Morphological study of the integument and corporal skeletal muscles of two psammophilous members of Scincidae (Scincus scincus and Eumeces schneideri)

Abstract

Sand deserts are common biotopes on the earth's surface. Numerous morphological and physiological adaptations have appeared to cope with the peculiar conditions imposed by sandy substrates, such as abrasion, mechanical resistance and the potential low oxygen levels. The psammophilous scincids (Lepidosauria) Scincus scincus and Eumeces schneideri are among those. S. scincus is a species frequently used to study displacement inside a sandy substrate. E. schneideri is a species phylogenetically closely related to S. scincus with a similar lifestyle. The aims of this study focus on the morphology of the integument and the muscular system. Briefly, we describe interspecific differences at the superficial architecture of the scales pattern and the thickness of the integument. We highlight a high cellular turnover rate at the level of the basal germinal layer of the epidermis, which, we suggest, corresponds to an adaptation to cutaneous wear caused by abrasion. We demonstrate the presence of numerous cutaneous holocrine glands whose secretion probably plays a role in the flow of sand along the integument. Several strata of osteoderms strengthen the skin. We characterize the corporal (M. longissimus dorsi and M. rectus abdominus) and caudal muscular fibers using immunohistochemistry, and quantify them using morphometry. The musculature exhibits a high proportion of glycolytic fast fibers that allow rapid burying and are well adapted to this mechanically resistant and oxygen-poor substrate. Oxidative slow fibers are low in abundance, less than 10% in S. scincus, but a little higher in E. schneideri.

Keywords: immunohistochemistry; integument; muscle fibers; sand dwelling; skinks.

FIGURE 1

Dorsal views of Scincus scincus (a) and Eumeces schneideri (b)

FIGURE 2

Integumentary morphology of Scincus scincus (left columns a–d) and Eumeces schneideri (right column e–h). All tissue sections were stained with Masson's Trichrome. (a,e) Low magnification images are from the body's dorsal region, whereas (b and f) illustrate sections from the body's ventral region. Numerous osteoderm layers, stained in orange, are present in the dermis of both species. Successive rigid osteoderm plates are articulated by hinges consisting of dense connective tissue. High magnification images (c and g) depict the epidermal layer of both species. The superficial alpha and beta keratin layers constituting the scale are clearly visible on (c), whereas (g) illustrates the presence of melanocytes in the superficial dermal stratum laxum. (d,h) The presence of holocrine glands between the folds of the skin superficial layers of holocrine glands in the junction between the folds of the skin superficial layers

FIGURE 3

(a) Boxplots representing the absolute thickness (in μm) of the body integument (epidermal and dermal layers including osteodermic stratum) of Eumeces schneideri and Scincus scincus. (b) Similar boxplots representing the skin in absolute thickness (in μm) but assessed separately for the dorsal and ventral regions of E. schneideri and S. scincus. (c) Boxplots representing the relative thickness of the body integument (epidermal and dermal layers including osteodermic strates) of Eumeces schneideri and Scincus scincus. These relative values were obtained from absolute thicknesses values corrected relating to differences in overall body size between the species. Data were analyzed using the nonparametric Wilcoxon test (for (a and c)) and Kruskal–Wallis test (for (b)) and significant difference was set at a p‐value <.01

FIGURE 4

Scanning electron micrographs of scales of Scincus scincus (left column a–c) and Eumeces schneideri (right column d–f). (a,b) Low magnification images illustrate the imbrication of the dorsal body scales of both species. Images (b and e), taken at high magnification, indicate the micro relief present on the surface of dorsal scales, whereas (c and f) focus on the microridges of the ventral scales. The white arrows indicate microridges

FIGURE 5

Boxplots representing the inter microridge spaces (in μm) of the scales of Eumeces schneideri and Scincus scincus. The measurements were taken from 9 scales (30 microridge values per body region) selected at random from the ventral and dorsal regions of both species. Data were analyzed using the nonparametric Kruskal–Wallis test and significant differences were set at a p‐value <.01

FIGURE 6

Immunohistochemical detection of dividing cells in the epidermal layer of Scincus scincus (a,b) and Eumeces schneideri (c,d). Anti‐proliferating cell nuclear antigen (PCNA) labeled cells are identified by a brown color in the nucleus (arrows)

FIGURE 7

Number of epidermal dividing cells labeled with the anti‐proliferating cell nuclear antigen (PCNA) antibody per microscopic field of 0.084 mm² for Eumeces schneideri and Scincus scincus, respectively. For each species (n = 3), 16 microscopic fields were selected at random from the head to the tail of each lizard. Data were pooled per species and presented as boxplots. Statistical analysis was performed using the nonparametric Wilcoxon test (significant differences was set at a p‐value <.01). The proliferation index is calculated by the ratio: number of PCNA positive nuclei per field/number of total nuclei per field and correspond to 37.7% for Scincus scincus and 64.7% for Eumeces schneideri

FIGURE 8

Illustration of the immunohistochemical method used for the typing of muscle fibers. The panel of four pictures corresponds to immunohistochemical labeling performed on serial sections of the muscles of Scincus scincus (a,b) and Eumeces schneideri (c,d), respectively. For each species, fast myosin was detected by a specific antibody (MY32 [Abcam]) in the first section (a,c) whereas slow muscle fibers were indicated in the following sections (b,d) by immunolabeling with specific anti‐slow myosin antibodies (NOQ 7.5.4D [Abcam]). On the left the pictures (a,c), correspond to the first section treated with antibodies raised against fast myosin, in which fast muscle fibers appear deeply stained in brown, intermediate fibers lightly stained in brown and slow muscle fibers negative (blue). In contrast, in the following sections (b,d), immunostained with antibodies raised against anti‐slow myosin only, slow muscle fibers are labeled in deep brown whereas intermediate and fast fibers are totally negative (blue color). The comparison between the consecutive sections (a vs. b and c vs. d) attests to the specificity of the immunolabeling and the clear‐cut identification of the three muscle fiber types