revealed a significant increase in the TEWL, digestion in the prickle cell layer of the epidermis, and slight bleeding in the papillary dermis and around hair follicles, observed 24 hours after proteolytic skin maceration, although there was no macroscopic erosion, and only dot‐like skin redness was observed. These results indicated that proteolytic skin maceration induces inner tissue damage caused by transdermally penetrated proteases. Next, the influence of bacteria inoculation on proteolytic skin maceration was examined. Bacteria inoculation was performed in the skin after the treatment of proteolytic skin maceration and normal skin. Bacteria‐inoculated proteolytic skin maceration caused expanded skin flares without macroscopic erosions, as found in IAD in the health care setting. Histological analysis revealed the formation of bacteria‐rich clusters within the papillary dermis and remarkable tissue damage around the clusters in the bacteria‐inoculated proteolytic skin maceration. These results suggest that the overload of digestive enzymes and intestinal bacterial flora due to faecal incontinence accelerated the inner tissue damage, although the outer skin showed only skin flares. IAD is usually diagnosed as contact dermatitis, and skin care for IAD is performed in accordance with that for contact dermatitis. However, the pathophysiological finding in the rat IAD model, bacterial clusters in the dermis, was different from the pathophysiology of contact dermatitis. The skin care for IAD should be based on the pathophysiology of IAD.

In our previous study,25 we focused only on the proteases because proteases were considered to have a significant influence on IAD development by accelerating the transdermal penetration of macromolecules.21, 22 The influence of lipidolytic enzymes, the major digestive enzymes in faeces as well as proteases,20 has not been examined in macerated skin. Faeces contained lipase and phospholipase A2, which are secreted from the pancreas. Lipase digests the triglycerides into free fatty acids and monoglycerides and phospholipase digests phospholipids into fatty acids.

Lipids have important roles in skin barrier function and structure. The sebum is a lipid layer, which covers the surface of the skin. It serves as a barrier against water. The sebum is mainly composed of triglycerides.26, 27, 28 Therefore, there is a possibility that lipase digests the sebum lipid, which may accelerate skin maceration via penetration of excessive water. The intercellular lipid lamellae in the stratum corneum are the main barrier for the diffusion of substances across the skin. The lamellae lipids are predominantly composed of 3 lipid classes: ceramides, cholesterol, and free fatty acids.29 The lamellae lipid may be digested by lipidolytic enzymes, resulting in impairment of the skin barrier function and acceleration of skin maceration. IAD development is considered to be accelerated in skin with an impaired lamellae lipid barrier. Lipids also contribute as a cell membrane. The plasma membrane, a bilayer of phospholipids, is important for the boundary between the inside and outside of the cell and for selective permeability.30 Phospholipase may digest the lipid bilayer, resulting in the degradation of cells and necrosis.

It has been reported that lipase treatment on the skin surface induced skin irritation macroscopically and skin barrier impairment.31, 32 However, the influence of lipidolytic enzymes on the tissue structure has not been revealed. Moreover, the influence of lipidolytic enzymes treated with proteases and/or skin maceration on the skin barrier impairment and tissue structure alteration has not been examined. We have two hypotheses: lipidolytic enzymes enhance the skin barrier impairment caused by skin maceration and cause tissue damage via transdermal penetration in macerated skin; furthermore, the combination of lipidolytic enzymes and proteases is considered to enhance the histological changes and skin barrier impairments in macerated skin compared with proteases only or lipidolytic enzymes only. We conducted experiments to examine the influence of lipidolytic enzyme‐associated skin maceration, termed “lipidolytic maceration”, on IAD development, especially on skin barrier impairment and tissue structural alteration. Experiments were conducted using two steps. First, histological examination was performed ex vivo, followed by skin physiological examination and histological examination performed in vivo.

2. METHODS

2.1. Reagents

Bovine serum albumin (BSA), a trypsin from porcine pancreas, α‐chymotrypsin from bovine pancreas, lipase from porcine pancreas, Dulbecco's modified Eagle medium (DMEM), and penicillin/streptomycin were purchased from Nacalai Tesque (Kyoto, Japan). Pentobarbital sodium was purchased from Kyoritsu Seiyaku (Tokyo, Japan). VectaStain ABC Kit and biotin‐conjugated anti‐mouse IgG antibody were purchased from Vector Laboratories (Burlingame, CA, USA), and 3,3′‐diaminobenzidine tetrahydrochloride (DAB) tablets were purchased from Wako Pure Chemical (Osaka, Japan). Dextran conjugated with FITC (FITC‐dextran) was purchased from Invitrogen (Carlsbad, CA, USA). Phospholipase A2 from porcine pancreas was purchased from Sigma‐Aldrich (St. Louis, MO, USA). Foetal bovine serum (FBS) was purchased from BioWest (Nuaille, France). Sheep anti‐FITC HRP was purchased from Southern Biotech (Birmingham, AL, USA).

2.2. Animal

Ten 6‐month‐old male Sprague‐Dawley rats (Japan SLC, Shizuoka, Japan), produced and maintained under conditions that were specifically pathogen‐free of various microorganisms, were fed a standard laboratory diet and filtered water ad libitum. The rats were individually housed in a temperature‐ and humidity‐controlled room (23°C ± 2°C temperature, 50% ± 10% humidity). The animal experiments were performed in accordance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals and were approved by the Animal Research Committee of The University of Tokyo (P‐12‐45).

2.3. Skin maceration treatment

A modified method of skin maceration treatment by agarose gel application25 was used. Approximately 100‐μL droplets of 1% agarose gel dissolved in 153 mM Tris‐HCl buffer (pH 7.4) were immersed in the following treatment solutions and gently shaken overnight in a cooler box: 153 mM Tris‐HCl buffer (pH 7.4); proteolytic solution (0.25% wt/vol trypsin and 0.40% wt/vol α‐chymotrypsin in Tris‐HCl buffer); lipidolytic solution (10 units/L lipases and 0.20% wt/vol phospholipase A2 in Tris‐HCl buffer); and proteolytic and lipidolytic solution (0.25% wt/vol trypsin, 0.40% wt/vol α‐chymotrypsin, 10 units/L lipases, and 0.20% wt/vol phospholipase A2 in Tris‐HCl buffer).

Appropriate pH of the treatment solution was determined by a preliminary test. We examined enzyme activities at pH 4.5 to 9.0 both in vitro and ex vivo. The activities of enzymes were the highest at pH 7.4, indicating that pH 7.4 was appropriate for an IAD model.

Skin maceration treatment was performed by applying the prepared agarose gel droplets on the dorsal rat skin 3 days after hair removal. We used 6‐month‐old rats, which were approximately the same age as the rats in the previous study.25 Aged skin causes the enhancement of the maceration‐induced disruption of the skin barrier function,17 and there is clinical evidence that IAD is problematic among aged patients.3, 8 Therefore, a 6‐month‐old rat was suitable as an aged model for the examination of the pathophysiology of IAD.

2.4. Ex vivo examination of the influence of lipidolytic enzymes on tissue structural alterations

Five rats were used in the ex vivo investigation to examine the influence of lipidolytic enzymes on the tissue structural alteration. The dorsal hair of the rats was removed using an electric shaver and depilatory cream under anaesthesia 3 days before agarose gel application. Full‐thickness dorsal skin was obtained from each rat and divided into 8 1× 1 cm2 pieces. The divided skin pieces were randomly assigned to 1 of the following 8 patterns of skin treatment: no treatment (NT), maceration (M), proteases without maceration (P), lipases without maceration (L), proteases and lipases without maceration (PL), lipases with maceration (LM), proteases with maceration (PM), and proteases and lipases with maceration (PLM).

Each skin piece was placed in 24‐well plates. The areas below the dermis of the skin pieces were soaked in a culture medium (10% FBS and 1% penicillin/streptomycin in DMEM). First, skin maceration treatment was performed in the M, PM, LM, and PLM groups. Agarose gels containing pH 7.4 Tris‐HCl buffer were applied on the surface of the skin pieces. Skin maceration treatment was performed for 2 hours, and then, the agarose gels were removed. The skin surface was wiped with paper cloth (PROWIPE; DAIO Paper Corp., Tokyo, Japan).

Next, the enzyme treatment was performed in the P, L, PL, LM, PM, and PLM groups. For the application of lipidolytic enzymes and/or proteases, agarose gels immersed in a proteolytic solution or lipidolytic solution or proteolytic and lipidolytic solution were applied on the skin pieces after skin maceration treatment. Application of enzymes was performed for 15 minutes, and then, the agarose gels were removed. The skin surface was wiped with paper cloth. According to previous research, agarose gel application for <30 minutes does not cause skin maceration.17 Therefore, the application of enzyme‐containing agarose gel for <30 minutes was considered to cause enzyme influences without skin maceration. The application time of enzymes was set as 15 minutes based on the results of a preliminary test, which demonstrated that the 15‐minute application of enzyme‐containing agarose gel was enough to cause histological changes in PM similar to those seen in the previous study.25

Each skin piece was fixed in 4% paraformaldehyde in phosphate buffer (pH 7.4) and used for the histological analysis via haematoxylin and eosin (HE) staining.

2.5. In vivo examination of the influence of lipidolytic enzymes on macerated skin and proteolytic macerated skin

Five rats were used for the in vivo experiment in the histological analysis via HE staining. Four areas in the dorsal skin of 5 rats (total 20 areas) were randomly divided into 5 groups: NT, M, PM, LM, and PLM. In the ex vivo experiment, all of the combinations of the three factors (maceration, proteases, and lipidolytic enzymes) were performed as skin treatment; skin treatments with enzymes only (P, L, and PL) were excluded. Maceration‐associated skin treatment (M, PM, LM, and PLM) and NT were adopted in the in vivo experiment to examine the influences of the enzymes on skin maceration because skin maceration is thought to be the essential condition to reproduce IAD from the results of the ex vivo experiments. Agarose gel was prepared for each treatment of the M, PM, LM, and PLM groups by soaking it in Tris‐HCl buffer (pH 7.4), proteolytic solution, lipidolytic solution, and proteolytic and lipidolytic solution, respectively. Agarose gel was also immersed in 100 μg/mL FITC‐dextran.

The dorsal hair of the rats was removed using an electric shaver and depilatory cream under anaesthesia 3 days before skin maceration treatment. Skin hydration level and TEWL were measured by Moisture Checker (Scalar, Tokyo, Japan) and VapoMeter (Delfin Technologies, Kupio, Finland), respectively, before and after skin maceration treatment. Baseline values of skin hydration and TEWL were measured, followed by application of FITC‐dextran‐containing agarose gel on each skin treatment area for 30 minutes. After the removal of FITC‐dextran‐containing agarose gel, the skin surface of each treatment area was wiped with paper cloth, followed by skin maceration treatment performed via attachment of agarose gel droplets containing each treatment solution for M, PM, LM, and PLM. Each agarose gel droplet was applied using a polyurethane film dressing (Tegaderm transparent dressing; 3 M, St. Paul, MN, USA) for 3 hours and 30 minutes. The NT area was air‐dried for 3 hours and 30 minutes. After removal of the agarose gel, the skin was air‐dried for 30 minutes, and skin hydration and TEWL were measured on each skin treatment area. Measurement was performed repeatedly, and 3 stable measurements were used for the analysis of skin hydration and TEWL. Measurements were performed after 30 minutes of air‐drying following agarose gel removal to avoid possible measurement failures when the skin surface is wet.33, 34 The appearance of the skin surface was recorded using a digital camera (LUMIX DMC‐FX‐60; Panasonic, Tokyo, Japan). Twenty‐four hours after removal of the agarose gels, skin appearance was again recorded by the digital camera, and then, full‐thickness skin tissue samples were collected under general anaesthesia. The collected tissues were immediately fixed in 4% paraformaldehyde in phosphate buffer (pH 7.4) and used for HE staining and immunohistochemistry for FITC.

The skin barrier function, as well as the TEWL value, was immunohistologically examined via the transdermal penetration of FITC‐dextran. The molecular size of dextran is 3 kDa, which is too large to penetrate the intact skin barrier. In the previous research on skin maceration, transdermal penetration of FITC‐dextran was used as the indicator of skin barrier impairment.17

2.6. Histological and immunohistological analyses

The collected tissue samples were fixed in 4% paraformaldehyde in phosphate buffer (pH 7.4), dehydrated with series of ethanol, cleansed with series of xylene, and then embedded in paraffin. Approximately 4 μm‐thick paraffin sections were used for HE staining to observe the histological structure and also for immunohistochemistry for FITC to observe the distribution of FITC‐dextran.

For the immunohistochemistry analysis, endogenous peroxidase activity in the sections was inactivated via incubation in 0.3% hydrogen peroxide/methanol for 30 minute. Antigen retrieval via autoclaving at 121°C for 15 minutes in 10 mM citrate buffer (pH 6.0) was performed, followed by incubation with HRP‐conjugated anti‐FITC antibody (diluted 1:200, 60 min), visualisation with DAB, and counterstaining with haematoxylin. The sections were observed using a microscope (BZ‐X700; KEYENCE Corp., Osaka, Japan).

Some HE‐stained tissue sections showed erythrocyte leakage from the blood vessels. In such cases, the distance from the skin surface to the deepest area of erythrocyte leakage (d, black arrows in Figure 1G and I) was measured with ImageJ software (NIH, Bethesda, MD, USA).

Figure 1.

Figure 1

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Histological findings of tissues affected by maceration and/or enzymes ex vivo. In the NT (A), M (B), P (C), L (D), PL (E), and LM (F) groups, no histological damage was observed, and erythrocytes (intense eosin staining) were observed inside the capillary blood vessels identified in the microvascular endothelium (haematoxylin staining), indicating intact blood vessels (black arrows). In the PM (G, H) and PLM (I, J) groups, erythrocytes leaked outside the capillary blood vessels in the papillary dermis (yellow arrowheads), indicating slight bleeding. The erythrocyte leakage from the capillary blood vessels was observed in a deeper area of the dermis in the PLM (I, J) group than in the PM (G, H) group. Black arrows in G and I show the distances from the skin surface to the deepest area of erythrocyte leakage (G, I). Each panel shows a representative experiment of 5 replicates. E: epidermis, D: dermis, bar = 100 μm

2.6.1. Statistical analysis

Differences in the depth of erythrocyte leakage were analysed using Student's t‐test. For the analysis of the differences of skin hydration and TEWL, multiple comparisons were performed by Tukey‐Kramer test. P values <.05 were considered statistically significant. Data were reported as means ± standard deviations (SDs).

3. RESULTS

3.1. Influence of lipidolytic enzymes on the tissue structure ex vivo

The ex vivo experiment was conducted to determine whether the lipidolytic enzymes cause tissue structural alterations and whether lipidolytic enzymes treated with skin maceration and/or proteases accelerate such alterations.

Figure 1 shows the histological findings of the ex vivo experiment. In the NT, M, P, L, PL, and LM groups, no histological damage was observed, and erythrocytes (intense eosin staining) were observed inside the capillary blood vessels identified in the microvascular endothelium (haematoxylin staining), indicating intact blood vessels (Figure 1, black arrows). In the PM and PLM groups, erythrocytes leaked outside the capillary blood vessels in the papillary dermis, indicating slight bleeding (Figure 1, yellow arrowheads). Distances from the skin surface to regions with erythrocyte leakage in the PLM group (221.2 ± 20.1 μm) were significantly greater than those in the PM group (119.3 ± 15.0 μm, P < .01).

3.2. Influence of lipidolytic enzymes in skin maceration on the tissue structure and skin barrier function in vivo

The in vivo experiment was conducted to determine whether the lipidolytic enzymes cause tissue structural alterations, whether such enzymes accelerate the skin barrier impairment caused by skin maceration, and whether the combination of lipidolytic enzymes and proteases accelerate the histological changes in macerated skin compared with proteases only.

The skin appearance (Figure 2) did not change 30 minutes after skin maceration treatment in all groups. Twenty‐four hours after skin maceration treatment, the NT, M, PM, and LM groups did not show any change in the skin appearance, but dot‐like redness was slightly observed in the PLM group (Figure 2, black arrowheads).

Figure 2.

Figure 2

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Macroscopic findings of rat dorsal skin affected by maceration with/without enzymes in vivo. The skin appearance did not change 30 minutes after skin maceration treatment in all groups. Twenty‐four hours after skin maceration treatment, dot‐like redness was slightly observed in the PLM group (black arrowheads), although the NT, M, PM, and LM groups did not show any change in the skin appearance. Each panel shows a representative experiment of 4 replicates

The skin hydration level, the indicator of skin maceration, and TEWL, the indicator of skin barrier impairment, were measured 30 minutes after removal of the agarose gels (Figure 3). Enzyme‐associated maceration tended to increase the levels of skin hydration (NT group, 17.8% ± 7.8%; M group, 22.2% ± 4.8%; PM group, 26.1% ± 5.7%; LM group, 29.1% ± 5.2%; PLM group, 29.3% ± 5.6%). There were no significant differences in skin hydration levels among M, PM, LM, and PLM groups. The TEWL value in the enzyme‐associated maceration significantly increased in the M group (33.8 ± 9.9 g/m2 h, P = .003), LM group (43.2 ± 19.0 g/m2 h, P = .01), and PLM group (56.8 ± 31.7 g/m2 h, P = .01) compared with that of the NT group (10.8 ± 2.8 g/m2 h). The PM group did not show a significant increase in TEWL, but a tendency to increase, compared with the NT group, was observed (PM group, 44.5 ± 30.6 g/m2 h, P = .06). There were no significant differences in TEWL levels among M, PM, LM, and PLM groups.

Figure 3.

Figure 3

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Skin hydration and transepidermal water loss (TEWL) levels on skin affected by maceration with/without enzymes in vivo. Skin hydration level, the indicator of skin maceration, and TEWL, the indicator of skin barrier impairment, were measured 30 minutes after removal of the agarose gels on each treatment area in 5 rats. For the analysis of the differences, a multiple comparison was performed by Tukey‐Kramer test. Enzyme‐associated maceration tended to increase the levels of skin hydration. The TEWL value in the enzyme‐associated maceration significantly increased in the M, LM, and PLM groups compared with the NT group. The PM group did not show a significant increase in TEWL, but a tendency to increase, compared with the NT group, was observed. * P < .05, ** P < .01

The histological analysis revealed that erythrocytes leaked outside the capillary blood vessels not only in the papillary dermis but also around the hair follicles (Figure 4, yellow arrowheads) in the PM and PLM groups. Distances from the skin surface to regions of erythrocyte leakage in the PLM group (251.5 ± 32.5 μm) tended to be greater than those in the PM group (207.4 ± 39.7 μm, P = .09). In the other groups (NT, M, and LM), no histological damage was observed, and intact blood vessels were observed in the papillary dermis and around the hair follicles. These results were consistent with the histological findings in the ex vivo examination.

Figure 4.

Figure 4

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Histological findings of tissues affected by maceration with/without enzymes in vivo. Histology was examined around papillary dermis (NT: A; M: D; LM: Q; PM: J, K; PLM: N, O) and around hair follicles (NT: B, C; M: E, F; LM: H, I; PM: L, M; PLM: P, Q). Erythrocytes (yellow arrowheads) leaked outside the capillary blood vessels not only in the papillary dermis but also around the hair follicles in the PM and PLM groups. In the other groups (NT, M, and LM), no histological damage was observed, and intact blood vessels were observed in the papillary dermis and around the hair follicles. These results were consistent with the histological findings in the ex vivo examination. Each panel shows a representative experiment of 4 replicates. E: epidermis, D: dermis, bar = 100 μm

The penetration of FITC‐dextran was visualised via immunohistochemistry for FITC (Figure 5). FITC‐dextran was distributed in the entire epidermis in the LM, PM, and PLM groups. The LM group especially showed a higher intensity of FITC‐positive signals in the epidermis, and the PLM group showed FITC‐positive signals in the epidermis more slightly than the LM and PM groups.

Figure 5.

Figure 5

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Distribution of FITC‐dextran in tissues affected by maceration with/without enzymes in vivo. Distribution of FITC‐dextran in tissues was examined in NT (A), M (B), LM (C), PM (D), and PLM (E) groups. No FITC‐dextran positive signal was observed in the epidermis of the NT (A) and M (B) groups. FITC‐dextran was distributed in the entire epidermis in the LM (C), PM (D), and PLM (E) groups. The LM group (C) especially showed a higher intensity of FITC‐positive signals in the epidermis, and the PLM group (E) showed FITC‐positive signals in the epidermis more slightly than the LM and PM groups. Each panel shows a representative experiment of 4 replicates. E: epidermis, D: dermis, bar = 100 μm

4. DISCUSSION

In this experiment, macerated skin, enzyme‐affected skin, and enzyme‐affected macerated skin were reproduced in ex vivo and in vivo rat skin. This is the first attempt to determine the influence of lipidolytic enzymes on the tissue structure and skin barrier function under the condition of skin maceration or proteolytic skin maceration.

From the findings of the ex vivo experiments, the enzymes caused erythrocyte leakage in the dermis when the skin was treated with enzymes with skin maceration. The skin treatment with enzymes only did not cause any histological change, including erythrocyte leakage. It is suggested that erythrocytes leak outside the capillary blood vessels when the transdermally penetrated enzymes degrade the blood vessel wall. Therefore, the erythrocyte leakage is considered one of the indicators of tissue damage. Both the ex vivo and in vivo experiments showed that skin treatments, including both of the skin maceration and proteases application (PM and PLM), caused erythrocyte leakage. It is speculated that skin barrier impairment caused by skin maceration accelerated the transdermal penetration of enzymes, and transdermally penetrated proteases degraded the blood vessels, which resulted in erythrocyte leakage.

Erythrocyte leakage was observed in a larger area in the PLM groups than in the PM groups, that is, the addition of lipidolytic enzymes to skin maceration with proteases enhanced erythrocyte leakage. One possibility is that the lipidolytic enzymes accelerate the damage in the blood vessels by their ability to degrade the lipids in the cell membrane. However, skin maceration and lipidolytic enzymes application (LM) did not cause bleeding. These results indicate the possibility that lipidolytic enzymes are the factors that accelerate the transdermal penetration of proteases via the disruption of the intercellular lipid lamellae.

In the in vivo experiment, skin physiological function, including the levels of skin maceration and skin barrier function, was examined after skin maceration treatment. The level of skin maceration did not show a significant increase in the enzyme‐affected skin maceration compared with that in the skin maceration only. Skin barrier function was evaluated by TEWL levels and penetration of FITC‐dextran. Although the level of TEWL did not show a significant increase in the enzyme‐affected skin maceration compared with that in the skin maceration only, the degree of FITC‐positive signals in the epidermis increased in the enzyme‐affected skin maceration compared with that in the skin maceration only. These results indicate the possibility that proteases and lipidolytic enzymes enhance skin barrier impairment in macerated skin. Skin barrier functions in normal healthy skin include tight junctions in the granular layer of the epidermis, not only the intercellular lipid lamellae in the stratum corneum. Lipidolytic enzymes can disrupt the intercellular lipid lamellae, whereas proteases can degrade the junctional protein between keratinocytes in the epidermis. Considering the different influence of each enzyme on skin barrier function, it is suggested that FITC‐dextran penetrated through the stratum corneum barrier and stayed in the epidermis in the LM group. Skin treatment of PLM is considered to allow FITC‐dextran to penetrate not only to the stratum corneum barrier but also to the epidermal barrier; this resulted in the finding that the FITC‐positive signals in the epidermis were slightly higher in the LM group than in the PLM group. Although the distribution of FITC‐dextran in the dermis could not be observed from the result of immunohistochemistry analysis for FITC in this study, FITC‐dextran is supposed to penetrate the dermis in the PLM group.

There were some differences between the results of the PM group in the previous study and those in this research.25 In our previous study, the appearance of PM skin showed dot‐like redness 24 hours after agarose gel removal; however, no macroscopic change was observed on the PM skin in this study. Slight increases in the skin hydration level and TEWL value were observed in this study compared with those in the previous study. In the previous study, the PM treatment increased the skin hydration level by twice that of the NT and the TEWL value by 5 times that of the NT. In this study, the PM treatment increased the skin hydration level by 1.5 times that of the NT and the TEWL value by 4.5 times that of the NT. The reason why the influence of the PM treatment was milder in this study compared with that in the previous study may be because of the difference in the skin maceration protocols. In the previous report, the PM treatment was performed via application of an agarose gel containing proteolytic solution for 4 hours. In this study, the FITC‐dextran‐containing agarose gel was first applied for 30 minutes, followed by the application of the agarose gel containing proteolytic solution for 3 hours and 30 minutes. The application time of the protease‐containing agarose gel was shorter than that in the previous report, which may result in milder reactions to the PM treatment. Although skin changes were not observed, erythrocyte leakage in the dermis occurred based on the histology examination results; therefore, it is suggested that the PM treatment was performed appropriately in this study.

There were several limitations in this study. The IAD model in this study was reproduced by a 1‐time application of water and enzymes. In the health care setting, repeated exposure to faeces is common among patients with incontinence, which is considered to result in more severe IAD and delayed healing owing to exposure to water and enzymes during the healing process. Moreover, the concentrations of each enzyme in the IAD reproduction treatment were as high as those in the solid faeces of healthy patients.19, 20 Because diarrhoea stool contains digestive enzymes in higher concentrations than solid feces1, the perineal skin of actual patients with faecal incontinence may be exposed to greater loads of enzymes than in our experimental conditions; such conditions might also result in more severe IAD than in the IAD model in this study. Additional examinations are required to examine the influence of repeated exposure to faeces and greater loads of enzymes on the pathophysiology of IAD.

5. CONCLUSION

To determine the influence of lipidolytic enzymes on IAD development, tissue structural alteration and skin barrier impairment caused by lipidolytic enzymes were examined. Lipidolytic enzymes did not cause tissue structural alterations but were considered to accelerate the transdermal penetration of proteases in the macerated skin by the impairment of skin barrier function, which enhanced tissue damage in the macerated skin. Lipidolytic enzymes in macerated skin are factors that accelerate of IAD development via skin barrier impairment, and proteases are the factors that trigger IAD development via tissue damage. These results indicate that advanced nursing care of perineal skin in patients with faecal incontinence is required because of the deleterious influence of lipidolytic enzymes and proteases.

ACKNOWLEDGEMENTS

This work was funded by a Grant‐in‐Aid for Fellows of Japan Society for the Promotion of Science (#24‐3913). The Japan Society for the Promotion of Science had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Mugita Y, Minematsu T, Nakagami G, Sanada H. Influence of digestive enzymes on development of incontinence‐associated dermatitis: Inner tissue damage and skin barrier impairment caused by lipidolytic enzymes and proteases in rat macerated skin. Int Wound J. 2018;15:623–632. 10.1111/iwj.12906

Funding information Japan Society for the Promotion of Science, Grant/Award number: 24‐3913

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