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Sulfur mustard (HD) is an alkylating and cytotoxic chemical warfare agent, which inflicts severe skin toxicity and an inflammatory response. Effective medical countermeasures against HD-caused skin toxicity are lacking due to limited knowledge of related mechanisms, which is mainly attributed to the requirement of more applicable and efficient animal skin toxicity models. Using a less toxic analog of HD, chloroethyl ethyl sulfide (CEES), we identified quantifiable inflammatory biomarkers of CEES-induced skin injury in dose- (0.05–2 mg) and time- (3–168 h) response experiments, and developed a CEES-induced skin toxicity SKH-1 hairless mouse model. Topical CEES treatment at high doses caused a significant dose-dependent increase in skin bi-fold thickness indicating edema. Histopathological evaluation of CEES-treated skin sections revealed increases in epidermal and dermal thickness, number of pyknotic basal keratinocytes, dermal capillaries, neutrophils, macrophages, mast cells, and desquamation of epidermis. CEES-induced dose-dependent increases in epidermal cell apoptosis and basal cell proliferation were demonstrated by the terminal deoxynucleotidyl transferase (tdt)-mediated dUTP-biotin nick end labeling and proliferative cell nuclear antigen stainings, respectively. Following an increase in the mast cells, myeloperoxidase activity in the inflamed skin peaked at 24 h after CEES exposure coinciding with neutrophil infiltration. F4/80 staining of skin integuments revealed an increase in the number of macrophages after 24 h of CEES exposure. In conclusion, these results establish CEES-induced quantifiable inflammatory biomarkers in a more applicable and efficient SKH-1 hairless mouse model, which could be valuable for agent efficacy studies to develop potential prophylactic and therapeutic interventions for HD-induced skin toxicity.
Sulfur mustard (bis (2-chloroethyl) sulfide or HD) is a major chemical warfare agent that was used during World War I and has since been used in a number of conflicts including the Iraq-Iran war in 1988. It continues to be a major threat for use in battlegrounds and terrorist actions against civilian and military targets due to its easy accessibility, inexpensive manufacture and storage, and lack of effective countermeasures against its toxicity (Noort et al., 2002; Saladi et al., 2006). HD has many debilitating effects including ocular and dermal injury, respiratory tract damage, reproductive and developmental toxicity, and gastrointestinal and hematological effects (Graham et al., 2005). HD, a potent vesicant agent, causes inflammation and extensive blistering of the skin, an important primary target organ because of its large surface area and sensitivity of frequently dividing basal cells (Dacre and Goldman, 1996; Graham et al., 2005; Wormser, 1991). Integument inflammation is manifested as erythema and edema which progresses to blister formation, ulceration, necrosis and desquamation of the epidermis that can lead to permanent residual effects in humans (Balali-Mood and Hefazi, 2006; Balali-Mood et al., 2005; Blaha et al., 2000b; Greenberg et al., 2006; Sabourin et al., 2002; Smith et al., 1996, 1997; Wormser et al., 2005). Histopathology of the skin following exposure to HD is characterized by edema, dermal infiltration of inflammatory cells, premature death of basal layer epidermal cells, and epidermal-dermal separation (Smith et al., 1995, 1998; Vogt et al., 1984).
Despite limitations, in vivo animal models, critical for investigating HD-caused skin toxicity in humans, have been useful in studying its pathogenesis. Studies conducted in different animal models suggest that the major event after HD exposure to skin is an inflammatory response indicated by skin edema, production of inflammatory cytokines such as tumor necrosis factor α (TNF-α) and interleukins (IL-8, IL-6, IL-1α, IL-1β), activation of nuclear transcription factor kappa B (NF-κB), induction of metalloproteinase-9 (MMP-9), and damage of mitochondria and cell nuclei together with cell death by apoptosis or necrosis (Arroyo et al., 2001; Blaha et al., 2000a,b; Dillman et al., 2004; Graham et al., 2005; Isidore et al., 2007; Kan et al., 2003; Paromov et al., 2007; Ricketts et al., 2000; Rogers et al., 2004; Rosenthal et al., 1998; Sabourin et al., 2000, 2002; Shakarjian et al., 2006; Wormser et al., 2005). Although, these studies reflect on the possible cellular mechanisms that could be involved in HD-caused skin inflammatory response, a complete knowledge of these mechanisms related to its dose- and time-related toxic response is lacking, due to the requirement of more applicable and efficient animal skin toxicity models. Several studies have focused on developing weanling pig, hairless guinea pig, rabbit, or various mouse species as useful animal models of HD-induced skin toxicity to investigate its pathogenesis, and find effective countermeasures that are not available to date (Blaha et al., 2000a,b; Brodsky et al., 2006; Cowan et al., 2003; Gross et al., 2006; Isidore et al., 2007; Paromov et al., 2007; Simpson and Lindsay, 2005). However, furred and larger animal models are inefficient and costly, and quantitative biomarkers reflecting inflammation and related pathogenic indicators of chloroethyl ethyl sulfide (CEES)–induced skin injury in simple and efficient rodent models are limited (Henemyre-Harris et al., 2008; Isidore et al., 2007; Paromov et al., 2007)
The major aims of the present study, therefore, were to establish CEES (HD analog)-induced quantifiable inflammatory biomarkers, and develop an efficient, and reliable experimental rodent model that is more relevant to human skin toxicity, and can be utilized to assess CEES-induced skin toxicity and pathogenesis. SKH-1 hairless mouse is a widely used experimental rodent model for dermal research, safety and efficacy testing, especially in the UV-induced skin damage (Anwar et al., 2008; Bell et al., 2007). CEES, a commercially available and widely used experimental alternative to HD due to its similar chemical properties, was employed in the present study (Han et al., 2004). Our study outcomes establish important quantifiable inflammatory biomarkers of CEES-induced skin toxicity in a more resourceful SKH-1 hairless mouse model that could help in understanding the related histopathological mechanisms. Our investigation suggests that most of the CEES-induced inflammatory responses peak from or after 9 h and begin to subside within 72–168 h of postexposure, and a time-dependent association of edema with the recruitment of inflammatory cells. Moreover, these inflammatory biomarkers and SKH-1 mouse model could be practically valuable for agent efficacy studies to eventually develop effective prophylactic and therapeutic countermeasures against HD-induced skin toxicity.
CEES was purchased from Sigma-Aldrich Chemical Co. (St Louis, MO) and dissolved in high-performance liquid chromatography grade acetone for animal treatment. Female SKH-1 hairless mice (4–5 weeks of age) were obtained from Charles River Laboratories (Wilmington, MA), and housed under standard conditions at the Center of Laboratory Animal Care, University of Colorado Denver, CO. The animals were acclimatized for one week before their use in experimental studies that were carried out according to the specified protocol approved by the IACUC of the University of Colorado Denver, CO.
To assess CEES-induced skin tissue injury, we conducted both dose- and time-response studies. In the dose study, CEES doses in the range of 0.05–2 mg in 200 μl of acetone/mouse were applied topically on to the dorsal skin as 0.5–6 mg/kg/animal CEES has been used in experiments demonstrating its toxic responses in the skin tissue (Chatterjee et al., 2004). Acetone, used as a vehicle here enhances the skin permeability to hydrophilic and amphipathic compounds, and has been successfully used as a vehicle for most of the skin topical treatments (Dhanalakshmi et al., 2004; Tsai et al., 2001). The blistering dose of HD (more toxic as compared with CEES) that caused acute effects in humans is reported to be about 40–100 μg/cm2 and 0.1–4.3 mg/m3 (Kehe et al., 2008). Briefly, a total of 50 mice were randomly divided into 10 groups of five mice each; different groups were (1) control-untreated, (2) 200 μl of acetone alone/mouse (vehicle control), (3) 0.05 mg CEES, (4) 0.1 mg CEES, (5) 0.25 mg CEES, (6) 0.4 mg CEES, (7) 0.5 mg CEES, (8) 1 mg CEES, (9) 1.5 mg CEES, and (10) 2 mg CEES. The time-response study employed 1 and 2 mg CEES doses, which were based on the results from the dose-response study, and the study time points were 3, 6, 9, 12, 24, 48, 72, and 168 h. Each treatment dose and time point had five mice each. At the end of each desired treatment, the mice were euthanized using CO2, and the dorsal skin was collected and either snap frozen using liquid nitrogen or fixed in formalin unless otherwise stated. The dorsal skin harvested was equidistance from the smallest part of the neck and the base of the tail and parallel to the vertebral line (Paus et al., 1999).
In the dose-response study, the dorsal skin bi-fold thickness measurements (mm) were taken at 6, 9, and 12 h after the control and CEES treatment using an electronic digital caliper (Marathon, Inc., Belleville, ON, Canada). In the time-response study, the skin bi-fold thickness was measured at various intervals between 3 and 168 h after control and CEES treatment. For edema measurements, the dorsal skin samples collected in the dose- and time-response studies were adhered, stretched on a foil and 10-mm punch biopsies were taken from upper half of the dissected skin parallel to the spine (Paus et al., 1999). The punch biopsies were weighed, and oven dried at 37°C for 24 h. After drying, the biopsies were reweighed and the wet/dry weight ratios were calculated.
The integument was fixed in 10% phosphate-buffered formalin, dehydrated in ascending concentrations of ethanol, cleared with xylene, and embedded in paraffin (Triangle Biomedical Sciences, Durham, NC). Serial sections (5 μm) of the paraffin-embedded tissue blocks were cut with a rotary microtome and processed for hematoxylin and eosin (H&E), and immunohistochemical staining.
Apoptotic cells were identified on H&E stained sections by virtue of their eosinophilic cytoplasm and dark purple pyknotic nuclei, and counted in randomly selected fields (×400 magnifications) of control and CEES-treated skin tissue sections. The apoptotic index was determined as number of apoptosis-positive cells ×100/total number of cells. The epidermal thickness (μm) was measured in H&E stained skin tissue sections randomly in at least five fields per tissue sample under a microscope using Axiovision Rel 4.5 software (Carl Zeiss, Inc., Germany; ×400 magnification).
Apoptotic cells were also detected using the DeadEnd Colorimetric the terminal deoxynucleotidyl transferase (tdt)-mediated dUTP-biotin nick end labeling (TUNEL) system (Promega, Madison, WI) following a modified version of the manufacturer's protocol. In brief, the tissue sections were deparaffinized and rehydrated, and permeabilized with proteinase K (30 μg/ml) for 1 h at 37°C. Endogenous peroxidase activity was quenched using 3% hydrogen peroxide in methanol (vol/vol) for 10 min. After thorough washing with 1× phosphate-buffered saline (PBS), sections were incubated with equilibration buffer for 10 min and then terminal deoxynucleotidyl transferase (tdt) reaction mixture was added and incubated at 37°C for 1 h. Negative controls were included for all sections, these being deficient for the tdt-reaction mixture step. The tdt-reaction step was terminated by immersing the sections in 2× saline sodium citrate buffer for 15 min, treating them with conjugated horseradish peroxidase streptavidin for 30 min at room temperature and, after repeated washings in PBS, incubating them with 3,3′-diaminobenzidine (DAB) for ~10 min to allow brown color development. Sections were mounted after dehydration, and TUNEL-positive cells were quantified in 10 randomly selected fields at ×400 magnification. The apoptotic index was calculated as the number of apoptotic cells ×100 total number of cells.
The paraffin-embedded sections were heat immobilized and deparaffinized using xylene, then rehydrated in a graded series of ethanol with a final wash in distilled water. Antigen retrieval was performed in 10 mmol/l citrate buffer (pH 6.0). Endogenous peroxidase activity was blocked by immersing the sections in 3% hydrogen peroxide in methanol (vol/vol). The sections were then incubated with mouse monoclonal anti-proliferative cell nuclear antigen (PCNA) antibody IgG2a (DAKO, Carpinteria, CA) or BM8 monoclonal F4/80 rat anti-mouse IgG2a antibody (Caltag, Invitrogen, Carlsbad, CA) in PBS for 2 h at 37°C in a humidity chamber. To control for nonspecific staining, negative staining controls were used in which sections were incubated with N-Universal negative control antibody (DAKO) under identical conditions to those used for the specific staining. After incubation with primary antibody, the sections were incubated with the appropriate secondary antibody for 1 h followed by incubation with conjugated horseradish peroxidase streptavidin (DAKO) in PBS for 30 min at room temperature in a humidity chamber. Visualization was accomplished by incubation in DAB working solution for 10 min at room temperature and counterstaining with diluted hematoxylin for 2 min followed by rinsing in Scott's water. Finally, sections were dehydrated and mounted for microscopic observation. The proliferating cells were quantified by counting the DAB-positive nuclei and the total number of cells in 10 randomly selected fields (×400 magnification). The proliferation index was determined as number of PCNA-positive cells ×100/total number of cells. The macrophages were quantified by counting their number per cm2 field in five randomly selected fields (×400 magnification).
The deparaffinized and rehydrated skin sections were immersed in toluidine blue (Sigma-Aldrich Chemical Co. St Louis, MO) working solution (5 ml of toluidine blue + 45 ml of 1% sodium chloride, pH 2.0–2.5) for 2–3 min. Next, the sections were washed in distilled water (three times), dehydrated, cleared in xylene and mounted for microscopic observation. The mast cells were quantified by counting their number per cm2 field in five randomly selected fields (×400 magnification).
The myeloperoxidase (MPO) activity was measured using a Fluorescent MPO Detection Kit from Cell Technology (Mountain View, CA), following vendor's protocol. In brief, approximately 100 mg of clean (with red blood cells removed) skin tissue samples were homogenized in 1× homogenization buffer (provided in kit) with N-ethylmaleimide (Sigma) using polytron PT 10-35 (Kinematica, AG, Bohemia, NY). The homogenates were centrifuged at 8000 × g at 4°C for 20 min and 1 ml of solubilization buffer was added to the pellet after removing the supernatant. This mixture was again homogenized and subjected to two cycles of freezing and thawing followed by centrifugation at 8000 × g for 20 min at 4°C. The supernatants were collected and protein concentration determined by the Lowry method (Bio-Rad DC protein assay kit, Bio-Rad laboratories, Hercules, CA). For the MPO assay, a reaction mixture was prepared by adding detection reagent, 20mM hydrogen peroxide, 0.1–1μM eosinophil peroxidase inhibitor (Sigma), and 1× assay buffer. In a black opaque 96-well plate, 50 μl of prepared sample (or MPO standards) plus 50 μl of reaction mixture were added. The plate was incubated at room temperature in the dark for 60 min, and the fluorescence in each well was measured by a fluorescent plate reader using 530–570 nm excitation and 590–600 nm emission. A standard curve of MPO was prepared by serially diluting MPO in 1× assay buffer with 20mM catalase inhibitor (3-amino-1, 2, 4-triazole). The blank control readings were subtracted from all the sample readings. The MPO activity was determined as mU/μg protein using the MPO standard curve.
The microscopic immunohistochemical and histopathologic analyses were performed using a Zeiss Axioscop 2 microscope (Carl Zeiss, Inc., Germany); photomicrographs were captured by a Carl Zeiss AxioCam MrC5 camera with the Axiovision Rel 4.5 software. The data were analyzed using SigmaStat software version 2.03 (Jandel Scientific Corp., San Raphael, CA) for statistical significance of difference between untreated control group versus the CEES-treated groups. Significance was determined by one-way analysis of variance (one-way ANOVA) followed by the Bonferroni t-test for multiple comparisons. p < 0.05 was considered statistically significant.
A dose-dependent increase in the skin bi-fold thickness was observed 6–12 h after CEES exposure (Fig. 1A). Six hours of exposure to higher CEES doses (1–2 mg) caused a significant (p < 0.001) increase in skin bi-fold thickness reaching 0.79 ± 0.01–1.11 ± 0.1 mm as compared with 0.67 ± 0.02 mm skin thickness in untreated control group (Fig. 1A). After 9 h of CEES exposure, the skin bi-fold thickness increased significantly in a dose-dependent manner and reached a maximum of 1.31 ± 0.02 mm at 2 mg (p < 0.001) dose compared with 0.58 ± 0.02 mm in the control group (Fig. 1A). After 12 h of CEES treatment, the skin bi-fold thickness also increased significantly (p < 0.001) in a dose-dependent manner, reaching a maximum of 1.2 ± 0.1 mm bi-fold thickness (Fig. 1A). There was not a significant increase in skin bi-fold thickness following exposure to lower doses of CEES (0.05–0.25 mg); nor was a significant difference observed in skin bi-fold thickness between untreated control and vehicle control groups at any of the study time points (data not shown). Next we investigated the time dependency of these increases in skin bi-fold thickness, which seemed to cause a biphasic response (Fig. 1B). At 3-h post-CEES application (2 mg dose) the skin bi-fold thickness significantly increased (p < 0.05) as compared with the untreated control group. A second increase was seen at 12-h post-CEES exposure (p < 0.001), and the bi-fold thickness remained elevated until 24 h (Fig. 1B) and then decreased from 48- to 168-h post-CEES exposure. Together, these results suggest that the CEES-induced increase in the skin bi-fold thickness could be used as a quantitative primary marker of skin edema.
Next, using the skin biopsies from the dorsal skin exposed to CEES, we measured skin tissue wet/dry weight as a second primary marker of edema. In the dose-response study (Fig. 1C), a significant increase (p < 0.05) in the wet/dry weight ratio was seen 12-h post-CEES exposure with a 2 mg dose demonstrating marked edema; however, lower CEES doses (0.05–1 mg) did not show a significant increase in wet/dry weight ratio (Fig. 1C). In the time-response study (Fig. 1D) we did not observe a significant increase in the wet/dry weight at any given time point following either 1 or 2 mg CEES treatment. Together, these results indicate that the wet/dry weight of CEES-exposed skin biopsies would not serve as a very reliable marker for CEES-induced skin edema.
Because a dose-dependent increase in the skin bi-fold thickness was detected following CEES exposure indicating edema and inflammation, we sought a histological explanation for this change. Representative H&E stained sections showed an increase in both the epidermal and dermal thickness following CEES exposure as compared with the control (acetone) treated skin samples (Figs. 2A–C). When compared with untreated control skin (Fig. 2D), CEES treatment at 1 and 2 mg doses increased epidermal thickness (Figs. 2E and 2F, respectively). With higher doses of 1.5 (data not shown) and 2 mg CEES, an increase in upper epidermal cell necrosis and desquamation was also noted together with a slight decrease in epidermal thickness as compared with 1 mg CEES dose (Figs. 2E and 2F, red arrows). Apart from an increased epidermal thickness after CEES exposure, an increased cytoplasmic swelling with a concomitant shrinkage or condensation in the nuclei of the epidermal cells was also observed (Fig. 2F, black arrows). In addition, paranuclear clearing was observed in the CEES-treated skin epidermis (Figs. 2E and 2F, red arrowheads). These morphologic changes were suggestive of apoptotic cell death of the epidermal cells. As compared with untreated control (Fig. 2G) and 1 mg CEES-treated skin tissue sections (Fig. 2H), the skin tissue sections from the animals treated with a 2 mg CEES dose also showed an increase in the number of red blood cells in the thickened dermal region (Figs. 2F and 2I, black arrowheads). A further histopathological change seen here and consistent with inflammation was an increased number of mast cells (Figs. 2H and 2I, green arrows); this effect is further demonstrated and quantified, as discussed later.
Due to the presence of pyknotic nuclei in some of the keratinocytes of the H&E stained CEES-exposed skin sections, we quantified the apoptotic cells in the epidermal layer of the skin. CEES exposure of skin for 12 h showed a significant increase in the number of apoptotic cells in a dose-dependent manner (Fig. 3A). Whereas the effect was evidenced at all CEES doses, higher doses (1–2 mg) induced 44–51% apoptotic cells (p < 0.001), as compared with 14% in the untreated control (Fig. 3A). A similar strong increase in apoptotic cells was also observed in the time-response study with maximum apoptotic cells (52 and 61%, p < 0.001) at 72-h post-CEES exposure at 1 and 2 mg doses, respectively (Fig. 3B).
The characteristic pyknotic keratinocytes detected by H&E staining were further examined by immunohistochemical evaluation using TUNEL staining. Qualitative microscopic observation of TUNEL-stained skin sections revealed a substantial increase in the number of TUNEL-positive cells in CEES-treated groups as compared with the untreated controls (Fig. 3C). Quantitative analysis of the TUNEL-stained 12-h CEES-exposed skin sections revealed a dose-dependent increase in the TUNEL-positive cells. Specifically, we observed 30–39% (p < 0.001) TUNEL-positive cells following CEES treatment at 0.5–2 mg doses compared with untreated controls recording 17% TUNEL-positive cells (Fig. 3D). A time-dependent increase in the TUNEL-positive cells in the skin epidermis was also recorded post-CEES exposure (Fig. 3E). Once again, maximum effect was observed at 72 h with 47 and 55% (p < 0.001) TUNEL-positive cells following CEES exposure at 1 and 2 mg doses, respectively (Fig. 3E). These TUNEL staining results confirmed the proapoptotic effect of CEES observed earlier on H&E stained skin sections.
Above we demonstrated that the skin bi-fold thickness increases in response to CEES, and further that H&E stained skin specimens appear to have an increase in the epidermal thickness. Accordingly, next we specifically measured the epidermal thickness to confirm the above observations. After 12 h of CEES exposure (Fig. 4A), a significant dose-dependent increase in the skin epidermal thickness was recorded even at a CEES dose of 0.1 mg (37 ± 1.5 μm; p < 0.05). This effect reached a maximum at 1 mg CEES dose (56 ± 1.3 μm; p < 0.05) before decreasing slightly at 2 mg dose (50 ± 4.4 μm; p < 0.001); the untreated control group measured 18 ± 1.7 μm epidermal thickness (Fig. 4A). In the time-response study (Fig. 4B), we observed a significant increase in the epidermal thickness starting at 3 h (p < 0.01), which peaked at 9 (p < 0.01) and 12 h (p < 0.01) post-CEES exposure at both 1 and 2 mg doses, and then started decreasing between 24- and 168-h post-CEES exposure.
The expression of PCNA, a DNA polymerase-δ, correlates with both cell proliferation and DNA damage repair (Warbrick, 2000). Following our observations showing that CEES causes an increase in skin bi-fold and epidermal (H&E sections) thickness together with skin injury and apoptotic cell death of the epidermal keratinocytes as described above, we hypothesized that CEES causes proliferation in the skin tissue as a mitogenic response as well as a repair response to replace the CEES-induced apoptotic cells. Accordingly, next we assessed cellular proliferation in CEES-exposed skin using PCNA immunostaining. Qualitative microscopic observation of PCNA-stained skin sections revealed a substantial increase in PCNA-positive cells in the basal epidermal layer of the CEES-treated groups (Fig. 4C). Quantification of PCNA-staining after 12 h of CEES exposure, revealed a dose-dependent increase in cell proliferation, which reached a maximum of 25–28% (p < 0.001) at 1–2 mg CEES doses, compared with 10% PCNA-positive cells in untreated controls (Fig. 4D). In the time-response study, within 3 h an increase in PCNA staining was observed following CEES treatments that persisted until 72 h (Fig. 4E).
H&E staining of the sections showed that CEES causes an increase in the number of darker stained vesiculated cells (Fig. 2). Toluidine blue staining confirmed that these were mast cells (Fig. 5A). Quantification of the toluidine blue–stained sections revealed a considerable dose-dependent increase in the number of mast cells in the dermal region of the CEES-treated skin sections as compared with the untreated controls (Fig. 5B). The dose-response results in Figure 5B show 32 ± 2 (p < 0.01), 67 ± 6 (p < 0.001), and 56 ± 3 (p < 0.001) mast cells/cm2 following 0.1, 1, and 2 mg CEES treatments, respectively, compared with 15 ± 1 mast cells in untreated controls. We also observed a time-dependent increase in mast cells that reached a maximum at 12-h post-CEES exposure (Fig. 5C). We hypothesize that the CEES-induced increase mast cell number indicates that these cells are key players in the CEES-caused skin injury and inflammation.
We also measured the MPO activity of neutrophils in the dorsal skin from our CEES-treated mice. Tissue MPO activity is an indicator of the neutrophil infiltration in sites where acute inflammation is present (Bradley et al., 1982). MPO is found in the azurophilic granules of polymorphonuclear leukocytes and is unique to neutrophils and monocytes (Loria et al., 2008). Following 12 h of CEES treatment, skin lysates showed a significant (p < 0.01 to 0.001) dose-dependent increase in the MPO activity at 1–2 mg doses (Fig. 5D). The MPO activity in CEES-treated skin tissue increased in a time-dependent manner from 3- to 24-h post-CEES exposure (Fig. 5E). As a function of treatment time, we observed 1.76 ± 0.24 to 4.3 ± 0.39 and 2.25 ± 0.11 to 5.68 ± 1.2 mU/μg protein activity with 1 and 2 mg CEES doses as compared with 1.11 ± 0.34 mU/μg protein activity in untreated controls, respectively (Fig. 5E). The MPO activity was seen to be significantly (p < 0.001) enhanced and maximal at 24-h post-CEES exposure with 2 mg dose, which thereafter decreased from 48 to 168 h (Fig. 5D). The MPO activity results indicate neutrophil infiltration following CEES-induced skin injury.
Because the CEES-induced skin inflammation contained increases in mast cells and neutrophils, the possible presence of macrophages at the site of injury was also investigated immunohistochemically. We used BM8 monoclonal antibody which reacts with the mouse F4/80 antigen, a macrophage-restricted cell surface glycoprotein (Schaller et al., 2002). Qualitative microscopic observation of F4/80-stained skin sections revealed a substantial increase of macrophages in the CEES-treated group as compared with untreated controls (Fig. 6A). Quantitative analysis of the stained sections in the time-response study (Fig. 6B) showed an increased number of macrophages in the CEES-treated skin tissue from 24 to 168 h compared with the control sections. Macrophages were absent from the skin of the untreated control group, whereas in the vehicle (acetone) control group only a very few were present. The number of macrophages per cm2 reached a maximum of 8.4 ± 1.4 at 1 mg and 14 ± 0.9 at 2 mg CEES doses, 48-h post-CEES exposure compared with 1.8 ± 0.4 macrophages in vehicle controls (Fig. 6B). The number of macrophages per cm2 were 6 ± 1.3 at 1 mg and 9.6 ± 1.6 at 2 mg CEES doses following 24 h of exposure and remained almost constant through 168 h; in all of these groups the number was significantly higher (p < 0.001) when compared with vehicle controls, indicating macrophages do infiltrate CEES-treated skin.
In our comprehensive approach to identify quantifiable and consistent biomarkers of HD-induced skin injury, and establish a more applicable and reproducible animal model for its pathogenesis and agent efficacy studies, the present study identifies important quantifiable inflammatory biomarkers of CEES-induced skin injury in SKH-1 hairless mouse model. HD's exposure to skin induces a marked cutaneous inflammatory response but evaluation of anti-inflammatory therapies have been largely based on qualitative pathological assessments (Dacre and Goldman, 1996; Graham et al., 2005; Paromov et al., 2007; Wormser, 1991). Skin edema, erythema, infiltration of inflammatory mediators, and dilated blood vessels are reported to be important markers of inflammation (Anwar et al., 2008). To study HD's cutaneous response, inefficient, and costly weanling and hairless guinea pigs as well as furred and ear rodent models have been developed (Dachir et al., 2002; Henemyre-Harris et al., 2008; Paromov et al., 2007; Ricketts et al., 2000). Though, the development of cytokine IL-6 as HD-induced inflammatory biomarker in more applicable hairless mice is reported, other inflammatory markers have not been established (Ricketts et al., 2000)
In the present study, using topical application of 0.05–2 mg CEES doses and 3- to 168-h study time points in hairless mice, we documented that CEES causes a measurable increase in the skin bi-fold thickness starting at our earliest study time point of 3-h post-CEES application. We also used skin punch biopsies from untreated control and CEES-exposed mice groups to measure the wet/dry weight ratio, which is an indicator of increased vascular permeability or edema associated with inflammation. Curiously, only 12 h exposure at 2 mg CEES dose, demonstrated a significant increase in edema suggesting that wet/dry weight ratio is not a very sensitive marker. Skin thickness and wet/dry weight measure have been used as important biomarkers for HD-related skin inflammation (Reid et al., 2000; Wormser et al., 2004; Zlotogorski et al., 1997). The present study shows that the measurement of skin bi-fold thickness was a better primary biomarker as compared with the wet/dry weight measure for CEES-caused skin edema.
Studies conducted in various animal models have shown that HD/CEES–exposed skin sections exhibit erythema and edema which progress to blister formation, ulceration, necrosis, desquamation of skin, and epidermal-dermal separation (Blaha et al., 2000b; Graham et al., 2006; Greenberg et al., 2006; Sabourin et al., 2002; Shakarjian et al., 2006; Smith et al., 1996, 1997; Wormser et al., 2005). Our findings show that CEES-induced increase in the bi-fold thickness was associated with increased epidermal and dermal thickness. Quantification of epidermal thickness could serve as a valuable biomarker of CEES-related inflammation. Epidermal thickening increased in a CEES dose-dependent manner that was maximum at 1 mg dose. At a CEES dose of 2 mg, the increased epidermal necrosis and desquamation resulted in some epidermal thinning as compared with lower doses of CEES. Furthermore, at the 12-h time point of maximal change there was a marked increase in epidermal thickness even following treatment with the relatively low 0.1 mg CEES dose. In comparison, the less sensitive bi-fold thickness and wet/dry weight measurements showed significant changes only with higher (0.5–2 mg) CEES doses. This could be due to the inclusion of both epidermal and dermal layers when examining bi-fold thickness and edema. Some previous studies have reported the presence of blisters and epidermal dermal separation after treatment of skin with high levels of exposure to HD (Greenberg et al., 2006; Sabourin et al., 2002; Shakarjian et al., 2006), but in our studies these changes were not conspicuous. The increase in dermal thickening after CEES treatment observed in our investigation could be due to liquid retention in the dermis in association with skin edema.
Apoptosis plays an important role in inflammation and is important to the wound healing process. HD/CEES-induced apoptosis of basal keratinocytes is suggested in reports by several groups (Graham et al., 2006; Kan et al., 2003; Kehe and Szinicz 2005), and was also evident during CEES-induced skin injury in our studies. Histological analysis of CEES-treated skin sections demonstrated that epidermal thickening was associated with cytoplasmic swelling of epidermal cells containing condensed nuclei and with paranuclear clearing of cells, which indicated pyknosis and impending apoptotic cell death. These findings were confirmed by TUNEL analysis, demonstrating a dose-dependent increase in apoptosis, and its time-dependent increase up to 72 h following CEES exposure. We speculate that the CEES-induced increase in apoptosis, observed in our study, is due to the DNA damaging effect of CEES as suggested by various other reports (Graham et al., 2005; Kehe and Szinicz, 2005; Paromov et al., 2007). Cell proliferation is an important compensatory phenomenon that is initiated following apoptosis (Graham et al., 2006). In normal human and mouse epidermis, cells are in constant turnover, where the stem cells divide and differentiate into keratinocytes that ultimately desquamate on the surface of skin; differentiated cells, thereby, are constantly replaced by the proliferating cells from the basal layer (Polakowska et al., 1994). PCNA, a subunit of DNA polymerase, plays a crucial role in DNA replication and damage repair, and serves as a biomarker of proliferation (Gu et al., 2005; Kim and Lee, 2008). PCNA staining used in our study demonstrates a CEES dose- and a time-dependent increase in the PCNA-positive cells. Within 3 h of CEES application, there was an increase in the appearance of PCNA-positive cells; this increase persisted until 72 h. These data suggest that CEES-damaged skin cells initiate DNA repair and cell proliferation very quickly after the CEES insult and continue this response during maximal apoptotic cell generation.
Histological analysis of CEES-treated integument revealed an increase in the number of mast cells and blood vessels in the dermal region. The increase in the number of blood vessels suggests an increase in the blood flow in the CEES-treated skin tissue. The mast cells are key players in response to inflammatory stimuli; their secreted molecules include inflammatory mediators such as histamine, proteases, chemotactic factors, cytokines, etc (Theoharides and Cochrane, 2004). We hypothesize that some of these molecules may be important therapeutic targets in HD-induced skin toxicity. In addition, release of several inflammatory cytokines such as TNFα, interleukins, and related molecules such as transcription factor NF-κB, expression of MMP-9 have been reported following HD treatment to skin (Paromov et al., 2007; Wormser et al., 2005). The data here suggest that further studies are needed to determine and characterize the role of inflammatory mediators that increase early following CEES exposure and prior to other gross tissue damage.
In our study, the CEES-related increase in mast cells was approximately coincident with an infiltration of neutrophils. Histologically, the presence of a neutrophil infiltration has been reported following HD exposure using a mouse ear model (Wormser et al., 2005). We chose to quantify neutrophils by the MPO assay, which has been widely used and validated as a means to quantify neutrophil infiltration in skin (Maruyama et al., 2005). A dose-dependent increase in skin content of MPO activity was observed, which was maximal at 24 h after CEES exposure. The increase in MPO activity coincides with increases in skin bi-fold thickness, apoptotic cell death and the presence of mast cells; all indicative of or associated with inflammation. Following an increase in the mast cells and neutrophils, macrophage numbers increased in the CEES-treated skin and stayed elevated between 24–168 h. The presence of macrophages within the injured skin is expected in order to phagocytose the apoptotic cells and other cell debris generated by CEES-induced skin inflammation.
The process of CEES-induced inflammation and cytotoxicity has also been attributed to its DNA damaging effects (Brodsky et al., 2006; Kehe et al., 2000; Matijasevic et al., 2001). Studies are underway in our laboratory to establish the extent to which DNA damage can be used as molecular marker of CEES-induced skin injury. Thus far, our investigation suggests that most of the CEES-related inflammatory responses peak from or after 9 h and begin to subside within 72–168 h of postexposure in SKH-1 mouse model, indicating that repair mechanisms possibly begin early after CEES exposure that will help in designing of the further mechanistic and efficacy studies.
In summary, we were able to establish increases in the skin bi-fold thickness, epidermal thickness (histological morphometry), apoptotic cell death (histological analysis and TUNEL staining), cell proliferation (PCNA staining), mast cells (toluidine blue staining), neutrophil infiltration (MPO activity), and presence of macrophages (F4/80 staining) as consistent quantifiable primary inflammatory biomarkers of CEES-induced skin injury in a SKH-1 hairless mouse model. These biological markers have been quantitatively developed in a single, efficient, reproducible, and practically valuable rodent model for the first time, and can be utilized to study the molecular mechanism of HD-induced skin cytotoxicity. Furthermore, these inflammatory markers could be efficiently used in therapeutic efficacy studies to develop a new generation of effective prophylactics and therapeutics for HD-induced skin injury.
National Institutes of Health (grant number U54 ES015678).
The study sponsor (NIH) had no involvement in the study design; collection, analysis and interpretation of data; the writing of the manuscript; and the decision to submit the manuscript for publication.
We declare that there are no conflicts of interest.