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Home > Books > Regenerative Biomaterials - Emerging Biomaterial Solutions to Aid Tissue Regeneration [Working Title]

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Sustainable Biomaterials: Turning Food Waste into Advanced Regenerative Technology

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Sandi G. Dempsey, D. Adam Young, Robert W.F. Veale and Barnaby C.H. May

Submitted: 05 March 2025 Reviewed: 12 May 2025 Published: 30 July 2025

DOI: 10.5772/intechopen.1010994

Regenerative Biomaterials - Emerging Biomaterial Solutions to Aid Tissue Regeneration IntechOpen
Regenerative Biomaterials - Emerging Biomaterial Solutions to Aid... Edited by Yvonne Reinwald

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Regenerative Biomaterials - Emerging Biomaterial Solutions to Aid Tissue Regeneration [Working Title]

Dr. Yvonne Reinwald

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Abstract

Biomaterials used in regenerative medicine have advanced significantly over the past 50 years, with ongoing developments to improve structure, biological properties, and compatibility for soft and hard tissue repair. This chapter explores the incorporation of sustainability concepts in the design of bioscaffolds for soft tissue repair, specifically focusing on ovine forestomach matrix as a case example of sustainable biomaterial development. Key factors discussed include material safety, ethical considerations, and cultural acceptance. Additionally, the chapter addresses the growing importance of sustainability in medical device design, highlighting the need for eco-friendly principles in material selection, manufacturing, and application. The integration of these principles aims to balance safety, cost-effectiveness, and global health outcomes in the development of medical technologies. The chapter reviews the evolution of biomaterial design for regenerative medicine, from first-generation bioscaffolds to more recent iterations, and evaluates the unique sustainability characteristics employed with ovine forestomach matrix as a next generation regenerative scaffold for soft tissue repair. Derived from the forestomach of pasture-raised sheep, the technology upcycles waste from the global food industry and utilizes sustainable manufacturing to lessen the carbon footprint and limit introduction of synthetic chemicals. With demonstrated clinical efficacy over the past 15 years and applications in more than 7 million patients worldwide, ovine forestomach matrix is positioned as a third generation bioscaffold that imparts significant clinical value while still achieving global sustainability goals.

Keywords

  • tissue repair
  • ovine forestomach matrix
  • sustainable biotechnology
  • circular economy
  • extracellular matrix
  • ovine forestomach matrix

1. Introduction

The use of biomaterials for tissue repair can be traced back hundreds of years to the use of both placental tissue and dermal allografts [1, 2]. Today, there is a growing number of natural and synthetic biomaterials manufactured from a variety of raw materials that are presented in different forms depending on their intended use in clinical practice [3, 4]. These biomaterials are used to restore tissue function and facilitate repair by way of both structural support to regenerating tissues and by boosting the patient’s own natural healing processes.

While there is now a large number of commercially available bioscaffolds available for the restoration and regeneration of missing or damaged soft tissues, little has been written about the sustainability of these products [5, 6]. Sustainability concepts in the design, manufacturability and clinical application of bioscaffolds can encompass many facets including the sustainability and environmental impact of raw material inputs, cultural and religious considerations for their use, and more broadly in the context of supporting the United Nations’ Sustainable Development Goals (SDGs). Ideally, sustainability objectives are balanced with clinical performance goals, such that any bioscaffold achieves its goal to improve human health outcomes without negatively impacting the environment, as outlined in Table 1. This chapter will explore sustainability concepts in regenerative biomaterials, with a focus on a sustainable next generation bioscaffold termed ovine forestomach matrix (OFM). Sustainability has been a key consideration in the development and clinical application of OFM-based devices and serves as a worked example of applying sustainability concepts to biomaterial development.

FactorConsiderations
Source raw materialsRenewable natural sources (e.g. animal or plant)
Petrochemical sources (non-renewable)
Local vs. national/global sourcing
Ethical practices at the source and through the supply chain
Supply chainFull traceability (e.g.‘farm to patient’)
In-house vs. outsourced production processes
Environmental impact [7]Carbon footprint (e.g. during manufacturing and shipping)
Water utilization
Impact on biodiversity of the raw materials and the bioscaffold manufacture
Circular Economy PotentialRaw materials are generated from or can utilize existing processes
Ethical Sourcing / Social AcceptanceAnimal welfare
Religious or cultural acceptability
Cost-effectivenessBioscaffold is affordable and accessible to patients
Bioscaffold addresses critical global healthcare needs

Table 1.

Sustainability considerations. Factors and considerations for sustainable design, manufacture, and use of biomaterials for regenerative medicine. Adapted from Kasoju et al. [5], Elfawy et al. [6] and Murphy et al. [7].

2. Evolutions of bioscaffold design: First-, second- and third- generation bioscaffolds

In the context of sustainable biomaterials, it is important to consider the evolution of these technologies in order to frame key considerations of sustainability and clinical performance (Figure 1 and Table 2). In clinical practice, biomaterials are utilized across a wide range of applications for the regeneration of either hard or soft tissues. This chapter will focus primarily on bioscaffolds designed for soft tissue regeneration. Biomaterials scaffolds are derived from a wide range of raw materials, which can be artificially manufactured from natural (e.g. collagen, elastin, alginate, hyaluronic acid) or synthetic polymers (e.g. polyurethane, polylactic acid, polyglycolic acid), or are derived from intact xenogeneic (animal) or allogeneic (human) tissues [11, 12].

Figure 1.

Schematic representation of first-, second- and third generation bioscaffolds for soft tissue regeneration. Adapted from Park et al. [8], Montoya et al. [9] and Davis et al. [10]. Created in https://BioRender.com.

StrengthsLimitations
First generation (‘bioinert’)
Highly tunable structure including pore size and fiber diameter
Rate of tissue resorption can be engineered
Potentially a low cost of goods		Do not provide any biology to aid tissue regeneration
May yield a pro-inflammatory response if non-native chemical motifs are included (e.g. chemical crosslinks)
May rapidly disintegrate (‘gel’) on contact with fluids
‘Resorbs’ rather than ‘remodels’ into the regenerating tissue
Second generation (‘bioactive’)
Provide biological components to aid tissue regeneration
Can be tailored to respond to specific signals in specialized applications
Undergo constructive remodeling		Structural features may limit cellular infiltration and proliferation
May include cellular components (e.g. DNA, lipids) of the source tissue
Reduced compatibility may decrease the persistence of the device
Third generation (‘bioresponsive’)
Mimics native tissue environment, ideal host for cell infiltration and cell-ECM dynamics
Responsive to local biological cues (e.g. cells, specific enzymes)
Promotes tissue integration and wound healing without destructive foreign body response		May persist longer as the bioscaffold integrates and remodels
Requires tightly controlled manufacturing process

Table 2.

Strengths and limitations of first-, second- and third generation bioscaffolds for soft tissue repair. Adapted from Park et al. [8], Montoya et al. [9] and Davis et al. [10].

The advent of bioscaffolds for soft tissue regeneration was marked by the development and commercialization of a synthetic biomaterial comprising crosslinked reconstituted collagen and chondroitin sulphate that was first developed for the treatment of full thickness burns [13]. Yannas and Burke described the mechanism of action of the bioscaffold as acting as a template “for the synthesis of new connective tissue and the formation of a “neodermis,“ while it is slowly biodegraded” [14]. These first-generation biomaterials are manufactured using additive (bottom-up) processes, whereby synthetic or naturally derived polymers are the raw materials for the manufacture of the bioscaffold. Additive synthetic processes, utilizing raw materials such as solubilized collagen (e.g. chemically extracted from animal tissues) or synthetic polymers (e.g. polyurethane), can be used to create complex matrix structures and enable a high degree of control over physical properties, such as pore size, persistence and strength [15]. Since the early synthetic first-generation bioscaffold, a growing number have since been commercialized, for example, from polyurethane [16], polygalactin 910/polydioxanone [17], and crosslinked collagen-elastin [18]. First-generation synthetic bioscaffolds are ‘bio-inert’ in that they provide a scaffold for cells to infiltrate and proliferate but lack additional biological components that are known to aid regeneration (Figure 1).

An alternative approach to bioscaffold design starts with a suitable intact xenogeneic or allogeneic tissue that undergoes a subtractive (top-down) manufacturing process [19]. Biomaterials made by subtractive processes begin with a complex intact tissue which is ‘decellularized’, to remove unwanted components, retaining only the desired extracellular matrix (ECM) and associated ECM components (e.g. signaling and adhesion proteins) [20]. Such biomaterials are referred to as ‘decellularized extracellular matrices’ (dECM). A large number of different tissues have been decellularized to generate innovative biomaterials for clinical use including whole organs, as well as soft tissues from mammals, fish and even plants [21, 22, 23].

Early pioneers in the field of dECM bioscaffolds sought to overcome the limitations of first-generation synthetic bioscaffolds by acknowledging that the ‘best’ bioscaffold had already been developed through a millennia of evolution – the ECM that exists in all soft tissues [24, 25]. This jump in our understanding came with the increased appreciation of the complex interplay between cells and the ECM in living tissues, referred to as “dynamic reciprocity” [26] which would be challenging to engineer with bottom-up manufacturing processes. dECM-based biomaterials manufactured by subtractive processing offer significant advantages over first-generation bioscaffolds for soft tissue regeneration. Unlike synthetic bioscaffolds, these materials consist of a “native” ECM that retains structural proteins, growth factors, and bioactive molecules necessary for supporting cell adhesion, migration, and differentiation [27]. In this way, second-generation bioscaffolds are considered ‘bioactive’ as they can deliver biological components that participate in tissue regeneration, rather than being a purely bioinert scaffold (Figure 1). Unlike the first-generation bioscaffolds that simply resorb into the regenerating tissue, these second-generation bioscaffolds undergo constructive remodeling characterized by an early transition to the M2 macrophage phenotype and deposition of host ECM as the dECM bioscaffold is resorbed into the regenerating tissue [28]. Key to the success of these materials is the removal of cellular components that reduces the risk of inflammation and rejection by the host, while retaining the structure and biology of the tissue ECM [29]. Xenogeneic cellular material (e.g. DNA and cellular lipids) has been shown to lead to a pro-inflammatory response, particularly by host macrophages [28, 30]. The process of decellularization is a delicate balance between the removal of unwanted cellular components, and the retention of desirable ECM structure, biology and compatibility. However, this is especially challenging at the scale of commercial manufacturing. While some early dECM-based bioscaffolds successfully retained many important ECM-associated components that aid tissue regeneration [31, 32, 33], tissue processing either damaged the structural features of the ECM and/or was ineffective in removal of unwanted cellular components leading to accelerated degradation (Table 2). Accordingly, a new generation of dECM biomaterials have been designed and developed to balance key requirements of bioscaffold design for soft tissue regeneration, namely, structure, biology and compatibility (Table 2 and Figure 1).

3. Ovine forestomach matrix: A third generation bioscaffold for soft tissue regeneration

Research in the field of dECM-based bioscaffolds has highlighted the importance of the raw tissue starting material, as well as the manufacturing process used for decellularization [34, 35, 36, 37]. Preserving the native biology and structure of the tissue ECM is critical to ensuring the body responds appropriately and remodels the bioscaffold and was a key tenet in the development of a bioscaffold derived from ovine forestomach tissue. Manufacturing OFM starts with sheep forestomach (‘rumen’) tissue that would otherwise be discarded from New Zealand’s agricultural industry (Figure 2). Forestomach tissue undergoes a process to gently separate the unwanted layers of the tissue, namely the muscle (tunica muscularis), and epithelial layers (lamina epithelialis, comprising a stratified squamous epithelium) [38]. The remaining tissue layer, termed the propria submucosa comprises two layers, a dense luminal layer, termed the lamina propria and a more open reticular abluminal layer, termed the tunica submucosa (Figure 2). Decellularization of the propria submucosa proceeds using mild detergents, chelating agents and salts, a process which has been highly optimized to preserve the delicate microarchitecture, biological components and compatibility of the ECM (Figure 3) [38].

Figure 2.

Production of ovine forestomach matrix (OFM) from ovine forestomach tissue, including key sustainability considerations of the tissue source. Created in https://BioRender.com.

Figure 3.

Schematic representation of the structural, biological and immune compatibility properties of ovine forestomach matrix. Created in https://BioRender.com.

3.1 Structural considerations

Prior structural studies have characterized OFM using differential scanning calorimetry (DSC) [39], scanning electron microscopy (SEM) [38], atomic force microscopy (AFM) [40], histology and immunohistochemistry [38], Sirius Red staining [41], small-angle x-ray scattering (SAXS) [41, 42], and micro computerized topography (MicroCT) [40]. These structural studies provide evidence that the scaffold architecture of OFM mimics the architecture of soft tissue ECM. One unique structural feature of OFM is the presence of acellular vascular channels that run through the propria submucosa and ultimately provide a template for the reformation of host vasculature in the process of angio-conduction [40]. Studies have shown that these vascular channels retain an intact endothelial basement membrane [38] that enables host endothelial cell attachment, migration and vessel formation. The open porous ‘native’ architecture of OFM enables rapid cell infiltration and proliferation, as evidenced by various in vivo models of soft tissue regeneration [43, 44, 45].

3.2 Biological considerations

Proteomic analysis of OFM, as well as traditional quantitative and qualitative assays, have identified numerous structural, adhesion, regulatory and signaling proteins that are present in the bioscaffold [38, 46]. These characterization studies provide evidence that the bioscaffold contains not just structural proteins (e.g. collagen), but a heterogeneous mixture of naturally occurring ECM components (e.g. elastin, fibronectin and decorin), or ECM-associated components (e.g. growth factors, regulatory proteins). Importantly, the functionality of these components has been preserved as demonstrated by the bioactivity of OFM in in vivo, ex vivo and in vitro studies [38, 43]. In this way, these important ECM-associated components are available to participate in tissue regeneration when used clinically. The improved functionality of the OFM versus first and second-generation bioscaffolds has been demonstrated in various in vitro and in vivo studies [43, 44, 45]. For example, in vivo studies have demonstrated that OFM-based devices facilitate improved cellular adhesion, infiltration and proliferation into the scaffold, with deposition of a dense capillary network within the regenerating tissue [38, 43]. In vitro studies have demonstrated that OFM stimulates significantly more endothelial migration and proliferation relative to a first-generation bioscaffold that lacks ECM-associated growth factors [43]. Additionally, OFM devices have been shown to give rise to a significantly more blood vessels in vivo relative to a second-generation dECM-based bioscaffold [43]. This may be attributed to the unique bioactivity of OFM combined with avascular blood vessels found in the material. Finally, in vitro and in vivo studies demonstrated that components present in OFM recruit progenitor cells [47].

3.3 Compatibility considerations

OFM-based devices undergo constructive remodeling, akin to the processes that underly normal tissue turnover in mammalian tissues [43]. This pro-healing or anti-inflammatory response is characterized by an early transition to the M2 macrophage phenotype, as opposed to the sustained pro-inflammatory M1 macrophage response that leads to scar tissue deposition [48]. Proteomic analysis of OFM has identified naturally occurring anti-inflammatory proteins within the material (e.g. tissue inhibitors of matrix metalloproteinases) [46], and further, studies have demonstrated that these components of OFM inhibit tissue proteases in vitro [49]. While cellular components (e.g. DNA and lipids) are often a characteristic of second-generation dECM bioscaffolds, studies have shown the optimized manufacturing process for OFM effectively removes residual ovine cellular components while retaining ECM structure and biological components [38]. Studies in animal models of wound healing [43], abdominal wall reconstruction [44, 45], and rotator cuff repair [50] have demonstrated that OFM elicits a pro-healing response, and ultimately undergoes constructive remodeling leading to functional tissue regeneration. For example, Street et al. [48] demonstrated that human primary dendritic cells were not activated on exposure to OFM. Dendritic cells are highly responsive to foreign materials, so a non-activation response by OFM indicates that it is well tolerated by the immune system, despite the presence of proteins from a different species.

3.4 OFM: A bioresponsive third generation bioscaffold

OFM can be classified as a third generation ‘bioresponsive’ material (Figures 1 and 3). Structurally, the complex architecture of the dECM is biomimetic, providing a strong foundation for cell infiltration and attachment, and it persists in vivo supporting long term remodeling and reconstruction [38, 43]. In terms of tissue compatibility, it has been shown that OFM does not illicit dendritic cell activation [50], or induce a foreign body response in animals and humans, but rather supports constructive remodeling and resolution of the inflammatory phase as demonstrated by the progression to angiogenesis and vascular bed formation [43, 51]. In terms of biology, OFM contains a diverse range of biological molecules including growth factors, proteoglycans and regulatory proteins. As OFM slowly remodels these molecules boost the regenerative response in the healing tissue. In addition to its composition, the material is responsive to cells and the tissue environment. Macrophage breakdown of OFM leads to the release of a decorin derived chemotactic factor, that recruits local progenitor cells [47]. This interaction shows that OFM is a responsive participant in the regenerative process. In addition, the specific inhibition of tissue proteases by OFM has an important impact on the chronic wound environment. It has been shown that inhibitors (serpins) are found in OFM [46] and that OFM can specifically inhibit relevant proteases which prevent the resolution of inflammation in chronic wounds, such as neutrophil elastase [49]. Thus, OFM responds to neutrophil overactivity in the environment by suppressing this key protease.

3.5 Clinical applications of OFM-based devices

OFM can be fabricated into a range of medical devices for applications in soft tissue repair across the human anatomy, including devices for wound care [52], multi-layered devices for complex plastic surgery and burns [40], and reinforced biologics for abdominal wall repair [42]. Additionally, the technology can be further augmented with processes developed to impregnate devices with anti-microbials [39], additional pro-healing bioactives [40], or as a cell-carrier for engineered fibroblasts [53].

OFM-based devices are used clinically in acute and chronic wound healing [54, 55], burns [56], traumatic wounds [57, 58], abdominal wall reinforcement [59, 60], and breast reconstruction [61]. Over 7 million OFM-based devices have been employed in clinical settings to date. Generally, OFM-based devices have been shown to be resilient and robust in the presence of bacterial contamination [62, 63, 64, 65], lead to rapid tissue regeneration in complex volumetric defects [57, 58] and restore functional tissue to the affected site long-term. Since bioscaffold performance may come at the expense of sustainability goals, it is important to also consider OFM in terms of its sustainability, environmental impact, and cultural implications.

4. Forestomach tissue: A unique tissue source for dECM bioscaffolds

Traditionally second-generation dECM bioscaffolds have been manufactured from porcine, human, or bovine tissues. However, forestomach tissue sourced from sheep offers a unique alternative raw material, with many advantages with respect to sustainability and performance of the resultant dECM bioscaffold. Ruminants, including sheep, have evolved a unique digestive system that enables them to extract nutrients from fibrous plant material [66, 67]. The forestomach consists of four compartments, the rumen, reticulum, omasum, and abomasum, which break down complex plant matter using mechanical force and microbial fermentation (Figure 4) [68, 69]. The tissues of the forestomach have evolved distinctive structural, biological and immunological properties [71] that are imparted to OFM.

Figure 4.

Schematic representation of forestomach tissue development from birth to maturation at ~16 weeks. Adapted from Wardrop and Coombe [70]. Created in https://BioRender.com.

4.1 Physical properties of forestomach tissue

Traditionally second-generation dECM bioscaffolds have been manufactured from porcine, human, or bovine tissues. However, forestomach tissue sourced from sheep offers a unique alternative starting raw material, with many advantages with respect to sustainability and performance of the resultant dECM bioscaffold. Ruminants, including sheep, have evolved a unique digestive system that enables them to extract nutrients from fibrous plant material [66, 67]. The forestomach consists of four compartments, the rumen, reticulum, omasum, and abomasum, which break down complex plant matter using mechanical force and microbial fermentation (Figure 4) [68, 69]. The tissues of the forestomach have evolved distinctive structural, biological and immunological properties [71] that are imparted to OFM.

The forestomach, even in lambs, is a relatively large muscular organ. For example, in lambs, the distended rumen has a volume of 10–15 L, making it possible to produce large continuous sheets of OFM [38]. The “rumination” process involves the physical mixing, churning and moving of cellulose-based ingestate to maximize fermentation by the forestomach microbiome [72]. Thus the propria submucosa has evolved to sustain these demanding mechanical stresses, and is an especially dense layer of ECM, referred to previously as a ‘condensed fibrous layer’ [73]. For example, studies have shown that OFM isolated from sheep forestomach is significantly stronger than dECM isolated from other mammalian gastrointestinal (GI) tissues or bladder [42].

The forestomach of ruminants has evolved to be highly vascular. Firstly, the microbial fermentation process is metabolically demanding and requires a constant supply of nutrients, oxygen and the removal of byproducts, like lactic acid [69]. Secondly, the rumen wall absorbs volatile fatty acids (VFAs) produced by the fermentation process [69, 74, 75, 76, 77]. VFAs are the primary energy source for ruminants and the dense vascular network of the forestomach is essential for transporting VFAs directly into the blood stream as an immediate energy source. Thirdly, fermentation within the forestomach generates a large amount of heat, and therefore vasculature within the tissue has an important role in thermoregulation [78]. The highly vascular nature of the forestomach tissue imparts some of these beneficial properties to OFM. For example, acellular vascular channels found in OFM are a unique structural property of the material [40]. Additionally, ECM components that form vasculature, like collagen IV, fibronectin, perlecan, thrombospondin and vitronectin are present in OFM [46]. Both fibronectin [79] and perlecan [80] containing biomaterials have been shown to stimulate angiogenesis.

4.2 Immune system adapted to support digestive flora

Commensal microbes in the forestomach environment are essential for the fermentation process [81]. The forestomach must therefore maintain a tolerant immune environment, where immune cells tolerate bacteria, fungi and protozoa that are essential for digestion [82]. The local immune response in each compartment of the ruminant stomach has distinct functions; the tissue of the rumen and reticulum is adapted to immune tolerance and microbial balance, while omasum and abomasum create barrier immunity and direct antimicrobial activity, respectively. If an infection occurs, the immune cells of this organ must recognize and respond to harmful pathogens. In the rumen, dendritic cells, regulatory T cells and macrophages maintain the “immune tolerance” and prevent an excessive activation of the immune system [83]. Key cytokines promote a non-inflammatory environment while cells can mount a “local inflammatory response” in the case of injury or infection or dysbiosis [84]. The ECM of the propria submucosa plays an important role in supporting immune cell trafficking and chemotaxis while providing a local microenvironment of immune tolerance by sequestering chemokines like IL-10 and TGFβ [83]. The dynamic response to microbial antigens specific to this tissue are built into the ECM of the forestomach, for example via integrins that act as immune cell attachment sites. An abundance of hyaluronic acid and heparan sulfate proteoglycans, unique to forestomach tissue helps to promote immune cell migration and modulate inflammatory signals [85, 86].

Clinical studies indicate that OFM is well tolerated in the presence of bacterial contamination, a feature that may be attributed to the unique composition of bacterial regulatory proteins in the tissue ECM that are retained after tissue decellularization [63, 87, 88].

4.3 Regenerative capacity

In most adult tissue, cells and ECM exist in a resting state, while in juvenile and developing animals, especially in utero, more dynamic interactions exist between cells and the ECM. Consequently, there is strong evidence that dECM-based bioscaffolds isolated from juvenile tissues have increased bioactivity versus dECM isolated from adult or mature tissues [89]. This is believed to result from an increase in the tissue concentration of ECM-associated bioactives (e.g. growth factors) resulting from the plasticity of developing tissues. For this reason, OFM is produced from tissues collected from animals between 6 to 12 months in age. Additionally, unlike many other mammalian tissues, forestomach tissue develops exponentially in neonatal animals at the onset of grass feeding. The forestomach is the fastest growing organ in these young animals with a drastic increase in size within the first 6 months of the animal’s life (Figure 4) [90]. During weaning, the capacity of the rumen changes from 30 to 70% of the entire gastrointestinal tract of the ruminant [91, 92]. The rapid growth and development of the forestomach can be attributed to a unique cell population and biology. The shift in cell population in the first 6 months of the animal’s life shows a distinctly dynamic tissue environment as a result of the rapid growth, exposure to microbes and physical forces [68, 93, 94].

The physical and structural changes, including the development of papillae and rapid epithelialization are coordinated by the rapidly shifting cell population [95, 96]. Importantly, the growing forestomach is a rich source of progenitor and stem cells, which are located in the basal layer and contribute to the continuous turnover of epithelial cells [68, 97, 98]. The stem cell niche within the developing forestomach is maintained by the structural scaffolding and molecular signals of the tissue ECM.

Tissue turnover of the rumen is higher than other parts of the GI tract, as evidenced by analysis showing high level of cell cycle processing genes. For example, a cluster analysis of forestomach tissue groups it closer to tonsil and skin than other parts of the GI tract [99]. Tissues like forestomach that have an important barrier function need to be able to turnover and regenerate quickly to recover from injury. The stratified squamous epithelium of forestomach, like skin, has an adaptable mitotic index, where damage can easily be replaced by basal cells resting in G2 phase [100]. Stratified squamous epithelium is composed of a basal layer containing progenitor cells to rapidly divide and replace damaged or missing cells from the epithelium [101]. Tissues such as the skin, oral cavity, and gut are considered ‘high turnover’ or dynamic because they undergo frequent and rapid renewal of their cells and ECM, in contrast to ‘low turnover’ or static tissues like bone and nerve [102]. Constant degradation and regeneration of dynamic tissue requires an ECM capable of supporting rapid cell division, ECM synthesis and efficient repair which differs from static tissue in several ways. In terms of composition, dynamic tissue ECM, like that found in forestomach, is characterized by higher levels of elastin, GAGs and proteoglycans and regulatory proteins. Structurally, ECM from static tissue is likely to contain more densely packed, crosslinked collagen fibers, while ECM from dynamic tissue is more malleable and open in structure to allow rapid regeneration by cell division, recruitment and access to the matrix for remodeling [102].

Evolution has driven the forestomach tissue and ECM of ruminants to evolve distinct tracts with respect to mechanical properties, vascularity, immune regulators, bacterial regulators, and capacity for renewal and regeneration that may in part account for some of the unique properties of OFM seen in vitro, in vivo and in clinical applications.

5. Sustainability of ovine forestomach tissue: An alternative tissue source for sustainable bioscaffolds

Ovine forestomach tissue, the raw material for the production of OFM, is sourced from free-range, pasture-raised sheep in Aotearoa New Zealand, where farming practices are deeply aligned with environmental sustainability, ecosystem health, and low-impact methods [75]. Advanced agricultural practices are integral to New Zealand’s economic stability and cultural identity, as the country’s economy relies heavily on its natural resources, including agriculture. Unlike many European and North American countries, New Zealand’s temperate climate and geography make pasture-based farming a cost-effective means to produce high-quality meat for human consumption. By pasture raising its animals, New Zealand agriculture avoids the need for indoor housing (‘barn raised’), or artificial rearing systems, resulting in lower labor costs and improved animal welfare practices. Animals raised on the pasture benefit from enhanced immune function, which leads to healthier and more productive tissues. The soft tissues of pasture-raised livestock, living in a low-stress environment, produce tissues with increased firmness, lightness of fat cover, favorable fatty acid composition, and a higher meat oxidative stability. In contrast, tissues from barn-raised animals suffer from higher tissue mass variability [75].

Because New Zealand’s’ livestock are raised on the pasture, the necessity for veterinary medicines is reduced versus barn-raised livestock [103]. This is especially important when considering that the use of antibiotics in animals raised for human consumption is an important concern for the World Health Organization in relation to antibiotic resistance [104]. A comparison of antibiotic concentrations present in global animal tissues in 2012 ranged from 3.8 to 341 active units per mg/kg biomass, with New Zealand livestock meat having the third lowest average concentration of 9.4 active units per mg/kg biomass [105]. A more recent 2024 study, has now put this average concentration at 5.8 active units per mg/kg biomass [106]. This is in stark contrast to the antibiotic usage seen in other countries where animals are typically barn-raised, putting livestock at greater risk of microbial infection and where antibiotics administration is more widespread. Additional supplements, such as the use of hormonal growth promotants (HGPs), are regulated and prohibited in New Zealand sheep farming. The impact of these agents on animal tissue and their safety for human consumption is contested [107, 108, 109, 110], but there is clear evidence that environmental run off from HGPs and antibiotics used in farming has a negative impact on the environment, especially waterways and on antibiotic resistance.

New Zealand’s pasture-based and sustainable farming practices not only support animal health but also contribute to biodiversity and environmental conservation. Pasture-based agricultural practices promote soil health, reduce the need for synthetic fertilizers and pesticides, and help mitigate soil degradation, water pollution, and habitat destruction [111]. Additionally, New Zealand’s grass-fed livestock require fewer resources and produce lower carbon emissions compared to barn-raised livestock, contributing to a more carbon-efficient model of farming [112]. For example, New Zealand’s sheep meat production has a carbon footprint of 6.01 kg CO2-e per kg, which is significantly lower than the international average of 14.2 kg CO2-e per kg [113]. Through regenerative agriculture and soil management, New Zealand also sequesters carbon, further reducing the environmental impact of its agricultural practices [114].

Pasture-based farming in New Zealand accounted for approximately 18.4 million lambs in 2024, with the majority of this animal tissue being supplied for human consumption. Ovine forestomach, the raw material for OFM, is typically discarded, with only a small proportion of this tissue being consumed as tripe. As a byproduct of New Zealand’s existing agricultural industry, re-purposing this otherwise discarded tissue for the production of OFM provides a sustainable option to alternate animal tissue sources of dECM-based bioscaffolds. Animals raised as a tissue source for medical applications are usually housed in specialized facilities to enable animals to be raised and managed as ‘closed-herds’. These husbandry practices require significant resources and generate large amounts of waste. With a focus on output rather than animal welfare, these facilities have a detrimental impact on the environment, raise ethical concerns for end-users and cause health problems for the animals [115, 116].

New Zealand pasture-raised lambs offer several unique sustainable attributes over traditional livestock sources of raw material, including reduced anti-biotic usage, reduced cost and improved animal welfare. Wild caught animals may provide a suitable alternative to pasture raised and closed herds for the production of tissues for medical applications. However, even tissues sourced from wild animals may place a burden on the environment. For instance, although a bioscaffold produced from North Atlantic cod fishing could be considered a sustainable resource, the burden on the environment related to the fishing industry is concerning due to the impact of overfishing, habitat destruction, bycatch and climate change impact [115]. Synthetic bioscaffolds are a greater burden on the environment as generally these raw materials originate exclusively or in part from the petrochemical and/or plastics industries where environmental impact and sustainability has been an on-going concern.

6. Ovine forestomach tissue: Disease transmission considerations

Tissue-derived bioscaffolds for soft tissue have traditionally been sourced from a variety of different mammals. The risks of microbial contamination (e.g. bacteria and yeast) of these products is controlled for by terminal sterilization, chemical disinfection and/or aseptic processing. However, all source tissues pose a potential safety risk for the end users due to the possibility of viral or prion transmission. There are notable examples of iatrogenic transmission of viral and prion disease to humans from tissue-derived medical devices, including the transmission of human immunodeficiency disease from human demineralized bone [117], and the transmission of Creutzfeldt-Jakob disease (CJD) from human dural grafts [118]. Since early reports of transmission of human disease from tissue-derived bioscaffolds, more stringent controls have been placed on manufacturing, including sourcing, diagnostic screening and geographical controls.

6.1 Viral transmission risk mitigation

Human (cadaver)-derived materials often carry a higher risk of viral contamination, particularly when tissue degradation has occurred, or improper storage conditions are used [119, 120]. For this reason, there is a heavy onus on manufacturers of cadaveric tissue derived bioscaffolds for screening diagnostics controls (e.g HIV, Hepatitis) to prevent contamination of medical devices [121]. The transmission of porcine viruses to humans is well documented and this risk is escalated for tissue derived from barn-raised, closed herd animals [122]. However, like cadaveric tissues, viral inactivation studies can be used to validate the manufacturing process used for animal tissues are effective at eliminating viral pathogens from finished medical devices. An additional control that is not uncommon in the medical device industry is to use closed-herds’ of pigs or cows that are specifically raised as a tissue source to produce tissue-derived bioscaffolds. Closed herds are effectively quarantined from external pathogens enabling an additional level of safety with respect to transmissible viral pathogens. Globally there are several ovine viruses that are known to cause disease in humans. For example, viral diseases such as Rift Valley fever, Crimean-Congo hemorrhagic fever, Nairobi sheep disease caused by Bunyaviridae family viruses originate from sheep, but are only known outside of New Zealand. Rabies virus is also detected in sheep, but again is considered exotic to New Zealand [123]. The most common viruses in New Zealand sheep, include ovine adenovirus, ovine herpes virus and papillomavirus, are benign to humans. Orf and parainfluenza type 3 (PI-3) have been reported in New Zealand sheep, and can be transmitted to at risk humans, such as farmers and meat packers, but only lead to localized resolving lesion [124]. Generally, the risk from ovine viruses is less than those from human, porcine or bovine tissues, and this risk is further reduced when considering the only ovine viruses known to exist in New Zealand.

6.2 Prion transmission risk mitigation

The misfolded prion protein is the causative agent in human CJD as well as bovine spongiform encephalopathy (BSE) in cows, scrapie disease in sheep and chronic wasting disease (CWD) in deer [125, 126]. The transmission of prion disease from cattle to humans and from humans to humans is well documented [127, 128]. Unlike common micro-organisms (e.g. viruses, bacteria, yeast/mold), inactivation of the infectious isoform of the prion protein requires strongly denaturing alkaline conditions which is not generally amenable to the manufacture of tissue-based bioscaffolds [129, 130, 131, 132, 133, 134]. These strongly denaturing inactivation conditions may be compatible with the isolation of solubilized collagen from animal tissues, by could not be used for tissue decellularization which requires significantly more gentle conditions to preserve ECM structure and biology. Therefore, risk mitigation steps focus on tissue sourcing controls (i.e. species, age and geographic location) rather than diagnostics or inactivation [135, 136].

Intra-species and inter-species transmission of the infectious prion protein is governed by a “species barrier” that determines the ability of prion disease to transmit [137]. For example, prion transmission from sheep to cows, and cows to humans has been reported, indicating that there is no species barrier for transmission between these species [138, 139]. However, transmission from sheep to humans has not been demonstrated, either clinically or in transgenic animal models of the disease [140]. While sheep prion disease (‘scrapie’), has been known since the eighteenth century [141], and humans have co-habited with, farmed and consumed sheep for centuries, the human population has been protected from sheep prion disease by the species barrier to transmission [140].

In addition to a species-barrier that protects humans from potential prion disease transmission form sheep-tissue derived bioscaffolds, an additional layer of protection is afforded by New Zealand’s ‘prion free’ status [127, 142, 143, 144, 145, 146]. Prion disease in New Zealand cattle (BSE) and sheep (scrapie) has not been reported [143]. This status is maintained by New Zealand’s unique geographical isolation, strict border controls that prohibit the import of animal tissues into New Zealand, livestock feeding practices that prohibit the feeding of animal tissues to livestock, and ongoing monitoring. Additionally, prion disease onset typically affects older animals [147], therefore, by sourcing tissues from younger animals, this further reduces prion transmission risk. Every animal that enters the food supply in New Zealand receives a pre-mortem veterinary inspection to ensure animals do not show clinical signs of disease and the high-volume meat processing segregates high risk animal tissues (e.g. brain and brain stem) from lower risk tissues (e.g. forestomach).

7. Religious and ethical considerations: An often-overlooked aspect of sustainability

Animals and animal-derived technologies, including biomaterials, have contributed greatly to modern medicine and scientific discovery. While these contributions have undoubtably advanced human health, the use of animals in scientific research and medical technologies may run counter to certain ethical, cultural and religious principles. For some these potential objections can lead to a potential ethical quandary that may result in mistrust of those involved in industries that draw from animal-based resources, including healthcare, businesses, scientists and medical professionals [148].

Many religious and cultural groups hold strong views on the use of mammalian tissues for food or for medical applications. Approximately 23% of the world’s population is Muslim, of which 83% prefer or require halal as a dietary standard. Judaism represents 0.2% of the world population and 35% of these individuals practice kosher food standards [149]. In terms of the medical use of mammalian tissues, many religions object to or are concerned with the use of human tissues, organs, and blood. This may also extend to stem cell research and genetic engineering. In respect to the use of animal tissues in medical products, many Hindus follow a strict vegetarian diet which extends to a prohibition on the use of such products [150]. Similarly, Buddhists may object to the use of bovine-derived medical devices, while strict Muslims may object to the use of porcine-derived materials. There is also a growing number of the global population, who prefer not to consume meat and animal products due to the impact these have on the environment. Vegetarianism and veganism are now practiced by 6% of individuals under 35 years old [151, 152]. The growing conscience of animal protections in medical research has guided the adaption of the three Rs principles to reduce the use and harm of animals in scientific research “replace, reduce, refine” [153]. Similarly, in agricultural practices there is a growing consumer demand for ethically sourced and raised meat products. As society becomes increasingly aware of the ecological impact of industrial animal farming and the ethical ramifications of animal use, there is pressure on all stakeholders to innovate responsibly.

In the context of animal-tissue derived bioscaffolds, sustainability principles emphasize the responsible use of natural resources while minimizing negative environmental and ethical impacts (Table 1). Ethical business practices in this field must align with broader sustainability goals, which include reducing waste, promoting resource efficiency, and ensuring that production methods do not exploit animals or harm ecosystems. Organizations involved in the production and use of animal-derived biomaterials have a responsibility to adopt transparent, sustainable practices that respect both the environment and animal welfare. A sustainable approach involves sourcing materials from responsible suppliers who adhere to ethical standards and prioritize humane treatment of animals. This includes ensuring that animals are raised in conditions that meet high welfare standards and that any tissue collection processes are conducted with minimal harm.

In addition to the responsible sourcing of animal tissues, companies must also consider the life cycle of the products they create. This includes evaluating the energy, water, and resource consumption involved in the production of animal-derived biomaterials and aiming to reduce carbon emissions and waste throughout the supply chain. Integrating sustainability principles into business operations not only enhances corporate social responsibility (CSR) but also contributes to the broader movement toward a circular economy, where materials are reused, and waste is minimized.

OFM is created by sourcing materials from carefully managed pasture-raised farming systems that adhere to New Zealand’s animal welfare standards, sustainable farming and ecological responsibility. The tissue is obtained from the meat processing industry which is highly regulated in terms of animal welfare, halal and kosher compliance and export compliance of international markets [111]. It is derived from parts of the animal that would otherwise go to waste thereby maximizing resource use and promoting sustainability. In contrast, intensive farming and barn raised animals such as pigs and cows may be perceived as more harmful in terms of ethical and religious views compared with pasture raised animals [103, 151, 152, 154].

8. Ovine forestomach matrix: A universal bioscaffold for soft tissue regeneration

The United Nations’ Sustainable Development Goals (SDGs) were adopted in 2015 and act as an international framework for achieving a more inclusive, equitable, and sustainable future. The SDGs address many aspects of sustainability relevant to bioscaffold development including environmental sustainability, innovation and responsible production. Of particular relevance to the development and clinical application of bioscaffolds is SDG-3 that aims to “Ensure healthy lives and promote well-being for all at all ages”. This goal encompasses a wide range of targets aimed at improving health outcomes, reducing mortality rates, and addressing global health challenges.

The global cost of health care was estimated by the WHO in 2020 at $9 trillion, with spending unequally distributed across the nations. For example, the 2022 global market for soft tissue bioscaffolds is estimated at $1.8 billion, with the primary markets being developed countries (e.g. USA, EU), reflecting the inequity of access to these types of products [155]. While advances in the fields of regenerative medicine and bioscaffolds could contribute significantly to the global burden of disease, widespread adoption of bioscaffolds has generally been limited due to the traditionally high cost of these technologies. For example, synthetic first-generation bioscaffolds cost up to $USD25.34/cm2, second-generation bioscaffolds including urinary bladder matrix (UBM) and foetal bovine dermal matrix (FBDM) cost up to $USD18.55/cm2 and $39.06/cm2, respectively [156]. A fish skin graft has been reported to cost $USD78.30/cm2 [156]. OFM-based medical devices have been commercialized for a wide range of different clinical applications in soft tissue repair and regeneration including wound care, plastic and reconstructive surgery, and abdominal wall reinforcement. As highlighted earlier, OFM is produced from a sustainable tissue source that is otherwise discarded from New Zealand’s agricultural industry. As such there is a low cost of goods associated with forestomach tissue, relative to other mammalian tissue sources (e.g. porcine, bovine, cadaveric). Key to the clinical utilization of OFM-based devices is their relatively low cost, and a commitment to democratize the use of these regenerative technologies. For example, the reported cost of an equivalent OFM-based device to those examples listed above, is up to $USD11.19/cm2 [156], representing a significant cost reduction and an increase in the potential for wider global adoption.

Widespread clinical adoption of new technology must consider affordability but also clinical performance in order to be sustainable. OFM-based devices while being priced responsibly, have also been shown to be clinically effective. For example, a large real world evidence (RWE) study comparing the healing of diabetic foot ulcers (DFU) using OFM compared to a first-generation bioscaffold comprising reconstituted collagen and chemically modified cellulose (collagen/ORC) showed that treatment with OFM significantly reduced the time to heal these wounds [54]. While OFM and the first-generation collagen/ORC bioscaffold are equivalently priced ($USD 8-12/unit), treatment of DFUs with OFM was shown to reduce the time to wound closure by up to 5.3 weeks. A more recent RWE study similarly compared healing outcomes between OFM- and collagen/ORC-treated venous leg ulcers (VLU) [157]. In this study, similar outcomes were observed with the time to wound closure of OFM-treated VLU being reduced by ~35% relative to VLU treated with collagen/ORC. The improved healing outcomes of OFM-treated wounds versus those achieved with a first-generation synthetic bioscaffold (collagen/ORC) are likely attributed to improved structure, biology and compatibility (Figure 3) of OFM. For example, in vitro studies have shown that OFM gives rise to significantly more endothelial cell migration, and proliferation versus collagen/ORC [43] and has a preserved matrix structure versus collagen/ORC [39]. Globally chronic non-healing wounds are estimated to affect ~1–2% of the population [158]. Managing non-healing wounds places an enormous burden on healthcare systems, health care providers, patients and their family. For example, patients are burdened with the pain, smell and discomfort of a non-healing wound, reduced mobility and normal activities, and compromised ability to work and support their families and communities. As such, bioscaffolds that accelerate wound healing and soft tissue regeneration offer great promise for global health when these advanced technologies are sustainable and cost-effective.

Post-operative complications, for example, infection, tissue necrosis, dehiscence or recurrence may in some instances place a larger burden on the patient and healthcare system versus the initial procedure. It is estimated that the cost of a surgical site infection (SSI) is $USD34,000 to the healthcare system per SSI, as well as a cost to the patient of increased hospital days, number of procedures and reduced income [159]. First-generation bioscaffolds, lacking additional biological components to aid tissue regeneration and vasculogenesis, have relatively high infection rates. For example, an early first-generation synthetic bioscaffold comprising crosslinked reconstituted collagen and chondroitin sulphate has reported infection rates of up to 9–14% [160], 40% [161], 35% [162], 33% [163] and 26% [164]. Similarly, a more contemporary synthetic bioscaffold comprising polyurethane has reported infection rates of 42% [165], 40% [166], and 39% [167]. In contrast, infection rates for third generation bioscaffolds, including OFM-based devices are typically low, even when these devices are used in soft tissue defects that may be contaminated. Another clinical limitation of first-generation synthetic devices is ‘graft loss’ whereby the bioscaffold or part thereof, fails to integrate with host soft tissue [168], requiring re-operation and explanation of the bioscaffold. For example, incidence of graft loss for synthetic first-generation bioscaffolds have been as high as 45% [168], 21% [169], 19% [170], 34% [165], and 30% [171]. Graft loss has not been reported for OFM-based devices, which most likely can be attributed to the functional cell adhesion proteins and growth factors that can orchestrate cell migration and proliferation.

Many second-generation bioscaffolds do not persist in the hostile environment of an inflamed or contaminated soft tissue defect and may require repeat applications that come at a cost to the patient and the healthcare system. OFM-based devices have been shown to be relatively robust in these types of soft tissue defects, most likely due to their intact matrix structure and anti-inflammatory components. For example, a prospective evaluation of OFM-based devices used across 130 complex lower extremity reconstructions reported a median product application of one, thus avoiding the need for repeat surgical procedures [156]. In contrast, some bioscaffolds have been reported to require up to seven applications in a similar clinical setting to reconstruct similar lower extremity defects [172].

Disease recurrence additionally places significant burden on healthcare systems, patients and care providers. Bioscaffolds have traditionally been used in abdominal wall reinforcement, often as an alternative to synthetic surgical meshes. OFM-based devices have been extensively used in hernia repair with reported hernia recurrence rates of 1.2% [173], 0% [174], 0% [88], 6% [175], 3% [176], 2.8% [177], 16% [178], 4% [177], and 3.6% [179]. For example, one study compared recurrence rates of ventral hernia repaired with either OFM-based devices or acellular dermal matrices, with reported recurrence rates of ~3% and up to ~30%, respectively [59].

As a low-cost, high-performance third generation biomaterial, OFM has the potential to improve the global standard of care, thus contributing to achieving universal health coverage by providing high-quality, affordable medical solutions.

9. Conclusion

Regenerative biomaterials have seen multiple iterations over the past five decades, progressing from simple synthesized scaffolds to more elegant and complex designs. As this field continues to mature, it is critical to consider sustainability concepts in the design of future materials. Factors such as source material availability, environmental impact of both the processing and waste materials, animal welfare for biologically-sourced materials, and affordability of the end product are critical sustainability considerations. This ensures that proposed solutions are suitable for clinical scale up and do not sacrifice cultural and environmental goals at the expense of minimal clinical improvements. Presented as a case review, this chapter provided an overview of OFM in the context of sustainable biomaterial design. Capitalizing on its sustainable sourcing, design and manufacture, the commercial use of OFM in clinical practice has directly contributed to global health sustainability goals by offering high-quality, accessible, affordable medical solutions to address critical challenges worldwide.

Acknowledgments

The authors wish to acknowledge Aroa Biosurgery Limited and all those who have advanced the understanding of ovine forestomach matrix.

Conflict of interest

SGD, RWFV, DAY and BCHM are employees and shareholders of Aroa Biosurgery Limited, manufacturer of ovine forestomach matrix.

References

  1. 1. Kaul H, Ventikos Y. Dynamic reciprocity revisited. Journal of Theoretical Biology. 2015;370:205-208
  2. 2. Marin E, Boschetto F, Pezzotti G. Biomaterials and biocompatibility: An historical overview. Journal of Biomedical Materials Research. Part A. 2020;108(8):1617-1633
  3. 3. Webber MJ, Khan OF, Sydlik SA, Tang BC, Langer R. A perspective on the clinical translation of scaffolds for tissue engineering. Annals of Biomedical Engineering. 2015;43(3):641-656
  4. 4. Litowczenko J, Wozniak-Budych MJ, Staszak K, Wieszczycka K, Jurga S, Tylkowski B. Milestones and current achievements in development of multifunctional bioscaffolds for medical application. Bioactive Materials. 2021;6(8):2412-2438
  5. 5. Kasoju N, Sunilkumar A. Convergence of tissue engineering and sustainable development goals. Biotechnology for Sustainable Materials. 2024;1(1):20
  6. 6. Elfawy LA, Ng CY, Amirrah IN, Mazlan Z, Wen APY, Fadilah NIM, et al. Sustainable approach of functional biomaterials-tissue engineering for skin burn treatment: A comprehensive review. Pharmaceuticals (Basel). 2023;16(5):701
  7. 7. Murphy E, Curran TP, Holden NM, O'Brien D, Upton J. Water footprinting of pasture-based farms; beef and sheep. Animal. 2018;12(5):1068-1076
  8. 8. Park HJ, Hong H, Thangam R, Song MG, Kim JE, Jo EH, et al. Static and dynamic biomaterial engineering for cell modulation. Nanomaterials (Basel). 2022;12(8):1377
  9. 9. Montoya C, Du Y, Gianforcaro AL, Orrego S, Yang M, Lelkes PI. On the road to smart biomaterials for bone research: Definitions, concepts, advances, and outlook. Bone Research. 2021;9(1):12
  10. 10. Davis R, Singh A, Jackson MJ, Coelho RT, Prakash D, Charalambous CP, et al. A comprehensive review on metallic implant biomaterials and their subtractive manufacturing. International Journal of Advanced Manufacturing Technology. 2022;120(3-4):1473-1530
  11. 11. Hodde J, Record R, Tullius R, Badylak S. Fibronectin peptides mediate HMEC adhesion to porcine-derived extracellular matrix. Biomaterials. 2002;23(8):1841-1848
  12. 12. Costa A, Naranjo JD, Londono R, Badylak SF. Biologic scaffolds. Cold Spring Harbor Perspectives in Medicine. 2017;7(9):a025676
  13. 13. Yannas IV, Burke JF. Design of an artificial skin. I. Basic design principles. Journal of Biomedical Materials Research. 1980;14(1):65-81
  14. 14. Burke JF, Yannas IV, Quinby WC Jr, Bondoc CC, Jung WK. Successful use of a physiologically acceptable artificial skin in the treatment of extensive burn injury. Annals of Surgery. 1981;194(4):413-428
  15. 15. Ausländer S, Ausländer D, Fussenegger M. Synthetic biology-the synthesis of biology. Angewandte Chemie (International Ed. in English). 2017;56(23):6396-6419
  16. 16. Lim P, Li H, Ng S. Novosorb(R) biodegradable temporising matrix (BTM) and its applications. Surgical Technology International. 2023;42:47-52
  17. 17. MacEwan MR, MacEwan S, Kovacs TR, Batts J. What makes the optimal wound healing material? A review of current science and introduction of a synthetic nanofabricated wound care scaffold. Cureus. 2017;9(10):e1736
  18. 18. Vasios IS, Makiev KG, Georgoulas P, Ververidis A, Drosos G, Tilkeridis K. Use of MatriDerm with Split-thickness skin graft in post-traumatic full-thickness wound defects in orthopedic cases: A case report and systematic review of the literature. Journal of the American Podiatric Medical Association. 2024;114(2):22-009
  19. 19. Knothe Tate ML. Top down and bottom up engineering of bone. Journal of Biomechanics. 2011;44(2):304-312
  20. 20. Badylak SF. The extracellular matrix as a biologic scaffold material. Biomaterials. 2007;28(25):3587-3593
  21. 21. Hodde J. Naturally occurring scaffolds for soft tissue repair and regeneration. Tissue Engineering. 2002;8(2):295-308
  22. 22. Zhu Y, Zhang Q, Wang S, Zhang J, Fan S, Lin X. Current advances in the development of decellularized plant extracellular matrix. Frontiers in Bioengineering and Biotechnology. 2021;9:712262
  23. 23. Li Y, Wu J, Ye P, Cai Y, Shao M, Zhang T, et al. Decellularized extracellular matrix scaffolds: Recent advances and emerging strategies in bone tissue engineering. ACS Biomaterials Science & Engineering. 2024;10(12):7372-7385
  24. 24. Lantz GC, Badylak SF, Hiles MC, Coffey AC, Geddes LA, Kokini K, et al. Small intestinal submucosa as a vascular graft: A review. Journal of Investigative Surgery. 1993;6(3):297-310
  25. 25. Dahms SE, Piechota HJ, Dahiya R, Lue TF, Tanagho EA. Composition and biomechanical properties of the bladder acellular matrix graft: Comparative analysis in rat, pig and human. British Journal of Urology. 1998;82(3):411-419
  26. 26. Bissell MJ, Hall HG, Parry G. How does the extracellular matrix direct gene expression? Journal of Theoretical Biology. 1982;99(1):31-68
  27. 27. Ostadi Y, Khanali J, Tehrani FA, Yazdanpanah G, Bahrami S, Niazi F, et al. Decellularized extracellular matrix scaffolds for soft tissue augmentation: From host-scaffold interactions to bottlenecks in clinical translation. Biomaterials Research. 2024;6(28):0071
  28. 28. Brown BN, Londono R, Tottey S, Zhang L, Kukla KA, Wolf MT, et al. Macrophage phenotype as a predictor of constructive remodeling following the implantation of biologically derived surgical mesh materials. Acta Biomaterialia. 2012;8(3):978-987
  29. 29. Mendibil U, Ruiz-Hernandez R, Retegi-Carrion S, Garcia-Urquia N, Olalde-Graells B, Abarrategi A. Tissue-specific decellularization methods: Rationale and strategies to achieve regenerative compounds. International Journal of Molecular Sciences. 2020;21(15):5447
  30. 30. Londono R, Dziki JL, Haljasmaa E, Turner NJ, Leifer CA, Badylak SF. The effect of cell debris within biologic scaffolds upon the macrophage response. Journal of Biomedical Materials Research. Part A. 2017;105(8):2109-2118
  31. 31. Badylak SF, Valentin JE, Ravindra AK, McCabe GP, Stewart-Akers AM. Macrophage phenotype as a determinant of biologic scaffold remodeling. Tissue Engineering. Part A. 2008;14(11):1835-1842
  32. 32. Dziki JL, Wang DS, Pineda C, Sicari BM, Rausch T, Badylak SF. Solubilized extracellular matrix bioscaffolds derived from diverse source tissues differentially influence macrophage phenotype. Journal of Biomedical Materials Research. Part A. 2017;105(1):138-147
  33. 33. Valentin JE, Stewart-Akers AM, Gilbert TW, Badylak SF. Macrophage participation in the degradation and remodeling of ECM scaffolds. Tissue Engineering. Part A. 2009;15(7):1687-1694
  34. 34. Tao M, Ao T, Mao X, Yan X, Javed R, Hou W, et al. Sterilization and disinfection methods for decellularized matrix materials: Review, consideration and proposal. Bioactive Materials. 2021;6(9):2927-2945
  35. 35. Crapo PM, Gilbert TW, Badylak SF. An overview of tissue and whole organ decellularization processes. Biomaterials. 2011;32(12):3233-3243
  36. 36. Gilbert TW. Strategies for tissue and organ decellularization. Journal of Cellular Biochemistry. 2012;113(7):2217-2222
  37. 37. Hussey GS, Dziki JL, Badylak SF. Extracellular matrix-based materials for regenerative medicine. Nature Reviews Materials. 2018;3(7):159-173
  38. 38. Lun S, Irvine SM, Johnson KD, Fisher NJ, Floden EW, Negron L, et al. A functional extracellular matrix biomaterial derived from ovine forestomach. Biomaterials. 2010;31(16):4517-4529
  39. 39. Karnik T, Dempsey SG, Jerram MJ, Nagarajan A, Rajam R, May BCH, et al. Ionic silver functionalized ovine forestomach matrix - a non-cytotoxic antimicrobial biomaterial for tissue regeneration applications. Biomaterials Research. 2019;23(6):17
  40. 40. Smith MJ, Dempsey SG, Veale RW, Duston-Fursman CG, Rayner CAF, Javanapong C, et al. Further structural characterization of ovine forestomach matrix and multi-layered extracellular matrix composites for soft tissue repair. Journal of Biomaterials Applications. 2021;36(6):996-1010
  41. 41. Sizeland KH, Wells HC, Kelly SJR, Nesdale KE, May BCH, Dempsey SG, et al. Collagen fibril response to strain in scaffolds from ovine forestomach for tissue engineering. ACS Biomaterials Science & Engineering. 2017;3(10):2550-2558
  42. 42. Floden EW, Malak SF, Basil-Jones MM, Negron L, Fisher JN, Lun S, et al. Biophysical characterization of ovine forestomach extracellular matrix biomaterials. Journal of Biomedical Materials Research. Part B, Applied Biomaterials. 2011;96(1):67-75
  43. 43. Irvine SM, Cayzer J, Todd EM, Lun S, Floden EW, Negron L, et al. Quantification of in vitro and in vivo angiogenesis stimulated by ovine forestomach matrix biomaterial. Biomaterials. 2011;32(27):6351-6361
  44. 44. Overbeck N, Beierschmitt A, May BCH, Qi S, Koch J. In-vivo evaluation of a reinforced ovine biologic for plastic and reconstructive procedures in a non-human primate model of soft tissue repair. Eplasty. 2022;22:e43
  45. 45. Overbeck N, Nagvajara GM, Ferzoco S, May BCH, Beierschmitt A, Qi S. In-vivo evaluation of a reinforced ovine biologic: A comparative study to available hernia mesh repair materials. Hernia. 2020;24(6):1293-1306
  46. 46. Dempsey SG, Miller CH, Hill RC, Hansen KC, May BCH. Functional insights from the proteomic inventory of ovine forestomach matrix. Journal of Proteome Research. 2019;18(4):1657-1668
  47. 47. Dempsey SG, Miller CH, Schueler J, Veale RWF, Day DJ, May BCH. A novel chemotactic factor derived from the extracellular matrix protein decorin recruits mesenchymal stromal cells in vitro and in vivo. PLoS One. 2020;15(7):e0235784
  48. 48. Anderson JM, Rodriguez A, Chang DT. Foreign body reaction to biomaterials. Seminars in Immunology. 2008;20(2):86-100
  49. 49. Negron L, Lun S, May BCH. Ovine forestomach matrix biomaterial is a broad spectrum inhibitor of matrix metalloproteinases and neutrophil elastase. International Wound Journal. 2012;11(4):392-397
  50. 50. Street M, Thambyah A, Dray M, Amirapu S, Tuari D, Callon KE, et al. Augmentation with an ovine forestomach matrix scaffold improves histological outcomes of rotator cuff repair in a rat model. Journal of Orthopaedic Surgery and Research. 2015;20(10):165
  51. 51. Dardano AN, Efimenko I, Florio T, Mahedia M, Klapper AM. Rapid revascularization following application of ovine forestomach matrix graft in complex facial and scalp trauma. Trauma Cases and Review. 2024;10(1):105
  52. 52. Veale RWF, Kollmetz T, Taghavi N, Duston-Fursman CG, Beeson MT, Asefi D, et al. Influence of advanced wound matrices on observed vacuum pressure during simulated negative pressure wound therapy. Journal of the Mechanical Behavior of Biomedical Materials. 2022;Feb(138):105620
  53. 53. Merrilees MJ, Falk BA, Zuo N, Dickinson ME, May BCH, Wight TN. Use of versican variant V3 and versican antisense expression to engineer cultured human skin containing increased content of insoluble elastin. Journal of Tissue Engineering and Regenerative Medicine. 2014;11(1):295-305
  54. 54. Bosque BA, Frampton C, Chaffin AE, Bohn GA, Woo K, DeLeonardis C, et al. Retrospective real-world comparative effectiveness of ovine forestomach matrix and collagen/ORC in the treatment of diabetic foot ulcers. International Wound Journal. 2022;19(4):741-753
  55. 55. Raizman R, Hill R, Woo K. Prospective multicenter evaluation of an advanced extracellular matrix for wound management. Advances in Skin & Wound Care. 2020;33(8):437-444
  56. 56. Al Mousa RH, Bosque BA, Dowling SG. Use of ovine forestomach matrix in the treatment of facial thermal burns. Wounds. 2022;34(4):e17-e21
  57. 57. Taarea R, Florence A, Bendixen B, Castater CA. Early experience with ovine forestomach matrix for the reconstruction of abdominal defects following emergency open abdomen surgery at a level 2 trauma center. Trauma Cases and Review. 2024;10(1):102
  58. 58. Cormican MT, Creel NJ, Bosque BA, Dowling SG, Rideout PP, Vassy WM. Ovine forestomach matrix in the surgical management of complex volumetric soft tissue defects: A retrospective pilot case series. Eplasty. 2023;23:e66
  59. 59. Sivaraj D, Henn D, Fischer KS, Kim TS, Black CK, Lin JQ, et al. Reinforced biologic mesh reduces postoperative complications compared to biologic mesh after ventral hernia repair. Plastic and Reconstructive Surgery. Global Open. 2022;10(2):e4083
  60. 60. DeNoto G 3rd, Ceppa EP, Pacella SJ, Sawyer M, Slayden G, Takata M, et al. 24-month results of the BRAVO study: A prospective, multi-center study evaluating the clinical outcomes of a ventral hernia cohort treated with OviTex(R) 1S permanent reinforced tissue matrix. Annals of Medicine and Surgery (Lond). 2022;83:104745
  61. 61. Sweitzer K, Arias-Camison R, Cafro C, Langstein H. The use of reinforced ovine mesh in implant breast reconstruction: Equivalent outcomes to human acellular dermal matrices and more cost-effective. Annals of Plastic Surgery. 2024;93(6):664-667
  62. 62. Chaffin AE, Buckley MC. Extracellular matrix graft for the surgical management of Hurley stage III hidradenitis suppurativa: A pilot case series. Journal of Wound Care. 2020;29(11):624-630
  63. 63. Chaffin AE, Dowling SG, Kosyk MS, Bosque BA. Surgical reconstruction of pilonidal sinus disease with concomitant extracellular matrix graft placement: A case series. Journal of Wound Care. 2021;30(Sup7):S28-S34
  64. 64. Hsu A, Schlidt K, D'Adamo CR, Bosque BA, Dowling SG, Wolf JH. Surgical management of perianal fistula using an ovine forestomach matrix implant. Techniques in Coloproctology. 2023;27(9):769-774
  65. 65. Chan X, Tan CHM, Chong CXZ, Sivarajah SS, Koh FH. Technical approach in the management of perianal fistula: Combining ovine extracellular matrix with endoanal ultrasound to review the surgical outcome. Annals of Coloproctology. 2025;41(1):93-96
  66. 66. House JK, Smith BP, Fecteau G, VanMetre DC. Assessment of the ruminant digestive system. The Veterinary Clinics of North America. Food Animal Practice. 1992;8(2):189-202
  67. 67. Singleton AG. Ruminant digestion. Nature. 1953;172(4384):834-836
  68. 68. Garcia A, Masot J, Franco A, Gazquez A, Redondo E. Histomorphometric and immunohistochemical study of the goat rumen during prenatal development. The Anatomical Record (Hoboken). 2012;295(5):776-785
  69. 69. Harfoot CG. Anatomy, physiology and microbiology of the ruminant digestive tract. Progress in Lipid Research. 1978;17(1):1-19
  70. 70. Wardrop ID, Coombe JB. The post-natal growth of the visceral organs of the lamb I. The growth of the visceral organs of the grazing lamb from birth to sixteen weeks of age. Journal of Agricultural and Food Chemistry. 1960;54(1):140-143
  71. 71. Hofmann RR. Evolutionary steps of ecophysiological adaptation and diversification of ruminants: A comparative view of their digestive system. Oecologia. 1989;78(4):443-457
  72. 72. Kitamura N, Yoshiki A, Sasaki M, Baltazar ET, Hondo E, Yamamoto Y, et al. Immunohistochemical evaluation of the muscularis mucosae in the ruminant forestomach. Anatomia, Histologia, Embryologia. 2003;32(3):175-178
  73. 73. Banks WJ. Applied Veterinary Histology. 3rd ed. Vol. xiii. St. Louis: Mosby-Year Book; 1993, 526 p. p
  74. 74. Zitnan R, Voigt J, Wegner J, Breves G, Schroder B, Winckler C, et al. Morphological and functional development of the rumen in the calf: Influence of the time of weaning. 1. Morphological development of rumen mucosa. Archiv für Tierernährung. 1999;52(4):351-362
  75. 75. Prache S, Schreurs N, Guillier L. Review: Factors affecting sheep carcass and meat quality attributes. Animal. 2022;16(Suppl. 1):100330
  76. 76. Malmuthuge N, Liang G, Guan LL. Regulation of rumen development in neonatal ruminants through microbial metagenomes and host transcriptomes. Genome Biology. 2019;20(1):172
  77. 77. Lane MA, RLT B, Jesse BW. Sheep rumen metabolic development in response to age and dietary treatments. Journal of Animal Science. 2000;78(7):1990-1996
  78. 78. Shephard RW, Maloney SK. A review of thermal stress in cattle. Australian Veterinary Journal. 2023;101(11):417-429
  79. 79. Palomino-Durand C, Pauthe E, Gand A. Fibronectin-enriched biomaterials, biofunctionalization, and proactivity: A review. Applied Sciences. 2021;11(24):12111
  80. 80. Kim HN, Elgundi Z, Lin X, Fu L, Tang F, Moh ESX, et al. Engineered short forms of perlecan enhance angiogenesis by potentiating growth factor signalling. Journal of Controlled Release. 2023;362:184-196
  81. 81. Cholewinska P, Czyz K, Nowakowski P, Wyrostek A. The microbiome of the digestive system of ruminants - a review. Animal Health Research Reviews. 2020;21(1):3-14
  82. 82. Taschuk R, Griebel PJ. Commensal microbiome effects on mucosal immune system development in the ruminant gastrointestinal tract. Animal Health Research Reviews. 2012;13(1):129-141
  83. 83. Chase CCL. Enteric immunity: Happy gut, healthy animal. The Veterinary Clinics of North America. Food Animal Practice. 2018;34(1):1-18
  84. 84. Chassaing B, Darfeuille-Michaud A. The commensal microbiota and enteropathogens in the pathogenesis of inflammatory bowel diseases. Gastroenterology. 2011;140(6):1720-1728
  85. 85. Mummert ME. Immunologic roles of hyaluronan. Immunologic Research. 2005;31(3):189-206
  86. 86. Frey H, Schroeder N, Manon-Jensen T, Iozzo RV, Schaefer L. Biological interplay between proteoglycans and their innate immune receptors in inflammation. The FEBS Journal. 2013;280(10):2165-2179
  87. 87. Simcock J, May BC. Ovine forestomach matrix as a substrate for single-stage split-thickness graft reconstruction. Eplasty. 2013;13:e58
  88. 88. Lake SP, Deeken CR, Agarwal AK. Reinforced tissue matrix to strengthen the abdominal wall following reversal of temporary ostomies or to treat incisional hernias. World Journal of Gastrointestinal Surgery. 2024;16(3):823-832
  89. 89. Sicari BM, Johnson SA, Siu BF, Crapo PM, Daly KA, Jiang H, et al. The effect of source animal age upon the in vivo remodeling characteristics of an extracellular matrix scaffold. Biomaterials. 2012;33(22):5524-5533
  90. 90. Guilloteau P, Zabielski R, Blum JW. Gastrointestinal tract and digestion in the young ruminant: Ontogenesis, adaptations, consequences and manipulations. Journal of Physiology and Pharmacology. 2009;60(Suppl. 3):37-46
  91. 91. Steele MA, Penner GB, Chaucheyras-Durand F, Guan LL. Development and physiology of the rumen and the lower gut: Targets for improving gut health. Journal of Dairy Science. 2016;99(6):4955-4966
  92. 92. Warner RG, Flatt WP, Loosli JK. Ruminant nutrition, dietary factors influencing development of ruminant stomach. Journal of Agricultural and Food Chemistry. 1956;4(9):788-792
  93. 93. Pokhrel B, Jiang H. Postnatal growth and development of the rumen: Integrating physiological and molecular insights. Biology (Basel). 2024;13(4):269
  94. 94. Yuan Y, Sun DM, Qin T, Mao SY, Zhu WY, Yin YY, et al. Single-cell transcriptomic landscape of the sheep rumen provides insights into physiological programming development and adaptation of digestive strategies. Zoological Research. 2022;43(4):634-647
  95. 95. Scala G, Corona M, Maruccio L. Structural, histochemical and immunocytochemical study of the forestomach mucosa in domestic ruminants. Anatomia, Histologia, Embryologia. 2011;40(1):47-54
  96. 96. Baldwin RL, Connor EE. Rumen function and development. The Veterinary Clinics of North America. Food Animal Practice. 2017;33(3):427-439
  97. 97. Yohe TT, Tucker HLM, Parsons CLM, Geiger AJ, Akers RM, Daniels KM. Short communication: Initial evidence supporting existence of potential rumen epidermal stem and progenitor cells. Journal of Dairy Science. 2016;99(9):7654-7660
  98. 98. Franco AJ, Masot AJ, Aguado MC, Gomez L, Redondo E. Morphometric and immunohistochemical study of the rumen of red deer during prenatal development. Journal of Anatomy. 2004;204(6):501-513
  99. 99. Xiang R, Oddy VH, Archibald AL, Vercoe PE, Dalrymple BP. Epithelial, metabolic and innate immunity transcriptomic signatures differentiating the rumen from other sheep and mammalian gastrointestinal tract tissues. PeerJ. 2016;4:e1762
  100. 100. Goodlad RA. Some effects of diet on the mitotioc index and the cell cycle of the ruminal epithelium of sheep. Quarterly Journal of Experimental Physiology. 1981;66(4):487-499
  101. 101. Hooper CE. Cell turnover in epithelial populations. The Journal of Histochemistry and Cytochemistry. 1956;4(6):531-540
  102. 102. Matsubayashi Y. Dynamic movement and turnover of extracellular matrices during tissue development and maintenance. Fly (Austin). 2022;16(1):248-274
  103. 103. Arnott G, Ferris CP, O'Connell NE. Review: Welfare of dairy cows in continuously housed and pasture-based production systems. Animal. 2017;11(2):261-273
  104. 104. Eurosurveillance Editorial. WHO member states adopt global action plan on antimicrobial resistance. Euro Surveillance. 2015;20(21)
  105. 105. Hillerton JE, Irvine CR, Bryan MA, Scott D, Merchant SC. Use of antimicrobials for animals in New Zealand, and in comparison with other countries. New Zealand Veterinary Journal. 2017;65(2):71-77
  106. 106. Hillerton JE, Bryan MA, Scott D. Consumption of antimicrobials for use in food-producing animals in New Zealand, a measure of progress in reduction from 2015 to 2022. New Zealand Veterinary Journal. 2024;18(Dec):1-8
  107. 107. Roy BC, Sedgewick G, Aalhus JL, Basarab JA, Bruce HL. Modification of mature non-reducible collagen cross-link concentrations in bovine m. gluteus medius and semitendinosus with steer age at slaughter, breed cross and growth promotants. Meat Science. 2015;Dec(110):109-117
  108. 108. Packer DT, Geesink GH, Thompson JM, Polkinghorne RJ, Ball AB, McGilchrist P. The impact of different hormonal growth promotants (HGP) on desmin degradation and collagen content of various muscles from pasture and feedlot finished steer carcasses. Meat Science. 2021;182:108615
  109. 109. Lopez-Baca MA, Contreras M, Gonzalez-Rios H, Macias-Cruz U, Torrentera N, Valenzuela-Melendres M, et al. Growth, carcass characteristics, cut yields and meat quality of lambs finished with zilpaterol hydrochloride and steroid implant. Meat Science. 2019;158:107890
  110. 110. Watson R. EU says growth hormones pose health risk. BMJ. 1999;318(7196):1442
  111. 111. Caradus JR, Goldson SL, Moot DJ, Rowarth JS, Stewart AV. Pastoral agriculture, a significant driver of New Zealand's economy, based on an introduced grassland ecology and technological advances. Journal of the Royal Society of New Zealand. 2023;53(3):259-303
  112. 112. McDowell RW, Herzig A, van der Weerden TJ, Cleghorn C, Kaye-Blake W. Growing for good: Producing a healthy, low greenhouse gas and water quality footprint diet in Aotearoa, New Zealand. Journal of the Royal Society of New Zealand. 2024;54(3):325-349
  113. 113. BLNZ. The carbon footprint of New Zealand sheepmeat and beef 2022. Available from: https://beeflambnz.com/knowledge-hub/PDF/factsheet-and-faqs-carbon-footprint-new-zealand-sheepmeat-and-beef.pdf
  114. 114. Moot DJ, Davison R. Changes in New Zealand red meat production over the past 30 yr. Animal Frontiers. 2021;11(4):26-31
  115. 115. Durant JM, Aarvold L, Langangen O. Stock collapse and its effect on species interactions: Cod and herring in the Norwegian-Barents seas system as an example. Ecology and Evolution. 2021;11(23):16993-17004
  116. 116. Gyles C. Industrial farm animal production. The Canadian Veterinary Journal. 2010;51(2):125-128
  117. 117. Simonds RJ, Holmberg SD, Hurwitz RL, Coleman TR, Bottenfield S, Conley LJ, et al. Transmission of human immunodeficiency virus type 1 from a seronegative organ and tissue donor. The New England Journal of Medicine. 1992;326(11):726-732
  118. 118. Bonda DJ, Manjila S, Mehndiratta P, Khan F, Miller BR, Onwuzulike K, et al. Human prion diseases: Surgical lessons learned from iatrogenic prion transmission. Neurosurgical Focus. 2016;41(1):E10
  119. 119. Greenwald MA, Kuehnert MJ, Fishman JA. Infectious disease transmission during organ and tissue transplantation. Emerging Infectious Diseases. 2012;18(8):e1
  120. 120. Greenwald MA, Kerby S, Francis K, Noller AC, Gormley WT, Biswas R, et al. Detection of human immunodeficiency virus, hepatitis C virus, and hepatitis B virus in postmortem blood specimens using infectious disease assays licensed for cadaveric donor screening. Transplant Infectious Disease. 2018;20(1):e12825
  121. 121. Moore MA, Samsell B, McLean J. Allograft Tissue Safety and Technology. Biologics in Orthopaedic Surgery. 2019:49-62. DOI: 10.1016/B978-0-323-55140-3.00005-9
  122. 122. Noordergraaf J, Schucker A, Martin M, Schuurman HJ, Ordway B, Cooley K, et al. Pathogen elimination and prevention within a regulated, designated pathogen free, closed pig herd for long-term breeding and production of xenotransplantation materials. Xenotransplantation. 2018;25(4):e12428
  123. 123. Worthington B, Owen K. Import Risk Analysis: Sheep and Goat Genetic Material. New Zealand: Ministry of Agriculture and Forestry (MAF) New Zealand; 2005
  124. 124. Buttner M, Rziha HJ. Parapoxviruses: From the lesion to the viral genome. Journal of Veterinary Medicine. B, Infectious Diseases and Veterinary Public Health. 2002;49(1):7-16
  125. 125. Pan KM, Baldwin M, Nguyen J, Gasset M, Serban A, Groth D, et al. Conversion of alpha-helices into beta-sheets features in the formation of the scrapie prion proteins. Proceedings of the National Academy of Sciences of the United States of America. 1993;90(23):10962-10966
  126. 126. Cohen FE, Prusiner SB. Pathologic conformations of prion proteins. Annual Review of Biochemistry. 1998;67:793-819
  127. 127. Will RG. Epidemiology of Creutzfeldt-Jakob disease. British Medical Bulletin. 1993;49(4):960-970
  128. 128. Turner ML, Ironside JW. New-variant Creutzfeldt-Jakob disease: The risk of transmission by blood transfusion. Blood Reviews. 1998;12(4):255-268
  129. 129. Vadrot C, Darbord JC. Quantitative evaluation of prion inactivation comparing steam sterilization and chemical sterilants: Proposed method for test standardization. The Journal of Hospital Infection. 2006;64(2):143-148
  130. 130. Race RE, Raymond GJ. Inactivation of transmissible spongiform encephalopathy (prion) agents by environ LpH. Journal of Virology. 2004;78(4):2164-2165
  131. 131. Peretz D, Supattapone S, Giles K, Vergara J, Freyman Y, Lessard P, et al. Inactivation of prions by acidic sodium dodecyl sulfate. Journal of Virology. 2006;80(1):322-331
  132. 132. Manuelidis L. Decontamination of Creutzfeldt-Jakob disease and other transmissible agents. Journal of Neurovirology. 1997;3(1):62-65
  133. 133. Fichet G, Antloga K, Comoy E, Deslys JP, McDonnell G. Prion inactivation using a new gaseous hydrogen peroxide sterilisation process. The Journal of Hospital Infection. 2007;67(3):278-286
  134. 134. Antloga K, Meszaros J, Malchesky PS, McDonnell GE. Prion disease and medical devices. ASAIO Journal. 2000;46(6):S69-S72
  135. 135. Prusiner SB. Prions. Proceedings of the National Academy of Sciences of the United States of America. 1998;95(23):13363-13383
  136. 136. Hewitt PE, Llewelyn CA, Mackenzie J, Will RG. Creutzfeldt-Jakob disease and blood transfusion: Results of the UK transfusion medicine epidemiological review study. Vox Sanguinis. 2006;91(3):221-230
  137. 137. Gajdusek DC, Gibbs CJ Jr. Slow, latent and temperate virus infections of the central nervous system. Research Publications - Association for Research in Nervous and Mental Disease. 1968;44:254-280
  138. 138. Scott MR, Peretz D, Nguyen HO, Dearmond SJ, Prusiner SB. Transmission barriers for bovine, ovine, and human prions in transgenic mice. Journal of Virology. 2005;79(9):5259-5271
  139. 139. Scott MR, Will R, Ironside J, Nguyen HO, Tremblay P, DeArmond SJ, et al. Compelling transgenetic evidence for transmission of bovine spongiform encephalopathy prions to humans. Proceedings of the National Academy of Sciences of the United States of America. 1999;96(26):15137-15142
  140. 140. Raymond GJ, Hope J, Kocisko DA, Priola SA, Raymond LD, Bossers A, et al. Molecular assessment of the potential transmissibilities of BSE and scrapie to humans. Nature. 1997;388(6639):285-288
  141. 141. Conmer T, Morborne H. A Letter to Dr Hunter, physican in York, concerning rickets in sheep. London1772
  142. 142. Will RG. Acquired prion disease: Iatrogenic CJD, variant CJD, kuru. British Medical Bulletin. 2003;66:255-265
  143. 143. MacDiarmid SC. Scrapie: The risk of its introduction and effects on trade. Australian Veterinary Journal. 1996;73(5):161-164
  144. 144. Hunter N, Cairns D. Scrapie-free merino and poll Dorset sheep from Australia and New Zealand have normal frequencies of scrapie-susceptible PrP genotypes. The Journal of General Virology. 1998;79(Pt 8):2079-2082
  145. 145. Davidson RM. Control and eradication of animal diseases in New Zealand. New Zealand Veterinary Journal. 2002;50(Suppl. 3):6-12
  146. 146. Bossers A, Harders FL, Smits MA. PrP genotype frequencies of the most dominant sheep breed in a country free from scrapie. Archives of Virology. 1999;144(4):829-834
  147. 147. Imran M, Mahmood S. An overview of animal prion diseases. Virology Journal. 2011;1(8):493
  148. 148. Enoch S, Shaaban H, Dunn KW. Informed consent should be obtained from patients to use products (skin substitutes) and dressings containing biological material. Journal of Medical Ethics. 2005;31(1):2-6
  149. 149. Eliasi JR, Dwyer JT. Kosher and halal: Religious observances affecting dietary intakes. Journal of the American Dietetic Association. 2002;102(7):911-913
  150. 150. Easterbrook C, Maddern G. Porcine and bovine surgical products: Jewish, Muslim, and Hindu perspectives. Archives of Surgery. 2008;143(4):366-370
  151. 151. Hagelin J, Hau J, Schapiro SJ, Suleman MA, Carlsson HE. Religious beliefs and opinions on clinical xenotransplantation--a survey of university students from Kenya, Sweden and Texas. Clinical Transplantation. 2001;15(6):421-425
  152. 152. Ritchie EMaH. What share of people say they are vegetarian, vegan, or flexitarian? OurWorldinData.org. 2022. Available from: https://ourworldindata.org/vegetarian-vegan
  153. 153. Louis-Maerten E, Rodriguez Perez C, Cajiga RM, Persson K, Elger BS. Conceptual foundations for a clarified meaning of the 3Rs principles in animal experimentation. Animal Welfare. 2024;33:e37
  154. 154. Eriksson A, Burcharth J, Rosenberg J. Animal derived products may conflict with religious patients' beliefs. BMC Medical Ethics. 2013;14:48
  155. 155. GVR. Scaffold Technology Market Size, Share & Trends Analysis Report By Type (Hydrogels, Polymeric Scaffolds), By Disease Type (Cancer, Neurology), By Application, By End-use, By Region, And Segment Forecasts, 2023 – 2030. 2025. Available from: https://www.grandviewresearch.com/industry-analysis/scaffold-technology-market
  156. 156. Lawlor JC, Bosque BA, Frampton C, Young DA, Martyka P. Limb salvage via surgical soft-tissue reconstruction with ovine forestomach matrix grafts: A prospective study. PRS Global Open. 2024;12(12):e6406
  157. 157. Aburn R, Chaffin AE, Bosque BA, Frampton C, Dempsey SG, Young DA, et al. Clinical efficacy of ovine forestomach matrix and collagen/oxidised regenerated cellulose for the treatment of venous leg ulcers: A retrospective comparative real-world evidence study. International Wound Journal. 2025;22(4):e70368
  158. 158. Zhu X, Olsson MM, Bajpai R, Jarbrink K, Tang WE, Car J. Health-related quality of life and chronic wound characteristics among patients with chronic wounds treated in primary care: A cross-sectional study in Singapore. International Wound Journal. 2022;19(5):1121-1132
  159. 159. Monahan M, Jowett S, Pinkney T, Brocklehurst P, Morton DG, Abdali Z, et al. Surgical site infection and costs in low- and middle-income countries: A systematic review of the economic burden. PLoS One. 2020;15(6):e0232960
  160. 160. Shakir S, Messa CA, Broach RB, Rhemtulla IA, Chatman B, D’Angelantonio A, et al. Indications and limitations of bilayer wound matrix-based lower extremity reconstruction: A multidisciplinary case-control study of 191 wounds. Plastic and Reconstructive Surgery. 2020;145(3):813-822
  161. 161. Branski LK, Herndon DN, Pereira C, Mlcak RP, Celis MM, Lee JO, et al. Longitudinal assessment of Integra in primary burn management: A randomized pediatric clinical trial. Critical Care Medicine. 2007;35(11):2615-2623
  162. 162. Mueller CK, Bader RD, Ewald C, Kalff R, Schultze-Mosgau S. Scalp defect repair: A comparative analysis of different surgical techniques. Annals of Plastic Surgery. 2012;68(6):594-598
  163. 163. Zhu B, Cao D, Xie J, Li H, Chen Z, Bao Q. Clinical experience of the use of Integra in combination with negative pressure wound therapy: An alternative method for the management of wounds with exposed bone or tendon. Journal of Plastic Surgery and Hand Surgery. 2021;55(1):1-5
  164. 164. Garwood CS, Kim PJ, Matai V, Steinberg JS, Evans KK, Mitnick CD, et al. The use of bovine collagen-glycosaminoglycan matrix for atypical lower extremity ulcers. Wounds. 2016;28(9):298-305
  165. 165. Austin CL, Draper B, Larson KW, Thompson SJ. Biodegradable temporising matrix: Use of negative pressure wound therapy shows a significantly higher success rate. Journal of Wound Care. 2023;32(3):159-166
  166. 166. Wagstaff MJ, Schmitt BJ, Caplash Y, Greenwood JE. Free flap donor site reconstruction: A prospective case series using an optimized polyurethane biodegradable temporizing matrix. Eplasty. 2015;15:e27
  167. 167. Lo CH, Brown JN, Dantzer EJG, Maitz PKM, Vandervord JG, Wagstaff MJD, et al. Wound healing and dermal regeneration in severe burn patients treated with NovoSorb(R) biodegradable temporising matrix: A prospective clinical study. Burns. 2022;48(3):529-538
  168. 168. Heimbach DM, Warden GD, Luterman A, Jordan MH, Ozobia N, Ryan CM, et al. Multicenter postapproval clinical trial of Integra® dermal regeneration template for burn treatment. The Journal of Burn Care & Rehabilitation. 2003;24(1):42-48
  169. 169. Greenhalgh DG, Hinchcliff K, Sen S, Palmieri TL. A ten-year experience with pediatric face grafts. Journal of Burn Care & Research. 2013;34(5):576-584
  170. 170. Hicks CW, Zhang GQ, Canner JK, Mathioudakis N, Coon D, Sherman RL, et al. Outcomes and predictors of wound healing among patients with complex diabetic foot wounds treated with a dermal regeneration template (Integra). Plastic and Reconstructive Surgery. 2020;146(4):893-902
  171. 171. Kidd T, Kolaityte V, Bajaj K, Wallace D, Izadi D, Bechar J. The use of NovoSorb biodegradable temporising matrix in wound management: A literature review and case series. Journal of Wound Care. 2023;32(8):470-478
  172. 172. Dorweiler B, Trinh TT, Dunschede F, Vahl CF, Debus ES, Storck M, et al. The marine Omega3 wound matrix for treatment of complicated wounds: A multicenter experience report. Gefasschirurgie. 2018;23(Suppl. 2):46-55
  173. 173. Ankney C, Banaschak C, Sowers B, Szotek P. Minimizing retained foreign body in hernia repair using a novel technique: Reinforced biologic augmented repair (ReBAR). Journal of Clinical Medical Research. 2021;3(4):1-11
  174. 174. Sawyer MAJ. New ovine polymer-reinforced bioscaffold in hiatal hernia repair. JSLS. 2018;22(4):e2018.00057
  175. 175. Parker MJ, Kim RC, Barrio M, Socas J, Reed LR, Nakeeb A, et al. A novel biosynthetic scaffold mesh reinforcement affords the lowest hernia recurrence in the highest-risk patients. Surgical Endoscopy. 2020;35(9):5173-5178
  176. 176. DeNoto G, Ceppa EP, Pacella SJ, Sawyer M, Slayden G, Takata M, et al. A prospective, single arm, multi-center study evaluating the clinical outcomes of ventral hernias treated with OviTex® 1S permanent reinforced tissue matrix: The BRAVO study 12-month analysis. Journal of Clinical Medicine. 2021;10(21):4998
  177. 177. Sivaraj D, Fischer KS, Kim TS, Chen K, Tigchelaar SS, Trotsyuk AA, et al. Outcomes of biosynthetic and synthetic mesh in ventral hernia repair. Plastic and Reconstructive Surgery. Global Open. 2022;10(12):e4707
  178. 178. DeNoto G. Bridged repair of large ventral hernia defects using an ovine reinforced biologic: A case series. Annals of Medicine and Surgery. 2022;75:103446
  179. 179. Goetz M, Jurczyk M, Junger H, Schlitt HJ, Brunner SM, Brennfleck FW. Semiresorbable biologic hybrid meshes for ventral abdominal hernia repair in potentially contaminated settings: Lower risk of recurrence. Updates in Surgery. 2022;74(6):1995-2001

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Sandi G. Dempsey, D. Adam Young, Robert W.F. Veale and Barnaby C.H. May

Submitted: 05 March 2025 Reviewed: 12 May 2025 Published: 30 July 2025

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