Rooster feathering, androgenic alopecia, and hormone-dependent tumor growth: what is in common? - PubMed (original) (raw)
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Rooster feathering, androgenic alopecia, and hormone-dependent tumor growth: what is in common?
Julie Ann Mayer et al. Differentiation. 2004 Dec.
Abstract
Different epithelial organs form as a result of epithelial-mesenchymal interactions and share a common theme modulated by variations (Chuong ed. In Molecular Basis of Epithelial Appendage Morphogenesis, 1998). One of the major modulators is the sex hormone pathway that acts on the prototype signaling pathway to alter organ phenotypes. Here, we focus on how the sex hormone pathway may interface with epithelia morphogenesis-related signaling pathways. We first survey these sex hormone-regulated morphogenetic processes in various epithelial organs. Sexual dimorphism of hairs and feathers has implications in sexual selection. Diseases of these pathways result in androgenic alopecia, hirsutism, henny feathering, etc. The growth and development of mammary glands, prostate glands, and external genitalia essential for reproductive function are also dependent on sex hormones. Diseases affecting these organs include congenital anomalies and hormone-dependent breast and prostate cancers. To study the role of sex hormones in new growth in the context of system biology/pathology, an in vivo model in which organ formation starts from stem cells is essential. With recent developments (Yu et al. (2002) The morphogenesis of feathers. Nature 420:308-312), the growth of tail feathers in roosters and hens has become a testable model in which experimental manipulations are possible. We show exemplary data of differences in their growth rate, proliferative cell population, and signaling molecule expression. Working hypotheses are proposed on how the sex hormone pathways may interact with growth pathways. It is now possible to test these hypotheses using the chicken model to learn fundamental mechanisms on how sex hormones affect organogenesis, epithelial organ cycling, and growth-related tumorigenesis.
Figures
Fig. 1
Male and female adult chickens. A) Photo of a male white leghorn. B) Photo of a female white leghorn. C) Left: male tail feather. Right: female tail feather. Both are from the midline.
Fig. 2
Hormone biosynthetic pathway. Enzyme names are indicated in green. Products are indicated in black.
Fig. 3
How does the growth of male tail feathers differ from female tail feathers? Do they a) grow at different rates or b) grow at the same rate for different durations? Black, male; gray, female.
Fig. 4
Growth kinetics of male and female tail feathers. Regenerative tail feather length after tail feathers were plucked (time 0). Note the male and female feathers grow at similar rates, but the male tail feathers grow for a longer period of time. The increased standard error detected in the male growth curves at later time points are attributed to differences in the growth period between the center (which grows longest) and lateral tail feathers.
Fig. 5
Schematic of feather follicle structure. Feathers initially form as a cylinder. The mesenchymal structures (shades of red) are the dermal papilla (dp) and the pulp. Blood vessels (bv) bring nutrients to the growing feather follicles. The epithelia (shades of blue) will form the feather. The epithelial localized growth zone (LoGZ) is near the base of the feather above the dermal papilla. Feather branching into barbs takes place in the differentiating ramogenic zone (rz). After differentiation, the pulp cells die, enabling the feather branches to open up.
Fig. 6
Molecular expression in male and female tail feathers. Longitudinal male and female tail feather sections (in regeneration) were stained for H&E, PCNA, Feather Keratin A (Ker A), Cytokeratin II (Cyto II) and β-catenin (Beta-Cat). The distributions of PCNA and β-catenin protein were determined using immunostaining. The distribution of Feather Keratin A and Cytokeratin II were determined using in situ hybridization. A high power view of the feather collar region is shown (H &E, PCNA, and Beta-Cat). Since there was no staining in the collar region for Ker A, the ramogenic zone is shown for Ker A and Cyto II with size bars = 250µm. A lower power view is shown in the upper insets with size bars = 0.5 mm and a higher power view is shown in the lower insets with size bars = 200µm.
Fig. 7
Morphological difference in male and female tail feathers. H&E staining of male (A) and female (B) feathers at the same height up from the base reveals differences in diameter, rachis width, and barb density. The barb density is distributed as a gradient in male rectrices but is constant in female feathers as seen in scanning EM photographs. In males the density is lowest near the distal tip (compare C, D), moderate in the middle (compare E, F) and highest at the base (compare G, H) as shown in scanning EM photographs. Overall, the rachis is wider at the proximal base of tail feathers, but tapers toward the distal end; however, the width of the male is larger than the width of the female feathers at equivalent locations. These differences are shown schematically for male (I) and female (J) tail feathers.
Fig. 8
Possible models of hormone effects. Model A, the body (contour) feather is the default type. Sex hormones influence whether a “male” or “female” feather will develop. Model B, the “female” is the default feather type. Androgens cause a transition to a “male” feather type. Model C, the “male” feather is the default and estrogen converts the feather to a “female” feather type.
Fig. 9
Sex hormones influence on feather diameter and length. The dermal papilla (red), LoGZ (light blue) and differentiation zone (DZ, or ramogenic zone, dark blue) are shown.
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