20.3.1 Autologous tissues

Autogenous tissues are mainly used as patches or in tubular forms, harvested from a wide range of body sites. Tracheal small defects can be successfully treated with graft repairs from fascia lata [21], auricular and costal cartilage [22], bronchial patch [23], pericardium [24], and aortic grafts [25]. Autologous grafts have also been used in combination with artificial material support, such as tantalum wires [26], polypropylene meshes (Marlex™) [27], and silicone, or polyethylene and polyurethane stents [28, 29]. However, due to the high frequency of tumour recurrence, this approach has been mainly abandoned for lateral tracheal incision [21]. Although tissue grafts can successfully repair small tracheal defects, large defects require vascularization to maintain tissue viability and reduce the risk of necrosis. Another approach is to harvest tissue flap from the patient, preserving or reanastomosing the blood supply. Therefore, this method provides independent vascularization and favourable wound healing conditions in comparison to graft materials. Due to the significant segment length, cartilage grafts, polymer rings, or meshes are required to increase the rigidity and stability of the flap tissue [30]. Approaches for vascularized autologous tracheal repair include skin flaps [31], pedicled periosteum [32], intercostal muscle patch [33], and broncus [34]. Although vascularized autologous flaps have generally been successful, the complexity and the multi-staged approach constrain the surgical technique, thus limiting the common use of this procedure [30].

20.3.2 Artificial airway constructs

Synthetic materials have been extensively used to generate artificial constructs as a means of temporary tracheal support and tracheal prosthesis for tissue replacement. The common drawbacks of artificial materials are their lack of integration with the host tissues. Therefore, problems of migration, dislodgement, infection, and obstruction together with lack of epithelialization generally arise [21]. To date, there are mainly two types of airway stents commercially available as tracheal replacements: silicone tubes and expandable metallic stents. Silicone tubes are flexible and are proposed as long-term placements and atraumatic insertions. In particular, the Tracheaobronzane® Dumon™ Silicone Stent, which has shown excellent clinical results, is composed of external studs to prevent dislodgment, and extremities that are designed to maximize airflow, thus reducing formation of granulation tissue [35, 36]. Another product currently available is the Dynamic™ (Y) stent, characterized by a Y-geometry (tracheal and bronchial limbs), C-shaped stainless steel struts to mimic the cartilaginous rings, and optimized internal surface to limit obstructions [37]. Although silicone-based stents are easy to customize and remove, they exhibit strong limitations arising from difficulty in placement, tendency to dislodge, and frequently occurring obstructions due to lack of epithelialization and low inner to outer diameter ratio [38].

Therefore, metallic stents have been introduced to ameliorate correct delivery and placement with less invasive surgical procedures, in order to reduce obstructions, and improve their stability [38]. Ultraflex™ Esophageal NG Stent System is a self-expandable tubular mesh currently in use, and is made of flexible single-strand Nitinol wire covered with polyurethane, to reduce tissue ingrowth [39]. In order to further prevent tumour and tissue growth within the wire mesh stents, Polyflex® Esophageal Stent have been developed based on a silicone-covered polyester wire mesh with an internal silicone coating [40]. Uncovered metallic stents can potentially improve epithelialization and integration with the surrounding tissue, but are difficult to remove. In comparison, coated stents are able to decrease tumour and granulation tissue ingrowth and to improve removability [30].

The full replacement of tracheal tracts was experimentally performed in large animal models with a wide range of materials in tubular form, such as stainless steel [41], Vitallium [42], polyethylene [43–45], silicone [46–49] polytetrafluoroethylene [50, 51], and their combinations [52–55]. Solid prostheses were also used in clinical settings, with silicone tubes being the most exploited. The Neville artificial trachea was composed of a silicone tube with two suturable fabric cuffs at the extremities to prevent granulation tissue and enhance a stable fixation [56]. However, silicone prostheses still displayed severe stenosis and infection at the interface of the prosthesis and tracheal epithelium [21]. Upon the failure of solid prosthesis due to obstructive granulation tissue, porous materials were designed to stimulate tissue ingrowth and eventually migration of adjacent tracheal epithelium. Due to insufficient structural support, experimental mesh prostheses have been frequently reinforced with wire, plastic rings, or coils. Moreover, they have been externally wrapped with other tissues (e.g. pericardium, fascia lata or dura mater) or biopolymers (e.g. fibrin or reconstituted collagen) to prevent air leakage. The most exploited materials for mesh production have been polypropylene [27, 57, 58], polyethylene terephthalate [51, 59], polyurethane [60],and polytetrafluoroethylene [61] in multiple combinations mainly based on collagen grafting [62] and polypropylene reinforcement [63]. Some of the porous meshes for tracheal replacement were also produced for clinical applications; in particular Marlex™ based prosthesis [27, 64]. Despite the porous structure, epithelium did not cover the entire luminal surface of the constructs. Therefore, the continued proliferation of scar tissue could not be controlled, frequently leading to obstruction and stenosis. In addition, the lack of coverage in large sectors of the meshes frequently results in bacterial colonization, and the subsequent infection of the prostheses [21].

20.3.3 Allotransplanted airway constructs

Biological scaffolds derived from allogenic and xenogenic tissues are commonly used in a variety of reconstructive surgeries and also adapted for tissue and organ replacement in regenerative medicine strategies. During the past decade, tissue decellularization techniques have been optimized to reduce deleterious in vivo effects of residual cellular material in order to circumvent immunosuppression post-treatment. Furthermore, new methodologies have attempted to minimize deterioration of extracellular matrix (ECM) native composition, structure, and mechanical performance imparted by the decellularization processes. The hypothesis of this tissue replacement approach is that ECM-derived scaffolds produced by decellularization can maintain or promote site-appropriate cell phenotypes during the process of cell repopulation, via exposure of the ligands and bioactive molecules necessary to lead cell populations to organize in functional structures [65]. The matrix should provide an inductive 3D biological template to stimulate the regeneration of a functional tissue through in vivo cell recruitment or exogenous provision. Decellularization techniques are generally a combination of physical, ionic, chemical, and enzymatic methodologies, which are more efficiently imparted by perfusion systems, in particular for vascularized tissues or organs [66].

In 2008, Macchiarini et al. repopulated a decellularised human donor windpipe in a dedicated bioreactor with autologous epithelial cells (ECs) and mesenchymal stem cell (MSC)-derived chondrocytes (Fig. 20.1). The bioreactor was designed to seed and culture both ECs and MSCs, spatially segregated in the luminal and external surfaces of the construct, respectively. After 4 days of dynamic culture, the graft was used to replace a compromised left main bronchus of a 30-year-old woman. The patient was immediately provided with a normal functioning airway without immunosuppressive drugs [67]. Despite the clinical success of this windpipe transplantation, several questions have been raised. The long-term effects of decellularization treatments on the morphological changes of the donor tracheal tissues have not been investigated. It is not indicated whether the tissue would eventually be replaced by newly formed cartilaginous tissues or it would progressively degrade and loses its original supporting role [68]. Moreover, the lack of an intrinsic blood supply can result in the unpredictable healing of the tissue [69]. The optimization of graft preservation might ultimately broaden the clinical application of tissue engineered products, in relation to the time required to prepare the antigen-free tissue, culture the committed cell population, and the complete maturation of the tissue [70]. Further in vitro studies have evaluated the potential intrinsic changes and degradation of decellularized human tissue scaffolds. During one-year storage in physiological solution, human decellularized tracheas displayed loss of ECM architecture and changes in mechanical properties. Such degradation phenomena can be further accelerated in vivo, impacting the clinical relevance of long-term tissue transplantation [71]. Remlinger et al. observed biochemical and mechanical degradation of porcine tracheal ECM after implantation in a dog model. In order to limit the scaffold deterioration, pre-seeding of the matrix with chondrocytes may compensate the significant degradation of ECM [72].

Reconstruction of long-segment tracheal defects was also performed via tracheal allotransplantation without any decellularization treatments, in order to maintain the blood supply [4]. Delaere et al. reported a successful tracheal allotransplantation after indirect revascularization of the graft in heterotopic position. A segment of donor trachea was placed in the recipient’s forearm for revascularization, circumferentially wrapped with patient’s own tissues for 4 months. Immunosuppressive therapy was administered until transplantation took place. The donor tracheal cartilage was surrounded by the host’s blood vessels and lined with an epithelial layer originating from the patient. The tracheal allograft was then placed in the proper anatomical position with an intact blood supply.

20.3.4 Tissue-engineered airway constructs

Over the last two decades, numerous efforts have been concentrated to develop functional tissue engineered tracheal replacement, exploiting several scaffolding materials, different cell sources, and diversified culture conditions.

Scaffold-free approaches aim to develop tissue-like constructs to promote and lead to tissue development. Tani et al. used rabbit auricular chondrocytes to produce an ECM sheet, which was then wrapped around a silicone tube and cultivated for 6 weeks under dynamic condition (Fig. 20.2) [73]. The silicone support was removed after the dynamic conditioning and the chondrocyte sheet demonstrated sufficient structural integrity to maintain the shape, but the mechanical strength was found to be equivalent only to 30% of that in the native trachea.

In comparison, Gilpin et al. produced a similar chondrocyte sheet under static culture followed by dynamic culturing for 8 weeks [74]. The scaffold-free sheets were used as anterior cartilage grafting in a rabbit animal model. Although the construct exhibited no signs of inflammation, the implants showed evidence of mechanical failure. The main advantage of a purely cell-based approach is the reduction in inflammation at the implantation site. However, structural and mechanical integrities are generally compromised and the extensive time in culture are not cost- and time-effective for clinical applications [19].

Vacanti et al. in 1994 proposed a biodegradable sheet of nonwoven PGA mesh seeded with calf chondrocytes, as circumferential replacement of rat trachea [75]. No implanted animals survived, probably due to the excessive rigidity of the construct and lack of vascularization. Further studies attempted to line the cartilaginous construct with respiratory epithelium, and revealed the complexity to guarantee a continuous epithelium without infection [76]. In order to improve the mechanical functionality of the scaffold, Kojima et al. developed a modular construct composed by ovine chondrocyte-seeded nonwoven PGA mesh, placed in the grooves of a helical Silastic® template and then covered with an ovine fibroblast-seeded mesh (Fig. 20.3) [77].

The implants were inserted in the neck of the sheep to allow maturation for 8 weeks and subsequently implanted to repair a circumferential defect. However, animal survival was limited to 7 days due to significant stenosis, probably as a consequence of inflammatory reaction induced by PGA, leading to high cellular content and low proteoglycan presence compared to normal tracheas. The same group further developed the implants by wrapping the PGA matrices around a silicone tube and seeded with ovine nasal chondrocytes, where harvesting requires a less invasive procedure and thus more suitable for clinical applications [78]. After subcutaneous implantation, no inflammatory reactions were reported, resulting in a matrix composition similar to native trachea but with significantly lower mechanical properties. Furthermore, Kojima et al. assessed the feasibility of co-culture of ECs and chondrocytes from the same tissue source and revealed, after 4 weeks, that only 60% of the internal surface was covered with epithelium. Recently, Jungebluth et al. performed a tracheobronchial transplantation with a stem-cell-seeded bioartifical nanocomposite made of polyhedral oligomericsilsesquioxane covalently bonded to poly-[carbonate-urea]-urethane [79]. The construct was first seeded under dynamic culturing for 36 hours and then implanted. At early time points, biopsy samples showed the presence of necrotic tissue associated with fungal contamination, while 1 month after surgery biopsies revealed large inflammation areas with initial signs of epithelialization. The ability to line tissue construct with a continuous epithelium under in vitro conditions is one of the main challenges for airway tissue engineering. This limitation is mainly due to the synthetic (non-biological) origin of biodegradable polymers that discourage EC growth and organization. In contrast, collagen modified constructs have been shown to promote ciliated epithelium lining under both in vitro and in vivo conditions [80, 81]. Natural trachea, on the other hand, has the capability to restore the epithelial mucosa when a small portion of tracheal tissue is removed [82].

Several natural polymer-based materials have been applied as tracheal tissue engineering: specifically hyaluronan and silk fibroin (SF)-derived materials. Henderson et al. developed hyaluronan-based scaffolds, which have been shown to support chondrocyte growth under dynamic culture conditions [83]. However, in vivo animal implantation demonstrated a nonspecific foreign body response [84]. In addition, silk fibroin has been used as a coating in rabbit trachea defect reconstruction. The results demonstrated good biocompatibility of the material, promoting fibroblast invasion and capillary vessel formation [85]. In order to take advantage of the biocompatibility of natural polymers, several groups have proposed different methods to conjugate natural and synthetic polymers. Komura et al. developed a three-layered construct consisting of a collagen sheet, a PGA mesh and a poly L-lactide/ε-caprolactone copolymer mesh, seeded with rabbit chondrocytes [86]. Three months after implantation in the cervical trachea, the structural integrity of the scaffold was maintained and epithelial regeneration occurred in the lumen. Similar work has been developed by Omori et al. based on a polyethylene Marlex™ mesh tube covered with collagen sponge [87]. The airway reconstruction of a small defect was performed on an elderly patient, inducing good epithelialization and patency for two years. Furthermore, Lin et al. proposed a composite construct composed of a poly(ε-caprolactone) tubular frame embedded with type II collagen sponge [88]. Subcutaneous implantation of the scaffold revealed a significant increase in the mechanical properties after 8 weeks, exceeding those of native tracheal tissue, and a matrix composition similar to a cartilaginous tissue.

The same group developed a bioreactor system to grow the tissue engineered trachea seeded with chondrocytes after a maturation period under static culture. An increase in cell proliferation, glycosaminoglycan and collagen content was observed under dynamic culture. After 2 months’ implantation into rabbit tracheal defects, poor epithelialization occurred [89]. However, this work revealed the central role of a bioreactor system to stimulate the maturation and to augment the functional and mechanical quality of tissue engineered constructs. Similarly, Kojima et al. developed a construct made of stromal cells seeded in PGA mesh inserted into helical grooves of a silicone tube, where the external surface was coated with transforming growth factor β2 to stimulate chondrogenic differentiation [90]. After 6 weeks of subcutaneous implantation, cartilage formation was observed, although the mechanical properties were not evaluated. In general, the rationale for the combination of different materials is to compensate for the limitations of the single constituent. However, the inherent complexity in producing composite, multilayered, and hybrid materials has so far limited the translation of these scaffolds into clinical settings [19].

20.3.5 Airway tissue models

Together with tissue engineered constructs for airway replacement, research has also driven the focus towards tissue models. The main objective of tissue models is to mimic the microstructure, geometry, and functionalities of the native tissue in vitro. In addition, 3D tissue models provide new possibilities for the study of complex physiological and pathophysiological processes in vitro [91]. Along this route, tissue engineered constructs might reduce the need for organ or tissue transplantations by supporting the development of therapies to prevent or cure diseases at the origin [92]. In comparison, in vivo animal models can offer the entire complexity of the biological milieu, but are lacking in the independent control of the system variables and with inherent differences between animal and human responses. Therefore, distinct effects of the biological environment can hardly be segregated in experimental in vivo conditions. In contrast, under in vitro conditions each control variable can be systematically isolated, but with a strong simplification of the experimental settings, where the biochemical and mechanical signals present in the real biological system are strongly limited.

Under this perspective, Agarwal et al. developed an in vitro wound-healing model comprising type I collagen gel in a planar geometry for the co-culturing of human fibroblasts, and bronchial ECs [93]. The study investigated the expression of ECM proteins during would-healing tissue contraction in response to the presence of epithelium and different wound types, characteristic of asthma. Choe et al. also proposed a protocol based on highly hydrated type I collagen gel to setup, maintain, and characterize a tissue engineered human bronchial mucosa model obtained over 2 weeks and to be used for basic physiology and pathophysiology studies (Fig. 20.4) [94]. Because of extensive cell-mediated contraction, the bronchial mucosa model was also implemented to study airway remodelling and cell–substrate interactions in planar geometry. Moreover, an in vitro airway tissue model is commercially available with the same geometry. EpiAirway™ originates from normal (i.e. non-immortalized) human-derived tracheal/bronchial ECs, cultured to generate a 3D, pseudo-stratified model [95].

This 3D tissue model has been used primarily to study respiratory infections, and drug delivery. In contrast to these previous approaches, Miller et al. developed a tissue engineered model of the bronchioles, incorporating the native cylindrical geometry of the tissue and the presence of both fibroblasts and smooth muscle cells (SMCs) within a highly hydrated type I collagen gel [96]. Due to the original tissue geometry, the experimental model was mechanically stimulated under physiological conditions in order to approximate the behaviour of the native tissue (e.g. airway remodelling). Generally, airway tissue models do not resemble the native environment in terms of both tissue-like architecture and the ability to provide physiological mechanical cues to the resident cells. In particular, previous studies did not aim to reproduce independently and in combination the sequential occurrence of the in vivo mechanical stimuli (i.e. shear stress and circumferential strain). Recently, a method to produce tubular dense collagen-based constructs was proposed to potentially meet the demands of the above challenges. In less than one hour, tubular dense collagen-based constructs (TDCCs) with native tissue equivalent fibrillar protein densities and mechanical properties were produced by the circumferential wrapping of plastically compressed dense collagen gel-based sheets around a cylindrical support [97] (Fig. 20.5).

This method has potential clinical applications by dramatically reducing the time allocated for scaffold production, eliminating the dependence on donor availability and cell-based matrix synthesis as well as potentially reducing the risk of detrimental in vivo effects that are associated with residual biological materials. Furthermore, a TDCC was used to investigate air smooth muscle cell (ASMC) responses under physiological pulsatile flow. The role of shear stress alone, and in combination with circumferential strain on the proliferation, alignment, and phenotype of ASMCs seeded in 3D, was underscored together with the effects of ASMC-mediated remodelling on matrix morphological and mechanical properties. By providing ASMCs with a physiologically equivalent 3D environment, and through in vitro mechanical stimulation, ASMCs exhibited their native orientation, maintained their contractile phenotype and enhanced the mechanical properties of the TDCC through matrix remodelling.

20.4.1 Type I collagen structure

Collagens share the characteristic three α-helical chain structure, composed of repeated sequence of the triplet Gly-X-Y, where X and Y are frequently proline and hydroxyproline, respectively [101]. The three α-helical chains are then organized and stabilized in a triple helical quaternary structure, which is the trademark of the collagen superfamily [102]. The lack of side chain in glycine allows the compaction of the collagen triple helix [103]. On the other hand, the side chains of X and Y residues are exposed providing the capacity for lateral interactions with ECM macromolecules, and resulting in the formation of various supramolecular level of organizations [104]. Only collagen types I, II, III, V, and XI can be organized in highly ordered supramolecular aggregates (i.e. fibrils) [105] (Fig. 20.6).

The hierarchical organization at several discrete levels of the collagen protein results in the distinct mechanical and structural properties of the fibres [107]. The existence of a monomeric building unit for the collagen fibre was proposed by Gross in 1956 and defined as tropocollagen [108]. Collagen fibrils have a distinctive suprastructure, identified with a periodicity of 67 nm (D-period), which is represented by the quarter-staggered arrangement of individual collagen monomers in fibrillar-array [104]. In particular, type I collagen is the major organic component in tendons, ligaments, bone, skin, cornea, and vessels [109], and in the form of fibres it is characterized by substructural organization as fibrils and fibrillar bundles, which have been previously correlated with the ultimate strength of biological tissues [106].

20.4.2 Type I collagen as a biomaterial

Type I collagen has been extensively used as a biomaterial, due to the excellent biological properties and processability in different forms [110]. Type I collagen is well established in the clinical use of various tissue replacements and devices, such as wound dressing, tissue augmentation, ophthalmic barriers, bioprosthetic heart valves, arteriovenus shunts, suture threads, haemostatic membranes, dura mater substitutes, nerve conduits, and drug delivery [111]. The use of type I collagen, in particular in tissue engineering provides numerous advantages [98, 107, 110, 112], such as its:

Type I collagen-based biomaterials are mainly produced from decellularization of collagen-rich tissues and reconstitution of solubilized collagen [113]. Decellularization processes generally require purification and removal of cellular components to reduce an immunogenic potential. Subsequent chemical fixation can be used to strengthen and increase the stability of the collagen matrix [114]. Modern extraction methods are based on three main principles of solubilization: either in acidic [115, 116], neutral salt [117], or proteolytic solutions [118]. The latter strongly reduces the self-assembling capacity of tropocollagen molecules into fibril aggregation, due to the cleavage of terminal telopeptide regions induced by the proteolytic treatment [119]. To reduce this effect, endogenous proteases can be inhibited during the acidic solubilization [120]. In general, acidic extraction with an additional pepsin solubilization step is the most effective technique. However, some telopeptides are cleaved or partially denatured [118, 121, 122]. Multiple protocols are widely used to solubilize collagen, and various collagen solutions are readily available at a large variety of concentrations from many different sources. Procollagen solutions are used to produce reconstituted collagen gels by restoring the pH, temperature, and ionic concentration to physiological levels. Procollagen polymerization in physiological conditions results in a protein concentration of less than 0.85 wt%, which generates highly hydrated collagen gels with low values of collagen fibrillar density (CFD) and a limited range of mechanical properties (i.e. strength of 4–6 kPa and elastic modulus of 20–30 kPa) [123, 124]. Collagen gels under tension generate nonlinear stress–strain behaviour similar to those of native tissues [124, 125]. At low strains, uniaxial tensile curves present a nonlinear phase, called the ‘toe’ region, which is followed by a linear phase, where the elastic modulus is calculated, and terminating with an exponential failure region [126]. In the unloaded configuration, the collagen gel network presents an isotropic structure. Under low stress, a rearrangement phase occurs where collagen fibrils orientate and align towards the direction of the stress. Once the majority of collagen fibrils are recruited in the direction of the stress, an alignment phase follows, where higher stresses are necessary to axially deform the fibrils [127].

The method of collagen gel preparation easily allows the direct injection of cells prior to gelling, thus enhancing seeding efficiency and the homogeneous distribution of cells within the scaffold [128]. Cellularized collagen hydrogels were initially described in 1962 [129], and have since been identified as an excellent 3D substrate for the adhesion, proliferation, and differentiation of numerous cell types [107]. The use of 3D substrate to study cellular responses, in comparison to 2D cultures, is theoretically founded on the systems-based scientific approach proposed by Weiss in 1959 [130]. Through this, the emphasis is placed on the cells in relationship with their external physical environment, and not limited to the cells as single entities. Moreover, since the texture and mechanical properties of 3D collagen matrices resemble the fibrous connective tissue environment, they have become well-established model systems to study cell behaviour in an in vitro tissue-like milieu [131]. Initially, fibroblasts were studied encased within 3D hydrogels under both restrained and free conditions to underscore the mechanism of cell-mediated gel contraction [132]. Afterwards, Bell et al. adopted fibroblast-seeded 3D hydrogels as tissue-like structures to model wound contraction under in vitro conditions [133]. The same collagen matrices seeded with fibroblasts were then proposed as skin-equivalents in 1981 [134], and these have become the concept at the base of the first allogeneic cell-based product (Apligraf®, Fig. 20.7), approved by the US FDA as skin graft and in 1998 was commercialized by Organogenesis [135]. Cellular collagen hydrogels have proven to be effective for the treatment of skin ulcers, burns and oral mucosal defects [136, 137]. Acellular collagen hydrogels are also currently in clinical use for dura mater replacement [138].

Although collagen exhibits excellent biological properties [140], the wider application of its hydrogels in tissue engineering is limited due to low mechanical strength [141] (Fig. 20.8a), and structural instability arising from cell-mediated contraction attributable to the mechanical tension imposed by the constituent cells [142] (Fig. 20.8b). Several approaches have been taken to overcome the mechanical and structural drawbacks of collagen-based hydrogels, including cell-induced gel densification resulting from collagen remodelling and subsequent increase in CFD, a process that can be unpredictable and time consuming [133, 143]. Collagen remodelling comprises a complex set of events where, in analogy to native ECM, is continuously digested by synchronous proteolytic degradation (activated by matrix metallo proteinases (MMPs)) and reassembled by the cells (through endogenous collagen production), eventually resulting in morphological and structural changes [144].

A variety of cross-linking methods, using either chemical [146–148], photochemical [149–151], or enzymatic [152, 153] have also been investigated. However each method imparts some degree of cytotoxicity, and can be technically limited by a high cost–benefit ratio for large-scale implementation [154].

Brown et al. developed a processing technique based on plastic compression of collagen hydrogels in order to generate scaffolds with increased CFD and subsequent improvement in the mechanical properties (i.e. strength and elastic modulus) [155, 156] (Fig. 20.9).

Briefly, collagen hydrogels are subjected to a compressive stress of 1 kPa (1A) for 5 minutes, resulting in the expulsion of more than 95% of the water content and a 40-fold increase in CFD from 0.3 to 12%. The application of plastic compression on collagen hydrogels produces dense collagen (DC) constructs (Fig. 20.9b) with controlled protein content and mesoscale properties, significantly enhancing the mechanical and structural performances of collagenous hydrogels (i.e. strength of 0.6 MPa and elastic modulus of 1.5 MPa) [155]. DC scaffolds have been shown to improve cell metabolic activity [157] and stimulate osteoblastic differentiation [158]. DC gels also demonstrated their potential as osteoid model for bone tissue engineering [159, 160], also in combination with other biopolymers (e.g. chitosan) [161], silk fibroin derived anionic polypeptides [162], and bioactive glasses (i.e. Bioglass® 45S5) [163, 164].

20.5.1 SMCs

SMCs are a functionally key component of a large variety of organs and tissues, part of the cardiovascular, respiratory, gastrointestinal, urinary, and reproductive apparatuses [168]. Generally, SMCs share a common structure and the active role in the regulation of the tissue or organ functions, while the inducing stimuli differ substantially depending on the origin of the tissue (e.g. vascular and respiratory tone). In fact, SMCs display a phenotypic plasticity to accommodate diverse functions in multiple tissues. They exhibit a multifunctional capacity for contraction, migration, proliferation, synthesis of ECM, and secretion of growth factors [169]. These features allow SMCs to regulate the lumen diameter of hollow organs, through a reversible transient contraction or a chronic reduction/enlargement of the lumen, due to structural remodelling (e.g. vascular aneurism, airway narrowing). Specifically for the airway wall, airway smooth muscle tone acts in concert with the cartilaginous rings and cooperate to strengthen the airways structure, preventing collapse during respiration [18]. ASMCs are frequently involved in pathological mechanisms related to airway diseases [170]; in particular they participate in the inflammatory process and remodelling involved in asthma, due to the ability to modulate their in vivo phenotype in response to variations in the external conditions [171].

In contrast to skeletal muscle cells, SMCs can simultaneously proliferate and express a set of contractile-lineage specific proteins [172]. In response to specific stimuli, SMCs can modulate their phenotype by suppressing the expression of certain contractile protein genes, converting from a differentiated contractile phenotype to a dedifferentiated synthetic phenotype [169]. The phenotypic modulation has been implicated in mechanisms underlying a number of pathological conditions, including atherosclerosis and restenosis post-angioplasty, and airway remodelling in asthma [173, 174].

A similar shift in SMC phenotype is also observed post-extraction from their native site. The subsequent expansion, and culture using traditional culture techniques results in a significant loss of cell functionality [175]. In particular, phenotypic plasticity is typically displayed in proliferative media (i.e. 5–10% serum content), which alters their differentiation state towards a synthetic phenotype. Moreover, low cell density has been associated with a down-regulation of the contractile protein gene expression due to reduced cell–cell contact. Specifically, the influence of serum and the reduction in cell–cell interactions have been demonstrated to be independent and cumulative [176–179].

The substrate plays a predominant role in the SMC phenotype regulation, through ECM components that modulate cell integrin-mediated signalling [180]. SMCs removed from their native site and cultured under conventional static conditions (i.e. tissue culture plastic) did not exhibit alignment along a preferential direction [181], together with a shift of phenotype in favour of a proliferative state [182]. In particular, 2D substrate made of highly hydrated type I collagen gel enhances the synthetic state of SMCs [183, 184]. The 3D culture of SMCs within highly hydrated type I collagen gel has also demonstrated the further reduction of the expression of contractile proteins in comparison with monolayers [180, 185].

20.5.2 MSCs

At the stage of human development, progenitor cells are present in a large variety of tissues, while in adults they are mostly prevalent in the bone marrow [186]. MSCs can be easily isolated from the bone marrow, cultured, and expanded under in vitro conditions. Moreover, they can generate a spectrum of specialized mesenchymal tissues, such as bone, cartilage, muscle, marrow stroma, tendon, ligament, fat, and a variety of other connective tissues (Fig. 20.11) [187]. In addition, MSCs produce a variety of bioactive factors, which can serve to structure regenerative microenvironments in case of tissue injury [186]. MSCs possess unique features that make them attractive tools for tissue engineering and regenerative medicine. For example, stem cells can replicate in culture while retaining the ability to differentiate into specific lineages, which is in contrast to primary culture where the availability and phenotypic plasticity of cells are limiting factors [188]. In addition, adult stem cells, including MSCs, display decreased immunogenic potential, which may facilitate allogenic transplantation in building damaged or neo-mesenchymal tissues [189]. Under this perspective, the ad-hoc development of 3D culture substrates might allow greater control over MSC fate and ultimately tissue architecture and function [190]. The continuous remodelling of ECM, which comprises degradation and synthesis, alters the elasticity and biochemical cues of the matrix that affect cell activity and differentiation [191]. Recent data has demonstrated that the culture substrate is not to be considered as an inert, solid-state environment. On the contrary, MSC differentiation state and responses reflect culture conditions through multiple physical mechanisms, such as the geometry at the micro- and nanoscale, elasticity, and external mechanical stimuli transferred from the substrate to the cells [192]. In particular, soft matrices (elastic modulus ~ 1 kPa) have been shown to stimulate MSCs towards a neuronal-like phenotype. Increasing in substrate stiffness determined a shift to a myogenic cell fate, whereas the stiffest matrices (Elastic modulus ~ 30–100 kPa) lead to osteoblastic differentiation, due to the similarities with collagenous bone [193, 194]. MSC chondrogenic lineage stimulation has been successfully achieved in vitro by supplementing culture medium with bioactive factors, including dexamethasone and transforming growth factor beta1 (TGF-β1) [195]. Specifically, TGF-β1 has been shown to induce chondrogenesis in MSC preparation as early as day 7, even without the presence of dexamethasone [196].

20.6.1 Dynamic stimulation of SMCs

SMCs in their native sites are constantly exposed to cyclic mechanical stresses as a result of airflow during tidal breathing or blood flow in the cardiac cycle. Such mechanical forces have been demonstrated to play a fundamental role in regulating SMC alignment, phenotype and contractile function [18, 180]. Cyclic mechanical stimuli are commonly applied to cultures of ASMCs adherent to 2D substrates, which is generally a silicone distensible membrane stretched at a controlled rate [197–200]. The need for more physiologically relevant substrates has led to 3D culture systems, in order to resemble the cell organization in their native environment. In particular, SMC orientation can be modulated by the type of mechanical stimuli applied. The stimulation of SMC within tubular structure generally relies on bioreactor systems able to generate cyclic circumferential strains and flow-induced shear stresses [201].

Cells directly exposed to physiological shear stress displayed an orientation parallel to the flux direction, which is an organization consistent with cell arrangement in vivo (e.g. ephitelium) [202, 203]. In addition, SMC orientation is dependent on the magnitude of cyclic stretch applied. SMCs exhibited an alignment parallel to the stress direction at low values of strain (2%) [203, 204], while at higher strains (> 5–10%) they arranged perpendicular to the stress in order to limit cellular damage due to the excessive strain [205–208]. Moreover, SMC orientation parallel to the direction of the stretch (i.e. circumferentially) has been reported only for cells seeded within compliant 3D tubular substrates (e.g. type I collagen hybridized with biodegradable elastomers) exposed to low values of strain [203, 204, 209, 210], as reported in Plate XII (between pages 354 and 355). In comparison SMCs, cultured and stimulated with pulsatile flow or cyclic stretching on 2D substrates (e.g. silicone), exhibited an arrangement perpendicular to the strain and independent of the extent of strain [206–208, 211–213]. Table 20.1 summarizes the main parameters previously reported to stimulate SMCs extracted from various sites.

20.6.2 Dynamic stimulation of MSCs

MSCs have been widely used in combination with bioreactor-based systems in order to stimulate in vitro growth and expansion, while preventing dedifferentiation [216–218]. Dynamic culture systems in 3D have been proposed as effective alternative to traditional 2D culture models, in order to gain better control of the culture conditions as well as to mimic the dynamic 3D architecture of the native environment. A large variety of bioreactors have been adopted for the culture, growth, and maturation of MSCs within 3D constructs, including fixed-beds [219], stirred suspension [220], rotating wall vessels [221], and the most exploited category, perfusion chambers [222]. Perfusion culture was developed in order to mimic the physiological supply of nutrients within the tissue. It is generally based on the steady flow of media over or though a cell-seeded construct [223]. Perfusion systems, in comparison to stirred or suspended mechanisms, determine a more uniform distribution of cells at the seeding stage. In addition, MSC expansion and differentiation in perfused 3D scaffolds have been shown to be the most effective [220], improving specifically structure, function, and biological properties of engineered cartilage and bone constructs [224, 225]. In the case of airway tracts, a dedicated bioreactor in a double-chamber rotating configuration for the culture of MSCs in a decellularized trachea was previously developed for the development of tissue engineered hollow organs (Fig. 20.12) [67, 226]. The bioreactor was designed in order to facilitate cell-seeding procedures, to enhance oxygenation of the culture medium and mass transport, and to stimulate the resident cells with hydrodynamic stimuli, which have been shown to promote cell metabolic activity and the differentiation process towards chondrogenic lineage.

Furthermore, numerous studies have shown that mechanical stimulation plays an important role in MSC differentiation towards chondrogenic, osteogenic, and contractile lineages [211, 227–229]. In particular, MSC contractile phenotype can be stimulated through cyclic mechanical strain, as summarized in Table 20.2. The application of mechanical stimuli also induces a preferential MSC alignment on 2D or within the 3D substrate. MSCs have been generally shown to align only in the direction perpendicular to the strain applied (parallel to the flow, in case of tubular geometry) [211, 230–233], in contrast with in vivo evidences of SMCs subjected to similar stimuli, which align parallel to the strain (i.e. circumferential direction) or in an helical pattern [227]. In addition, the extent of cyclic strain would influence the alignment of MSC in order to prevent cell body damages for strain greater than 5% [230–233].