Mechanisms of lamellar collagen formation in connective tissues

Biomaterials 97 (2016) 74e84

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Review

Mechanisms of lamellar collagen formation in connective tissues Samaneh Ghazanfari a, b, c, d, *, Ali Khademhosseini c, d, e, f, g, Theodoor H. Smit a, b a

Department of Orthopedic Surgery, VU University Medical Center, Amsterdam, The Netherlands MOVE Research Institute, VU University, Amsterdam, The Netherlands c Biomaterials Innovation Research Center, Division of Biomedical Engineering, Department of Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, MA, USA d Harvard-Massachusetts Institute of Technology, Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, MA, USA e Wyss Institute for Biologically Inspired Engineering, Harvard University, Boston, MA, USA f Department of Bioindustrial Technologies, College of Animal Bioscience and Technology, Konkuk University, Hwayang-dong, Gwangjin-gu, Seoul, Republic of Korea g Department of Physics, King Abdulaziz University, Jeddah, Saudi Arabia b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 29 January 2016 Received in revised form 29 March 2016 Accepted 20 April 2016 Available online 27 April 2016

The objective of tissue engineering is to regenerate functional tissues. Engineering functional tissues requires an understanding of the mechanisms that guide the formation and evolution of structure in the extracellular matrix (ECM). In particular, the three-dimensional (3D) collagen fiber arrangement is important as it is the key structural determinant that provides mechanical integrity and biological function. In this review, we survey the current knowledge on collagen organization mechanisms that can be applied to create well-structured functional lamellar tissues and in particular intervertebral disc and cornea. Thus far, the mechanisms behind the formation of cross-aligned collagen fibers in the lamellar structures is not fully understood. We start with cell-induced collagen alignment and strain-stabilization behavior mechanisms which can explain a single anisotropically aligned collagen fiber layer. These mechanisms may explain why there is anisotropy in a single layer in the first place. However, they cannot explain why a consecutive collagen layer is laid down with an alternating alignment. Therefore, we explored another mechanism, called liquid crystal phasing. While dense concentrations of collagen show such behavior, there is little evidence that the conditions for liquid crystal phasing are actually met in vivo. Instead, lysyl aldehyde-derived collagen cross-links have been found essential for correct lamellar matrix deposition. Furthermore, we suggest that supra-cellular (tissue-level) shear stress may be instrumental in the alignment of collagen fibers. Understanding the potential mechanisms behind the lamellar collagen structure in connective tissues will lead to further improvement of the regeneration strategies of functional complex lamellar tissues. © 2016 Elsevier Ltd. All rights reserved.

Keywords: Collagen Arrangment Lamellar structure Liquid crystal behavior Functional tissue

1. Introduction The functionality of a lamellar tissue -such as the annulus fibrosus of the intervertebral disc (Fig. 1A) and the cornea (Fig. 1B) whose collagen fibers are organized in layers with alternating orientation-is connected with its structure [1]. Although enormous progress has been made in tissue engineering, no engineered tissue has entered routine clinical practice yet mainly because they do not have the appropriate structure and function of their native

* Corresponding author. Gustav Mahlerlaan 3004, 1081 LA, Amsterdam, The Netherlands. E-mail address: [email protected] (S. Ghazanfari). http://dx.doi.org/10.1016/j.biomaterials.2016.04.028 0142-9612/© 2016 Elsevier Ltd. All rights reserved.

counterparts [4,5]. During embryonic development, collagen fibers in many tissues, such as the cornea, assemble in consecutive layers with alternating orientation [6,7]. Reconstruction of such a complex native-like structure requires control over the fibrous arrangement of collagen and other matrix components. Disordered structures such as tumors or fibrotic tissue are disfunctional because tissue function strongly depends on ECM fiber arrangement [8]. Creating such structures thus is an essential aspect of functional tissue engineering. Collagen is the main fibrous and structural protein of the ECM and a major determinant of mechanical properties of connective tissues [1,9,10]. Collagen structural arrangement results in long term in vivo biomechanical stability of engineered tissues [1,11], but the mechanisms that create collagen anisotropy are not well

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Fig. 1. (A) Upper, schematic illustration of the intervertebral disc composed of a gel-like center called nucleus pulposus (NP) and lamellar layers of collagen fibers with alternating directions called annulus fibrosus (AF). Lower, 3D representation of the ovine AF using optical coherence tomography. Individual images on the right side with circled numbers (taken from approximately the location of corresponding numbers in the left image) show how collagen orientation is changed in depth. Scale bar represents 500 mm. Reprinted from Han et al. [2], with permission from John Wiley and Sons, copyright 2015. (B) Collagen orientation changes with depth in chicken cornea. The arrows indicate the two principal directions. Reprinted from Boote et al. [3], with permission from Investigative ophthalmology & visual science, copyright 2011.

understood. Therefore, we will review the underlying mechanisms behind collagen fiber alignment to create functional lamellar tissues in vitro. Many researchers have suggested that cells are the driving force behind matrix anisotropy [12e18]. Cells generate traction forces during tissue development, and the exerted force on the ECM is generally largest along the main axis of the cells [12e14]. The resulting mechanical stress applied to the tissue is considered to be responsible for the (re-)organization of collagen fibers [15e18]. Also, cell-generated forces can collectively create supra-cellular or tissue-level stresses, which affect alignment of the cells and collagen fibers as they are often aligned with the maximum shear stress [19,20]. Other researchers have suggested that preferred alignment occurs due to the strain-stabilization behavior in which fibers under mechanical strain are supported against remodeling through enzymatic degradation. Thus, collagen fibrils can assemble into specific patterns even in an a-cellular context [21,22]. These two mechanisms can describe simple one layer anisotropic alignment of collagen fibers. However, it is not clear how multiple-layers of collagen fibers stack together in lamellae-like tissues. Another mechanism called liquid crystalline behavior has been presented as a potential underlying mechanism behind multilayered cross-aligned tissues [23e27]. Liquid crystal is a state of material in between the liquid and the crystal in which the material can flow like a fluid and has orientational order like a solid. It is as yet unclear under which specific conditions this state happens in vivo and how to replicate that in a physiological condition in vitro. It is also not fully understood yet which mechanism or maybe combination of mechanisms are creating the multi-layered structural arrangement of collagen in native tissues. The aim of this review paper is to provide the current thinking on the potential of these mechanisms to establish a self-assembled arrangement of collagen fiber network in vitro. We first provide an overview on collagen fibrillogenesis and the important components controlling individual collagen fibril formation. Then, we review the mechanisms involved in suprafibrillar and tissue-specific collagen arrangement which are less understood.

2. Collagen fibrillogenesis In vitro, collagen fibril self-assembly from collagen molecules is a spontaneous and entropy-driven procedure caused by the loss of solvent from the collagen solution [28]. Purified collagen molecules in acidic solutions can bind to each other and assemble to fibrils at a certain concentration, temperature and pH with no other molecules involved [29]. The structure of the fibrils formed in vitro depends on environmental factors like buffer composition and temperature [1]. It was shown, for instance, that phosphate is an essential element in the formation of well periodically banded collagen fibrils, and fibrils formed at lower temperatures (~20
C) had a larger diameter than those formed at higher temperatures [30]. Although in vitro relevant parameters can be controlled, replicating structures with a unique morphology and defined size remains a challenge. Self assembly of proteins such as collagen can also be guided by external forces such as electric field to create structures with a unique morphology in vitro [31]. Fibril formation in vivo or by cultured cells is much more complex since about 50 molecules are known to interact with the fibrillary collagen molecules [29,32]. Collagen fibril formation needs regulators, such as fibronectin and collagen binding integrins, as well as nucleators, like collagen V and XI [29]. Collagen fibril assembly was completely inhibited in human smooth muscle cell cultures by inhibition of fibronectin assembly and the presence of an anti-a5b1 integrin antibody [33]. An anti-a2b1 integrin antibody inhibited the assembly of collagen fibrils in the same study [33]. Ledger et al. showed localization of fibronectin and procollagen in the secretory pathways of cultured fibroblasts meaning that these proteins interact within the cells before secretion in the ECM [34]. Thus, in vivo, other factors are in place to control what is otherwise a protein self-assembly process. Collagen V, as a collagen fibril nucleator, is an important regulator of collagen fibril diameter [35e37]. Collagen V is codistributed with collagen I in the same fibrils [35]. Analysis on collagen fibril structure in the cornea of mice with targeted deletion of Col5a1 gene showed that fibril diameter was increased and the density of the fibrils was decreased as compared to those seen in

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the wild-type mice [38]. Moreover, collagen V is essential for the assembly of collagen I in vivo since mice lacking collagen V resulted in embryonic lethality due to the lack of fibril formation in the mesenchyme [39]. Collagen XI is also required to nucleate the thin (~16 nm) collagen fibril assembly that exists in cartilage and cornea [40,41]. Collagen XI mutations in mice revealed a tremendous effect on cartilage morphogenesis due to the lack of thin collagen fibrils [42]. In addition to nucleator and organizer components, there are many other proteins and molecules that regulate the assembly and structure of collagen fibrils [29,43]. For instance, lysyl oxidase that enzymatically cross-links collagen molecules is essential for fibril formation and regulates its fibril shape [44]. Inhibition of lysyl oxidase hinders the development of tendon and causes shape irregularity of the collagen fibrils [44]. Lysyl oxidase does not influence fibril diameter in the cornea, but proved essential for the lamellar organization of collagen [45]. Tissue transglutaminase-2 is required for the self-assembly of the thin fibrils in the cornea [45]. Other important proteins that regulate the spatial arrangement and size of collagen fibrils in the cornea stroma are proteoglycans, which bind to specific receptor sites [46,47]. Mice with deficiency in lumican, a type of proteoglycan, showed abnormal fibril structure in the developing cornea [48]. Decorin and dermatan sulfate play a role in the slippage of collagen fibrils over each other [49,50] and in collagen fibrillar structural arrangement [51], respectively. Still many proteins and molecules affect fibril formation, but there are other mechanisms involved in governing the suprafibrillar and tissue-specific collagen fiber organization which we address in the following section. 3. Collagen suprafibrillar assembly mechanisms in vitro and in vivo 3.1. Cell-induced collagen alignment Cells actively remodel their collagen matrix by synthesis, degradation [52e54] and (re-) organization [55]. They generate traction forces during tissue development, and the resulting mechanical stress applied to the matrix, largest along the main axis of the cells, is responsible for reorganization of the fibers [12e18]. It has been shown for instance in the developing tissues (e.g. tendon) [56,57] that cell traction forces generated by actin fibers align with the extracellular collagen fibers. Collagen is deposited in parallel arrangement through plasma membrane protrusions, which are cell membrane transport pathways, called fibripositors (fibril depositors) [58]. Fibripositors align with the deposited collagen in the ECM and with the actin fibers inside the cells. This appeared as an essential feature in matrix production, because perturbation of actin fibers leads to a quick loss of fibripositors and thus disruption of deposited collagen fibers parallel alignment [58]. Moreover, the 3D reconstruction of scanning electron microscopy images of the developing cornea revealed that cell orientation determines the direction of the secreted collagen fibrils [59]. Thus, actin fiber and fibroblast-like cell alignment co-operate in matrix alignment. Cells are able to sense changes in the mechanical environment (e.g. strain) which is transmitted from the ECM to the intracellular environment through adhesion receptors (e.g. integrin receptors, mechanosensitive ion channels and tyrosine kinases) that connect to the cytoskeleton [60e65]. Eventually, cells respond to the stimulus via actin fibers and subsequently affect matrix alignment. In 2D, cells show stretch-avoidance behavior in response to uniaxial cyclic loading such that they align perpendicular to the stretch direction which minimizes the applied stress [66e70]. On the other hand, cells show no preferred orientation when exposed to biaxial cyclic loading [71].

Cells behave differently in a 3D environment as shown in Fig. 2. In a static 3D collagen gel, fibrin gel or a fast degrading scaffold, seeded cells and the produced collagen matrix align in the constrained direction [31,72e75]. Cellular contractile stresses in the constrained direction may be responsible for this preferred alignment [14,76,77]. On the other hand, cells and their stress (actin) fibers show stress-avoidance response at the tissue surface when a biaxially constrained tissue is exposed to uniaxial cyclic loading. Cells and stress fibers align biaxially in the core of the tissue, regardless of the tissue being stretched or not [78]. Summarizing, it is evident that in vivo mechanical loading, either uniaxial (e.g. in tendon) or biaxial (e.g. in cornea due to the internal ocular pressure or in blood vessels due to the pulsatile blood flow) plays a major role in tissue (re)structuring. Collagen fibers align in a preferred direction under mechanical stress and subsequently dictate the alignment of cells, but cells themselves are also sensitive to mechanical loading and can align collagen fibers. 3.2. Strain-stabilization The degradation of collagen fibers is dependent on matrix metalloproteinases (MMPs) which are a family of zinc-dependent proteolytic enzymes. MMP activity is regulated by tissue inhibitors of MMPs (TIMPs) [21]. Strain-stabilization theory suggests that collagen fibers in the presence of degrading enzymes, such as MMPs, constitute a smart complex in which the fibers under mechanical strain are protected against degradation [21,22]. In other words, specific fiber alignment is created by selectively removing unloaded or less loaded fibers, which results in a phenomenon called strain stabilization [79,80]. MMPs and TIMPs are usually expressed at low level in adult tissues and under normal physiological condition. However, they are upregulated during remodeling and under pathological conditions [81]. Furthermore, the balance of collagen synthesis and degradation is lost with aging by decreasing the ratio of collagen synthesis to degradation, which leads to collagen-related diseases like arthritis [82e84]. By applying uniaxial dynamic loading to tendon in vitro, enzymatic degradation slows down [85]. A computational model developed by Hadi et al. showed that fibers in the central region of a dogbone align transversely to the loading direction and therefore experience more isotropic load and undergo more decay than those aligned in the stretch direction [86]. Flynn et al. showed that only strained fibrils in a micro-network of reconstituted (in vitro assembled) collagen I and in the presence of a collagen cleaving enzyme were preserved (Fig. 3A) [21]. Ruberti et al. demonstrated that by applying strain to a natural collagen network of cornea, the fibers under tension were preserved while the fibers under less or no tension were degraded (Fig. 3B) [79]. The molecular mechanism of collagen degradation is poorly understood but important to explore. There is a theory that suggests MMPs unwind the helical structure of collagen fibrils by binding to it. However, another theory suggests that collagen fiber thermally unwinds locally by itself in proximity of the cleavage site before MMPs bind to it. Collagen contains imino-rich and iminopoor regions. Imino-rich regions are located preceding and imino-poor regions are located following the cleavage sites. Also, the melting point of collagen is related to its imino acid content, meaning that imino-poor regions have less stability and more flexibility which leads to better recognition of collagenase [85]. The thermal stability of collagen fibrils increases when exposed to tensile load due to the loss of configurational entropy of collagen molecules [79,87]. Bhole et al. suggested that by applying tension to collagen fibrils, the enzymatic binding sites become less exposed due to the geometrical changes of collagen monomers and three alpha chains [88]. However, Adhikari et al. showed that straining

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Fig. 2. Schematic view of cells and collagen orientation in uniaxially or biaxially 3D constrained constructs in the presence of static or dynamic loading. Collagen fibers orient in the direction of the constraint in both static (A) and cyclically (B) strained tissues when the construct is uniaxially constrained. However, when the construct is biaxially constrained, the collagen fibers align biaxially in statically strained (C) and in the core of cyclically strained (D) tissues, but align perpendicularly to the strain direction at the surface of cyclically strained tissues (E).

may speed up collagen degradation, but the rate of collagen trimmer, proteolysis, by collagenase did not seem to be dependent on force [89]. The application of force presumably turns the single molecule into a state more susceptible to enzymatic degradation. Chang and Buehler suggested that there are in fact two vulnerable situations of the collagen molecule cleavage sites: micro unfolding conformation (where mechanical strain can slow down the degradation rate) and unwinding conformation (where mechanical strain can speed up the cleavage rate) [85]. In 1977, Huang and Yannas discovered that enzymatic degradation of reconstituted collagen fibers first decreases and then increases with increased straining. With straining up to 4%, degradation rate decreases because the porosity of the collagenous construct decreases; as a consequence, the diffusion rate of the enzyme through the fibers decreases. But when the strain increases above a certain level, degradation rate increases because few fibers disrupt and a sudden stress applied to the neighboring fibers induce the breaking down of more fibers [90]. Based on this, Ghazanfari et al. suggested that the minimum degradation rate may explain the preferential collagen alignment of biaxially loaded samples [91]. To investigate that, they applied a range of uniaxial strains to a decellularized pericardium, which contains isotropic collagen matrix. They found that the degradation rate decreases and then increases with increasing uniaxial strains. Then, by

applying different sets of biaxial strains, they observed that the fibers aligned in the direction of the strain with a lower degradation rate. Thus, collagen orientation can be modulated by straincontrolled enzymatic degradation, but future studies are needed to better understand the molecular mechanisms associated with this observation. 3.3. Liquid crystal phasing Liquid crystallinity is a physical state between liquid and solid phases which refers to alignment and layeredness [24]. It has been suggested that collagen fibers can arrange based on liquid crystal assembly as it appears that a layered structure with alternating alignment principally forms during morphogenesis [23e27,92]. Collagen assembly in native tissues, such as the cornea, strongly resembles collagen fiber arrangement observed in the liquid crystalline phase [93e99]. This means that a 3D hierarchical assembly of collagen molecules can also develop in an a-cellular context [100]. However, in vitro, the liquid crystalline behavior of collagen happens only at low pH as there is a strong repulsion between the negatively charged collagen monomers [101,102]. Increasing the pH after liquid crystal formation of collagen molecules elevates the hydrophobic interaction of collagen molecules, which initiates fibril formation without changing the molecular orientation. This

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Fig. 3. Microscopy images of reconstituted (A) and natural (B) collagen alignment in the presence of uniaxial strain and a degrading enzyme. (A) Stretching reconstituted collagen fibrils over 100% of their initial length in the presence of a degrading enzyme by micropipettes. Fibrils in the path of applied strain are preserved. However, non-strained fibrils are degraded (Bar ¼ 10 mm). Reprinted from Flynn et al. [21]. (B) Strips of cornea subjected to a degrading enzyme (1), uniaxial strain (2), a degrading enzyme and uniaxial strain (3). There is no preferential collagen alignment in 1 and there is a slight change in the fiber alignment in 2. However, there is a clear preferential alignment in 3 (original mag 200). Reprinted from Ruberti et al. [79], with permission from Elsevier, copyright 2005.

happens because a given amino acid from one triple helix will be exposed to many amino acids of adjacent triple helices [101e103]. Indeed, collagen fibril morphology is highly dependent on the pH of the environment: increasing the pH of the medium from about neutral (6.9e7.4) to (7.6e8.1) results in an increase of fibril diameter from 20 nm to 200 nm in chick corneal epithelium [104]. A schematic representation of different forms of collagen selforganization (aligned/nematic, twisted/cholesteric and isotropic) is shown in Fig. 4. Collagen molecules tend to align in the same direction in the nematic phase and towards different directions in the isotropic phase. They only organize into layers with alternating orientation in cholesteric phase, but this process is not yet well understood. Collagen structure strongly depends on collagen concentration (Fig. 5). Liquid crystal behavior appears at the concentration between 40 and 80 mg/ml [97]. By increasing the collagen concentration to about 80 mg/ml, the fibril arrangement changes towards

a dense cholesteric arrangement [100]. This concentration is similar to procollagen concentration in secretory vesicles of the cells, which is about several tens of mg/ml [106]. These molecules are secreted in the ECM in nematic order (all in the same direction without positional order) [106]. Many studies have shown that collagen fibrillogenesis occurs from acid soluble collagen in vitro by bringing a highly concentrated collagen solution to neutral pH, which lowers the repulsive forces, either in the presence of sodium hydroxide or ammonia vapor [103,107,108]. In order to mimic such liquid crystal-like structures for tissue regeneration applications, we need to get insight into how these structures can be created in vitro. Collagen concentration of acid soluble solution can be increased by dialysis, evaporation and injection methods in vitro (Fig. 6) [109]. These methods are briefly explained in this section.

Fig. 4. Schematic representative of different forms of collagen molecules assembly which are aligned (nematic) (A), twisted (cholesteric) (B) and isotropic (C). Collagen molecules are highly aligned in the nematic phase and not aligned in the isotropic phase. In the cholesteric phase, the collagen molecules orientation rotates between consecutive layers. Reprinted from Nassif et al. [105], with permission from American Chemical Society, copyright 2010.

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Fig. 5. Effect of collagen concentration and the pH on collagen fibrils arrangement and morphology. (A) Phase transition of collagen solution prepared in acetic acid from isotropic to cholesteric at different concentrations. Reprinted from Gobeaux et al. [96], with permission from American Chemical Society, copyright 2007. (B) Transformation of the liquid crystal to a stabilized gel by neutralization; continuous cholesteric twist of collagen monomers transform into a discrete cholesteric twist after stabilization. Reprinted from Besseau et al. [100], with permission from Elsevier, copyright 1995. (C) Variation of width of consecutive layers of collagen fibrils, fibril diameter and rotational angle of fibrils in each layer with collagen concentration. Fiber width decreased, the number of major collagen fiber orientations in half a pitch (P/2) increased and P/2 decreased with an increase in collagen concentration. Reprinted from Mosser et al. [26], with permission from Elsevier, copyright 2006.

Fig. 6. Schematic illustration of current methods for obtaining concentrated collagen gel. (A) Dialysis, (B) Injection and dialysis, (C) Evaporation, (D) Injection and evaporation.

3.3.1. Dialysis With dialysis, low concentration collagen solution at acidic pH is exposed to a high osmolarity solution, such as PEG, separated by a dialysis tube. With osmolaric flow, a high concentration state can be reached [102]. To initiate fibrillogenesis, pH is increased to

neutral. It was observed that collagen fibers align in the plane of the flattened gel in the most concentrated area (margin of the gel) and the orientation was slowly changed to the perpendicular direction away from the margin (Fig. 7). Co-secretion of glycosaminoglycans and collagen by chondrocytes in vivo may have the same effect as

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structures were either nematic (at 20 mg/ml) or precholestric (at 40 mg/ml) (Fig. 8) [103]. The concentration of collagen cannot increase more due to the high viscosity of the solution. This approach results in a liquid crystal-like collagen structure in vitro. However, evaporation is not considered as a potential mechanism in vivo.

Fig. 7. Polarizing micrograph including quantitative plots of average fiber orientation in 60  60 mm pixels. The orientation slightly changes by increasing the distance from the margin of the gel. Reprinted from Knight et al. [102], with permission from John Wiley and Sons, copyright 1998.

using PEG in that they both absorb water and initiate liquid crystalline formation [102]. 3.3.2. Evaporation In this approach, collagen solution with low concentration and low pH is gradually concentrated by evaporation of the solvent [26,92,100,102,103,109,110]. After reaching a high concentration (above 20 mg/ml), the pH is increased to neutral to initiate fibrillogenesis. Depending on the final collagen concentration,

Fig. 8. Transmission electron microscopy (TEM) images of the nematic (A) and precholestric (B) phase of collagen solutions. Collagen fibrils are aligned toward the same direction in the nematic phase (A), while the fibrils orientation rotated periodicaly in the precholestric phase. Reprinted from Besseaua et al. [103], with permission from Elsevier, copyright 2002.

3.3.3. Injection Here low concentration collagen solution at acidic pH is gradually injected into a chamber. This creates a concentration gradient due to the evaporation of the solution, which is highest at the surface [109]. When collagen reaches a concentration above 45 mg/ ml, collagen molecules assemble into a cornea-like plywood structure, with collagen orientation changing by 90
between consecutive layers (2e10 mm half pitch) [97]. Abrupt changes in collagen orientation in e.g. cornea may be due to molecules like sugars, proteoglycans, or glycosaminoglycans, which by making hydrogen bonds can disassemble large collagen fibrils, just as acidic pH does. These molecules disrupt and mask the water hydrogen bonds, which are between opposing helices, and allow collagen to change from an attractive regime to a repulsive one so that collagen fibrils can disassemble [97,111,112]. 3.3.4. Injection and dialysis Wang et al. combined injection and dialysis to obtain reproducible and dense homogeneous structures comparable to native connective tissues [109]. This is not achievable by each individual method, because of the limited collagen solution volume. In order to form dense fibrillar collagen matrices and initiate fibrillogenesis, the pH was increased from acidic pH to a range of 9e10. Similar to other studies, aligned or helicoidal geometries formed depending on the collagen concentration [109]. 4. Discussion Cells are important regulators of collagen matrix anisotropy: they modulate synthesis and degradation of the collagen fibers under mechanical stress [113e115]. Furthermore, they may secrete collagen fibers via fibripositors in the direction of their main axis, or realign existing collagen fibers towards a preffered direction by contraction via actin fibers [11e18,55,57,58,116e118]. There is another mechanism, called stain-stabilization, in which mechanical load affects the enzymatic degradation of collagen fibers and therefore modulate their alignment without the interaction of cells [79,80,90,91]. Thus, collagen fiber alignment can be regulated by mechanical stress and subsequently induce the alignment of cells thereof through contact guidance. The combination of cell-induced collagen orientation and strain-stabilization may explain the anisotropic alignment of a single layer of collagen fibers in vivo; however, the mechanism of layer formation in collagenous tissues remains obscure. Thus far, the layered structure of collagen fibers has been explained with the liquid crystal theory. It is well documented that in vitro, purified soluble collagen molecules assemble into fibrillar collagen by raising pH from acidic, where repulsion between charged collagen molecules is large enough to avoid the molecule aggregation, to neutrality or beyond [103,107,108]. However, in vivo, the transition from molecular to fibrillar collagen happens by enzymatic cleavage of N- and C-terminal propeptides which induces a decrease in the solubility of collagen molecules [27,119,120]. High concentration of collagen is essential in order to reach the liquid crystal phase. Interestingly, the concentration in which the liquid crystalline structure is formed in vitro is comparable to concentration of procollagen in secretory vesicles of the cells in vivo [106]. The concentration of collagen may increase by the presence of ECM proteins that absorb water, for instance glycosaminoglycans

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Fig. 9. Effect of micro patterned surface, Reprinted from Singh et al. [130], with permission from Royal Society of Chemistry, copyright 2014 (A) and scaffold fibers, Reprinted from de Jonge et al. [127], with permission from Mary Ann Liebert, Inc., copyright 2014 (B) on collagen fiber alignment stained for collagen (green) (A,B) and cell cytoplasm (red) (B). Double arrow indicate the orientation of the substrate pattern (A). Scale bar represents 100 mm (A) and 50 mm (B). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

[102]. Although there are explanation as to how the liquid crystal phasing can occur in vivo, there is little evidence for this. Lamellar bone is often presented as a typical example of a biological tissue with a liquid crystal-like structure. However, bone essentially differs from soft tissues like cornea and annulus fibrosus both in formation and structure. While the layered structure of cornea and annulus fibrosus form almost instantly during early embryonic development, lamellar bone formation is a slow remodeling process in adults taking weeks to complete. Biological clocks, inducing oscillations in development and physiology, have been suggested to control the incremental structure formation in bone [121]. Furthermore, there is doubt regarding the presence of an alternating collagen fiber orientation in lamellar bone, routinely shown in text books. Marotti et al. showed that the lamellar bone essentially consists of alternating dense acellular and loose cellular lamellae instead of alternating alignment of collagen fibers [122]. This sheds some doubt on the suggested similarities between lamellar bone and liquid crystal structure observed in dense collagen solutions in vitro [25,26,95,98e100,110]. The mechanisms discussed in this review could mimic the underlying mechanism of in vivo collagen organization and can be used in in vitro model systems to create engineered tissues with a native-like collagen structure. However, there are other methods which can be used to control collagen organization in vitro. For example, collagen and cell alignment can be guided by topological cues, referred to as “contact guidance”, like microgrooves or fibrous scaffolds (Fig. 9) [123e132]. These patterned layers of cells and collagen matrices can be then stacked together to create a lamellar structure. The effect of contact guidance on cell and collagen alignment is predominant over mechanical loading [125,131]. Wang et al. showed that cells align in the substrate micro-grooves direction regardless of the loading direction, as microgrooves limit the cell orientation [125]. Further, de Jonge et al. showed that collagen fibers orient in the direction of the scaffold fibers in vitro [127]. They also demonstrated that collagen fiber orientation changed towards the constraint direction when scaffold fibers were degraded [127]. The impact of contact guidance cues (stent struts) on the collagen fiber alignment of the adjacent tissue in vivo was also evaluated. It appears that the effect of contact guidance on collagen alignment was predominant over other mechanical loading under physiological condition in vivo [128,130]. It was also suggested that the collagen structure of the primary stroma may

play a contact guidance role to direct the orientation of the secondary stroma in the cornea [58]. This mechanism can be used to regenerate tissues with comparable mechanical properties and collagen anisotropy as native tissues. Further, this mechanism can be employed to create a scaffold with appropriate template for in situ tissue engineering that can guide the formation of collagen matrix in a desired direction. It must be emphasized, however, that cells used in the in vitro studies discussed here are much more mature than the cells present in early development where cornea and annulus fibrosus originate; this is reflected in their capacity to condensate [133] and in their contractile capacity [134]. This difference also applies to the cells that form lamellar bone, as compared to cells that from the cornea or the annulus fibrosus. In conclusion, the presented collagen fiber assembly mechanisms provide fundamental information for designing new tissue engineering strategies to mimic the arrangement of the basic structural component of connective tissues. Cells may assemble the structure of lamellar collagen, but collagen fibers also may selfassemble into lamellar structures, in particular at high densities. The mechanical stress field is a strong determinant of orientation, both for the cells and for the collagen fibers. Understanding the origin of lamellar, alternating aligned collagen structures is important for the tissue engineering of functional tissues like cornea and annulus fibrosis. Acknowledgments S. Ghazanfari and T.H. Smit acknowledge financial support from a ZonMW-VICI grant 918.11.635 (The Netherlands). References [1] P. Fratzl, in: Collagen: Structure and Mechanics, Springer Science & Business Media, 2008. [2] S.K. Han, C.W. Chen, J. Wierwille, Y. Chen, A.H. Hsieh, Three dimensional mesoscale analysis of translamellar cross-bridge morphologies in the annulus fibrosus using optical coherence tomography, J. Orthop. Res. 33 (2015) 304e311. [3] C. Boote, A. Elsheikh, W. Kassem, C.S. Kamma-Lorger, P.M. Hocking, N. White, C.F. Inglehearn, M. Ali, K.M. Meek, The influence of lamellar orientation on corneal material behavior: biomechanical and structural changes in an avian corneal disorder, Invest. Ophthalmol. Vis. Sci. 52 (2011) 1243e1251. [4] L.G. Griffith, G. Naughton, Tissue engineeringecurrent challenges and expanding opportunities, Science 295 (2002) 1009e1014. [5] N.L. Nerurkar, D.M. Elliott, R.L. Mauck, Mechanical design criteria for intervertebral disc tissue engineering, J. Biomech. 43 (2010) 1017e1030.

82

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[6] C. Cintron, H. Covington, C.L. Kublin, Morphogenesis of rabbit corneal stroma, Invest. Ophthalmol. Vis. Sci. 24 (1983) 543e556. [7] E.D. Hay, J.P. Revel, Fine Structure of the Developing Avian Cornea, Karger, Basel, 1969. [8] M.E. Nimni, Collagen: structure, function, and metabolism in normal and fibrotic tissues, Semin. Arthritis Rheum. 13 (1983) 1e86. [9] P. Fratzl, K. Misof, I. Zizak, G. Rapp, H. Amenitsch, S. Bernstorff, Fibrillar structure and mechanical properties of collagen, J. Struct. Biol. 122 (1998) 119e122. [10] S. Fazaeli, S. Ghazanfari, V. Everts, T.H. Smit, J.H. Koolstra, The contribution of collagen fibers to the mechanical compressive properties of the temporomandibular joint disc, Osteoarthr. Cartil. (2016), http://dx.doi.org/10.1016/ j.joca.2016.01.138 pii: S1063-4584(16)00157-6. [11] M. Santin, Strategies in Regenerative Medicine: Integrating Biology with Materials Design, Springer Science & Business Media, 2009. [12] J. Hirai, T. Matsuda, Venous reconstruction using hybrid vascular tissue composed of vascular cells and collagen: tissue regeneration process, Cell Transpl. 5 (1996) 93e105. [13] R.J. Klebe, H. Caldwell, S. Milam, Cells transmit spatial information by orienting collagen fibers, Matrix 9 (1989) 451e458. [14] V.H. Barocas, R.T. Tranquillo, An anisotropic biphasic theory of tissue equivalent mechanics: the interplay among cell traction, fibrillar network deformation, fibril alignment, and cell contact guidance, J. Biomech. Eng. 119 (1997) 137e145. [15] S.L.M. Dahl, M.E. Vaughn, L.E. Niklason, An ultrastructural analysis of collagen in tissue engineered arteries, Ann. Biomed. Eng. 35 (2007) 1749e1755. [16] M. Chiquet, Regulation of extracellular matrix gene expression by mechanical stress, Matrix Biol. 18 (1999) 417e426. [17] M. Chiquet, L. Gelman, R. Lutz, S. Maier, From mechanotransduction to extracellular matrix gene expression in fibroblasts, Biochim. Biophys. Acta 1793 (2009) 911e920. [18] M.A. van Vlimmeren, A. Driessen-Mol, C.W. Oomens, F.P. Baaijens, An in vitro model system to quantify stress generation, compaction, and retraction in engineered heart valve tissue, Tissue Eng. C. Methods 17 (2011) 983e991. [19] M. Eastwood, V. Mudera, D. Mcgrouther, R. Brown, Effect of precise mechanical loading on fibroblast populated collagen lattices: morphological changes, Cell Motil. Cytoskelet. 40 (1998) 13e21. [20] J.H.C. Wang, P. Goldschmidt-Clermont, J. Wille, F.C.P. Yin, Specificity of endothelial cell reorientation in response to cyclic mechanical stretching, J. Biomech. 34 (2001) 1563e1572. [21] B.P. Flynn, A.P. Bhole, N. Saeidi, M. Liles, C.A. Dimarzio, J.W. Ruberti, Mechanical strain stabilizes reconstituted collagen fibrils against enzymatic degradation by mammalian collagenase matrix metalloproteinase 8 (MMP8), PLoS ONE 5 (2010) e12337. [22] H. Birkedal-Hansen, W. Moore, M. Bodden, L. Windsor, B. Birkedal-Hansen, A. DeCarlo, J. Engler, Matrix metalloproteinases: a review, Crit. Rev. Oral Biol. Med. 4 (1993) 197e250. [23] D.W.L. Hukins, J. Woodhead-Galloway, Liquid crystal model for the organization of molecules in collagen fibrils, Biochem. Soc. Trans. 6 (1978) 238e239. [24] N.S. Murthy, Liquid crystallinity in collagen solutions and magnetic orientation of collagen fibrils, Biopolymers 23 (1984) 1261e1267. [25] M.M. Giraud-Guille, Liquid crystallinity in condensed type I collagen solutions: a clue to the packing of collagen in extracellular matrices, J. Mol. Biol. 224 (1992) 861e873. [26] G. Mosser, A. Anglo, C. Helary, Y. Bouligand, M.M. Giraud-Guille, Dense tissue-like collagen matrices formed in cell-free conditions, Matrix Biol. 25 (2006) 3e13. [27] P.D.S. Peixoto, A. Deniset-Besseau, M. Schanne-Klein, G. Mosser, Quantitative assessment of collagen I liquid crystal organizations: role of ionic force and acidic solvent, and evidence of new phases, Soft Matter 7 (2011) 11203e11210. [28] K.E. Kadler, D.F. Holmes, J.A. Trotter, J.A. Chapman, Collagen fibril formation, Biochem. J. 316 (1996) 1e11. [29] K.E. Kadler, A. Hill, E.G. Canty-Laird, Collagen fibrillogenesis: fibronectin, integrins, and minor collagens as organizers and nucleators, Curr. Opin. Cell Biol. 20 (2008) 495e501. [30] B.R. Williams, R.A. Gelman, D.C. Poppke, K.A. Piez, Collagen fibril formation, optimal in vitro conditions and preliminary kinetic results, J. Biol. Chem. 253 (1978) 6578e6585. [31] R. de la Rica, E. Mendoza, L.W. Chow, K.L. Cloyd, S. Bertazzo, H.C. Watkins, J.A. Steele, M.M. Stevens, Self-assembly of collagen building blocks guided by electric fields, Small 10 (2014) 3876e3879. [32] G.A. Di Lullo, S.M. Sweeney, J. Korkko, L. Ala-Kokko, J.D. San Antonio, Mapping the ligand-binding sites and disease-associated mutations on the most abundant protein in the human, type I collagen, J. Biol. Chem. 277 (2002) 4223e4231. [33] S. Li, C. Van Den Diepstraten, S.J. D'Souza, B.M.C. Chan, J.G. Pickering, Vascular smooth muscle cells orchestrate the assembly of type I collagen via alpha2beta1 integrin, RhoA, and fibronectin polymerization, Am. J. Pathol. 163 (2003) 1045e1056. [34] P.W. Ledger, N. Uchida, M.L. Tanzer, Immunocytochemical localization of procollagen and fibronectin in human fibroblasts: effects of the monovalent ionophore, monensin, J. Cell Biol. 87 (1980) 663e671. [35] D.E. Birk, J.M. Fitch, J.P. Babiarz, T.F. Linsenmayer, Collagen type I and type V

[36]

[37] [38]

[39]

[40]

[41]

[42]

[43] [44]

[45]

[46]

[47]

[48]

[49] [50]

[51]

[52]

[53]

[54]

[55]

[56]

[57]

[58]

[59]

[60] [61]

are present in the same fibril in the avian corneal stroma, J. Cell Biol. 106 (1988) 999e1008. D.E. Birk, J.M. Fitch, J.P. Babiarz, K.J. Doane, T.F. Linsenmayer, Collagen fibrillogenesis in vitro: interaction of types I and V collagen regulates fibril diameter, J. Cell Sci. 95 (1990) 649e657. D.E. Birk, Type V collagen: heterotypic type I/V collagen interactions in the regulation of fibril assembly, Micron 32 (2001) 223e237.
on, W.G. Cole, R.J. Wenstrup, F. Young, A.R. Slomovic, F. Segev, E. He D.S. Rootman, D. Whitaker-Menezes, I. Chervoneva, D.E. Birk, Structural abnormalities of the cornea and lid resulting from collagen V mutations, Invest. Ophthalmol. Vis. Sci. 47 (2006) 565e573. R.J. Wenstrup, J.B. Florer, E.W. Brunskill, S.M. Bell, I. Chervoneva, D.E. Birk, Type V collagen controls the initiation of collagen fibril assembly, J. Biol. Chem. 279 (2004) 53331e53337. D.R. Keene, J.T. Oxford, N.P. Morris, Ultrastructural localization of collagen types II, IX, and XI in the growth plate of human rib and fetal bovine epiphyseal cartilage: type XI collagen is restricted to thin fibrils, J. Histochem. Cytochem. 43(995) 967e979. Y. Li, D.A. Lacerda, M.L. Warman, D.R. Beier, H. Yoshioka, Y. Ninomiya, J.T. Oxford, N.P. Morris, K. Andrikopoulos, F. Ramirez, A fibrillar collagen gene, Col11a1, is essential for skeletal morphogenesis, Cell 80 (1995) 423e430. R. Seegmiller, F.C. Fraser, H. Sheldon, A new chondrodystrophic mutant in mice. Electron microscopy of normal and abnormal chondrogenesis, J. Cell Biol. 48 (1971) 580e593. R.P. Mecham, The Extracellular Matrix: an Overview, Springer-Verlag, Germany, 2011. A. Herchenhan, F. Uhlenbrock, P. Eliasson, M. Weis, D. Eyre, K.E. Kadler, S.P. Magnusson, M. Kjaer, Lysyl oxidase activity is required for ordered collagen fibrillogenesis by tendon cells, J. Biol. Chem. 290 (2015) 16440e16450. L. Wang, P.C. Uhlig, E.F. Eikenberry, H. Robenek, P. Bruckner, U. Hansen, Lateral growth limitation of corneal fibrils and their lamellar stacking depend on covalent collagen cross-linking by transglutaminase-2 and lysyl oxidases, respectively, J. Biol. Chem. 289 (2014) 921e929. A.J. Quantock, R.D. Young, Development of the corneal stroma, and the collagen-proteoglycan associations that help define its structure and function, Dev. Dyn. 237 (2008) 2607e2621. J.E. Scott, M. Haigh, ‘Small’-proteoglycan:collagen interactions: keratan sulphate proteoglycan associates with rabbit corneal collagen fibrils at the ‘a’ and ‘c’ bands, Biosci. Rep. 5 (1985) 765e774. S. Chakravarti, G. Zhang, I. Chervoneva, L. Roberts, D.E. Birk, Collagen fibril assembly during postnatal development and dysfunctional regulation in the lumican-deficient murine cornea, Dev. Dyn. 235 (2006) 2493e2506. J.B. Bard, M.K. Bansal, A.S. Ross, The extracellular matrix of the developing cornea: deposition and function, Development 103 (1988) 195e205. G.D. Pins, D.L. Christiansen, R. Patel, F.H. Silver, Self-assembly of collagen fibres. Influence of fibrillar alignment and decorin on mechanical properties, Biophys. J. 73 (1997) 2164e2172. R.A. Hahn, D.E. Birk, b-D xyloside alters dermatan sulfate proteoglycan synthesis and the organization of the developing avian corneal stroma, Development 115 (1992) 383e393. T.D. Nguyen, R. Liang, S.L. Woo, S.D. Burton, C. Wu, A. Almarza, M.S. Sacks, S. Abramowitch, Effects of cell seeding and cyclic stretch on the fiber remodeling in an extracellular matrix-derived bioscaffold, Tissue Eng. Part A 15 (2009) 957e963. B. Kim, J. Nikolovski, J. Bonadio, D.J. Mooney, Cyclic mechanical strain regulates the development of engineered smooth muscle tissue, Nat. Biotechnol. 17 (1999) 979e983. U.A. Stock, D. Wiederschain, S.M. Kilroy, D. Shum-Tim, P.N. Khalil, J.P. Vacanti, J.E. Mayer Jr., M.A. Moses, Dynamics of extracellular matrix production and turnover in tissue engineered cardiovascular structures, J. Cell. Biochem. 81 (2001) 220e228. N. de Jonge, F.M. Kanters, F.P. Baaijens, C.V. Bouten, Strain-induced collagen organization at the micro-level in fibrin-based engineered tissue constructs, Ann. Biomed. Eng. 41 (2013) 763e774. C. Gealy, A.J. Hayes, R. Buckwell, R.D. Young, B. Caterson, A.J. Quantock, J.R. Ralphs, Actin and type I collagen propeptide distribution in the developing chick cornea, Invest. Ophthalmol. Vis. Sci. 50 (2009) 1653e1658. E.G. Canty, T. Starborg, Y. Lu, S.M. Humphries, D.F. Holmes, R.S. Meadows, A. Huffman, E.T. O'Toole, K.E. Kadler, Actin filaments are required for fibripositor-mediated collagen fibril alignment in tendon, J. Biol. Chem. 281 (2006) 38592e38598. E.G. Canty, Y. Lu, R.S. Meadows, M.K. Shaw, D.F. Holmes, K.E. Kadler, Coalignment of plasma membrane channels and protrusions (fibripositors) specifies the parallelism of tendon, J. Cell. Biol. 165 (2004) 553e563. R.D. Young, C. Knupp, C. Pinali, K.M.Y. Png, J.R. Ralphs, A.J. Bushby, T. Starborg, K.E. Kadler, A.J. Quantock, Three-dimensional aspects of matrix assembly by cells in the developing cornea, PNAS 111 (2014) 687e692. F. Sachs, C.E. Morris, Mechanosensitive ion channels in nonspecialized cells, Rev. Physiol. Biochem. Pharmacol. 132 (1998) 1e77. A.J. Banes, M. Tsuzaki, J. Yamamoto, T. Fischer, B. Brigman, T. Brown, L. Miller, Mechanoreception at the cellular level: the detection, interpretation, and diversity of responses to mechanical signals, Biochem. Cell. Biol. 73 (1995) 349e365.

S. Ghazanfari et al. / Biomaterials 97 (2016) 74e84 [62] A.M. Malek, S. Izumo, Mechanism of endothelial cell shape change and cytoskeletal remodeling in response to fluid shear stress, J. Cell. Sci. 109 (1996) 713e726. [63] B. Geiger, J.P. Spatz, A.D. Bershadsky, Environmental sensing through focal adhesions, Nat. Rev. Mol. Cell. Biol. 10 (2009) 21e33. [64] J.G. Lock, B. Wehrle-Haller, S. Stromblad, Cell-matrix adhesion complexes: master control machinery of cell migration, Semin. Cancer Biol. 18 (2008) 65e76. [65] A.L. Berrier, K.M. Yamada, Cell-matrix adhesion, J. Cell. Physiol. 213 (2007) 565e573. [66] I. Mills, C.R. Cohen, K. Kamal, G. Li, T. Shin, W. Du, B.E. Sumpio, Strain activation of bovine aortic smooth muscle cell proliferation and alignment: study of strain dependency and the role of protein kinase A and C signaling pathways, J. Cell. Physiol. 170 (1997) 228e234. [67] K. Kurpinski, J. Park, R.G. Thakar, S. Li, Regulation of vascular smooth muscle cells and mesenchymal stem cells by mechanical strain, Mol. Cell. Biomech. 3 (2006) 21e34. [68] S. Ghazanfari, M. Tafazzoli-Shadpour, M.A. Shokrgozar, Effects of cyclic stretch on proliferation of mesenchymal stem cells and their differentiation to smooth muscle cells, Biochem. Biophys. Res. Commun. 388 (2009) 601e605. [69] S. Ghazanfari, M. Tafazzoli-Shadpour, M.A. Shokrgozar, N. Amirizadeh, E.J. Rangraz, Morphological changes of mesenchymal stem cells by cyclic stretch, Int. Conf. Biomed. Eng. Inf. 1 (2008) 743e747. [70] S. Ghazanfari, M. Tafazzoli-Shadpour, M.A. Shokrgozar, N. Haghighipour, N. Amirizadeh, Analysis of alterations in morphologic characteristics of mesenchymal stem cells by mechanical stimulation during differentiation into smooth muscle cells, Yakhteh Med. J. 12 (2010) 73e80. [71] R. Kaunasa, S. Usamia, S. Chien, Regulation of stretch-induced JNK activation by stress fiber orientation, Cell. Signal. 18 (2006) 1924e1931. [72] R. Gauvin, R. Parenteau-Bareil, D. Larouche, H. Marcoux, F. Bisson, A. Bonnet, F.A. Auger, S. Bolduc, L. Germain, Dynamic mechanical stimulations induce anisotropy and improve the tensile properties of engineered tissues produced without exogenous scaffolding, Acta Biomater. 7 (2011) 3294e3301. [73] M.R. Neidert, R.T. Tranquillo, Tissue-engineered valves with commissural alignment, Tissue Eng. 12 (2006) 891e903. [74] M.P. Rubbens, A. Driessen-Mol, R.A. Boerboom, M.M. Koppert, H.C. van Assen, B.M. TerHaar Romeny, F.P. Baaijens, C.V. Bouten, Quantification of the temporal evolution of collagen orientation in mechanically conditioned engineered cardiovascular tissues, Ann. Biomed. Eng. 37 (2009) 1263e1272. [75] S. Ghazanfari, A. Driessen-Mol, G.J. Strijkers, F.P.T. Baaijens, C.V.C. Bouten, The evolution of collagen fiber orientation in engineered cardiovascular tissues visualized by diffusion tensor imaging, PLOS ONE 10 (2015) e0127847. [76] A.L.F. Soares, C.W.J. Oomens, F.P.T. Baaijens, A computational model to describe the collagen orientation in statically cultured engineered tissues, Comput. Methods Biomech. Biomed. Engin. 17 (2014) 251e262. [77] V. Barocas, T. Girton, R. Tranquillo, Engineered alignment in media equivalents: magnetic prealignment and Mandrel compaction, J. Biomech. Eng. 120 (1988) 660e666. [78] J. Foolen, V.S. Deshpande, F.M.W. Kanters, F.P.T. Baaijens, The influence of matrix integrity on stress-fiber remodeling in 3D, Biomaterials 33 (2012) 7508e7518. [79] J.W. Ruberti, N.J. Hallab, Strain-controlled enzymatic cleavage of collagen in loaded matrix, Biochem. Biophys. Res. Commun. 336 (2005) 483e489. [80] A.P. Bhole, B.P. Flynn, M. Liles, N. Saeidi, C.A. Dimarzio, J.W. Ruberti, Mechanical strain enhances survivability of collagen micronetworks in the presence of collagenase: implications for load-bearing matrix growth and stability, Philos. Trans. A. Math. Phys. Eng. Sci. 367 (2009) 3339e3362. [81] R.P. Verma, C. Hansch, Matrix metalloproteinases (MMPs): chemicalbiological functions and (Q)SARs, Bioorg. Med. Chem. 15 (2007) 2223e2268. [82] T.E. Cawston, A.J. Wilson, Understanding the role of tissue degrading enzymes and their inhibitors in development and disease, Best. Pract. Res. Clin. Rheumatol. 20 (2006) 983e1002. [83] P.K. Mays, R.J. McAnulty, J.S. Campa, G.J. Laurent, Age-related changes in collagen synthesis and degradation in rat tissues. Importance of degradation of newly synthesized collagen in regulating collagen production, Biochem. J. 276 (1991) 307e313. [84] R.H. Pearce, B.J. Grimmer, M.E. Adams, Degeneration and the chemical composition of the human lumbar intervertebral disc, J. Orthop. Res. 5 (1987) 198e205. [85] S.W. Chang, M.J. Buehler, Molecular biomechanics of collagen molecules, Mater. Today 17 (2014) 70e76. [86] M.F. Hadi, E.A. Sander, J.W. Ruberti, V.H. Barocas, Simulated remodeling of loaded collagen networks via strain-dependent enzymatic degradation and constant-rate fiber growth, Mech. Mater. 44 (2012) 72e82. [87] C.A. Miles, M. Ghelashvili, Polymer-in-a-box mechanism for the thermal stabilization of collagen molecules in fibers, Biophys. J. 76 (1999) 3243e3252. [88] A.P. Bhole, B.P. Flynn, M. Liles, N. Saeidi, C.A. Dimarzio, J.W. Ruberti, Mechanical strain enhances survivability of collagen micro networks in the presence of collagenase: implications for load-bearing matrix growth and stability, Philos. Trans. A Math. Phys. Eng. Sci. 367 (2009) 3339e3362. [89] A.S. Adhikari, E. Glassey, A.R. Dunn, Conformational dynamics accompanying the proteolytic degradation of trimeric collagen I by collagenases, J. Am.

83

Chem. Soc. 134 (2012) 13259e13265. [90] C. Huang, I.V. Yannas, Mechanochemical studies of enzymatic degradation of insoluble collagen fibers, J. Biomed. Mater. Res. 11 (1977) 137e154. [91] S. Ghazanfari, A. Driessen-Mol, C.V.C. Bouten, F.P.T. Baaijens, Modulation of collagen fiber alignment by strain-controlled enzymatic degradation, Acta Biomater. 35 (2016) 118e126. [92] M.M. Giraud-Guille, Liquid crystalline phases of sonicated type I collagen, Biol. Cell. 67 (1989) 97e101. [93] J.E. Kirkwood, G.G. Fuller, Liquid crystalline collagen: a self-assembled morphology for the orientation of mammalian cells, Langmuir 25 (2009) 3200e3206. [94] A.C. Neville, Biology of Fibrous Composites, Cambridge University Press, New York, 1993. [95] M.M. Giraud-Guille, L. Besseau, R. Martin, Liquid crystalline assemblies of collagen in bone and in vitro systems, J. Biomech. 36 (2003) 1571e1579. [96] F. Gobeaux, E. Belamie, G. Mosser, P. Davidson, P. Panine, M.M. Giraud-Guille, Cooperative ordering of collagen triple helices in the dense state, Langmuir 23 (2007) 6411e6417. [97] P.D.S. Peixoto, A. Deniset-Besseau, M. Schmutz, A. Anglo, Achievement of cornea-like organizations in dense collagen I solutions: clues to the physicochemistry of cornea morphogenesis, Soft Matter 9 (2013) 11241e11248.
e, C. Catania, P. Legriel, G. Pehau[98] Y. Wang, T. Azaïs, M. Robin, A. Valle Arnaudet, F. Babonneau, M.M. Giraud-Guille, N. Nassif, The predominant role of collagen in the nucleation, growth, structure and orientation of bone apatite, Nat. Mater. 11 (2012) 724e733. [99] M.M. Giraud-Guille, G. Mosser, E. Belamie, Liquid crystallinity in collagen systems in vitro and in vivo, Curr. Opin. Colloid Interface Sci. 13 (2008) 303e313. [100] L. Besseau, M.M. Giraud-Guille, Stabilization of fluid cholesteric phases of collagen to ordered gelated matrices, J. Mol. Biol. 251 (1995) 197e202. [101] D.P. Knight, D. Feng, M. Stewart, Structure and function of the salachian egg case, Biol. Rev. 71 (1996) 81e111. [102] D.P. Knight, L. Nash, X.W. Hu, J. Haffegee, M.W. Ho, In vitro formation by reverse dialysis of collagen gels containing highly oriented arrays of fibrils, J. Biomed. Mater. Res. 41 (1998) 185e191. [103] L. Besseaua, B. Coulombb, C. Lebreton-Decosterb, M. Giraud-Guillec, Production of ordered collagen matrices for three-dimensional cell culture, Biomaterials 23 (2002) 27e36. [104] J.B.L. Bard, D.J.S. Hulmes, I.F. Purdom, A.S. Ross, Chick corneal development in vitro: diverse effects of pH on collagen assembly, J. Cell Sci. 105 (1993) 1045e1055. [105] N. Nassif, F. Gobeaux, J. Seto, E. Belamie, P. Davidson, P. Panine, G. Mosser, P. Fratzl, M.M. Giraud Guille, Self-assembled collagenapatite matrix with bone-like hierarchy, Chem. Mater. 22 (2010) 3307e3309. [106] R. Martin, J. Farjanel, D. Eichenberger, A. Colige, E. Kessler, D.J.S. Hulmes, M. Giraud-Guille, Liquid crystalline ordering of procollagen as a determinant of three-dimensional extracellular matrix architecture, J. Mol. Biol. 301 (2000) 11e17. [107] W.D. Comper, A. Veis, The mechanism of nucleation for in vitro collagen fibril formation, Biopolymers 16 (1977) 2113e2131. [108] Y. Bouligand, J.P. Denefle, J.P. Lechaire, M. Maillard, Twisted architectures in cell free assembled collagen gels: study of collagen substrates used for cultures, Biol. Cell. 54 (1985) 143e162. [109] Y. Wang, J. Silvent, M. Robin, F. Babonneau, A. Meddahi-Pelle, N. Nassif, M.M. Giraud Guille, Controlled collagen assembly to build dense tissue-like materials for tissue engineering, Soft Matter 7 (2011) 9659e9664. [110] M.M. Giraud-Guille, Cholesteric twist of collagen in vivo and in vitro, Mol. Cryst. Liq. Cryst. Inc. Nonlinear Opt. 153 (1987) 15e30. [111] N. Kuznetsova, S.L. Chi, S. Leikin, Sugars and polyols inhibit fibrillogenesis of type I collagen by disrupting hydrogen-bonded water bridges between the helices, Biochemistry 37 (1998) 11888e11895. [112] S. Leikin, D.C. Rau, V.A. Parsegian, Temperature-favoured assembly of collagen is driven by hydrophilic not hydrophobic interactions, Nat. Struct. Biol. 2 (1995) 205e210. [113] D. MacKenna, S.R. Summerour, F.J. Villarreal, Role of mechanical factors in modulating cardiac fibroblast function and extracellular matrix synthesis, Cardiovasc. Res. 46 (2000) 257e263. [114] R.T. Prajapati, M. Eastwood, R.A. Brown, Duration and orientation of mechanical loads determine fibroblast cyto-mechanical activation: monitored by protease release, Wound Repair Regen. 8 (2000) 238e246. [115] R.T. Prajapati, B. Chavally-Mis, D. Herbage, M. Eastwood, R.A. Brown, Mechanical loading regulates protease production by fibroblasts in threedimensional collagen substrates, Wound Repair Regen. 8 (2000) 226e237. [116] F.H. Silver, J.W. Freeman, G.P. Seehra, Collagen self-assembly and the development of tendon mechanical properties, J. Biomech. 36 (2003) 1529e1553. [117] R.K. Sawhney, J. Howard, Slow local movements of collagen fibers by fibroblasts drive the rapid global self-organization of collagen gels, J. Cell Biol. 157 (2002) 1083e1091. [118] C. Guidry, F. Grinnell, Contraction of hydrated collagen gels by fibroblasts: evidence for two mechanisms by which collagen fibrils are stabilized, Coll. Relat. Res. 6 (1987) 515e529. [119] D.J. Prockop, D.J.S. Hulmes, in: P.D. Yurchenco, D.E. Birk, R.P. Mecham (Eds.), Extracellular Matrix Assembly and Structure, 47, Acad. Press, San Diego, 1994.

84

S. Ghazanfari et al. / Biomaterials 97 (2016) 74e84

[120] D.J. Prockop, A.L. Sieron, S.W. Li, Procollagen N-proteinase and procollagen Cproteinase. Two unusual metalloproteinases that are essential for procollagen processing probably have important roles in development and cell signaling, Matrix Biol. 16 (1998) 399e408. [121] T.G. Bromage, R.S. Lacruz, R. Hogg, H.M. Goldman, S.C. McFarlin, J. Warshaw, W. Dirks, A. Perez-Ochoa, I. Smolyar, D.H. Enlow, A. Boyde, Lamellar bone is an incremental tissue reconciling enamel rhythms, body size, and organismal life history, Calcif. Tissue Int. 84 (2009) 388e404. [122] G. Marotti, M. Ferretti, C. Palumbo, The problem of bone lamellation: an attempt to explain different proposed models, J. Morphol. 274 (2013) 543e550. [123] X.F. Walboomers, W. Monaghan, A.S. Curtis, J.A. Jansen, Attachment of fibroblasts on smooth and microgrooved polystyrene, J. Biomed. Mater. Res. 46 (1999) 212e220. [124] C. Oakley, N.A. Jaeger, D.M. Brunette, Sensitivity of fibroblasts and their cytoskeletons to substratum topographies: topographic guidance and topographic compensation by micromachined grooves of different dimensions, Exp. Cell Res. 234 (1997) 413e424. [125] J.H. Wang, E.S. Grood, The strain magnitude and contact guidance determine orientation response of fibroblasts to cyclic substrate strains, Connect. Tissue Res. 41 (2000) 29e36. [126] W.A. Loesberg, X.F. Walboomers, J.J. van Loon, J.A. Jansen, The effect of combined cyclic mechanical stretching and microgrooved surface topography on the behavior of fibroblasts, J. Biomed. Mater. Res. A 75 (2005) 723e732. €ntjens, F.P. Baaijens, C.V. Bouten, [127] N. de Jonge, J. Foolen, M.C. Brugmans, S.H. So

[128]

[129]

[130]

[131]

[132]

[133]

[134]

Degree of scaffold degradation influences collagen (re)orientation in engineered tissues, Tissue Eng. Part A 20 (2014) 1747e1757. S. Ghazanfari, A. Driessen-Mol, B. Sanders, P.E. Dijkman, S.P. Hoerstrup, F.P.T. Baaijens, C.V.C. Bouten, In-vivo collagen remodeling and maturation in the vascular wall of decellularized stented tissue engineered heart valves, Tissue Eng. Part A. 21 (2015) 2206e2215. S. Ghazanfari, A. Driessen-Mol, S.P. Hoerstrup, F.P.T. Baaijens, C.V.C. Bouten, Collagen matrix remodeling in stented pulmonary arteries after trans-apical heart valve replacement, Cells Tissues Organs 201 (2016) 159e169. S. Singh, S.B. Bandini, P.E. Donnelly, J. Schwartz, J.E. Schwarzbauer, A cellassembled, spatially aligned extracellular matrix to promote directed tissue development, J. Mater. Chem. B 2 (2014) 1449e1453. € ller, W.W. Ahmed, T. Wolfram, A.M. Goldyn, K. Bruellhoff, B.A. Rioja, M. Mo J.P. Spatz, T.A. Saif, J. Groll, R. Kemkemer, Myoblast morphology and organization on biochemically micro-patterned hydrogel coatings under cyclic mechanical strain, Biomaterials 31 (2010) 250e258. J. Wu, Y. Du, S.C. Watkins, J.L. Funderburgh, W.R. Wagner, The engineering of organized human corneal tissue through the spatial guidance of corneal stromal stem cells, Biomaterials 33 (2012) 1343e1352. D.D. Klumpers, D.J. Mooney, T.H. Smit, From skeletal development to tissue engineering: lessons from the micromass assay, Tissue Eng. Part B. Rev. 21 (2015) 427e437. C.E. Murry, G. Keller, Differentiation of embryonic stem cells to clinically relevant populations: lessons from embryonic development, Cell 132 (2008) 661e680.