Title: Development of human skin equivalents mimicking skin aging : contrast between papillary and reticular fibroblasts as a lead

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The handle http://hdl.handle.net/1887/47933 holds various files of this Leiden University dissertation

Author: Janson, David

Title: Development of human skin equivalents mimicking skin aging : contrast between papillary and reticular fibroblasts as a lead

Issue Date: 2017-04-19

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Development of human skin equivalents mimicking skin aging: contrast between papillary

and reticular fibroblasts as a lead

David Janson

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ISBN: 978-9462956247

Printed by Proefschriftmaken.nl || Uitegeverij BOXPress

The research described in this thesis was performed at the Department of Dermatology of the Leiden University Medical Center.

The research was financially supported by CHANEL Parfum Beauté, France.

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Development of human skin equivalents mimicking skin aging:

contrast between papillary and reticular fibroblasts as a lead

Proefschrift

ter verkrijging van

de graad van Doctor aan de Universiteit Leiden, op gezag van Rector Magnificus prof.mr. C.J.J.M. Stolker

volgens besluit van het College voor Promoties te verdedigen op woensdag 19 april 2017

klokke 13:45

Door

David Janson geboren te Stompwijk

2 mei 1986

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Co-promotor Dr. A. El Ghalbzouri

Leden

promotiecommissie

Prof. Dr. M. H. Vermeer

Prof. Dr. J. Schalkwijk

Radboud University, Nijmegen

Prof. Dr. S. Gibbs

VU University Medical Centre, Amsterdam Prof. Dr. J. A. Bouwstra

Leiden Amsterdam Centre for Drug Research, Leiden

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Chapter 1 Introduction

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The study of skin aging can occur on roughly three levels: in cell culture (in vitro), in animal models and in human subjects (in vivo). Most in vitro research on skin aging is performed on monolayer populations of skin cells. These monolayer cultures lack the structural complexity of the skin and can not represent the interactions between different cell types and molecules that the skin consists of. Traditionally, animal models could be used to bridge the gap between the relative simplicity of the culture dish and the in vivo human skin. However, more and more it becomes clear that the skin of most laboratory- animals is not very similar to human skin. In addition, the use of animal experimentation is becoming more restricted for obvious ethical reasons. For example, animal experiments are banned for testing of cosmetic ingredients in the EU (1). This has led to a need for different models in order to translate basic skin aging research to the in vivo human skin.

Complex in vitro tissue models, such as human skin equivalents (HSEs), could be such a model. HSEs are in vitro tissue cultures of skin cells that have a high similarity to in vivo skin morphology and function. However, currently there is no HSE model available that is tailored to represent skin aging in vitro. Therefore, the main goal of this thesis was to contribute to the development of an in vitro HSE model that mimics characteristics of skin aging.

In the following introduction, the skin aging process and its properties will be discussed first. Thereafter, the HSEs will be introduced and their application to skin aging research will be described.

Skin aging

Skin aging is typically divided in two types: intrinsic- and extrinsic skin aging. The most common cosmetic changes caused by skin aging are caused by extrinsic skin aging (2).

These include wrinkle formation, pigmentary irregularities and elastosis. Ultraviolet (UV) light is the most important external cause of extrinsic aging. Other external factors are for example electromagnetic radiation (such as infrared), ozone and tobacco smoke (3).

Intrinsic aging can not be related to any external factors, but is related to the general organismal deterioration with the passage of time. Intrinsic skin aging occurs throughout the entire skin, but due to the dominant effect of external factors, the effects of intrinsic aging on skin are only observed in parts of the body not often exposed to external factors.

These parts are often called unexposed skin, which is based on our habit to cover them with textiles. Characteristics of intrinsically aged skin are for example thin, translucent skin, sagginess, dryness, hair greying and hair loss (4). The focus of this thesis, and therefore also of the rest of this introduction, is on intrinsic skin aging.

Skin aging not only causes cosmetic, but also functional changes. There is a general decline in all skin functions, such as thermo- and water regulation, immune response and

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INTRODUCTION

regenerative capacity (5). This leads to an increase in skin conditions that, although rarely fatal, often impact the quality of life in people affected by these conditions and present a burden on healthcare. The spectrum of disorders of aged skin contains a whole range of morbidities, ranging from idiopathic conditions such as general (or senile) dry skin and itching, impaired wound healing and hypothermia to an increased incidence of specific skin diseases such as dermatitis and eczema (6).

Histology of intrinsic skin aging

The skin consists of two compartments: the epidermis and the dermis. The epidermis is a layer of cells, mainly consisting of keratinocytes, which constantly divide and differentiate, eventually dying and getting sloughed off. It forms a barrier against the outside environment that offers protection from harmful substances, pathogens, UV radiation and dehydration. The dermis consists mostly of extracellular matrix (ECM). It gives structure and elasticity to the skin. The most common cell in the dermis is the fibroblast, which produces and maintains the ECM. In addition, one can find blood vessels and several skin appendages, such as hairs, (smooth) muscles, sebaceous glands and sweat glands in the dermis (7).

Sections of (intrinsically) aged skin are characterized by general atrophy, especially in the dermis. Therefore, the histological picture of aged skin can be summarized quite concisely: the number/volume of most skin components is decreased and the ones that remain look abnormal compared with young skin (3). Since the goal of this thesis is to mimic some changes of aged skin in HSEs, some specific characteristics of aged skin need to be defined. The characteristics of importance to this thesis are discussed below.

The most characteristic property of aged skin is the disappearance of epidermal ridges;

a flattening of the dermal-epidermal junction (8). In contrast to photoaged skin, the interfollicular epidermal thickness is hardly affected in intrinsic aging. The number of proliferating keratinocytes in aged skin is not decreased, but the proliferation speed decreases, which leads to an overall reduction in proliferation (9). A consequence of this is slower epidermal turnover, which in turn could lead to the loss of the rete ridges.

The differentiation process of keratinocytes appears not to be affected by age directly.

However, the increased prevalence of dry skin in elderly people hints at an increased predisposition to aberrations in terminal differentiation, cornification and desquamation of keratinocytes (10, 11). Other cell types in the epidermis, mainly the Langerhans cells and melanocytes, decrease in number during skin aging (12).

Since the dermis is a more static tissue than the epidermis, alterations and aberrations can have longer effects. For example, the half-life of collagen, an important constituent of ECM, is 15 years in the skin (13). Therefore, changes in ECM components and homeostasis

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Figure 1: Histological characteristics of intrinsic skin aging. The most common and obvious characteristic of aged skin is the loss of the rete ridges (arrows); a flattening of the dermal-epidermal junction. The amount of collagen is reduced and the matrix is more disorganized. In addition, skin appendages and blood vessels decrease in number and become more fragile. SC: stratum corneum, Epi: epidermis, PD: papillary dermis, RD:

reticular dermis. Scale bars: 50µm.

can have significant and long lasting effects on the skin. One of the most characteristic changes in aging skin is a decrease in collagen (type I) production by the fibroblasts (14).

This leads to a general atrophy of the dermal ECM and is believed to be one of the major culprits of the detrimental effects of skin aging. Another important component of the ECM is the elastic fibre network, which consists of long bundles of the elastin protein and confers elastic properties to the skin. The half-life of elastin is very long: longer than the human lifespan (15). During intrinsic skin aging the elastic fibre network is damaged and gradually lost (8, 16). In photoaging, the production of elastin is increased in response to UV. However, the elastin is also damaged, which causes accumulation of non-degradable elastin; the aforementioned solar elastosis. A histological picture of intrinsic skin aging is shown in figure 1.

Fibroblasts and skin aging

As experimentally confirmed in HSEs, the dermis populated and maintained by fibroblasts is crucial to skin appearance and mechanical properties, and clearly affects epidermal development and homeostasis (17). Hence, the studies in this thesis are based on the

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INTRODUCTION

premise that the fibroblasts are pivotal in skin aging. We therefore studied two processes involving dermal fibroblasts that may be important drivers of skin aging: fibroblast senescence and loss of papillary fibroblasts in the dermis. Both processes are described below and depicted in figure 2.

Senescence

The term cellular senescence was originally coined to describe the state that normal (i.e.

untransformed), primary cells enter when they lose their replicative capacity in vitro.

After a certain number of population doublings most cells will stop dividing permanently (18). Later it was discovered that cell senescence is part of the cellular stress response (19). Basically, after a significant amount of genotoxic stress cells will block their cell cycle progression. Based on the severity of damage and the cell’s ability to repair damage, three options are possible: the cell can repair the damage and re-enter the cell cycle, go into senescence or remove itself by apoptosis. Due to its role in genotoxic stress response, senescence is seen as a tumor suppressive mechanism (20, 21).

It appears as if senescence is an intermediate between complete recovery and apoptosis, but this may be an oversimplification. Besides losing their replicative ability, senescent fibroblasts undergo a number of changes. These include alterations in morphology, secretory phenotype and ECM production. These changes are suspected to contribute to aging (22, 23).

Based on the in vitro alterations that occur in senescent cells, it is inferred that senescence plays a role in in vivo aging (24, 25). The number of senescent cells is increased in aged skin (and other tissues as well) (26-28). Senescent cells appear to contribute to aging in the skin by creating a more pro-inflammatory environment and by promoting matrix degradation. However, the exact role and the extent of the contribution of senescent cells in aging are not completely clear.

Papillary and reticular fibroblasts

The dermis can be divided in two morphologically distinct parts. The upper part, called papillary dermis, is characterized by a high cell density and loose matrix. The lower part, called reticular dermis, is characterized by a low cell density and dense matrix (7). The fibroblasts from these layers look and behave differently in culture (29). For example, compared with reticular fibroblasts, papillary fibroblasts show a leaner and more spindle- shaped morphology and show increased proliferation. (30-32).

Since papillary fibroblasts are closer to the epidermis, it is assumed that papillary fibroblasts are better able to support the epidermis. For example, they differ in the secretion of a number of growth factors and papillary fibroblasts are better at stimulating

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Figure 2: Proposed model of the role of fibroblasts in skin aging. Skin aging is characterized by a general atrophy of the skin and its cellular components. The most common characteristic of skin aging is flattening of the dermal-epidermal junction. This is correlated to a reduction of the papillary dermis. In this thesis two mechanisms involved in skin aging were investigated in HSEs: (I.): the decrease in the papillary dermis and papillary fibroblasts and (II.): the appearance of senescent fibroblasts in aged skin.

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INTRODUCTION

epidermal (terminal) differentiation in HSEs (33, 34).

A study by Mine et al. showed that papillary fibroblasts are more affected by aging than reticular fibroblasts (35). During skin aging, the rete ridges and part of the papillary dermis are lost, which leads to a relative increase of the reticular dermis and reticular fibroblasts.

The main hypothesis of this thesis is that these changes in the dermis have significant effects on the entire skin, because reticular and papillary fibroblasts have different effects on skin homeostasis; the (relative) increase of reticular fibroblasts and the decrease of papillary fibroblasts will alter skin homeostasis.

Myofibroblasts and Transforming growth factor-β

Transforming growth factor-β(TGF-β) is an important regulator in normal skin homeostasis. Its two main effects are stimulation of matrix formation and inhibition of epidermal proliferation (36, 37). If the skin is injured, TGF-βrecruits immune cells, stimulates the production of new ECM and stimulates the closure of the wound (38). An important step in this process is the differentiation of fibroblasts to myofibroblasts, which is also induced by TGF-β.

Myofibroblasts are known for their role in tissue remodelling; they play an essential role during wound healing (39). In the wound, the first action of the recruited myofibroblasts is to attach to the damaged dermis and contract, thus closing the wound as much as possible. Then the damaged tissue needs to be regenerated, to which the myofibroblast contributes considerably by generating ECM components. After regeneration is completed, myofibroblasts should disappear from the healed tissue. If the healing process is not properly terminated, myofibroblasts can remain in healed tissue.

Improperly stopped healing processes can lead to fibrotic diseases, such as liver and lung fibrosis and hypertrophic scar formation. Myofibroblasts are strongly implicated in these disorders (40).

Reticular fibroblasts share some characteristics with myofibroblasts, such as increased contractility and formation of dense matrix. However, it is not known whether reticular fibroblasts represent a precursor state to myofibroblasts, have a “mild” myofibroblast-like phenotype or are not related to myofibroblasts at all.

The number of alpha smooth muscle actin (α-SMA, a myofibroblast marker) positive fibroblasts is higher in reticular populations than in papillary populations in vitro. How- ever, the percentage of positive fibroblasts, even in reticular populations, is minimal (<

5%). Expression ofα-SMA is not found in dermal fibroblasts in vivo (41).

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Human Skin equivalents

There are several ways to perform experiments with skin. The experimental systems differ in complexity, ranging from monolayer cultures of skin cells to in vivo studies (both in humans and in animal models). Human skin equivalents (HSEs) fit in between both ends of this spectrum. HSEs are more complex than monolayer cultures and more representative of the tissue morphology of in vivo skin. While HSEs lack the sheer complexity of in vivo skin, which contains a multitude of different cell types and is influenced by many hormonal, metabolic, nervous and immune processes, HSEs are not hampered by the ethical and practical constraints of in vivo skin. HSEs allow the study of processes in a dedicated, specially designed experiment.

HSEs can be generated in several ways. They can consist of only an epidermis or both a dermal and epidermal part: a full thickness equivalent. Most HSEs only contain fibroblasts and keratinocytes, but other cell types, such as melanocytes, vascular cells and several types of immune cells can be included as well (42). HSEs can be used to model several skin diseases, for example skin cancer, skin infections and skin irritation (43-50).

The main type of HSE used in this thesis is the Fibroblast Derived Matrix (FDM) HSE (17). In this type of HSE fibroblasts are seeded in a transwell and stimulated to produce ECM. This combination of ECM and fibroblasts is then used as a dermal matrix onto which the keratinocytes are seeded. In most full-thickness HSEs the dermal part contains or consists of artificial components, for example a bovine or rattail collagen type I gel seeded with fibroblasts. The FDM HSE more clearly represents the dermis of human skin than other types of full-thickness equivalents, since the ECM is generated by the fibroblasts themselves.

Aims

The main goal of this thesis was to contribute to the development of an in vitro HSE model that mimics characteristics of skin aging. In chapter 2, we started by examining if FDM HSEs can be representative of in vivo skin and used for studies on skin aging. For this purpose, FDM HSEs were treated with a known anti-aging compound: Bio Marine Complex (51). The effects of this compound on in vivo skin collagen production and keratinocyte proliferation are well described and thus allowed us to show the similarities between the response of HSEs and in vivo skin to this compound.

After showing the validity of HSEs as a model for skin aging in vitro, several alterations were made to the composition and culture process of FDM HSE, with a focus on the fibroblasts. The alterations were based on the two processes involved in skin aging discussed above: the appearance of senescent fibroblasts and the loss of papillary fibroblasts

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INTRODUCTION

in aged skin. We hypothesized that alterations to the HSE culture process that mimic these processes can introduce features of skin aging in HSEs.

Serial passage is commonly used to induce replicative senescence in vitro and is often used as a model for aging in in vitro monolayer cultures (52, 53). In chapter 3, HSEs were generated with long-cultured (and often passaged) fibroblasts and compared with skin equivalents generated with short-cultured fibroblasts of a few passages to determine whether the inclusion of senescent fibroblasts introduces features of aged skin to the HSE.

The remainder of the chapters focusses on papillary and reticular fibroblasts. Unlike senescent fibroblasts, there is no clear definition for these populations in monolayer culture yet. Therefore, we first tried to identify markers for both fibroblasts populations in chapter 4. In chapter 5 and 6, we investigated if papillary fibroblasts can differentiate into reticular fibroblasts. In chapter 5 the fibroblasts were serially passaged and in chapter 6 TGF-β1 was used to induce differentiation. In both chapters 5 and 6 HSEs were generated with papillary and reticular fibroblasts. Based on the results obtained in these HSEs a final study was performed to elucidate how the different fibroblast populations interact with keratinocytes in vitro. This is described in chapter 7. The results from these four chapters led to an extension of the hypothesis of Mine et al. regarding the role of the fibroblast populations in skin aging (35). The future perspectives of the presented research and discussion concerning our hypotheses are presented in chapter 8.

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18. Hayflick L. The limited in vitro lifetime of human diploid cell strains. Exp Cell Res 1965: 37:614- 636.

19. Toussaint O, Dumont P, Remacle J et al. Stress-induced premature senescence or stress-induced senescence-like phenotype: one in vivo reality, two possible definitions? ScientificWorldJournal 2002: 2:230-247.

20. Collado M, Serrano M. The power and the promise of oncogene-induced senescence markers.

Nat Rev Cancer 2006: 6:472-476.

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22. Campisi J, d’Adda di Fagagna F. Cellular senescence: when bad things happen to good cells. Nat Rev Mol Cell Biol 2007: 8:729-740.

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the dark side of tumor suppression. Annu Rev Pathol 2010: 5:99-118.

24. Erusalimsky JD, Kurz DJ. Cellular senescence in vivo: its relevance in ageing and cardiovascular disease. Exp Gerontol 2005: 40:634-642.

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129:467-474.

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27. Ressler S, Bartkova J, Niederegger H et al. p16INK4A is a robust in vivo biomarker of cellular aging in human skin. Aging Cell 2006: 5:379-389.

28. Waaijer ME, Parish WE, Strongitharm BH et al. The number of p16INK4a positive cells in human skin reflects biological age. Aging Cell 2012: 11:722-725.

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30. Schafer IA, Pandy M, Ferguson R, Davis BR. Comparative observation of fibroblasts derived from the papillary and reticular dermis of infants and adults: growth kinetics, packing density at confluence and surface morphology. Mech Ageing Dev 1985: 31:275-293.

31. Azzarone B, Macieira-Coelho A. Heterogeneity of the kinetics of proliferation within human skin fibroblastic cell populations. J Cell Sci 1982: 57:177-187.

32. Harper RA, Grove G. Human skin fibroblasts derived from papillary and reticular dermis:

differences in growth potential in vitro. Science 1979: 204:526-527.

33. Pageon H, Zucchi H, Asselineau D. Distinct and complementary roles of papillary and reticular fibroblasts in skin morphogenesis and homeostasis. Eur J Dermatol 2012.

34. Sorrell JM, Baber MA, Caplan AI. Site-matched papillary and reticular human dermal fibroblasts differ in their release of specific growth factors/cytokines and in their interaction with keratinocytes. J Cell Physiol 2004: 200:134-145.

35. Mine S, Fortunel NO, Pageon H, Asselineau D. Aging alters functionally human dermal papillary fibroblasts but not reticular fibroblasts: a new view of skin morphogenesis and aging. PLoS ONE 2008: 3:e4066.

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37. Chambers RC, Leoni P, Kaminski N, Laurent GJ, Heller RA. Global expression profiling of fibroblast responses to transforming growth factor-beta1 reveals the induction of inhibitor of differentiation-1 and provides evidence of smooth muscle cell phenotypic switching. Am J Pathol 2003: 162:533-546.

38. Gosain A, DiPietro LA. Aging and wound healing. World J Surg 2004: 28:321-326.

39. Hinz B. Formation and function of the myofibroblast during tissue repair. J Invest Dermatol 2007: 127:526-537.

40. Hinz B, Phan SH, Thannickal VJ, Galli A, Bochaton-Piallat ML, Gabbiani G. The myofibroblast:

one function, multiple origins. Am J Pathol 2007: 170:1807-1816.

41. Rajkumar VS, Howell K, Csiszar K, Denton CP, Black CM, Abraham DJ. Shared expression of phenotypic markers in systemic sclerosis indicates a convergence of pericytes and fibroblasts to a myofibroblast lineage in fibrosis. Arthritis Res Ther 2005: 7:R1113-R1123.

42. van den Bogaard EH, Tjabringa GS, Joosten I et al. Crosstalk between keratinocytes and T cells in a 3D microenvironment: a model to study inflammatory skin diseases. J Invest Dermatol 2014: 134:719-727.

43. Auxenfans C, Fradette J, Lequeux C et al. Evolution of three dimensional skin equivalent models reconstructed in vitro by tissue engineering. Eur J Dermatol 2009: 19:107-113.

44. Kim DS, Cho HJ, Choi HR, Kwon SB, Park KC. Isolation of human epidermal stem cells by adherence and the reconstruction of skin equivalents. Cell Mol Life Sci 2004: 61:2774-2781.

45. Lacroix S, Bouez C, Vidal S et al. Supplementation with a complex of active nutrients improved dermal and epidermal characteristics in skin equivalents generated from fibroblasts from young or aged donors. Biogerontology 2007: 8:97-109.

46. Thakoersing VS, Gooris GS, Mulder A, Rietveld M, El GA, Bouwstra JA. Unraveling barrier properties of three different in-house human skin equivalents. Tissue Eng Part C Methods 2012:

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48. Commandeur S, Sparks SJ, Chan HL et al. In-vitro melanoma models: invasive growth is determined by dermal matrix and basement membrane. Melanoma Res 2014: 24:305-314.

49. Haisma EM, de BA, Chan H et al. LL-37-derived peptides eradicate multidrug-resistant Staphylococcus aureus from thermally wounded human skin equivalents. Antimicrob Agents Chemother 2014: 58:4411-4419.

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50. van D, V, Alloul-Ramdhani M, Danso MO et al. Knock-down of filaggrin does not affect lipid organization and composition in stratum corneum of reconstructed human skin equivalents.

Exp Dermatol 2013: 22:807-812.

51. Kieffer ME, Efsen J. Imedeen in the treatment of photoaged skin: an efficacy and safety trial over 12 months. J Eur Acad Dermatol Venereol 1998: 11:129-136.

52. de Magalhaes JP. From cells to ageing: a review of models and mechanisms of cellular senescence and their impact on human ageing. Exp Cell Res 2004: 300:1-10.

53. Rattan SI. Cell Senescence In Vitro. eLS. John Wiley & Sons, Ltd, 2012.

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Chapter 2 Marine derived nutrient improves epidermal and dermal structure and prolongs the lifespan of

reconstructed human skin equivalents

Marion Rietveld

¹

, David Janson

¹

, Rachida Siamari

¹

, Jana Vicanova

²

, Maja Troest Andersen

³

, Abdoelwaheb El Ghalbzouri

¹

1

Department of Dermatology, Leiden University Medical Center, Leiden, The Netherlands

2

DermData, Prague, Czech Republic

3

Ferrosan A / S, Soeborg, Denmark

Journal of Cosmetic Dermatology, 2012, 11, 213-222

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Abstract

Imedeen is a cosmeceutical that provides nutrients to the skin. One of its active ingredients is the Marine Complex (MC). The aim of this study was to evaluate whether MC affects skin morphogenesis differently in female and male human skin equivalents (HSEs).

HSEs were established with cells obtained from female or male donors between 30 and 45 years of age and cultured for 7 or 11 weeks in the presence or absence of MC.

Using immunohistochemistry, we examined early differentiation by keratin 10 expression, (hyper)proliferation by keratin 17 and Ki67, and basement membrane composition by laminin 332 and collagen type VII. In addition, the expression of collagen type I and the secretion of pro-collagen I were measured.

MC strongly increased the number of Ki67 positive epidermal cells in female HSEs.

In the dermis, MC significantly stimulated the amount of secreted pro-collagen I and increased the deposition of laminin 332 and collagen type VII. Furthermore, MC prolonged the viable phase of HSEs by slowing down its natural degradation. After 11 weeks of culturing, the MC treated HSEs showed higher numbers of viable epidermal cell layers and a thicker dermal extracellular matrix compared to controls. In contrast, these effects were less pronounced in male HSEs.

The MC nutrient positively stimulated overall HSE tissue formation and prolonged the longevity of both female and male HSEs. The ability of MC to stimulate the deposition of basement membrane and dermal components can be used to combat human skin aging in vivo.

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MARINE DERIVED NUTRIENT IMPROVES EPIDERMAL AND DERMAL STRUCTURE

Introduction

On macroscopic level, human skin is the best available biomarker of aging. During this process, the skin loses its structural and functional characteristics. The most pronounced modifications occur in the dermis, where a decrease in the deposition of extracellular matrix (ECM) components has been reported (1). This lower production leads to a disorganized and imbalanced ratio of ECM components that affects the dermal-epidermal junction (DEJ) and eventually the epidermal compartment. At the basement membrane zone, the DEJ flattens with aging, which results in a decreased synthesis of basement membrane proteins such as laminins and collagens, leading to weaker attachment of the epidermis to the dermis (2). In turn, this leads to lower keratinocyte cell activity reflected by thinning of the epidermis by 10–50% between the age of 30 and 80 years (3, 4). In addition to keratinocytes, the numbers of melanocytes and Langerhans cells also diminish in the epidermis (2). Furthermore, studies have shown that during aging, the number of dermal papillae decreases per surface area (5). These dermal-epidermal alterations lead to wrinkles, discoloration, telangiectasia and elastosis (6).

As the skin is a hormone sensitive organ, different hormones play a major role in the structure and function of both male and female skin (7). For example, male skin is thicker than female skin and contains more collagen and elastin. Furthermore, the structure and maintenance of male skin differs from female skin due to larger pores and a higher production of sebum (8). Despite these differences, the outcome of intrinsic aging processes is similar between women and men.

The cellular processes that contribute to skin aging (e.g. cell senescence) can be studied in mouse models or in conventional two-dimensional monolayer cell cultures.

In the latter model, the communication between the dermal and epidermal compartment cannot be studied, while this interaction is of crucial importance in controlling processes leading to proper skin homeostasis. Mouse models do not represent the human skin environment and are therefore less suitable to study human skin aging. In contrast to mouse models and monolayer cell cultures, reconstructed human skin equivalents (HSEs) contain a microenvironment that is highly similar to that of in vivo human skin, which makes them an attractive tool to study human skin aging in vitro. HSEs are three- dimensional culture systems engineered by seeding human keratinocytes onto a three- dimensional dermal matrix populated with human fibroblasts. After cell attachment, the culture is first kept under submerged conditions to allow keratinocyte proliferation.

Thereafter, the HSE is cultured at the air-liquid (A/L) interface to air-expose the epidermal compartment and to further induce keratinocyte proliferation and differentiation. Under these conditions, a HSE is formed that shows high similarity with the native tissue from which it was derived (9, 10). The HSEs are also easy to modulate, which allows generation

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of skin models consisting of epidermal and dermal cells isolated from different types of donors.

Over the last decade, our understanding of skin aging has increased and a large number of cosmeceuticals has been documented, claiming to prevent, counteract or reverse degenerative alterations in skin aging (11). One of such cosmeceuticals is Imedeen^TM, an oral skin care product. The aim of this skincare product is to provide a combination of nutrients to the skin from within to improve the overall structure, appearance and the quality of the skin. Imedeen’s main ingredients include vitamin C, zinc and, the subject of this study, Marine Complex (MC). MC consists of fish protein polysaccharides similar to skin ECM (12). Several studies have proven the effects of ImedeenTM ingredients at a cellular level (13, 14). Human studies have also demonstrated clinically visible and objectively measurable effects on aging female skin (12, 15). The present study was designed to investigate the effects of MC, one of the active ingredients of ImedeenTM, in HSEs generated with cells obtained from either middle aged female or male skin donors. We examined whether MC affects epidermal morphology, basement membrane composition, pro-collagen type I secretion and whether observations from earlier in-vivo studies in middle aged females could be mimicked in these skin models (14, 15). Finally, we also evaluated whether MC improves the quality of HSEs after a prolonged culture period.

Materials and Methods

Cell culture

Cultures of primary normal human epidermal keratinocytes (NHEK) and primary normal human dermal fibroblasts (NHDF) were established from human mammary skin cells obtained from female donors aged 35, 40 and 45 years, and from human abdominal skin surgery from male donors aged 31, 35, 42 and 45 years old. Keratinocytes and fibroblasts were isolated as described earlier (16). In short, human dermis and epidermis were separated through overnight incubation of the skin with dispase II (Roche Diagnostics Nederland B.V., Almere, The Netherlands). Epidermis was incubated with trypsin (BD Biosciences, Breda, The Netherlands) at 37°C for 15 minutes. After trypsin inactivation, the cells were filtered with a 70µm cell strainer (BD Biosciences, Breda, The Netherlands), and cultured in keratinocyte medium at 37°C and 7.3% CO2 until subconfluency. Keratinocyte medium consisted of 3 parts Dulbecco’s modified Eagle’s medium (DMEM, Gibco/Invitrogen, Breda, The Netherlands) and 1 part Ham’s F12 medium (Gibco/Invitrogen) supplemented with 5% fetal bovine serum (FBS, HyClone/Greiner, Nürtingen, Germany), 0.5µM hydrocortisone, 1µM isoproterenol, 0.1µM insulin (Sigma-

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Aldrich, Zwijndrecht, The Netherlands), 100 U/mL penicillin and 100µg/mL streptomycin (Invitrogen). Fibroblasts were isolated from the dermis with collagenase II (Invitrogen) and dispase II (ratio 3:1 and 3 ml/g dermis) at 37°C for 2 hours. The cells were filtered with a 70µm cell strainer, and cultured in fibroblast medium at 37°C and 5 % CO2 until subconfluency. Fibroblast medium consisted of DMEM supplemented with 5% FBS, 100 U/mL penicillin and 100µg/mL streptomycin. Early passage (passages 2–3) were used for experiments.

Fibroblast-derived matrix

Fibroblast-derived matrices (FDMs) were generated as described earlier (16). In short, 400.000 fibroblasts obtained from either male or female donors were seeded into a polyester permeable support (6 well plates with 0.4 µM pore size Transwell inserts, Corning Incorporated, Schiphol-Rijk, The Netherlands) in the presence or absence of MC (figure 1). The FDMs were supplemented with 5% FBS, penicillin (100 IU/mL) and streptomycin (100 µg/mL), 50 µM ascorbic acid phosphate (Sigma-Aldrich). Culture medium was refreshed twice a week. FDMs were generated with fibroblasts obtained from either female or male donors.

Reconstructed humans skin equivalent

FDMs were seeded with 500.000 NHEKs (passage 1-2) per HSE obtained from either female (f-HSE) or male (m-HSE) donors. Cultures were incubated overnight in keratinocyte medium as described above, but with reduced FBS (1%), 53µM selenious acid, 10mM L-serine, 10µM L-carnitine, 1µM dL-α-tocopherol-acetate, 250µM ascorbic acid phosphate, 12µM bovine serum albumin and a lipid supplement containing 25µM palmitic acid, 15µM linoleic acid and 7µM arachidonic acid (Sigma-Aldrich). The HSEs were then cultured at the air-liquid interface in supplemented keratinocyte medium as described above, but without FBS. Culture medium was refreshed twice a week. After 7 and 11 weeks of air-exposed culture, the HSEs were processed for analysis. To one part of the 7-week old HSEs, MC was added and these HSEs were cultured for an additional 4 weeks at the air-liquid interface (Figure 1).

Supplementation of Imedeen Marine Complex

HSEs were supplemented twice a week with MC at a final concentration of 70 µg/mL starting at two different time points: at the start of the air-exposed culture phase or after 7 weeks of culturing (Biomarine Complex batch number: 199888).

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RNA isolation

RNA was isolated from monolayer fibroblast cultures, treated with MC (70 ug/mL) or untreated controls, using the RNEasy kit (Qiagen, Venlo, The Netherlands), according to the manufacturer’s instructions. RNA was extracted after 60 minutes and 24 hours.

Q-PCR analyses

cDNA was generated of 1µg RNA using the iScript cDNA synthesis kit (BioRad, Veenen- daal, The Netherlands) according to manufacturer’s instructions. PCR reactions were based on the SYBR Green method (BioRad) and consisted of 2x Sybr Green Mastermix, 1ng cDNA template and 500 nM of forward and reverse primers. The PCRs were run on the CFX384 system (BioRad). The PCR cycles were: 3,5 minutes at 95 C to activate the polymerase, 45 cycles of 10 sec 95 C and 30 sec 60 C, followed by the generation of a melt curve. Primers were checked before on dilution series of normal fibroblasts cDNA.

Reference genes were analyzed with the GeNorm method (17). Expression analysis was performed with the BioRad Software (CFX Manager) and was based on the delta delta Ct method with the reference genes that were stably expressed in the GeNorm analysis. The primers are listed in Table 1.

Target Sequence Forward Sequence Reverse

Col1A1 ATGTTCAGCTTTGTGGACCTCCGG CGCAGGTGATTGGTGGGATGTCT

EI24 TTCACCGCATCCGTCGCCTG GAGCGGGTCCTGCCTTCCCT

SND1 CGTGCAGCGGGGCATCATCA TGCCCAGGGCTCATCAGGGG

Table 1: Primers used in this study. SND1 was used as reference gene.

Morphological and immunohistochemical analysis

HSE cultures were washed in PBS, one half was snap-frozen in liquid nitrogen while the other half was fixed in 4% formaldehyde and paraffin embedded (FFPE). Global morphological analysis was performed on 5 µM FFPE sections through staining with haematoxylin and eosin. Immunohistochemical analysis was either performed on FFPE sections or cryosections. 5µm FFPE sections were rehydrated through xylene and ethanol.

5µm cryosections were fixed with acetone. Primary antibodies used are shown in Table 2.

Following incubation with the primary antibody, sections were stained with avidin-biotin- peroxidase system (GE Healthcare, Buckinghamshire, UK), as described by manufacturer’s instructions. Staining was visualized with 3-amino-9-ethylcarbazole (AEC). All sections were counterstained with haematoxylin.

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Antigen (antibody clone) Supplier

Collagen Type I Sigma

Keratin 17 (KS6.KA12)/( CK-E3) Sanbio, Novus

Keratin 10 (DE-K10) Abcam

Collagen VII (PHM12)/(LH7.2) Chemicon

Laminin 332 (BM165) Kind gift from Dr. A. Aumailly

Ki67 (MIB1) DAKO

Table 2: Primary antibodies used in this study

Protein determination by enzyme-linked immunosorbent assay

Secreted human pro-collagen type I in the culture medium of HSEs was quantified by enzyme-linked immunosorbent assay (MetraCICP ELISA kit; Quidel Corporations, CA, USA). Culture medium of the HSEs was collected at weeks 1, 3, 5, 7, 9 and 11. Measure- ments and data analysis were performed according to the manufacturer’s procedure.

Estimation of proliferation index

The proliferation index was calculated as the number of Ki67 positive basal keratinocytes divided by the total number of basal keratinocytes counted. A minimum of 50 basal cells was counted in three areas of sections of two different samples at a magnification of 200x.

The resulting data are expressed as the mean of the two independent experiments ± SD.

Statistics

Each culture was performed in duplicate. Statistical significance was determined using the two-tailed Student’s t-test.

Results

Marine Complex increased expression of collagen type I in human fibroblasts

To evaluate whether MC affects FDM formation and epidermal morphogenesis, we first evaluated its effect on monolayer cultures of fibroblasts isolated from two different female and male donors. After 1 and 24 hrs of supplementation we determined the expression of collagen type I by quantitative PCR. As shown in figure 2, addition of 70µg/ml MC to human female fibroblasts resulted in a significant, two-fold increase of collagen type I mRNA after 24 hrs. This increase was less pronounced in male fibroblasts (data not shown).

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Figure 1: Scheme illustration supplementation of MC at the start of the cultures and addition to 7 week old skin cultures. The cultures were processed for analyses on week 7 and 11. The FDMs serving as control are not included in this scheme.

Figure 2: Q-PCR analyses showing increased average expression of collagen type I mRNA in female fibroblasts (two donors) after stimulation with MC (white bars) compared to untreated controls (black bars). The normalized fold expression is based on the reference genes SND1and EI24. Error bars show SEM. *p<0.05.

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Marine Complex increased number of viable cell layers

Next, we determined the effect of MC on epidermal morphology in f-FDMs and m- FDMs (figure 3). For this purpose, HSEs were cultured for either 7 weeks (figure 3a, b) or 11 weeks (figure 3c, d), in the presence or absence of MC. To evaluate whether MC is able to reactivate the epidermis after 7 weeks, we supplemented MC to 7-week old HSEs and cultured them for an additional 4 weeks (figure 1). As shown in figure 3, HSEs supplemented with MC and cultured for 7 or 11 weeks showed significantly more viable cell layers in f-HSEs (around 8 layers ± 1), compared to untreated f HSEs.

In m-HSEs, this effect was also significant in 7-week old cultures but less pronounced in 11-week old m-HSEs (figure 3A, 3B), (around 7 layers ± 1). When the results of the different donors were combined, the effect of MC was more pronounced (figure 3C, 3D).

Irrespective of the conditions, in all experiments a nicely differentiated epidermis was formed containing all strata including stratum basale, stratum spinosum and stratum corneum. Supplementation of MC to HSEs significantly increased the number of cell layers in 7-week old cultures (figure 3C). The effect of MC was still significant in 11-week old f-HSEs but was less pronounced in m-HSEs (figure 3D). Addition of MC in week 7 also significantly increased the number of cell layers in both f-HSEs and m-HSEs.

Epidermal morphogenesis was not altered by Marine Complex

To examine whether MC influences the differentiation program we evaluated the expression of early differentiation marker keratin 10 (K10) in HSEs using immunohistochemistry.

K10 was expressed in all suprabasal cell layers irrespective of the presence or absence of MC, indicating that the early differentiation program occurs normally as in native tissue.

The activation and hyperproliferation associated marker keratin 17 (K17) was absent in all HSEs, indicating that MC does not induce epidermal stress. Epidermal cell proliferation was clearly affected by the presence of MC as demonstrated by the ki67 marker. In f-HSEs, supplementation of MC resulted in an increased number (± 8 cells/ 100 basal cells) of proliferating cells compared to the controls (figure 4A). In m-HSEs this effect was also significant, but less pronounced (figure 4B). However, the number of proliferating basal cells in m-HSEs was significantly increased in 7-week old cultures supplemented with MC (figure 5, black bars). A similar effect was observed in the 11-week old cultures (figure 5, white bars). In addition, when MC was added from week 7 on and cultured for an additional 4 weeks, the increase of proliferating cells was still significant compared to the control (figure 5, dashed bar). In f-HSEs, the effect of MC on cell proliferation was more pronounced (data not shown).

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Figure 3: Shown are cross sections of one representative donor of HSEs generated with female (A) or male (B) cells that have been cultured in the absence (-) or presence (+) of BioMarine Complex (MC). Irrespective of the gender of the donors, HSEs that have been cultured for 7 weeks (a, b) show more viable cell layers when they are supplemented with MC at the beginning of the culture. Similar results were found when HSEs were cultured for 11 weeks (c, d) in the absence or presence of MC. Supplementation of MC to HSEs from the seventh week on and cultured for an additional 4 weeks (e) resulted in more viable cell layers compared to a and c; (Scale bar: 20 µm). The numbers of viable cell layers are reflected in panels C (f-HSE) and D (m-HSE). Cell layers were counted randomly in three different areas of the cross sections. Black bars represent the average of four female donors while white bars represent the average of four male donors. Supplementation of MC to HSEs significantly increased the number of cell layers in 7 week old cultures (C); * p<0.05. The effect of MC was still significant in 11-week old female cultures but was less pronounced in male HSEs (D). However, when adding MC from week 7 on, it also significantly increased the number of cell layers in both female and male HSEs; *p<0.05.

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Figure 4: Shown are cross sections of one representative donor of HSEs generated with female (f-HSE)(A) or male (m-HSE) (B) cells that have been cultured in the absence (- ) or presence (+) of BioMarine Complex (MC). Irrespective of the gender of the donors or MC supplementation, HSEs that have been cultured for 11 weeks express the early differentiation marker K10 in all suprabasal layers. The hyperproliferation associated marker K17 was absent irrespective of absence or presence of MC. In HSEs supplemented with MC, the number of proliferative cells (Ki67 positive cells) in the basal layer was increased. This increase was more pronounced in f-HSEs. (Scale bar: 50µm).

Figure 5: Graph shows the effect of MC on cell proliferation in m-HSE. The number of Ki67 positive cells present per 100 basal cells were counted in three different areas of the cross sections. Black bars represent the average number of Ki67 positive cells after 7 weeks of culture while white bars represent the 11-week old cultures. Supplementation of MC to f-HSEs significantly increased the number of Ki67 positive cells both in 7- and 11-week old cultures. * p<0.05. The effect of MC was still significant when MC was added from week 7 on (dashed bar):*p<0.05. Results represent the average of 4 donors.

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Figure 6: Shown are cross sections of one representative donor of HSEs generated with female (f-HSE)(A) or male (m-HSE) (B) cells that have been cultured in the absence (-) or presence (+) of BioMarine Complex (MC) for 11 weeks. In f-HSEs, supplementation of MC clearly increased the deposition of laminin 332 and collagen type VII at the dermal-epidermal junction, while in m-HSEs, supplementation of MC only increased the deposition of collagen type VII. Collagen type I deposition was present in all dermal compartments irrespective of the absence or presence of MC. (Scale bar: 20µm).

Marine Complex affects basement membrane formation

Next, we evaluated the effect of MC on basement membrane formation in HSEs. For this purpose, we analyzed the two basement membrane proteins laminin 332 and collagen type VII. Immunohistochemical analyses demonstrated nice deposition of these proteins at the dermal-epidermal junction. In addition, a clear effect of MC on the expression of both proteins was observed (figure 6). In f-HSEs cultured for 11 weeks in the presence of MC, the deposition of laminin 332 and collagen type VII was higher compared to the controls. When MC was supplemented from week 7 on and cultured for an additional 4 weeks, a similar effect was seen for both proteins. Interestingly, in m-HSEs, this increased deposition was only observed for collagen type VII and not for laminin 332.

Marine Complex increases pro-collagen type I deposition in f-FDMs

Collagen type I was expressed throughout the dermal compartment of both f-HSEs and m-HSEs (figure 6). As direct quantification of the collagen type I cannot be performed within the context of the dermis, we chose to measure the collagen type I precursor pro- collagen type I in the culture medium of HSEs as a measure of ECM formation. The effect of MC on the dermal compartment was evaluated by measuring the released pro-collagen

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Figure 7: Graph shows the effect of BioMarine Complex (MC) on pro-collagen type I deposition in f-HSE (A) and m-HSE (B). White bars represent the HSEs without MC and black bars represent HSEs supplemented with MC. Pro-collagen type I was measured in medium collected from the HSEs during the culture period (weeks 1 to 11). As shown in graph A, addition of MC significantly increased the deposition of pro-collagen type I in f-HSEs, while in m-HSEs no difference was observed. * p<0.05. Results represent the average of two donors.

type I by the fibroblasts in the culture medium of the HSEs collected in weeks 1, 3, 5, 7, 9 and 11 (figure 7A). Supplementation of MC significantly increased the deposition of pro- collagen type I in f-HSEs. This increase peaked in week 7 (866 ng/mL ± 121). In 11-week cultures, the secretion of pro-collagen decreased to almost 368 ng/mL (± 68). In m-HSEs, this observation was not found, as the basal secretion already started at 829 ng/mL (± 30) in week 1 and decreased to around 548 ng/mL (±70) in week 11 (figure 7B).

Discussion

Most human skin aging research has been performed either in vivo) (animal models or skin biopsies) or in monolayer cell cultures. Animal models are not only constrained by obvious ethical limitations and high costs, but also misrepresent the human skin microenvironment. Monolayer cultures allow more experimental flexibility but do not reflect the human in vivo) situation. Human skin equivalents (HSEs) fill the gap between these two different research models by allowing great experimental flexibility and placing experiments in the context of human skin tissue. Several studies have already showed that epithelial-mesenchymal interactions are of crucial importance for the overall homeostasis of the skin (18, 19). In addition, studies also showed that the microenvironment in which the fibroblasts are embedded affects epidermal morphogenesis (19, 20).

In our present study, we investigated whether supplementation of Marine Complex (MC), the active ingredient of ImedeenTM, affects the overall architecture of human skin including epidermis, basement membrane and dermis in both female and male

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generated HSEs. An earlier in-vivo study on ImedeenTM demonstrated positive effects (e.g. the decrease of fine lines, increased dermal volume and a smoother skin) in female subjects (14). In this study we have used fibroblast-derived matrix (FDM) as a dermal equivalent instead of classical rat-tail collagen based HSEs. FDMs are more suitable for studies in which factors secreted by fibroblasts are crucial for the overall skin morphology.

Furthermore, they can be cultured for longer periods of time compared to the rat-tail collagen based HSEs (16).

HSEs generated with keratinocytes and fibroblasts obtained from female skin (f-HSE) or male skin (m-HSE) were supplemented with MC by two different approaches. In the first approach, MC was added from the start of the cultures and analyzed after 7 and 11 weeks. In the second approach, we evaluated whether MC can reactivate epidermal proliferation of "mature" 7-week old HSEs. For this purpose, MC was added from week 7 on and cultured for an additional 4 weeks, resulting in a total air-exposed culture period of 11 weeks. Histological analyses showed that regardless of the approach and gender of the donors, MC improved epidermal morphology by increasing the number of viable cell layers. This was observed in 7- and 11-week old cultures but also in cultures that had been supplemented with MC from week 7 on, which had only been cultured in the presence of MC for 4 weeks. This suggests that MC reactivates epidermal turnover in long-term cultured HSEs. These observations were supported by the fact that the early differentiation program (assessed by Keratin 10 staining) was not altered by MC, regardless of the approach, donor gender and culture period. Addition of MC also did not induce the expression of the hyper-proliferation associated marker K17. This indicates the presence of a normalized epidermis, as the "epidermal stress protein" K17 is not expressed in healthy skin but present in skin that has been wounded or activated (e.g. after UV- irradiation, addition of growth factors) (21). Obviously, increased epidermal thickness in MC treated HSEs could be explained by the presence of proliferating cells in the basal cell layers. Therefore, we examined the expression of Ki67-postive cells in all cultures. In m-HSEs, a significant increase in cell proliferation was observed in 7-week old cultures but also in 11-week old cultures that have been supplemented with MC for 4 weeks. In f-HSEs, the presence of MC significantly increased basal cell proliferation in all cultures compared to their controls. This clearly indicates that MC acts on the proliferation program while leaving the differentiation program unaffected. One might hypothesize that MC may function as a specific growth factor that keeps epidermal turnover in balance and maintaining epidermal homeostasis. Especially when we compared 7-week with 11- week old HSEs, a constant epidermal thickness was observed in the presence of MC. These observations also raise the question whether MC could serve as a compound maintaining stem cells in the epidermal cell layers. It is known that most HSEs cultured up to 8 weeks consist of only 1 or 2 viable cell layers. This might be explained by a very rich environment

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containing growth factors and serum, in which the keratinocytes hyperproliferate and deplete their proliferative capacity within 8 weeks, or that the microenvironment lacks ingredients that maintain the stem cell compartment in the epidermal layers (20). Further research is necessary to elucidate whether MC has a role in these complex processes.

As stated earlier, interactions between the dermal and epidermal compartment are important for overall homeostasis of the skin. At the dermal epidermal junction (DEJ) of the basement membrane, various proteins are involved in signal transduction from one compartment to another. Collagen type VII is the principal constituent of the anchoring fibrils, attachment structures that play a major part in stabilizing the association of the basement membrane zone to the underlying papillary dermis. Laminin 332 is a major component of the actin filaments, and has been found to directly bind collagen type VII (22). Hence, both collagen type VII and laminin 332 are important for the tight connection of the epidermis and dermis. Besides their importance for the integrity of the DEJ, both laminin 332 and collagen type VII provide a special trans-basement-membrane route for the transmission of information from the dermis to the basal keratinocytes and vice versa, indirectly regulating keratinocyte proliferation and supporting cell migration during wound healing (23). The effect of MC on the expression of these proteins was quite remarkable, especially in f-HSEs, as the deposition of both laminin 332 and collagen type VII was increased at the DEJ of these models. This was observed in MC supplemented cultures of both 7 and 11 weeks. Interestingly, in m-HSEs similar observations were only found for the deposition of collagen type VII, while the deposition of laminin 332 remained unaffected by MC. This curious difference might be explained by the fact that fibroblasts and keratinocytes isolated from male and female skin likely differ in their sensitivity to hormone or hormone-like factors present in their microenvironment. Further analyses on the deposition of other DEJ proteins such as collagen type IV or integrin-subunits α6ß4 might shed new light on the role of MC in basement membrane formation. Finally, we evaluated whether MC activated fibroblasts to secrete collagen. Since the dermal matrix of FDMs consists solely of fibroblasts that secrete their own extracellular matrix, we evaluated the secretion of pro-collagen type I in medium collected throughout the culture period of HSEs. In f-HSEs, pro-collagen type I was increased during the whole culture period of 11 weeks, but was only significantly increased in weeks 3, 5 and 11 compared to the control samples. In m-HSEs, pro-collagen type I secretion remained unaltered by the presence of MC. The differences obtained in f-HSEs and m-HSEs might be explained by the fact that male skin differs in some aspects from female skin. Male skin is known to have thicker epidermis and dermis, the fibroblasts secret more collagen and elastin, it contains larger pores and produces more sebum. These parameters are partly directed by the hormones present in the skin. In addition, one can assume that the concentration of MC added to m-HSEs was too low to overcome its effect. Possibly, the fibroblasts had

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already a basal level of collagen production/activation that could not be improved by MC at a concentration of 70µg/mL.

In conclusion, the present study shows that the active ingredient of Imedeen, Marine Complex (MC) acts on the overall organization of HSEs. The basement membrane and the dermis of f-HSEs were more affected by MC than in m-HSEs. However, this might be explained by differences in male and female skin in vivo). The epidermis of both f-HSEs and m-HSEs were stimulated by MC, without activating the stress and hyperproliferation pathways. A remarkable observation was that MC was able to reactivate 7-week old HSEs that were already in a "mature" state. MC also increased the deposition of the basement membrane proteins collagen type VII in f-HSEs and m-HSEs and laminin 332 in f-HSEs.

Furthermore, this study clearly showed that MC increased pro-collagen type I secretion in f-HSEs. Although some differences between f-HSEs and m-HSEs were observed, this work demonstrates the "anti-aging" potential of MC. The nutricosmetic ingredient MC seems to fight natural aging by counteracting on cellular aging-associated processes observed in human skin.

Acknowledgements

We would like to thank Ir. Suzan Commandeur and Dr. Maria Ponec of the Department of Dermatology, Leiden University Medical Center (LUMC), Leiden, The Netherlands for carefully reading the manuscript. The work was sponsored by Ferrosan A/S Denmark.

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130:604-608.

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9. Bell E, Ehrlich HP, Sher S et al. Development and use of a living skin equivalent. Plast Reconstr Surg 1981: 67:386-392.

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11. Glaser DA. Anti-aging products and cosmeceuticals. Facial Plast Surg Clin North Am 2004:

12:363-72, vii.

12. Kieffer ME, Efsen J. Imedeen in the treatment of photoaged skin: an efficacy and safety trial over 12 months. J Eur Acad Dermatol Venereol 1998: 11:129-136.

13. Lacroix S, Bouez C, Vidal S et al. Supplementation with a complex of active nutrients improved dermal and epidermal characteristics in skin equivalents generated from fibroblasts from young or aged donors. Biogerontology 2007: 8:97-109.

14. Vicanova J, Bouez C, Lacroix S, Lindmark L, Damour O. Epidermal and dermal characteristics in skin equivalent after systemic and topical application of skin care ingredients. Ann N Y Acad Sci 2006: 1067:337-342.

15. Skovgaard GR, Jensen AS, Sigler ML. Effect of a novel dietary supplement on skin aging in post- menopausal women. Eur J Clin Nutr 2006: 60:1201-1206.

16. El Ghalbzouri A, Commandeur S, Rietveld MH, Mulder AA, Willemze R. Replacement of animal- derived collagen matrix by human fibroblast-derived dermal matrix for human skin equivalent products. Biomaterials 2008: 30:71-78.

17. Vandesompele J, De PK, Pattyn F et al. Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biol 2002:

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18. Maas-Szabowski N, Shimotoyodome A, Fusenig NE. Keratinocyte growth regulation in fibroblast cocultures via a double paracrine mechanism. J Cell Sci 1999: 112 ( Pt 12):1843-1853.

19. Stark HJ, Willhauck MJ, Mirancea N et al. Authentic fibroblast matrix in dermal equivalents normalises epidermal histogenesis and dermoepidermal junction in organotypic co-culture.

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20. Muffler S, Stark HJ, Amoros M et al. A stable niche supports long-term maintenance of human epidermal stem cells in organotypic cultures. Stem Cells 2008: 26:2506-2515.

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21. McGowan KM, Coulombe PA. Onset of keratin 17 expression coincides with the definition of major epithelial lineages during skin development. J Cell Biol 1998: 143:469-486.

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Chapter 3 Effects of serially passaged fibroblasts on dermal and epidermal morphogenesis in human skin

equivalents

David Janson

¹

, Marion Rietveld

¹

, Rein Willemze

¹

, Abdoelwaheb El Ghalbzouri

¹

1

Department of Dermatology, Leiden University Medical Center, Leiden, The Netherlands

Biogerontology, 2013, 14(2):131-40

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Abstract

Serial passaging has a profound effect on primary cells. Since serially passaged cells show signs of cellular aging, serial passaging is used as an in vitro model of aging. To relate the effect of in vitro aging more to in vivo aging, we generated human skin equivalents.

We investigated if human skin equivalents generated with late passage fibroblasts show characteristics of aged skin when compared with human skin equivalents generated with early passage fibroblasts.

Late passage fibroblasts had enlarged cell bodies and were more often positive for myofibroblast markerα-smooth muscle actin, senescence associatedβ-galactosidase and p16 compared with early passage fibroblasts. Skin equivalents generated with late passage fibroblasts had a thinner dermis, which could partly be explained by increased matrix metalloproteinase-1 secretion. In equivalents generated with late passage fibroblasts epidermal expression of keratin 6 was increased, and of keratin 10 slightly decreased.

However, epidermal proliferation, epidermal thickness and basement membrane formation were not affected.

In conclusion, compared with human skin equivalents generated with early passage fibroblasts, human skin equivalents generated with late passage fibroblasts showed changes in the dermis, but no or minimal changes in the basement membrane and the epidermis.

Title: Development of human skin equivalents mimicking skin aging : contrast between papillary and reticular fibroblasts as a lead

The handle http://hdl.handle.net/1887/47933 holds various files of this Leiden University dissertation

Author: Janson, David