Abstract
A variety of surgical and non-surgical approaches have been used to address the impacts of nervous system injuries, which can lead to either impairment or a complete loss of function for affected patients. The inherent ability of nervous tissues to repair and/or regenerate is dampened due to irreversible changes that occur within the tissue remodeling microenvironment following injury. Specifically, dysregulation of the extracellular matrix (i.e., scarring) has been suggested as one of the major factors that can directly impair normal cell function and could significantly alter the regenerative potential of these tissues. A number of tissue engineering and regenerative medicine-based approaches have been suggested to intervene in the process of remodeling which occurs following injury. Decellularization has become an increasingly popular technique used to obtain acellular scaffolds, and their derivatives (hydrogels, etc.), which retain tissue-specific components, including critical structural and functional proteins. These advantageous characteristics make this approach an intriguing option for creating materials capable of stimulating the sensitive repair mechanisms associated with nervous system injuries. Over the past decade, several diverse decellularization methods have been implemented specifically for nervous system applications in an attempt to carefully remove cellular content while preserving tissue morphology and composition. Each application-based decellularized ECM product requires carefully designed treatments that preserve the unique biochemical signatures associated within each tissue type to stimulate the repair of brain, spinal cord, and peripheral nerve tissues. Herein, we review the decellularization techniques that have been applied to create biomaterials with the potential to promote the repair and regeneration of tissues within the central and peripheral nervous system.
Keywords: Decellularization, Nervous system, Acellular scaffolds, Hydrogels, TBI, SCI, PNI
1. Introduction
1.1. Clinical and economic burden of CNS/PNS injury
The nervous system is divided into the central (CNS) and peripheral (PNS) regions, which work in tandem to efficiently control homeostatic function in response to internal and external stimuli. More specifically, the CNS is composed of the brain and spinal cord, which initiate a biochemical or physical response based upon the information received from the expansive network of nerves that make up the PNS. A disruption in this intricate network can result in moderate to severe mental or physical impairment that can lead to a significant clinical and economic burden.
Traumatic injury represents a vast majority of non-recoverable CNS injuries at nearly 80% with degenerative diseases and stroke/ischemia contributing to the remaining 20% [1]. Spinal cord injuries (SCIs) are primarily caused by vehicle and falling accidents representing nearly 70% of all cases of SCI in the United States [2]. In 2016, the number of total affected patients suffering from traumatic brain (TBI) and spinal cord injuries (SCI) reached greater than 82 million people globally [3].
Peripheral nerve injury is also commonplace, with peripheral nerve injuries affecting more than 20 million Americans with a high prevalence among younger individuals [4-6]. These injuries have an economic impact of more than $18 billion per year. The impact of peripheral nerve injury depends largely upon the type and severity of injury as well as the location of injury [7,8]. Each year in the United States, more than 600,000 surgical procedures are performed to address nerve injuries in the upper extremities alone [9]. Despite significant advances in surgical technique, and as will be discussed in greater detail below, results and the prospects for full functional recovery are generally considered poor with fewer than 50% of patients reporting satisfactory motor function following surgical reconstruction [7,10].
1.2. CNS injury
Patients that experience traumatic injury to the brain or spinal cord undergo some degree of neuronal dystrophy; however, this type of prognosis, in the most severe cases can be fatal. Survivors of these injuries are typically exposed to a variety of complications ranging from loss of sensation to complete paralysis. Depending on the severity of the injury or disease state, the quality of life is poor due to a lack of effective treatments capable of reversing this pathology.
An injury to central nervous tissues initiates a cascade of events at both the cellular and molecular level [11]. These changes significantly alter the normal environmental conditions and ultimately leaves the tissues in an impermissible state for axon regeneration, inhibiting full functional recovery [12]. It is currently understood that the events post-injury occur in two distinct states: a primary and secondary lesion [1]. The primary lesion can be classified as the direct result of traumatic impact, and the tissue response varies dramatically based upon the type of incident that inflicts damage. Initial signs of primary damage can include cerebral edema and contusions, disruption of nerve fibers, hematomas and neuronal cell death [13]. In the secondary lesion, astrocytic reactivity occurs, which initiates morphological changes as well as a rapid migration of astrocytes to the perimeter of the lesion site, where cyst and scar formation prevents axon sprouting [14]. As a safeguard mechanism for uninjured neurons, reactive astrocytes cordon off the secondary lesion, subsequently resolving the inflammatory response but preventing axonal regrowth within the area [15,16]. Astrocytic phenotypes are known to modulate along a spectrum in accordance with the severity of the injury and can directly determine the extent of neuronal functional loss [17,18].
Previous attempts at treating the crippling effects of these injuries have been largely unsuccessful to date, as the variables causing the regenerative inhibition are multifactorial. The glial scarring, present post-injury, effectively eliminates the potential for axonal regrowth in the CNS tissues. This newly formed physical barrier accompanied by its abnormal molecular composition poses a significant challenge for researchers looking for ways to expose its vulnerabilities and promote functional recovery.
1.3. PNS injury
Peripheral nervous system injuries (PNI) are the result of disruption in the intricate feedback network, spanning from the peripheral organs to the CNS. Patients suffering from a traumatic PNI can experience either a decrease or a complete loss of motor and sensory function. Mild to moderate cases of PNI are often treatable, as peripheral nerves offer a significantly greater regenerative potential than that of their CNS counterpart; however, the prognosis for PNI recovery is directly dependent upon the extent of damage that is incurred by the nerves. In order to simplify the severity of these injuries, Sunderland et al. delineates between them using five distinct degree categories, summarized in Fig. 1.
Fig. 1.
Sunderland’s Five Degrees of Nerve Trauma.
The superior regenerative potential of the peripheral nervous system is in large part thanks to the supportive environment generated after nerve injury [19]. Neuronal cell bodies of the PNS within the CNS and dorsal root ganglion (DRG) possess the ability to shift from a transmitting state to a pro-repair phenotype to regrow after injury. In contrast, where the environment inhibits recovery after CNS injury, the PNS has complex mechanisms that support regeneration at the site of injury [12,20-23]. Axonotmesis and neurotmesis injuries to the PNS elicit a response from multiple cell types both at the site of injury and distally to support regenerating axons towards their original targets [24,25]. Denervated axons distal to the injury are degraded by local Schwann cells (SCs) followed by the infiltration of macrophages in a process known as Wallerian degeneration [26]. This process begins within two days of the injury, lasting approximately two weeks [27]; however, the macrophage population does not peak until 2–3 weeks post-injury. The presence of macrophages plays an additional role of supporting SCs, fibroblasts and promoting angiogenesis through the release of cytokines and mitogenic signals [24,28-30]. Triggered by inflammatory macrophages during the first few days after injury, SCs de-differentiate and take on a repair-supportive phenotype [19,22,28]. Beyond phagocytosing myelin and axonal debris, these de-differentiated SCs proliferate and secrete neurotrophic factors, including NGF and BDNF to support axon regeneration [19,28,31]. Ultimately, the dedifferentiated SCs migrate along new, axially aligned vasculature promoted by macrophages guiding axons towards their distal targets [24,29].
Despite these mechanisms for PNS regeneration, actual patient outcomes after peripheral nerve injury remain varied, with factors such as severity and location of the injury impacting functional outcomes. Mild axonotmesis injuries recover better than more severe axonotmesis and neurotmesis injuries [23]. These injuries are subject to misalignment of motor and sensory nerves as they regrow randomly towards distal targets [32,33]. Large distances from injury to the end organ also plays a significant role in poor outcomes [23,34]. The rate of axon growth is limited to around 1 mm/day [7,32]. However, phenotypic changes favoring repair in neurons and SCs are not static and decline with prolonged axotomization and denervation [22,34]. Functional recovery is rare if reinnervation is not established within a window of 12–18 months, mainly due to atrophy of the target muscle and neuromuscular junctions [7].
1.4. Considerations for restoring function in CNS/PNS injury
Developing successful therapies requires a deep understanding of the tissue environments they will co-inhabit. Therefore, it is important to identify the primary mediators responsible for the onset and persistence of disease to employ directed strategies that will promote functional recovery. In the nervous system, there are several factors that inhibit normal tissue repair, such as glial cells, ECM molecules, and macrophages. However, these elements can also contribute to the remodeling of the injury site under the appropriate conditions. In order to understand their impact, we have included a brief overview of some of the main components involved in nervous system injuries and how decellularized ECM (dECM) materials may provide a therapeutic solution for reversing the resulting inhibitory microenvironment.
1.4.1. Role of glial cells as regulators of tissue repair
The regenerative capacity of the central and peripheral nervous system is inherently different in response to injury. This could be attributed to varying functional characteristics of the supporting cells, or neuroglia, that co-inhabit these tissues. The neuroglia residing in the CNS are astrocytes, oligodendrocytes, microglia, and ependymal cells, while the neuroglia of the PNS are composed of satellite cells and SCs. Each of these cells are vitally important; however, in the case of repair and regenerative potential of the nervous tissues, astrocytes and SCs appear to play the most pivotal role in CNS and PNS respectively.
Unique to the CNS, astrocytes are the primary glial cells that are responsible for responding to damage [16,35]. Most pathologies resulting in functional loss in the neurons of the CNS directly result from a dysfunction of the damage control pathways of astrocytes. Prior to injury, astrocytes are critical for maintaining homeostasis [36]. Specifically, they are responsible for providing mechanical support [37], regulating water and ion transport [38], supporting synaptic function [39], aiding detoxification [40], glucose storage and metabolic activity [41]. In response to injury, astrocytes undergo dynamic phenotypic transformation, increased proliferation and upregulated expression of glial fibrillary acidic protein (GFAP). These cells also play a role in the secretion of pro-inflammatory cytokines and other inhibitory ECM molecules, which together allow for glial scarring to persist and reduce the potential for regeneration. Although these cells are primarily seen as negative regulators of repair, they can also be activated toward a constructive remodeling phenotype [42,43]. Astrocytes can alter their morphology and trigger an increase in antioxidants and soluble growth factors, which ultimately provides more favorable conditions for CNS tissue repair.
In the PNS, the SCs reside in areas surrounding axons and are active contributors to the myelination of axons, enhance structural support, and are key regulators of metabolic and synaptic function [44]. In response to traumatic PNI, resilient SCs retain viability during axon degeneration and promote the release of essential neurotrophic factors to initiate nerve regeneration. Like astrocytes, the SCs can actively modulate their phenotype in response to damage. These characteristic changes allow SCs to transition to a de-differentiated state, which leads to an increased expression of several regenerative genes along with the release of neurotrophic factors, adhesion and ECM-associated molecules [45]. These environmental changes that are initiated by SCs heavily contribute to the inherent regenerative potential that exists within the PNS.
1.4.2. Extracellular matrix remodeling
The extracellular matrix (ECM) is a network of secreted proteins that play a critical role in the maintenance of tissue homeostasis in each organ system throughout the body [46]. There is a dynamic interplay that exists between the tissue-resident cells and their surrounding ECM, which ultimately dictates the specificity, phenotype, and function of a tissue. Therefore, the ECM of both the central and peripheral nervous systems each require their own unique structural and functional characteristics, which allow them to accommodate the responsibilities of their host cells.
The ECM comprises approximately 20% of the total CNS tissue volume and primarily consists of hyaluronan, chondroitin sulfate proteoglycans, and tenascins [47]. Also, ECM molecules, such as heparan sulfate proteoglycans, laminins, collagen, and fibronectin combine to form the basement membrane and reinforce the blood-brain-barrier (BBB) [48-50]. Irregular ECM deposition and composition result in the presence of damage-associated molecular patterns (DAMPs), which can trigger an unpredictable inflammatory response [51,52]. Perineuronal nets (PNNs) are a unique group of dense ECM complexes that surround neurons and regulate axonal growth [53,54]. PNNs are believed to be involved with CNS plasticity [55-58] and could be a target for stimulating functional repair post-injury [59-62].
Of the ECM components in the CNS, chondroitin sulfate proteoglycans (CSPGs) have been identified as a key constituent of the PNN involved in the injury response [59,60,62]. These CSPGs have been shown to play an inhibitory role in neuronal repair after CNS injury; however, the digestion of the perineuronal net, via degradation of CSPGs, can reverse this response and promotes axonal sprouting [59,60,62]. Motor and sensory recovery in a rat spinal cord hemisection model has also shown functional gains as a direct result of digesting CSPGs [60]. Therefore, targeted changes in ECM composition could be used as a potential therapeutic strategy for functional restoration following traumatic injury.
In contrast to CNS ECM, PNS ECM is predominantly composed of collagens, laminin, fibronectin, and proteoglycans, which is more typical of the matrix organization observed in the majority of tissues [20,63-66]. In response to injury, peripheral nerves undergo Wallerian degeneration, which removes damaged or dysfunctional axons at the distal end as well as excess myelin in preparation to support Schwann cell proliferation and neurotrophin secretion [67]. This repair response features an upregulation of de novo matrix production to facilitate nerve regeneration.
Several in vitro studies, testing dorsal root ganglia (DRG) have documented a marked improvement of axon elongation and sprouting, as well as enhanced SC morphology in the presence of ECM molecules, specifically laminin [68-70]. However, although laminin outperformed its ECM counterparts during exposure to DRG cultures, other neuronal cell types may prefer alternative ECM molecules, or a combination thereof, to maximize growth potential [71]. Similar to the CNS, axon regeneration and outgrowth during DRG cultures can be inhibited by sulfated proteoglycans [72]. Additionally, ECM ultrastructure and mechanical properties may be a predictive measure of the regenerative capacity of neurite extension [73,74].
The pronounced role of the ECM in PNS axon regeneration has equally been demonstrated in vivo. At the most basic level, acellular muscle grafts can stimulate axon growth through a porous basal lamina, composed primarily of laminin and fibronectin [75]. Also, the enzymatic digestion of inhibitory CPSGs predictably reverse stunted axons and initiate a repair response [76,77]. The addition and combination of ECM molecules to nerve guides has also been shown to improve the regenerative outcomes in long gap models [78-80]. Although ECM composition plays a significant role in axon regeneration, ECM patterning and spatial congruence are necessary for axon guidance. Therefore, in order to mimic the success seen in autologous nerve grafts, it is imperative to combine both ECM composition and structural alignment to encourage axon repair.
1.4.3. Immune cell modulation and macrophage involvement
Immune cell activity is imperative for the initiation of tissue repair and remodeling in response to both CNS and PNS injury. Although there are several immune cells involved in nervous tissue repair, the macrophage may be one of the most critical, due to its diverse set of functions. There is significant evidence highlighting the importance of these cells in response to tissue-specific injuries within the nervous system: SCI [81-87], TBI [88-91], and PNI [92-94].
Tissue-resident microglia are the predominant macrophage population within the CNS and are phenotypically distinct from circulating monocytes that appear during injury and inflammation [95-97]. In an unperturbed state, microglia primarily assist with debris clearance through phagocytosis, antigen presentation, and the secretion of immunomodulatory molecules. In response to CNS injury, astrocytes release a combination of pro-inflammatory cytokines and chemokines, along with the production of reactive oxygen species and nitric oxide [98-101]. Together, these microenvironmental cues can act as a beacon to summon immune cells, such as circulating monocytes and activate resident microglia to assist in the repair of damaged tissue. The activation of microglia in response to injury, which is heavily influenced by CD4+ T cells, leads to phenotypic selection, increased proliferation, as well as the spontaneous production of neuroprotective stimulatory molecules [102-104]. Although this response can have a positive impact on tissue repair, it has been suggested that this hypertrophic state can lead to neurotoxicity with the potential of propagating neurodegenerative disease [103,105-107].
Comparable to the microglia of the CNS, the PNS offers its own distinct set of tissue-resident endoneurial macrophages, which play a crucial role in early tissue maintenance post-injury [108]. Macrophage activation and response in the PNS is directly related to severity, and the recruitment of hematogenous macrophages is triggered well after initial axotomy during axonal breakdown [109,110]. Infiltrating monocytes assist resident macrophages and SCs for efficient debris clearance, demyelination, and remyelination after Wallerian degeneration [111-113]. Also, macrophages influence SC proliferation and their migration along regenerative bridges [29].
As with most injured or diseased tissue, macrophage phenotype can predict long term regenerative outcomes [114]. Thus, the development of regenerative medicine-based interventions must consider incorporating the ability to modify these phenotypic signatures to enhance their therapeutic potential. Direct intervention with the local delivery of pro-regenerative cytokines has been shown as a method to alter macrophage phenotype, leading to improved SC infiltration and accelerated axon growth after PNI [30]. However, this macrophagemodifying mechanism has also been observed after dECM derivatives are introduced [115].
Both CNS and PNS injuries elicit a robust macrophage response that can either support or disrupt tissue repair based upon the molecular profile that exists within each microenvironment. In recent years, several groups have studied the therapeutic potential of dECM materials and their impact on the host response after injury [114,116,117]. A resounding theme shared between these studies is the inherent ability of the ECM to shift the macrophage phenotype from a pro-inflammatory to a pro-remodeling state [115]. Therefore, the implementation of dECM materials may be of interest for reversing the pro-inflammatory conditions observed within the nervous system niche post-injury.
2. Methods of decellularization treatments
Decellularization has become a popular technique for obtaining acellular scaffolds through physical and/or chemical processing. Careful considerations must be made when determining the appropriate decellularization method, such as differences in tissue sources, the desired end-product morphology, and the target location for treatment. Several popular methods have been created and applied to successfully decellularize tissues, with success often measured by mitigating an immunogenic response. These methods have broadly employed mechanical delamination of tissue layers, freeze/thaw cycles, enzymatic and detergent-based washing steps, and perfusion/agitation to ensure maximum infiltration and removal of cellular components. Due to the sensitive nature of brain, spinal cord, and peripheral nerve tissues, there have been several attempts at finding an adequate combination of treatments to preserve the mechanical integrity of the scaffolds as well as the specific molecular profiles that accompany each unique niche. Additionally, some methods have prepared dECM from tissues sourced outside of the intended target area (e.g., paravertebral muscle matrix in spinal cord) to achieve desirable outcomes [118].
2.1. Detergent-based treatment
A variety of detergents have been used to disrupt cell-cell and cell-matrix bonds, while preserving the integrity of the matrix, resulting in decellularized tissues that are nonimmunogenic. At the level of whole organs, detergents perfused in low concentrations have seen success by accessing cellular attachments throughout the bulk of the organ through the vasculature. While perfusion is a viable strategy for whole organs, tissues can alternatively be separated before decellularization through mechanical delamination and subsequently exposed to detergents in an agitated, static, or high-pressure environment.
Detergents are frequently used among the various decellularization methods applied to nervous tissues, but the specific type or combination of detergents is still up for debate. Overall, these chemicals can be divided into three categories: ionic (subdivided into cationic and anionic), nonionic, and zwitterionic/amphoteric. While all classes of detergents have shown variable success in disruption of cell-cell and cell-matrix attachments, they can also cause changes within matrix-matrix adhesions. These effects can specifically alter growth factor content as well as mechanical properties of the native ECM structure as described in Table 1 [174]. For example, SDS is a well-accepted and effective detergent capable of reducing DNA content within decellularized tissues. However, it is also known to significantly diminish desirable matrix components along with biomolecules, such as growth factors and hormones, which are quintessential elements of a bioactive scaffold.
Table 1.
Decellularization Methods for Nervous System Applications.
| Category | Class | Examples | Tissue Types | Applications | References | ||
|---|---|---|---|---|---|---|---|
| CNS | PNS | CNS | PNS | ||||
| Detergent | Ionic | Sodium dodecyl sulfate (SDS) | Paravertebral muscle, cerebellum, whole brain, sciatic nerve, tibial nerve, femoral nerve, median nerve, caudia equine | ✓ | ✓ | [118-120] | [121-126] |
| Sodium deoxycholate (SDC) | Spinal cord, sciatic nerve, optic nerve, whole brain, cerebral cortex, sciatic nerve, tibial nerve, femoral nerve, caudia equine | ✓ | ✓ | [15,127-141] | [138,142-148] | ||
| Triton X-200 | sciatic nerve, tibial nerve, femoral nerve, human peripheral nerve | ✓ | N/A | [121,142,149-157] | |||
| Nonionic | Triton X-100 | Paravertebral muscle, spinal cord, sciatic nerve, optic nerve, whole brain, cerebral cortex, cerebellum, sciatic nerve, tibial nerve, femoral nerve, median nerve, caudia equine | ✓ | ✓ | [15,118,120,127-141,146,147,158] | [121,122,124,138,143-148,158-162] | |
| Zwitterionic | Sulfobetaine-10 (SB-10) | sciatic nerve, tibial nerve, femoral nerve, human peripheral nerve | ✓ | ✓ | [163] | [121,142,149-157] | |
| Sulfobetaine-16 (SB-16) | sciatic nerve, tibial nerve, femoral nerve, human peripheral nerve | ✓ | ✓ | [163] | [121,142,149-157] | ||
| CHAPS | sciatic nerve, tibial nerve, femoral nerve | ✓ | N/A | [158] | |||
| Chemical | Peracetic acid | UBM, whole brain, spinal cord, optic nerve, sciatic nerve | ✓ | ✓ | [127,128,132,134,136,137,139,164-167] | [122,126,142,145-147,161,168-170] | |
| EDTA | whole brain, spinal cord, optic nerve, cerebellum, sciatic nerve | ✓ | ✓ | [120,128,134,136,137,139] | [145,159,160] | ||
| Enzyme | Trypsin | whole brain, spinal cord, optic nerve, cerebellum, sciatic nerve | ✓ | ✓ | [120,128,134,136,137,139] | [145,159,160] | |
| DNase | sciatic nerve | ✓ | ✓ | [127,129,131,135,136,138] | [138,147,159-161,171] | ||
| RNase | sciatic nerve | ✓ | ✓ | [131,138] | [138,147,159,160] | ||
| Pancreatin | sciatic nerve | ✓ | [138] | N/A | |||
| Chondroitinase ABC | sciatic nerve, human peripheral nerve | ✓ | N/A | [149,153,155-157,172,173] | |||
In comparison to other commonly used detergents, SDS is considered to be one of the most potent and disruptive chemical agents. However, when used at high concentrations and long incubation times, it can significantly reduce sensitive bioactive components within the tissues. Triton X-100, a non-ionic detergent, has been used as a more ECM-friendly alternative to SDS. Comparisons made between these two detergents have shown improved collagen retention with the use of Triton X-100, at the cost of less effective DNA removal [175]. To compensate for this, Triton X-100 has often been combined with sodium deoxycholate (SDC), an ionic detergent shown to have greater biocompatibility than SDS [176]. Detergent optimization is a critical process that is required to obtain an ECM scaffold with minimized matrix disruption, adequate antigen removal, and limited cytotoxicity. Further removal of undesirable biological components, such as dsDNA and other cellular components, can be aided by the inclusion of enzymatic treatment.
2.2. Enzymatic-based treatment
While detergents are classified by their charge distribution, enzymatic decellularization agents (nucleases, collagenases, and proteases) are organized by the specific structures that they target. These enzymes are utilized to modify and remove antigenic material to decrease immunogenicity, while minimally contributing to any residual cytotoxicity. Various enzymes used in decellularization methods are listed in Table 1; however, enzymes are often included as supplements to detergents in order to enhance antigen removal, as shown in Tables 2 and 3. Trypsin/EDTA, commonly used in cell culture practices, are used together prior to detergent application to both cleave and disrupt cell-ECM adhesions. While this enzyme alone could decellularize a tissue entirely, it has been shown to compromise the structural integrity of the ECM with the prolonged exposure that would be required for complete decellularization. Nucleases, specifically DNases and RNases, are used to aid in the removal of nucleic acids where the detergent combination would not meet the threshold for acceptable levels of residual DNA content.
Table 2.
Decellularization Methods for Central Nervous System (CNS) Applications.
| Application | Study model | Source Tissue | Additional Components |
Post Processing | Decellularization Method | Notable Results | References | |||
|---|---|---|---|---|---|---|---|---|---|---|
| # | Step | Time | RPM | |||||||
| SCI | in vitro | Rat spinal cord | N/A | N/A | Mechanical tissue separation Tissue segmented | Good ECM structure preservation. Constructs contained laminin, collagen, and fibronectin and show increased neural cell adhesion and proliferation. | [130] | |||
| 2x | 1% Triton X-100 | 3 h | yes* | |||||||
| 1% Deoxycholate Stored at 4 °C in PBS | 3 h | yes* | ||||||||
| in vitro | Rat spinal cord | Genipin crosslinker and glutaraldehyde crosslinker | Lyophilized, immersed in crosslinker solutions, lyophilized again, and sliced into discs for culturing | Mechanical tissue separation Tissue segmented | Genipin was equivalent in structural properties and superior in biocompatibility to glutaraldehyde. Degradation of the genipin crosslinked scaffold was around 20% in trypsincontaining buffer over 14 days. Genipen increased proliferation and ECM secretion. | [15,130] | ||||
| 2x | 1% Triton X-100, 1 h media changes | 3 h | 100 | |||||||
| 1% Deoxycholate, 1 h media changes Lyophilization | 3 h | 100 | ||||||||
| 5 mg/mL crosslinker immersion at 37 °C Lyophilization Cobalt-60 irradiation for sterilization Stored dry at −20 °C | 24 h | |||||||||
| Rat spinal cord hemisection up to 8 weeks | Rat thoracic spinal cord | human umbilical cord MSCs | Freeze dried and gamma irradiated | Mechanical tissue separation | hUCB-MSC addition to ECM scaffolds showed increased motor function recovery. Increases in axon sprouting, myelination, and oligodendrocyte migration were observed. | [130,133] | ||||
| 2x | Tissue segmented 1% Triton X-100 | 3 h | yes* | |||||||
| 1% Deoxycholate | 3 h | yes* | ||||||||
| Freeze-dried Gamma irradiation | 24 h | |||||||||
| Rat spinal cord hemisection | Rat paravertebral muscle | N/A | N/A | 1x | 3% Triton X-100 | 2d | Parallel and linear axonal spouting was observed and axon survival was increased. | [118,181] | ||
| 0.1% SDS | 2d | |||||||||
| PBS | 1d | |||||||||
| Subcutaneous rat implant model up to 1 week | Rat sciatic nerve | N/A | N/A | Method 1: Detergent | For methods 1 and 2, nuclear content was eliminated but some myelin was detected. Immunogenicity was lower than untreated sciatic nerve in methods in all 3 methods with method 3 showing the least MHC II presentation. Mechanical properties of normal sciatic nerve were preserved. | [138] | ||||
| 2x | 3% Triton X-100 | 12 h | ||||||||
| 4% Deoxycholate Stored at 4 °C in PBS | 24 h | |||||||||
| Method 2: Detergent/Enzyme | ||||||||||
| 0.5% Triton X-100 | 48 h | yes* | ||||||||
| DNase and RNase at 37 °C | 12 h | |||||||||
| Method 3: Enzyme | ||||||||||
| Hypotonic solution rinse at 4 °C | 12 h | |||||||||
| Frozen at −80 °C | 6 h | |||||||||
| Thawed at 37 °C | 30 m | |||||||||
| 0.05% pancreatin | 6 h | |||||||||
| DNase and RNase | 12 h | |||||||||
| in vitro | Rat spinal cord | N/A | Triton X-100 Deoxycholate | Varied | This study observed effects of agitation on decellularization and found 120 RPM to be optimal for spinal cord. | [141] | ||||
| Varied | ||||||||||
| Rat spinal cord hemisection model up to 14 days | Rat spinal cord | bFGF and HP (heparin modified poloxamer) | lyophilized and mixed with bFGF and HP | 1x | 1% triton X-100 at 4 °C | 12 h | 120 | Inclusion of HP-bFGF increases PC12 cell survival by 50%. bFGF reduced glial scarring and increased recovery. | [140] | |
| 8 h | 120 | |||||||||
| 4.0% sodium deoxycholate at 4 °C | ||||||||||
| 10x | 0.01% PBS elution rinse at 4 °C | 10m | 60 | |||||||
| in vitro, PC12 cell line up to 7 days in culture | Porcine brain | N/A | Comminuted into < 1 mm particles, pepsin/HCl solubilized, gelled via neutralization and dilution | 1x | Freeze/Thaw cycle at −80 °C Longitudinal quartering of tissue | 16 h | This study observed the effects of various nerve tissue sources. Brain derived ECM scaffolds showed significantly greater neurite extension length. Otherwise spinal cord, brain, and UBM showed equivalent benefits. | [128,134,139] | ||
| 1x | Type 1 water at 4 °C | 16 h | 60 120 for brain/200 for spinal cord/nerve | |||||||
| Porcine spinal cord | 0.02% Trypsin/0.05% EDTA at 37 °C | 1 h | ||||||||
| 3% Triton X-100 | 1 h | |||||||||
| 1.0 M Sucrose | 15 m | |||||||||
| Porcine optic nerve | 4% Deoxycholate | 1 h | ||||||||
| 1% Peracetic acid in 4% ethanol Lyophilized and stored dry | 2 h | |||||||||
| Rat acute SCI up to 8 weeks post-injury | Porcine spinal cord UBM | hMSCs | Same gelation process as Crapo et al. 2012 above | Same method as Crapo et al. 2012 above with additional steps. | This study compared SC ECM with UBM hydrogels. Equivalence was found in most categories. hMSCs added to SC ECM found to have little effect. SC ECM and UBM both found to significantly modulate the immune response. | [128,137] | ||||
| Digested 1.0 mg/mL pepsin, 0.01 M HCl Neutralized 0.1 M NaOH, 10x PBS | ||||||||||
| in vitro with rat SCs, in vivo rat SCI for 14 days | Rat sciatic nerve | SCs | Lyophilized, digested, applied as hydrogel | SB-10 in 50 mM sodium/10 mM phosphate buffer 0.14% Triton X-200/0.06 mM SB-16 in 50 mM sodium/10 mM phosphate 200 uL chondroitinase ABC at 37 °C | 16 h | The ECM hydrogel supported axon growth, SC survival, and axon myelination. Locomotive recovery was increased, and no chronic immune response was observed. | [150,151,163] | |||
| TBI | Mouse CCI model up to 35 days | Porcine brain | N/A | Lyophilized and reconstituted with PBS | Same method as Crapo et al. 2012 above. | ECM showed reduced glial scarring post-injury as well as reduced neurodegeneration and reduced lesion volume. | [128,139] | |||
| Controlled cortical impact rat model up to 26–28 days. | Porcine UBM | NSCs cultered in UBM hydrogel prior to injection | Same gelation process as Crapo et al. 2012 above | Mechanical separation of the luminal surface 1 M saline rinse | UBM hydrogels reduced CNS tissue loss and mitigated losses in memory, cognition, and motor faculties. UBM and UBM + NSCs showed significant benefits, with reduction of loss in memory and cognition being significant with the inclusion of NSCs. | [166,167] | ||||
| 1x | 0.1% Peracetic acid/4% ethanol Lyophilization | 2 h | ||||||||
| Solubiliztion in 1 mg/mL pepsin/0.01 M HCl Frozen at −20 °C for storage | 2d | |||||||||
| Chick embryo chorioallantoic membrane | Whole rat brain | N/A | 2x | Antibiotic/Antimycotic rinse | The observed angiogenic response was comparable to FGF-2 application. Little to no inflammatory response was seen. | [135,182] | ||||
| 4x | Freeze/Thaw cycle 2x | 4% Sodium deoxycholate | 1 h | |||||||
| 2000 kU DNase I in 1 M Na CL | 15 m | |||||||||
| Storage at 4 °C in PBS | ||||||||||
| in vitro | Whole Rat Brain | N/A | Lyophilized, mixed with gelatin, electrospun, and then genipin crosslinked | Antibiotic/Antimycotic rinse | A cytocompatible scaffold was produced that, when cultured with allogeneic rat mSCs, showed enhanced differentiation potential. | [127,135] | ||||
| 4x | Freeze/thaw cycle | |||||||||
| 1x | 1% Triton X-100 | 1 h | 60 | |||||||
| 4.0% deoxycholate | 1 h | 60 | ||||||||
| 2000 KU DNase in 1 M NaCl Stored in PBS 1% antibiotic/antimycotic | 1 h | 60 | ||||||||
| In vitro characterization with NSCs and subcutaneous and intracranial implantation in a rat model up to 4 weeks | Mouse Cerebellum | N/A | N/A | Transcardial 1% SDS infusion (pre-harvest) | 5 m | Scaffolds retained neurosupportive proteins and increased migration and proliferation of NSCs in coculture. When compared to UBM intracranial implants, advantages were seen in cerebellum-derived ECM scaffolds. | [120] | |||
| 1x | 1% SDS | 1 h | 100 | |||||||
| 0.02% Trypsin/0.05% EDTA | 30 m | 60 | ||||||||
| 1% Triton X-100 | 1 h | 100 | ||||||||
| 1 M Sucrose | 15 m | 60 | ||||||||
| Antibiotic Rinse Stored in PBS | 3d | |||||||||
| In vitro with iPS cells | Halved rat whole brain | N/A | 3x | 0.1% SDS in PBS with 1% antibiotic/antimycotic | 24 h | Decellularized brain scaffolds induced neural differentiation pathways when cultured with iPS cells. | [119] | |||
| 1x | Centrifugation in 50 mL conical tubes | 5 m | 10,000 | |||||||
| 10x | PBS solution centrifugation (SDS removal) | 5 m | 10,000 | |||||||
| 3D in vitro culturing | Fetal and young adult porcine brain | N/A | Lyophilized and gelled following various manufacturer instructions for Hydromatrix, Puramatrix, and HyStem C | Mechanical separation of the dura mater | Neural network formation was enhanced in the presence of fetal and young adult ECM compared to collagen alone in 3D culture. | [128,136] | ||||
| 1x | 0.05% Trypsin/EDTA with 0.2% DNase at 37 °C | 1 h | ||||||||
| 3% Triton X-100 with 0.2% DNase | 1 h/30 m | |||||||||
| 1 M Sucrose | 15 m | |||||||||
| 4% sodium deoxycholate | 1 h/15m | |||||||||
| 0.1% peracetic acid/4% ethanol Lyophilization and storage at −20 °C | 2 h | |||||||||
| in vitro 3D culturing with NSCs up to 7 weeks | Mouse brain 1.5 mm sections | N/A | N/A | 1x | 4% sodium deoxycholate | 14 h | 150 | NSCs labeled with GFP showed continued growth over 7 weeks cultured on brain ECM sections, and, when exposed to mitogenic stimuli, retained their NSC phenotype. | [129] | |
| 40 kU/mL DNase | 1 h | 150 | ||||||||
| deionized water 150 | 4h | |||||||||
| 3% Triton X-100 | 2 h | |||||||||
| 150 40 kU/mL DNase 2x | 1 h | 150 | ||||||||
| deionized water 150 | 4h | |||||||||
| 4% sodium deoxycholate 150 | 12 h | |||||||||
| 40 kU/mL DNase | 1 h | 150 | ||||||||
| deionized water | 2 h | 150 | ||||||||
| 3% Triton X-100 | 2 h | 150 | ||||||||
| 40 kU/mL DNase | 1 h | 150 | ||||||||
| in vitro | Rat cerebral cortex | Genipin crosslinker | — | Freeze/thaw Triton X-100 Deoxycholate DNase and RNase Genipin crosslinker | Genipen increased pore sizes of scaffolds. Fibronectin and basement membrane were weakly retained but laminin was strongly retained. | [131] | ||||
| Stroke | Rat stroke model up to 14 days and 12 weeks | Porcine UBM | N/A | Same gelation process as Crapo et al. 2012 above | 1x | Mechanical delamination 0.1% Peracetic acid/4% ethanol Debris removal via PBS rinses Lyophilization Solubilization in 1 mg/mL pepsin/0.01 M HCl Frozen at −20 °C for storage | 2 h | 300 | Injection of the UBM hydrogel increased cell proliferation and density at the injury site interface with some penetration. However, at 12 weeks there were no significant differences in scar formation. | [164,165] |
| Parkinson's | Parkinson's Disease model rats up to 20 days | Rat brain | bFGF | N/A | Transcardial PBS perfusion during harvest | Sustained release of bFGF from ECM scaffolds increased cell survival and behavioral recovery. | [127,132] | |||
| 1x | 3% Triton X-100 | 12 h | 120 | |||||||
| 4% deoxycholic acid sodium salt | 24 h | 120 | ||||||||
| 0.1% peracetic acid/4% ethanol Stored in PBS 1% antibiotic/antimycotic | 3 h | 120 | ||||||||
Table 3.
Decellularization Methods for Peripheral Nervous System (PNS) Applications.
| Application | Study model | Source Tissue | Additional Components |
Post Processing | Decellularization Method | Notable Results | References | |||
|---|---|---|---|---|---|---|---|---|---|---|
| # | Step | Time | RPM | |||||||
| Transection and gap | In vitro analysis of decellularization | Rat sciatic nerve | N/A | N/A | Sondell et al. 1998 Method | Direct method comparison: Sondell’s method disrupted the nerve ECM more than the Hudson method. This disruption could be from shrinkage/swelling cycles when applying deoxycholate/water. Hudson’s method produced more demyelination and improved adhesive ratios. | [143,144,146,150-152,154] | |||
| distilled water | 7 h | |||||||||
| 2x | 3% Triton X-100 | 12 h | ||||||||
| 4% Sodium deoxycholate distilled water | 24 h | yes* | ||||||||
| Hudson et al. 2004 Method | ||||||||||
| distilled water | 7 h | |||||||||
| 125 mM sulfobetaine-10 | 15 h | yes* | ||||||||
| 50 mM phosphate and 100 mM sodium | 15 m | |||||||||
| 0.14% Triton X-200, 0.6 mM sulfobetaine-16, 10 mM phosphate and 50 mM sodium | 24 h | yes* | ||||||||
| 3x | 50 mM phosphate and 100 mM sodium | 5 m | ||||||||
| 125 mM sulfobetaine-10 | 7 h | yes* | ||||||||
| 50 mM phosphate and 100 mM sodium | 15 m | |||||||||
| 0.14% Triton X-200, 0.6 mM sulfobetaine-16, 10 mM phosphate and 50 mM sodium | 15h | |||||||||
| 3x | 50 mM phosphate and 100 mM sodium | 15 m | ||||||||
| 1.5 cm rat sciatic nerve defect | Rat sciatic nerve | N/A | N/A | Distilled water | 7 h | yes* | Decellularized similarly to Hudson’s method with increased ECM preservation. Outperformed Hudson’s method in nerve histomorphometry and muscle mass. Outperformed by autograft. | [158] | ||
| 1 M Sodium chloride | 15 h | yes* | ||||||||
| 2.5 nM Triton X-100 | 24 h | yes* | ||||||||
| 1X PBS | 15 m | yes* | ||||||||
| Distilled water | 7 h | yes* | ||||||||
| 1 M Sodium chloride | 15h | yes* | ||||||||
| 100 nM CHAPS | 24 h | yes* | ||||||||
| 5x | 1X PBS | 5 m | yes* | |||||||
| In vitro characterization and effect on PC12 cells | Rat sciatic nerve | N/A | axially aligned channels | 2x | distilled water | 7 h | Decellularization significantly increased pore size and created axially aligned channels. In vitro work showed enhanced PC12 cell migration both in number and depth into the graft. | [147] | ||
| 3% Triton X-100 | 12 h | yes* | ||||||||
| 2% deoxycholic acid in 50 nM Trizma base | 24 h | yes* | ||||||||
| DNase and RNase (5 U/ml) in 10 nM magnesium chloride | 12 h | |||||||||
| 0.1% peracetic acid | 1 h | |||||||||
| 3.5 cm Rat sciatic nerve gap defect | Rat sciatic nerve | N/A | N/A | DMEM 10% FBS 2% pen/strep/amph PBS | 2wks | yes* | Nerve autograft outperformed both the detergent free and detergent decellularized grafts. Only nerve autograft and detergent free grafts showed muscular recovery. | [186] | ||
| 1wk | yes* | |||||||||
| In vitro characterization and subcutaneous implantation | Rat sciatic nerve | Apoptosis assisted | N/A | DMEM with camptothecin (5 or 10uM) 37 °C | 1-3d | 14 | The apoptosis-assisted method maintained basal lamina microarchitecture with decreased cytotoxicity. | [171] | ||
| 4X PBS | 24 h | |||||||||
| 2X PBS | 30 m | |||||||||
| 3x | 1X PBS | 30 m | Also, inflammatory infiltrate and stromal remodeling decreased compared to autograft control. | |||||||
| 75 U/mL DNase | 36 h | |||||||||
| 2x | 1X PBS | 30 m | ||||||||
| In vitro characterization | Rat sciatic nerve | N/A | N/A | 2x | 10 mM TRIS-HCl; pH 8.0 | yes* | Reduced DNA content by greater than 95% while preserving collagen, laminin, and fibronectin. | [126] | ||
| 0.1% SDS with 10 kIU/mL | yes* | |||||||||
| 3x | 1X PBS | 30 m | yes* | |||||||
| 1 U/mL Benzonase | 3 h | yes* | ||||||||
| 1.5 M NaCl | yes* | |||||||||
| 0.1% peracetic acid in PBS | 3 h | yes* | ||||||||
| 3x | 1X PBS at 4C | 30 m | yes* | |||||||
| 1X PBS at 4C | 48 h | yes* | ||||||||
| 1 cm and 1.5 cm rat sciatic nerve defect models | Rat sciatic nerve and rabbit median nerve | N/A | N/A | 1X PBS | Allogeneic and xenogeneic tissue sources were used and compared in vivo to isograft control up to 24 weeks. Many axons and Schwann cells were shown migrating into the bridge, with few macrophages present. Both tissue sources were inferior to isograft control, contradicting previous results from the group in Wakimura et al. 2015. | [124,162] | ||||
| 1% SDS | 24 h | yes* | ||||||||
| Distilled water | 30 m | |||||||||
| 1% Triton X-100 | 1 h | yes* | ||||||||
| Distilled water | 30 m | |||||||||
| 100 U/mL penicillin-G, 100 ug/mL streptomycin, 0.25 ug/mL amphotericin B | 7d | |||||||||
| In vitro characterization | Rat sciatic nerve | N/A | N/A | 2x | 50 nM Tris buffer | 12 h | Comparisons were made to Sondell’s and Hudson’s methods. The proposed method showed a better removal of DNA content and preservation of ECM components. | [146,150,151,159,160,187,188] | ||
| 1% Triton X-100 in 50 nM Tris buffer at 4 °C | 24 h | |||||||||
| HBSS | 24 h | 24 | ||||||||
| 2x | 40,000 U/L DNase, 20 mg/L RNase, 100 mg/L Trypsin at 37C | 45 m | yes* | |||||||
| 1% Triton X-100 in 50 nM Tris buffer at 4 °C | 12 h | |||||||||
| 3x | HBSS at 4 °C | |||||||||
| 1.5 cm rat sciatic nerve defect | Porcine sciatic nerve | N/A | Processed into an ECM hydrogel | distilled water | 7 h | Resembling Sondell’s method, this study reduced washing steps to preserve ECM needed for gelation. The hydrogel was found to be superior using the altered method, but inferior to autograft control. | [189] | |||
| 3% Triton X-100 | 12 h | |||||||||
| 4% Sodium deoxycholate | 24 h | yes* | ||||||||
| 3x | distilled water | 15 m | yes* | |||||||
| Ethanol:Dichloromethane (1:2) distilled water | 24 h | |||||||||
| 1.5 cm rat sciatic nerve defect model | Porcine sciatic nerve | N/A | Processed into an ECM hydrogel | Distilled water | 14 h | 300 | The decellularized nerve ECM hydrogel was used as a lumen filler for a rat model of nerve gap injury. It showed increased regenerative bridge formation followed by enhanced M2 macrophage and Schwann cell migration and involvement. | [145] | ||
| 0.02% Trypsin/0.05% EDTA at 37 °C | 1 h | 300 | ||||||||
| 3% Triton X-100 | 1 h | 300 | ||||||||
| Distilled water | 300 | |||||||||
| 1 M Sucrose | 15 m | 300 | ||||||||
| 4% Sodium deoxycholate | 1 h | 300 | ||||||||
| 0.1% Peracetic acid/4% ethanol | 2 h | 300 | ||||||||
| 1X PBS | 15 m | 300 | ||||||||
| 2x | Distilled water | 15 m | 300 | |||||||
| 1X PBS | 15 m | 300 | ||||||||
| 1.5 cm rat sciatic nerve defect model | Rat sciatic nerve | N/A | N/A | Distilled water | 6 h | Method altered detergent combinations from Hudson’s method to reduce residual myelin. Comparing these two methods to the nerve autograft, the modified method had reduced immunogenicity and remyelination more comparable to autograft controls. However, morphologic assessment of axon regrowth showed no difference between the two methods. | [142] | |||
| 125 mM sulfobetaine-10 | 12 h | yes* | ||||||||
| 3x | 1X PBS | 10m | ||||||||
| 0.14% Triton X-200, 0.6 mM sulfobetaine-16 | 24 h | yes* | ||||||||
| 1X PBS | 10 m | |||||||||
| Distilled water | 10 m | |||||||||
| 125 mM sulfobetaine-10 | 12 h | yes* | ||||||||
| 4% Sodium deoxycholate | 24 h | yes* | ||||||||
| 3x | 1X PBS | 2 h | ||||||||
| 0.1% Peracetic acid | 3 h | |||||||||
| 3x | 1X PBS | 1 h | ||||||||
| Rabbit tibial nerve | Rabbit tibial and femoral nerves | N/A | N/A | Method 1: Hudson Method (control) | These methods limit hands-on time to prevent contamination and necessity of sterilization. Methods 2 and 3 were less harsh than Hudson’s method with small amount of DNA fragments and Schwann cells still present. Method 2 produced uneven decellularization towards the center of the graft. Method 3 did remove most of the myelin, axons, and DNA residue to a satisfactory degree with ECM preservation. | [121] | ||||
| 2x | 125mM SB-10, 10 mM phosphate, 50 mM sodium | 15 h | yes* | |||||||
| 0.14% Triton X-200, 0.6 mM SB-16, 10 mM phosphate, 50 mM sodium | 24 h | yes* | ||||||||
| Method 2: Triton X-100 based Method | ||||||||||
| 125 mM SB-10, 0.2% Triton X-100, 1% penicillin–streptomycin | 48 h | yes* | ||||||||
| Sonication at 40 Hz | 2m | |||||||||
| Method 3: Triton X-100 and SDS method | ||||||||||
| 125 mM SB-10, 0.2% Triton X-100, 1% penicillin–streptomycin freeze–thaw cycle | 48 h | |||||||||
| 3x | 1X PBS | 30 m | ||||||||
| 0.25% SDS and sonicated for 5 m every 30 m at 40 Hz | 3 h | |||||||||
| rat trigeminal, infraorbital sciatic nerve transection model | Porcine fetal urinary bladder matrix | N/A | Pressed and lyophilized into a sheet. ETO sterilized | 8x | Nanopure water | 30 m | The method uniquely included a chamber that cycled between ambient pressure and 0 psi at a rate of 1 min/cycle. The wrap enhanced epi- and endoneurial organization and increased angiogenesis; however, the number of axons and myelination were unchanged. | [161] | ||
| 3% Triton X-100 | 30 m | |||||||||
| 3 M NaCl | 30 m | |||||||||
| DNase | 1 h | |||||||||
| 0.1% Peracetic acid/4% ethanol 1X PBS | 1 h | |||||||||
| 3x | Nanopure water | |||||||||
| Rat sciatic nerve gap model looking at immunogenicity and mechanical properties | Rat sciatic nerve | N/A | N/A | Method 1: Sondell method (control) | The freeze-thaw method combined with enzymatic degradation showed the lowest immunogenicity of all methods tested. Also, fewer inflammatory cells were present and less necrosis seen at the downstream muscle with less MHC-II presentation compared to all other methods and native tissue. All decellularization techniques had similar mechanical properties to the native nerve. | [138] | ||||
| distilled water | 12 h | yes | ||||||||
| 2x | 3% Triton X-100 | 12 h | ||||||||
| 4% Sodium deoxycholate | 24 h | |||||||||
| Method 2: Low detergent method | ||||||||||
| ultrapure water | 1 h | yes | ||||||||
| 0.5% Triton X-100 | 48 h | yes | ||||||||
| ultrapure water | 48 h | |||||||||
| DNase and RNase at 37 °C | 12 h | |||||||||
| Method 3: Freeze-thaw method | ||||||||||
| hypotonic solution at 4 °C | 12 h | |||||||||
| hypotonic solution at −80 °C | 6 h | |||||||||
| hypotonic solution at 37 °C | 30 m | |||||||||
| 0.05% pancreatin | 6 h | |||||||||
| DNase and RNase at 37 °C | 12 h | |||||||||
| 1.5 cm rat sciatic nerve defect model | Rat cauda equina | N/A | Decellularized cauda equina allograft embedded within chitosan conduit | 3x | 1X PBS | Cauda equina source tissue used to test less harsh method because it lacks epineurium. This tissue required half the time (12 h) in SDS to achieve similar decellularization results to a sciatic nerve control (24 h). Cauda equina grafts performed better than other groups and similarly to autograft. | [123] | |||
| 0.5% SDS at 16 °C penicillin (100 U/mL) and streptomycin (0.1 mg/mL) in PBS | 12 h | 40 | ||||||||
| 10x | ||||||||||
| Rat sciatic nerve | 1X PBS | 12 h | ||||||||
| 10x | 1X PBS | |||||||||
| In vitro analysis | Porcine cauda equina | N/A | Homogenized cauda equina ECM with PLGA to form an electrospun scaffold | distilled water | Method incorporated insoluble proteins of the cauda equina ECM into an aligned, electrospun PLGA material. This incorporation of ECM proteins enhanced neurite outgrowth of dorsal root ganglion cells compared to PLGA material alone. | [148] | ||||
| 3% Triton X-100 | 2 h | yes | ||||||||
| 4% Sodium deoxycholate 1X PBS rinse Homogenized Centrifugation steps to isolate ECM precipitate | 2 h | yes | ||||||||
| In vitro analysis | Rat sciatic nerves | N/A | Nerve ECM coated onto PCL based conduits | distilled water | 15 m | Polydopamine (PDA) was coated onto polycaprolactone (PCL) conduits to successfully increase attachment of peripheral nerve ECM as a surface modification. This was shown to have potentially beneficial changes in Schwann cell behavior when cultured on the material in vitro. | [122] | |||
| 0.5% Triton X-100 in 1 M NaCl distilled water | 2 h | yes | ||||||||
| 1% SDS in 1 M NaCl distilled water, solution changed daily | 2d | yes | ||||||||
| 0.1% peracetic acid and 4% EtOH in 1 M NaCl | 4 h | |||||||||
| distilled water lyophilized | 4 h | |||||||||
| 1.0 cm rat sciatic nerve defect model | Rat sciatic nerve | Acellular nerve graft augmented with additional NGF or GDNF | N/A | 3x | Liquid nitrogen | 2m | Comparison of chondroitinase ABC treated acellular nerve grafts to NGF or GDNF imbibed grafts. NGF significantly enhanced motor axon outgrowth and number of myelinated axons. Sensory nerve outgrowth was significantly stunted. Dorsal root ganglion outgrowth in the presence of chondroitinase ABC and NGF showed no effect on sensory neurite outgrowth. | [172,173] | ||
| Thawed in water bath at 37 °C | 2 m | |||||||||
| 2 U/mL chondroitinase ABC in PBS at 37 °C | 16 h | |||||||||
| Incubated with NGF (10ug/mL) or GDNF (20 ug/mL) solution Rinsed in lactated Ringer solution | 1 h | |||||||||
| 2x | Rinsed in lactated Ringer solution | |||||||||
| Clinical study of digital nerve gaps > 2.5 cm | Human peripheral nerve | N/A | gamma irradiated | distilled water | 7 h | Applying the Avance decellularized nerve graft from Axogen, 86% of patients recovered meaningful function, matching autograft repair (60–80%). When studied in large diameter repairs, Avance decellularized graft was not limited in this regard and presented similar motor and sensory recovery as the gold-standard. In sensory repair applications, patients regained sensory function in 15 out of 16 cases with the Avance graft. | [149,153,155-157] | |||
| 125 mM sulfobetaine-10 | 15 h | yes* | ||||||||
| 50 mM phosphate and 100 mM sodium | 15 m | |||||||||
| Clinical study of large diameter cable repairs | 0.14% Triton X-200, 0.6 mM sulfobetaine-16, 10 mM phosphate and 50 mM sodium | 24 h | yes* | |||||||
| 3x | 50 mM phosphate and 100 mM sodium | 5 m | ||||||||
| 125 mM sulfobetaine-10 | 7 h | yes* | ||||||||
| Clinical study of trigeminal nerve repair | 50 mM phosphate and 100 mM sodium | 15 m | ||||||||
| 0.14% Triton X-200, 0.6 mM sulfobetaine-16, 10 mM phosphate and 50 mM sodium | 15 h | |||||||||
| 3x | 50 mM phosphate and 100 mM sodium Chondroitinase ABC at 37 °C | 15 m | ||||||||
| Crush | rabbit sciatic nerve crush model, clinical model of recurrent cubital tunnel syndrome | Porcine Small intestinal submucosa | N/A | Pressed and lyophilized into a sheet. ETO sterilized | Distilled water Delaminate tunica mucosa and tunica muscularis externa | Scaffold promoted vascularization and remodeling at implantation site. The product, AxoGuard Nerve Protector from AxoGen, showed enhanced grip and pinch strength as well as reduced pain levels for patients with chronic cubital tunnel syndrome. | [169,170] | |||
| 0.1% Peracetic acid/4% ethanol Distilled water | 2 h | |||||||||
2.3. Mechanical based methods
Another major goal of decellularization methods is to maintain the native ECM structure and composition. Therefore, a balance must be found between sufficient removal of immunogenic material and preservation of the ECM. Mechanical methods such as physical delamination or multiple freeze-thaw cycles can effectively remove dense cell regions or lyse cells, allowing for less harsh chemical treatments. While mechanical methods alone have been used successfully in a few tissues, they are often used in conjunction with chemical or enzymatic methods because they are ineffective at clearing genetic material from a scaffold after cell lysis. Excessive physical decellularization methods can also disrupt the natural ECM ultrastructure and alter mechanical properties.
Freeze-thaw cycles can cause cell lysis while still maintaining mechanical integrity as well as ECM matrix components such as collagen and GAGs. However, freeze-thaw cycles alone are insufficient at removing genetic material after cell lysis in most cases [177]. Similarly, the use of high hydrostatic pressure is another mechanical method that can be beneficial if used under specific conditions. Pressures greater than 600 MPa have been applied to tissues to destroy cell membranes [178,179]. For both, a combination of water or PBS washes and DNase I are used to break down and wash away fragments while removing any remaining genetic material; however, high pressures can denature or deform ECM proteins [180].
2.4. Post-processing of decellularized scaffolds
Decellularization yields an acellular scaffold, which, for many applications, provides a suitable framework to support recellularization for both in vitro and in vivo studies. When retention of native tissue architecture is desired, acellular scaffolds are often the final product of the decellularization process. However, post-processing techniques have been applied by many groups to enhance these scaffolds and address their clinical target. These techniques can include lyophilization, milling, and digestion of the dECM to create an injectable hydrogel [128] as well as crosslinking with glutaraldehyde or genipin to enhance structural integrity [15,127,131].
Hydrogel formation has been approached as a more flexible alternative to the application of unprocessed dECM. Solubilized dECM has been shown to create a homogeneous gel-like structure while preserving tissue-specific matrix and cell-secreted molecules. dECM hydrogels also form at physiologic conditions, which make them ideal for applications that require the delivery of cargo (i.e., cells, growth factors, or hormones). Hydrogel ECM concentrations can most often be used as a predictor of mechanical stiffness post-formation, with an increase in ECM concentration attributing to a higher elastic modulus. Although dECM hydrogels have shown promise for tissue-specific applications, it remains unclear if tissue-matching improves functional outcomes. For example, as seen in Tukmachev et al., spinal cord dECM versus urinary bladder matrix (UBM) showed little difference in healing response [137].
While ECM concentrations can slightly modulate the stiffness of the resulting hydrogel, crosslinkers have been implemented to increase the mechanical strength of dECM products. Nervous tissues, specifically CNS, have a weak ECM structure when decellularized. To remedy this, groups have implemented glutaraldehyde or genipin crosslinkers to bolster the structural properties [15,127,131]. A noted disadvantage of glutaraldehyde is its cytotoxic characteristics, leading more researchers to implement genipin, which provides the same structural advantages, but greater cellular proliferation and survival [15].
3. Decellularized materials for CNS applications
Applications of dECM to the CNS have included tissue sources such as spinal cord [15,128,130,133,134,137,140,141], brain [119,120,127-129,131,132,135,136,139], peripheral nerve [128,138], UBM [137,164-167], and paravertebral muscle [118]. Methods applied to these various tissue sources along with their target sites for dECM application are summarized for CNS application in Table 2. These dECM materials, are being used to treat functional deficits as a result of traumatic SCI, TBI, stroke [164,165], and Parkinson’s Disease [132]. Compared to other strategies, decellularized constructs show low immunogenicity, promotion of cellular repopulation, and promotion of angiogenesis into the healing site [130,135]. Each target treatment presents its own challenges based on differences in the innate microenvironments present within CNS tissue defects.
3.1. Spinal cord injury
To address SCI, decellularized scaffolds have been sourced from spinal cord (SC dECM), paravertebral muscle, sciatic nerve, optic nerve, brain, and UBM (Table 2). Predominantly, decellularized scaffolds applied to SCI have been created using SC dECM via combinations of physical, detergent, enzymatic, and chemical-based methods. While there is no optimal standard for decellularizing spinal cord to date, the current methods being employed overlap significantly in detergent selection while adding other steps to optimize the retention of bioactive factors and 3D tissue architecture.
There is much overlap between many of the methods applied for dECM tissues for SCI and can be generalized to a set of steps that most groups have employed for SCI (Table 2). First, in the generalized method, the tissue is physically separated upon excision from fatty and connective tissues before further processing [15,128,130]. Optionally, a freeze/thaw cycle can be included before the application of enzymes and detergents. Enzymes disrupting cell adhesion, such as trypsin/EDTA, are applied, followed by detergents, which often include one of, or a combination of, Triton X-100, SDS, and SDC. Following the dissolution of cell membranes with detergents, DNases and RNases are optionally used to degrade nuclear fragments. After the final rinses, SC dECM is processed according to each group’s post-processing procedures. Some groups have elected to agitate tissues throughout their procedures at variable speeds. Yin et al. and Wang et al. studied the inclusion of agitation in detergent-based methods consisting of Triton X-100 and SDC [125,141]. They found that 120 RPM was an optimal agitation speed for CNS structural preservation while also assisting in the clearance of cell fragments during the decellularization process [141].
Additives to SC dECM were incorporated in some studies which included: bioactive factors, crosslinking agents, and/or seeded cells. Use of exogenous bioactive factors, seen in Xu et al., included bFGF with heparin poloxamer (HP) added to their SC dECM and they reported an increase in cell survival in vitro by 50% as well as reductions in glial scarring in vivo [140]. They showed that the inclusion of both bFGF and HP contributed to enhancing bioactivity and reinforcing the 3D structure [140]. Use of crosslinking agents, seen in a study by Jiang et al., included both glutaraldehyde and genipin crosslinkers following application of detergents to increase the poor mechanical strength of the CNS ECM [15,183]. Their results showed that enhanced structural integrity was achieved as a result of crosslinking post-decellularization, significantly decreasing degradation rates, increasing the tensile strength two-fold and the modulus nearly three-fold over uncrosslinked structures [15]. When they compared glutaraldehyde to genipin directly, they found that the biocompatibility was significantly higher in the SC dECM crosslinked with genipin, while mechanical properties remained statistically unchanged [15]. This agrees historically with studies showing that genipin is a superior crosslinker in biological applications over glutaraldehyde with reduced cytotoxicity and increased biocompatibility [184,185]. Lastly, two studies included seeded cells in their final dECM constructs. Human MSCs (hMSCs) were used in both Liu et al. as well as Tukmachev et al., showing a significant increase in motor function over the SC dECM application alone in the former, and no statistical difference was observed in comparison to SC dECM in the latter [133,137]. These outcomes could be attributed to differences in hMSC source or variations in the decellularization method, among other possibilities. Overall, SC dECM constructs showed low immunogenicity, retention of bioactive factors, and the ability to enhance spinal cord regeneration. Each study prioritized different functional outcomes in SCI however, with some groups focusing on reducing the glial scar, while the others promoting axonal elongation.
The strategies discussed above all employ SC dECM to treat SCI, but other tissue sources can also be used. Zhang et al. decellularized paravertebral muscle tissue with SDS, a protein-protein disruptor, to remove cellular fragments [118]. This strategy was found to promote parallel and linear axonal sprouting across the lesion site and increasing axonal survival when compared to a no treatment group [118]. Furthermore, Crapo et al. and Tukmachev et al. compared decellularized UBM to brain and SC dECM to examine if tissue-specific ECM had an advantage over ECM from other tissue sources [128,137]. It was determined in Tukmachev et al. that there were no significant differences between SC dECM and UBM in spinal cord injuries in rats [137]. However, they did report that both SC dECM and UBM had increased vascularity, neurite extension, and macrophage polarization capabilities compared to the control group [137].
3.2. Traumatic brain injury
TBI, when compared to SCI, has less frequently applied decellularization based treatment methods. Of these methods, only tissues sourced from brain or UBM have been decellularized for use in the injured brain site (Table 2). The strategies employed to decellularize brain tissues are vastly similar to those used for SC dECM and rely primarily on Triton X-100 and SDC based methods (Table 2) [120,127,129,131,132,135,136,139]. Alternatively, SDS has also been used in lieu of Triton/SDC in some brain dECM methods [119,120], with Zhu et al. perfusing SDS transcardially prior to harvest [120]. Lastly, enzymes have been used more prevalently in brain dECM methods to enhance removal of nuclear fragments, as seen in Table 2 [120,127,129,131,135,136,139]. While some of the SC dECM methods have been directly compared to one another, there is no equivalent study for brain dECM. It is not known if specific detergent/enzyme combinations convey any advantages over other methods through direct comparison.
Post-processing modifications to brain dECM have included the use of crosslinking agents (genipin) however, the addition of exogenous factors have yet to be as explored. Crosslinking of constructs occurred following decellularization to enhance the structural integrity of brain dECM [127,131]. Brain dECM has been shown to have an angiogenic response comparable to that of FGF-2 [135], enhanced neural lineage differentiation potential in vitro with MSCs and iPS cells [119,127], improved formation of neural network architecture [136], the ability to retain neural differentiation phenotypes when mitogenic stimuli were introduced [129], and the ability to increase cellular adhesion and proliferation in culture [131].
3.3. Stroke
Recently, a stroke model employing a UBM hydrogel was explored in two studies by Ghuman et al. [164,165]. In short, UBM was decellularized using a standardized mechanical delamination and chemical-based method consisting of peracetic acid and PBS rinses to remove antigenic materials (Table 2, Stroke section). In the stroke applications, Ghuman et al. found that UBM hydrogel injection caused increased cellular proliferation, preferentially near the periphery of the cavity, but with some cellular penetration into the bulk of the injected material [165]. Furthermore, M2 macrophage polarization was shown to be increased in the presence of the UBM gel, indicating constructive remodeling, and more neural progenitors were observed as gel concentration was increased to include more UBM [165]. However, despite these results, a subsequent animal model revealed that, while cavity size was reduced, scar formation was not significantly affected by the UBM hydrogel [164]. These results highlight the importance of regulating the poorly maintained regenerative environment existing after CNS injury and demonstrates the future work needed to bridge this gap.
3.4. Parkinson’s disease
Lastly, one group has recently begun development and testing of a brain dECM approach for a Parkinson’s Disease therapy [132]. This strategy incorporates exogenous bFGF, similarly to Xu et al. above in SCI applications, as well as a detergent/chemical-based method to decellularize brain tissue. Their results showed that implementation of a brain dECM scaffold increased neuroprotection in vivo [132]. In addition, they also showed an increase in cell survival and a subsequent improvement in behavioral recovery up to twenty days post-treatment in a Parkinson’s rat model [132].
4. Decellularized materials for PNS applications
Most applications of dECM targeting PNS regeneration have sought to improve nerve grafting by generating decellularized allografts [125,126,138,142,146,147,158,171,186] or xenografts [138]. Others have processed decellularized nerve [145] or urinary bladder sources [161] to produce hydrogels or wraps to augment existing repair procedures. While the methods vary, they often utilize a combination of ionic and non-ionic detergents and sometimes enzymatic processes to decellularize the peripheral nerve tissues.
In order to study PNI, researchers have developed two primary models: transection (or gap) and crush. The transection or gap injury model aims to address the impacts of severe PNI, where a segment of the nerve is completely severed and removed, to mimic the physiological limit upon which natural nerve regrowth is not possible. The crush injury model represents a mild injury prognosis, where the nerve is expected to make a full recovery post-injury. Each model provides a deeper insight into the critical factors necessary to assist nerve regeneration for PNI patients.
4.1. Transection and gap injury
Serious trauma to the peripheral nervous system resulting in a full transection requires coaptation surgery and is often further complicated by retraction of the nerve stumps [190]. If nerve ends cannot be coapted directly without tension, the current gold standard repair involves the use of an autologous nerve graft to bridge the gap. Often using the sural nerve as donor tissue, the autologous nerve graft procedure results in donor site morbidity. As an alternative, decellularization techniques have been utilized to create acellular nerve grafts from allogeneic and xenogeneic sources that emulate the same structure and ECM composition of the autologous nerve graft.
Sondell et al. was one of the first investigators to decellularize peripheral nerve for use as an allograft [146]. Rat sciatic nerves were treated using Triton X-100 and SDC (Table 3). This process was repeated once, and then the graft was washed in distilled water and stored in PBS. Early understanding of peripheral nerve decellularization was enhanced by Hudson et al., who compared the effects of many different detergents on cellular clearance and ECM preservation. An anionic detergent, Triton X-200 was found to be the most effective decellularization agent and combined with two amphoteric detergents to further increase effectiveness [150,151]. This method was more effective at decellularizing the peripheral nerve tissue while better preserving the ECM when compared to the Sondell et al. method [143,144,150,151]. This foundation led to a wide range of work optimizing the methods, structure, and composition of decellularized nerve grafts.
Despite the advantages of the detergents Hudson et al. method identified, variations of the Sondell et al. method using Triton X-100 and SDC are still being investigated [145,147,148]. Sridharan et al. used a modified Sondell et al. method to decellularize rat sciatic nerves but imparted an axially aligned channel structure through controlled freezing [147]. The study hypothesized that native pore size of basal laminal tubes were not ideal for nerve regeneration and sought to increase the pore size to enhance axon penetration. They showed that their methods were able to effectively decellularize the tissue and significantly increase pore size while keeping the channels axially aligned. Briefly, the sciatic nerves were placed into insulating molds and stood upright, perpendicular to the freeze-dryer plate. The freezing rate was controlled to a constant rate of 1 °C/min to a final temperature of −30 °C. This was held steady for one hour before warming to −10 °C to lyophilize for 24 h. In vivo work has not been published; however, in vitro work showed enhanced PC12 cell migration both in number and depth into the graft.
Remnant detergents can be cytotoxic to infiltrating cells, so investigators have tried reducing or removing the detergent load [138,158,171-173,186]. Kim et al. devised a method that used minimal detergent concentrations to decellularize rat sciatic nerves [158]. This method decellularized the nerve to a similar degree to the Hudson method but displayed better ECM preservation. In a rat sciatic nerve gap model, the method without the detergents outperformed the Hudson method in nerve histomorphometry and muscle mass, but ultimately the nerve autograft controls showed greater recovery.
Several decellularization methods have been developed that use no detergents to create acellular grafts [138,171-173,186]. Both Wang et al. and Boyer et al. explored the use of mechanical and enzymatic decellularization methods to replace detergent based methods. Wang et al. performed a single freeze-thaw cycle and used DNase and RNase to finish the decellularization of rat sciatic nerves. This method produced less inflammation in vivo compared to the Sondell et al. method and a light detergent method which showed expression of MHC-II within the scaffolds [138]. To enhance removal of CSPGs, a known inhibitor of regrowth, Boyer et al. used a method that combined multiple freeze-thaw cycles with chondroitinase ABC. The reduction of CSPGs with chondroitinase ABC was shown to increase axon growth into critically long grafts [173]. Boyer et al. augmented the scaffold with NGF and found enhanced motor neuron survival and growth [172]. Cornelison et al. used camptothecin, hypertonic PBS solution, and 75 U/ml DNase to decellularize sciatic nerves without the use of detergents in comparison to the Hudson method [171]. Their apoptosis-assisted decellularization method was able to successfully decellularize the tissue while maintaining basal lamina microarchitecture while being less cytotoxic than detergent decellularization methods. The apoptosis-assisted method also produced less inflammatory infiltrate and stromal remodeling compared to the autograft control. Vasudevan et al. developed a decellularization method that utilized a yet undescribed technique [186]. Rat sciatic nerves were cultured in DMEM with 10% fetal bovine serum and 2% penicillin/streptomycin/amphotericin for two weeks under constant agitation. After the two-week culture period, the nerves were transferred to a new tube containing only PBS and kept for one-week under constant agitation. The abrupt termination of nutrient supply was expected to aid in decellularization. In a rat sciatic nerve gap model, histologic results indicated that nerve autografts outperformed both the detergent-free and detergent decellularized grafts. Only nerve autograft and detergent-free grafts showed functional recovery in muscle force testing.
Another approach to replacing the nerve autograft is nerve guidance conduits. While outcomes of current nerve guidance conduits are often inferior to the nerve autograft, researchers have investigated numerous approaches to improve these devices. The inclusion of ECM is one such strategy. Prest et al. generated a hydrogel from a decellularized sciatic nerve and used it as a lumen filler for a nerve guidance conduit [145]. Canine sciatic nerves were decellularized using a method adapted from a spinal cord decellularization protocol [128]. Briefly, the nerves were soaked in distilled water for 14 h followed by a series of agitated washes as shown in Table 3. The resultant decellularized graft was then powdered and digested in pepsin and 0.01 M HCl and neutralized to a pH of 7.4 to form a hydrogel. The use of the decellularized nerve ECM hydrogel as a lumen filler for a rat model of nerve gap injury showed increased regenerative bridge formation followed by enhancing the M2 macrophage phenotype as well as SC migration and involvement. However, it is unclear how this method would perform in comparison to the autograft controls.
A transected nerve that does not experience retraction can often be directly coapted. These nerve injuries receive far less research attention; however, some work has been done on materials to protect the anastomosis site, providing structural support and positive cues to the site of injury. Ren et al. explored the use of a fetal UBM wrap to repair no gap ocular nerve transections [161]. UBM from a fetal porcine source was frozen and thawed. Dense cellular layers of the urinary bladder were mechanically delaminated leaving the tunica mucosa and tunica propria. This was done as previously described [191]. However, Ren et al. used a vacuum chamber to rinse the bladder in sequential washes of nanopure water, 3% Triton X-100, and 3 M NaCl. The vacuum-assisted method was hypothesized to be more gentle than traditional agitation. Fetal UBM decellularized with this method possessed more dsDNA than UBM generated with agitation but was still significantly reduced from native. Used in a rat model of trigeminal, infraorbital nerve transection and direct coaptation, the fetal UBM showed enhanced angiogenesis, but the number of axons, myelination, and functional response were unaltered at 28 days post-surgery.
4.2. Crush injury model
Nerve injuries resulting from a crush or compression injury to the peripheral nervous system can result in neurapraxia or minor axonotmesis injuries, Sunderland’s first and second-degree injuries (Fig. 1). These types of injuries often fully recover in most patients and as such research into additional benefits of ECM use is limited. In cases of minor neurapraxia injuries, full recovery occurs within days, however for more severe neurapraxia or axonotmesis injuries recovery can take weeks to months. While outcomes are generally good for these patients, a limited number of ECM materials are still used to improve results in some cases.
The AxoGen® AxoGuard Nerve Protector, made from multiple porcine small intestine submucosal ECM (SIS) layers, is used clinically to wrap and protect injured peripheral nerves. This ECM sheet is used to shield the repair site from surrounding tissue, minimizing soft tissue attachments while allowing for diffusion of nutrients through the material. The SIS material has been used in a number of applications, including PNI [168]. As previously described, porcine SIS is commonly decellularized by mechanical delamination. Papatheodorou et al. examined the use of SIS in a retrospective clinical study of recurrent cubital tunnel syndrome. Outcomes for patients following decompression procedures is good, but scar tissue formation can cause recurrence of symptoms. Using the AxoGuard SIS Nerve wrap, patients had improved function accompanied by reduced pain and follow-up of 2–5 years did not reveal any recurrence of symptoms [170].
5. Regulatory pathways for dECM clinical translation
In general, any new biomedical therapy must be evaluated during pre-clinical animal studies prior to conducting clinical trials in human patients. Several dECM-based materials have been successfully commercialized and therefore may be used as a template to simplify the regulatory process [192]. Historically, most dECM products declare as 510(k) medical devices or biologics with the Food and Drug Administration (FDA). Products classified as medical devices or biologics must satisfy several pre-clinical and clinical requirements that unequivocally demonstrates safety and efficacy. After the pre-clinical studies have occured, either an Investigational Device Exemption (IDE), for medical devices, or an Investigational New Drug (IND) application, for biologics, must be approved by the FDA before proceeding into clinical studies, which can occur in several phases.
Each phase level is designed to carefully examine specific effects of the product after treating a set of eligible patients. Phase I generally consists of a small cohort of test subjects and is primarily focused on evaluating material safety and identifying any potential side effects. In Phase II, the number of test subjects increases and further examines the safety and efficacy. Phase III is used to directly compare the experimental material to similar products with a much larger test population to determine its effectiveness and compile safe-use information. The data obtained from the varying phase levels are then used to support a Biological Product License (BLA) or a Premarket Approval (PMA) application. Once the application is approved, the product can be marketed, as it is considered to have satisfied the FDA requirements.
Currently, only a few dECM products for nerve specific applications have made it to clinical trials. One of these products is the Avance® Nerve Graft from AxoGen, which is a decellularized human allograft used to treat PNI. AxoGen has completed two clinical studies for this product in addition to two ongoing studies. The first completed pilot study examined the effects of the Avance Nerve Graft on neurovascular bundle recovery in prostate cancer patients. The second pilot, CHANGE, was a prospective randomized controlled trial comparing Avance to hollow nerve conduits in the treatment of a nerve gap. Their graft showed a significant improvement in sensory function 6–12 months after treatment [193]. The success of the completed studies has allowed AxoGen to continue with their current ongoing clinical studies to gather more data to support their BLA submission. The RANGER study is a large multi-center retrospective study, consisting of up to 5000 test subjects, to assess patient outcomes up to 36 months post-treatment for PNI [194-196]. Additionally, the RECON study is a multi-center randomized prospective trial that will further evaluate the effectiveness of the Avance Nerve Graft. Other AxoGen products, such as the AxoGuard Nerve Protector, have shown improved preliminary clinical outcomes for the treatment of recurring cubital tunnel syndrome [170].
6. Challenges of using dECM materials to direct neural cell fate
Pre-clinical and clinical success of dECM materials in various disease models have made them an intriguing option for their application in the treatment of nervous system injuries. Although they appear to have many advantages, there are still several challenges remaining for the appropriate design and implementation of these materials. Below, we highlight some of the major limiting factors that must be considered to maximize the effectiveness of this emerging approach.
Tissue selection, based upon both source and age, can have a significant impact on dECM materials [197]. Site-specific tissues or organs harvested from exclusive regions are inherently different, as their microenvironments are designed to promote a specific cell function. Therefore, these native tissues have unique mechanical, biochemical and topographical properties that can illicit vastly different cellular responses. The age of the source tissue has been implicated in temporal changes to ECM composition, molecular profiles and mechanical properties, such as an increase in fibrosis, tissue stiffening and inflammation with aging [197]. Furthermore, if tissues are non-human, there are additional concerns related to species-specific differences that could negatively impact dECM material integration. For example, xenogeneic transplantation poses a risk of host immune rejection and incompatible homology [198]. The ECM composition of the sourced tissue must also be factored into the selection process. Specifically, the ECM proteins and proteoglycans that comprise healthy native tissues. As discussed above, CNS and PNS tissues have vastly different ECM contents and therefore, may require matching the appropriate ECM constituents to improve remodeling outcomes. Another related element to ECM organization is the material stiffness, porosity and fiber orientation, which have all been shown to impact cell fate [199].
Once the ideal source of tissue has been selected, the decellularization method, reagents and post-decellularization treatments can further enhance or deplete critical biological signatures, such as tissue-specific growth factors and hormones. The use of enzymatic digestion to form hydrogel-based dECM materials additionally alters ultrastructural and mechanical properties. Particularly, stiffness and local ECM fiber organization within a hydrogel can change dramatically in comparison to native tissues. After dECM materials are implanted, they undergo a natural degradation process, with degradation times varying as a function of concentration and material composition. In the case of CNS and PNS injury, scarring has been identified as a major barrier to repair. Therefore, dECM products must be carefully designed to optimize degradation times to promote regeneration of the damaged site.
Next generation dECM materials for nervous system injuries will require an in depth understanding of the diseased tissue and the ideal conditions that will initiate robust tissue repair. The goal is to create biomaterials that mimic healthy brain, spinal cord and peripheral nerve tissues. To accomplish this, researchers must consider testing a range of decellularization methods and techniques that preserve both the structural and functional components of the tissues. Tissue selection and processing should be carefully determined as well as the effects of dECM supplementation with growth factors and stem cells. Isolation and enrichment of the soluble fraction from decellularized nervous system tissues may also help stimulate functional regeneration. Finally, the addition of crosslinking agents and advancements in 3D printing may allow for improved tunability of the mechanical, topographical and structural properties of dECM materials with user-specified micropatterning and porosity to help direct neural cell fate.
7. Conclusion
An increasing number of disease models and therapies are incorporating dECM and its derivatives, due to their inherent ability to promote constructive remodeling of tissue [168,200]. The process of decellularization uses various treatments, including exposure to chemical agents, enzymes and/or mechanical means to strip away cells and lipids from a desired tissue leaving behind a non-immunogenic and functional scaffold [201]. The acellular scaffolds that remain offer a unique material consisting of cell-secreted structural ECM proteins, growth factors, hormones and other bioactive molecules that can provide a permissible environment to promote repair and remodeling after injury [202,203]. The application of this technique has shown promise for creating biologically active materials that may specifically support functional repair in the nervous system.
While regenerative capability in the CNS is poor, the application of decellularized constructs in the spinal cord have seen more success than in brain-specific applications [125]. Decellularized CNS ECM based scaffolds have shown the ability to modulate host cell phenotypes, differentiation, migration, proliferation, and other factors that can contribute to the constructive remodeling of CNS tissues [125]. Previous attempts to increase axonal elongation through the injury site have included the application of growth factors and material implantation, but these methods alone showed either random and aberrant axonal growth, or little functional return when used independently [130]. Since CNS injury often results in cavitation and scarring, therapies that bridge the lesion site are imperative [118]. dECM derived scaffolds can provide both a guidance substrate and a carrier of sequestered bioactive factors to promote normal axonal sprouting [130,140,204].
The application of dECM may also provide an alternative therapy to stimulate peripheral nerve regeneration. A significant advantage of using decellularization to generate acellular nerve grafts is the avoidance of donor-site morbidities, which are a direct consequence of using nerve autografts [205,206]. Additionally, the antigenic material removed from allografts during this process eliminates the need for immunosuppression after surgery [207]. Clinically, acellular allografts have shown the ability to improve both motor and sensory function in small nerve gaps (~3.2 cm on average) [208]. However, currently there is not a singular superior protocol for obtaining an ideal allograft, as chemically-distinct decellularization protocols in the literature have yielded varying results when compared directly [155]. This finding emphasizes the importance of selecting the appropriate reagents and treatments in order to optimize peripheral nerve recovery after injury. With improved knowledge of the explicit factors required to regenerate these tissues, decellularization methods may be tuned to support improved recovery in critical nerve defects.
Footnotes
Supplementary data to this article can be found online at https://doi.org/10.1016/j.ymeth.2019.07.023.
References
- [1].Bauchet L, et al. , Strategies for spinal cord repair after injury: a review of the literature and information, Ann. Phys. Rehabil. Med 52 (4) (2009) 330–351. [DOI] [PubMed] [Google Scholar]
- [2].Univ. of Alabama Birmingham, Facts and Figures at a Glance, National Spinal Cord Injury Statistical Center, 2018. [Google Scholar]
- [3].James SL, Global, regional, and national burden of traumatic brain injury and spinal cord injury, 1990–2016: a systematic analysis for the Global Burden of Disease Study 2016, Lancet Neurol. 18 (1) (2019) 56–87. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [4].Kouyoumdjian JA, Peripheral nerve injuries: a retrospective survey of 456 cases, Muscle Nerve 34 (6) (2006) 785–788. [DOI] [PubMed] [Google Scholar]
- [5].Lundborg G, Richard P, Bunge memorial lecture. Nerve injury and repair–a challenge to the plastic brain, J. Peripheral Nerv. Syst. JPNS 8 (4) (2003) 209–226. [DOI] [PubMed] [Google Scholar]
- [6].Taylor CA, et al. , The incidence of peripheral nerve injury in extremity trauma, Am. J. Phys. Med. Rehabil 87 (5) (2008) 381–385. [DOI] [PubMed] [Google Scholar]
- [7].Grinsell D, Keating CP, Peripheral nerve reconstruction after injury: a review of clinical and experimental therapies, Biomed. Res. Int 2014 (2014) 698256. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [8].Siemionow M, Brzezicki G, Chapter 8: current techniques and concepts in peripheral nerve repair, Int. Rev. Neurobiol 87 (2009) 141–172. [DOI] [PubMed] [Google Scholar]
- [9].Brattain K, Analysis of the Peripheral Nerve Repair Market in the United States, Minneapolis, MN. [Google Scholar]
- [10].Ruijs AC, et al. , Median and ulnar nerve injuries: a meta-analysis of predictors of motor and sensory recovery after modern microsurgical nerve repair, Plast. Reconstr. Surg 116 (2) (2005) pp. 484–94; discussion 495–6. [DOI] [PubMed] [Google Scholar]
- [11].Fitch MT, Silver J, CNS injury, glial scars, and inflammation: inhibitory extracellular matrices and regeneration failure, Exp. Neurol 209 (2) (2008) 294–301. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [12].Gaudet AD, Popovich PG, Extracellular matrix regulation of inflammation in the healthy and injured spinal cord, Exp. Neurol 258 (2014) 24–34. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [13].Maas AI, et al. , Moderate and severe traumatic brain injury in adults, Lancet Neurol. 7 (8) (2008) 728–741. [DOI] [PubMed] [Google Scholar]
- [14].Fitch MT, et al. , Cellular and molecular mechanisms of glial scarring and progressive cavitation: in vivo and in vitro analysis of inflammation-induced secondary injury after CNS trauma, J. Neurosci 19 (19) (1999) 8182–8198. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [15].Jiang T, et al. , Preparation and characterization of genipin-crosslinked rat acellular spinal cord scaffolds, Mater. Sci. Eng. C Mater. Biol. Appl 33 (6) (2013) 3514–3521. [DOI] [PubMed] [Google Scholar]
- [16].Sofroniew MV, Astrocyte barriers to neurotoxic inflammation, Nat. Rev. Neurosci 16 (5) (2015) 249–263. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [17].Sofroniew MV, Molecular dissection of reactive astrogliosis and glial scar formation, Trends Neurosci. 32 (12) (2009) 638–647. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [18].Sofroniew MV, Vinters HV, Astrocytes: biology and pathology, Acta Neuropathol. 119 (1) (2010) 7–35. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [19].Clements MP, et al. , The wound microenvironment reprograms Schwann cells to invasive mesenchymal-like cells to drive peripheral nerve regeneration, Neuron 96 (1) (2017) 98–114.e7. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [20].Allodi I, et al. , Specificity of peripheral nerve regeneration: interactions at the axon level, Prog. Neurobiol 98 (1) (2012) 16–37. [DOI] [PubMed] [Google Scholar]
- [21].Gonzalez-Forero D, Moreno-Lopez B, Retrograde response in axotomized motoneurons: nitric oxide as a key player in triggering reversion toward a dedifferentiated phenotype, Neuroscience 283 (2014) 138–165. [DOI] [PubMed] [Google Scholar]
- [22].Jessen KR, Mirsky R, The repair Schwann cell and its function in regenerating nerves, J. Physiol 594 (13) (2016) 3521–3531. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [23].Scheib J, Hoke A, Advances in peripheral nerve regeneration, Nat. Rev. Neurol 9 (12) (2013) 668–676. [DOI] [PubMed] [Google Scholar]
- [24].Cattin AL, Lloyd AC, The multicellular complexity of peripheral nerve regeneration, Curr. Opin. Neurobiol 39 (2016) 38–46. [DOI] [PubMed] [Google Scholar]
- [25].Roballo KCS, Bushman J, Evaluation of the host immune response and functional recovery in peripheral nerve autografts and allografts, Transpl. Immunol (2019). [DOI] [PubMed] [Google Scholar]
- [26].Cashman CR, Hoke A, Deficiency of adaptive immunity does not interfere with Wallerian degeneration, PLoS One 12 (5) (2017) e0177070. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [27].Eder M, et al. , Neurographic course Of Wallerian degeneration after human peripheral nerve injury, Muscle Nerve 56 (2) (2017) 247–252. [DOI] [PubMed] [Google Scholar]
- [28].Caillaud M, et al. , Peripheral nerve regeneration and intraneural revascularization, Neural Regen. Res 14 (1) (2019) 24–33. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [29].Cattin A-L, et al. , Macrophage-induced blood vessels guide schwann cell-mediated regeneration of peripheral nerves, Cell 162 (5) (2015) 1127–1139. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [30].Mokarram N, et al. , Effect of modulating macrophage phenotype on peripheral nerve repair, Biomaterials 33 (34) (2012) 8793–8801. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [31].Reuss BM, Glial Growth Factors (2009) 819–825. [Google Scholar]
- [32].Griffin MF, et al. , Peripheral nerve injury: principles for repair and regeneration, Open Orthopaedics J. 8 (2014) 199–203. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [33].Navarro X, Functional evaluation of peripheral nerve regeneration and target reinnervation in animal models: a critical overview, Eur. J. Neurosci 43 (3) (2016) 271–286. [DOI] [PubMed] [Google Scholar]
- [34].Sulaiman W, Gordon T, Neurobiology of peripheral nerve injury, regeneration, and functional recovery: from bench top research to bedside application, Ochsner J. 13 (1) (2013) 100–108. [PMC free article] [PubMed] [Google Scholar]
- [35].Pekny M, Nilsson M, Astrocyte activation and reactive gliosis, Glia 50 (4) (2005) 427–434. [DOI] [PubMed] [Google Scholar]
- [36].Ikezu T, Gendelman HE, Neuroimmun. Pharmacol (2017). [Google Scholar]
- [37].Kacem K, et al. , Structural organization of the perivascular astrocyte endfeet and their relationship with the endothelial glucose transporter: a confocal microscopy study, Glia 23 (1) (1998) 1–10. [PubMed] [Google Scholar]
- [38].Nagelhus EA, et al. , Aquaporin-4 in the central nervous system: cellular and subcellular distribution and coexpression with KIR4.1, Neuroscience 129 (4) (2004) 905–913. [DOI] [PubMed] [Google Scholar]
- [39].Hertz L, Zielke HR, Astrocytic control of glutamatergic activity: astrocytes as stars of the show, Trends Neurosci. 27 (12) (2004) 735–743. [DOI] [PubMed] [Google Scholar]
- [40].Bak LK, et al. , The glutamate/GABA-glutamine cycle: aspects of transport, neurotransmitter homeostasis and ammonia transfer, J. Neurochem 98 (3) (2006) 641–653. [DOI] [PubMed] [Google Scholar]
- [41].Wiesinger H, et al. , Metabolic pathways for glucose in astrocytes, Glia 21 (1) (1997) 22–34. [DOI] [PubMed] [Google Scholar]
- [42].Eddleston M, Mucke L, Molecular profile of reactive astrocytes–implications for their role in neurologic disease, Neuroscience 54 (1) (1993) 15–36. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [43].Silver J, Miller JH, Regeneration beyond the glial scar, Nat. Rev. Neurosci 5 (2) (2004) 146–156. [DOI] [PubMed] [Google Scholar]
- [44].Corfas G, et al. , Mechanisms and roles of axon-Schwann cell interactions, J. Neurosci 24 (42) (2004) 9250–9260. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [45].Jessen KR, Mirsky R, Why do schwann cells survive in the absence of axons? Ann. N. Y. Acad. Sci 883 (1) (1999) 109–115. [DOI] [PubMed] [Google Scholar]
- [46].Brown BN, Badylak SF, Extracellular matrix as an inductive scaffold for functional tissue reconstruction, Transl. Res 163 (4) (2014) 268–285. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [47].Bignami A, et al. , Hyaluronic acid and hyaluronic acid-binding proteins in brain extracellular matrix, Anat. Embryol 188 (5) (1993) 419–433. [DOI] [PubMed] [Google Scholar]
- [48].Almutairi MM, et al. , Factors controlling permeability of the blood-brain barrier, Cell Mol. Life Sci 73 (1) (2016) 57–77. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [49].George N, Geller HM, Extracellular matrix and traumatic brain injury, J Neurosci Res 96 (4) (2018) 573–588. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [50].Rutka JT, et al. , The extracellular matrix of the central and peripheral nervous systems: structure and function, J. Neurosurg 69 (2) (1988) 155–170. [DOI] [PubMed] [Google Scholar]
- [51].Bianchi ME, DAMPs, PAMPs and alarmins: all we need to know about danger, J. Leukoc Biol 81 (1) (2007) 1–5. [DOI] [PubMed] [Google Scholar]
- [52].Piccinini AM, Midwood KS, DAMPening inflammation by modulating TLR signalling, Mediat. Inflamm 2010 (2010). [DOI] [PMC free article] [PubMed] [Google Scholar]
- [53].Galtrey CM, et al. , Distribution and synthesis of extracellular matrix proteoglycans, hyaluronan, link proteins and tenascin-R in the rat spinal cord, Eur. J. Neurosci 27 (6) (2008) 1373–1390. [DOI] [PubMed] [Google Scholar]
- [54].Garcia-Alias G, Fawcett JW, Training and anti-CSPG combination therapy for spinal cord injury, Exp. Neurol 235 (1) (2012) 26–32. [DOI] [PubMed] [Google Scholar]
- [55].Edgerton VR, et al. , Plasticity of the spinal neural circuitry after injury, Annu. Rev. Neurosci 27 (2004) 145–167. [DOI] [PubMed] [Google Scholar]
- [56].Nudo R, Adaptive plasticity in motor cortex: implications for rehabilitation after brain injury, J. Rehabil. Med 35 (2003) 7–10. [DOI] [PubMed] [Google Scholar]
- [57].Rossignol S, et al. , Adaptive changes of locomotion after central and peripheral lesions, Can. J. Physiol. Pharmacol 82 (8–9) (2004) 617–627. [DOI] [PubMed] [Google Scholar]
- [58].Wolpaw JR, Tennissen AM, Activity-dependent spinal cord plasticity in health and disease, Annu. Rev. Neurosci 24 (2001) 807–843. [DOI] [PubMed] [Google Scholar]
- [59].Bradbury EJ, et al. , Chondroitinase ABC promotes functional recovery after spinal cord injury, Nature 416 (6881) (2002) 636–640. [DOI] [PubMed] [Google Scholar]
- [60].Caggiano AO, et al. , Chondroitinase ABCI improves locomotion and bladder function following contusion injury of the rat spinal cord, J. Neurotrauma 22 (2) (2005) 226–239. [DOI] [PubMed] [Google Scholar]
- [61].Galtrey CM, Fawcett JW, The role of chondroitin sulfate proteoglycans in regeneration and plasticity in the central nervous system, Brain Res. Rev 54 (1) (2007) 1–18. [DOI] [PubMed] [Google Scholar]
- [62].Massey JM, et al. , Chondroitinase ABC digestion of the perineuronal net promotes functional collateral sprouting in the cuneate nucleus after cervical spinal cord injury, J. Neurosci 26 (16) (2006) 4406–4414. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [63].Chernousov MA, et al. , Regulation of Schwann cell function by the extracellular matrix, Glia 56 (14) (2008) 1498–1507. [DOI] [PubMed] [Google Scholar]
- [64].Deumens R, et al. , Repairing injured peripheral nerves: bridging the gap, Prog. Neurobiol 92 (3) (2010) 245–276. [DOI] [PubMed] [Google Scholar]
- [65].Kemp SW, et al. , Growth factor and stem cell enhanced conduits in peripheral nerve regeneration and repair, Neurol. Res 30 (10) (2008) 1030–1038. [DOI] [PubMed] [Google Scholar]
- [66].Zhang Y, et al. , A nerve graft constructed with xenogeneic acellular nerve matrix and autologous adipose-derived mesenchymal stem cells, Biomaterials 31 (20) (2010) 5312–5324. [DOI] [PubMed] [Google Scholar]
- [67].Gao X, et al. , The role of peripheral nerve ECM components in the tissue engineering nerve construction, Rev. Neurosci 24 (4) (2013) 443–453. [DOI] [PubMed] [Google Scholar]
- [68].Palm SL, Furcht LT, Production of laminin and fibronectin by Schwannoma cells: cell-protein interactions in vitro and protein localization in peripheral nerve in vivo, J. Cell Biol 96 (5) (1983) 1218–1226. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [69].Plantman S, et al. , Integrin-laminin interactions controlling neurite outgrowth from adult DRG neurons in vitro, Mol. Cell. Neurosci 39 (1) (2008) 50–62. [DOI] [PubMed] [Google Scholar]
- [70].Wood MD, Willits RK, Applied electric field enhances DRG neurite growth: influence of stimulation media, surface coating and growth supplements, J. Neural Eng 6 (4) (2009) 046003. [DOI] [PubMed] [Google Scholar]
- [71].Guan W, et al. , Sensory neuron subtypes have unique substratum preference and receptor expression before target innervation, J. Neurosci 23 (5) (2003) 1781–1791. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [72].Ughrin YM, et al. , Multiple regions of the NG2 proteoglycan inhibit neurite growth and induce growth cone collapse, J. Neurosci 23 (1) (2003) 175–186. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [73].Balgude AP, et al. , Agarose gel stiffness determines rate of DRG neurite extension in 3D cultures, Biomaterials 22 (10) (2001) 1077–1084. [DOI] [PubMed] [Google Scholar]
- [74].Willits RK, Skornia SL, Effect of collagen gel stiffness on neurite extension, J. Biomater. Sci. Polym. Ed 15 (12) (2004) 1521–1531. [DOI] [PubMed] [Google Scholar]
- [75].Hall S, Axonal regeneration through acellular muscle grafts, J. Anat 190 (Pt 1) (1997) 57–71. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [76].Krekoski CA, et al. , Axonal regeneration into acellular nerve grafts is enhanced by degradation of chondroitin sulfate proteoglycan, J. Neurosci 21 (16) (2001) 6206–6213. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [77].Udina E, et al. , Rolipram-induced elevation of cAMP or chondroitinase ABC breakdown of inhibitory proteoglycans in the extracellular matrix promotes peripheral nerve regeneration, Exp. Neurol 223 (1) (2010) 143–152. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [78].Bailey SB, et al. , The influence of fibronectin and laminin during Schwann cell migration and peripheral nerve regeneration through silicon chambers, J. Neurocytol 22 (3) (1993) 176–184. [DOI] [PubMed] [Google Scholar]
- [79].Chen YS, et al. , Peripheral nerve regeneration using silicone rubber chambers filled with collagen, laminin and fibronectin, Biomaterials 21 (15) (2000) 1541–1547. [DOI] [PubMed] [Google Scholar]
- [80].Labrador RO, et al. , Influence of collagen and laminin gels concentration on nerve regeneration after resection and tube repair, Exp. Neurol 149 (1) (1998) 243–252. [DOI] [PubMed] [Google Scholar]
- [81].David S, et al. , Macrophage and microglial plasticity in the injured spinal cord, Neuroscience 307 (2015) 311–318. [DOI] [PubMed] [Google Scholar]
- [82].David S, Kroner A, Repertoire of microglial and macrophage responses after spinal cord injury, Nat. Rev. Neurosci 12 (7) (2011) 388–399. [DOI] [PubMed] [Google Scholar]
- [83].Gensel JC, Zhang B, Macrophage activation and its role in repair and pathology after spinal cord injury, Brain Res. 1619 (2015) 1–11. [DOI] [PubMed] [Google Scholar]
- [84].Kong X, Gao J, Macrophage polarization: a key event in the secondary phase of acute spinal cord injury, J. Cell Mol. Med 21 (5) (2017) 941–954. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [85].Shechter R, et al. , Infiltrating blood-derived macrophages are vital cells playing an anti-inflammatory role in recovery from spinal cord injury in mice, PLoS Med. 6 (7) (2009) e1000113. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [86].Shechter R, et al. , Recruitment of beneficial M2 macrophages to injured spinal cord is orchestrated by remote brain choroid plexus, Immunity 38 (3) (2013) 555–569. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [87].Shechter R, et al. , The glial scar-monocyte interplay: a pivotal resolution phase in spinal cord repair, PLoS One 6 (12) (2011) e27969. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [88].Corps KN, et al. , Inflammation and neuroprotection in traumatic brain injury, JAMA Neurol. 72 (3) (2015) 355–362. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [89].Donat CK, et al. , Microglial activation in traumatic brain injury, Front. Aging Neurosci 9 (2017) 208. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [90].Kumar A, et al. , Microglial/macrophage polarization dynamics following traumatic brain injury, J. Neurotrauma 33 (19) (2016) 1732–1750. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [91].Mishra SK, et al. , Early monitoring and quantitative evaluation of macrophage infiltration after experimental traumatic brain injury: a magnetic resonance imaging and flow cytometric analysis, Mol. Cell. Neurosci 78 (2017) 25–34. [DOI] [PubMed] [Google Scholar]
- [92].Chen P, et al. , Role of macrophages in Wallerian degeneration and axonal regeneration after peripheral nerve injury, Acta Neuropathol. 130 (5) (2015) 605–618. [DOI] [PubMed] [Google Scholar]
- [93].Rosenberg AF, et al. , In vivo nerve-macrophage interactions following peripheral nerve injury, J. Neurosci 32 (11) (2012) 3898–3909. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [94].Tomlinson JE, et al. , Temporal changes in macrophage phenotype after peripheral nerve injury, J. Neuroinflam 15 (1) (2018) 185. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [95].Ginhoux F, et al. , Fate Mapping Analysis Reveals that Adult Microglia Derive from Primitive Macrophages 330 (2010) 841–845. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [96].Katsumoto A, et al. , Ontogeny and functions of central nervous system macrophages, J. Immunol. (Baltimore, Md. 1950) 193 (6) (2014) 2615–2621. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [97].Prinz M, et al. , Ontogeny and homeostasis of CNS myeloid cells, Nat. Immunol 18 (4) (2017) 385–392. [DOI] [PubMed] [Google Scholar]
- [98].Brosnan CF, et al. , Reactive nitrogen intermediates in human neuropathology: an overview, Dev. Neurosci 16 (3–4) (1994) 152–161. [DOI] [PubMed] [Google Scholar]
- [99].Gendelman HE, et al. , An experimental model system for HIV-1-induced brain injury, Adv. Neuroimmunol 4 (3) (1994) 189–193. [DOI] [PubMed] [Google Scholar]
- [100].Meeuwsen S, et al. , Cytokine, chemokine and growth factor gene profiling of cultured human astrocytes after exposure to proinflammatory stimuli, Glia 43 (3) (2003) 243–253. [DOI] [PubMed] [Google Scholar]
- [101].Van Wagoner NJ, et al. , Interleukin-6 (IL-6) production by astrocytes: autocrine regulation by IL-6 and the soluble IL-6 receptor, J. Neurosci 19 (13) (1999) 5236–5244. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [102].Grunblatt E, et al. , Neuroprotective strategies in Parkinson's disease using the models of 6-hydroxydopamine and MPTP, Ann. N. Y. Acad. Sci 899 (2000) 262–273. [DOI] [PubMed] [Google Scholar]
- [103].Liu B, et al. , Role of nitric oxide in inflammation-mediated neurodegeneration, Ann. N. Y. Acad. Sci 962 (2002) 318–331. [DOI] [PubMed] [Google Scholar]
- [104].Morale MC, et al. , Estrogen, neuroinflammation and neuroprotection in Parkinson's disease: glia dictates resistance versus vulnerability to neurodegeneration, Neuroscience 138 (3) (2006) 869–878. [DOI] [PubMed] [Google Scholar]
- [105].Kadiu I, et al. , Mononuclear phagocytes in the pathogenesis of neurodegenerative diseases, Neurotox. Res 8 (1–2) (2005) 25–50. [DOI] [PubMed] [Google Scholar]
- [106].Liu B, Hong JS, Role of microglia in inflammation-mediated neurodegenerative diseases: mechanisms and strategies for therapeutic intervention, J. Pharmacol. Experim. Therap 304 (1) (2003) 1–7. [DOI] [PubMed] [Google Scholar]
- [107].Yoles E, Schwartz M, Degeneration of spared axons following partial white matter lesion: implications for optic nerve neuropathies, Exp. Neurol 153 (1) (1998) 1–7. [DOI] [PubMed] [Google Scholar]
- [108].Mueller M, et al. , Macrophage response to peripheral nerve injury: the quantitative contribution of resident and hematogenous macrophages, Lab. Invest 83 (2) (2003) 175–185. [DOI] [PubMed] [Google Scholar]
- [109].Griffin JW, et al. , Macrophage systems in peripheral nerves. A review, J. Neuropathol. Experim. Neurol 52 (6) (1993) 553–560. [DOI] [PubMed] [Google Scholar]
- [110].Lunn ER, et al. , Absence of wallerian degeneration does not hinder regeneration in peripheral nerve, Eur. J. Neurosci 1 (1) (1989) 27–33. [DOI] [PubMed] [Google Scholar]
- [111].Beuche W, Friede RL, The role of non-resident cells in Wallerian degeneration, J. Neurocytol 13 (5) (1984) 767–796. [DOI] [PubMed] [Google Scholar]
- [112].Beuche W, Friede RL, Myelin phagocytosis in Wallerian degeneration of peripheral nerves depends on silica-sensitive, bg/bg-negative and Fc-positive monocytes, Brain Res. 378 (1) (1986) 97–106. [DOI] [PubMed] [Google Scholar]
- [113].Prineas JW, Demyelination and remyelination in recurrent idiopathic polyneuropathy. An electron microscope study, Acta Neuropathol. 18 (1) (1971) 34–57. [DOI] [PubMed] [Google Scholar]
- [114].Brown BN, et al. , Macrophage phenotype as a predictor of constructive remodeling following the implantation of biologically derived surgical mesh materials, Acta Biomater. 8 (3) (2012) 978–987. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [115].Meng FW, et al. , Solubilized extracellular matrix from brain and urinary bladder elicits distinct functional and phenotypic responses in macrophages, Biomaterials 46 (2015) 131–140. [DOI] [PubMed] [Google Scholar]
- [116].Brown BN, et al. , Macrophage phenotype and remodeling outcomes in response to biologic scaffolds with and without a cellular component, Biomaterials 30 (8) (2009) 1482–1491. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [117].Valentin JE, et al. , Macrophage participation in the degradation and remodeling of extracellular matrix scaffolds, Tissue Eng. Part A 15 (7) (2009) 1687–1694. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [118].Zhang XY, et al. , Chemically extracted acellular muscle: a new potential scaffold for spinal cord injury repair, J. Biomed. Mater. Res. A 100 (3) (2012) 578–587. [DOI] [PubMed] [Google Scholar]
- [119].DeQuach JA, et al. , Decellularized porcine brain matrix for cell culture and tissue engineering scaffolds, Tissue Eng. Part A 17 (21–22) (2011) 2583–2592. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [120].Zhu T, et al. , An acellular cerebellar biological scaffold: preparation, characterization, biocompatibility and effects on neural stem cells, Brain Res. Bull 113 (2015) 48–57. [DOI] [PubMed] [Google Scholar]
- [121].Boriani F, et al. , A novel technique for decellularization of allogenic nerves and in vivo study of their use for peripheral nerve reconstruction, J. Biomed. Mater Res. A 105 (8) (2017) 2228–2240. [DOI] [PubMed] [Google Scholar]
- [122].Chen CC, et al. , The physicochemical properties of decellularized extracellular matrix-coated 3d printed poly(epsilon-caprolactone) nerve conduits for promoting schwann cells proliferation and differentiation, Materials (Basel) 11 (9) (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
- [123].Sun X, et al. , Acellular Cauda Equina allograft as main material combined with biodegradable chitin conduit for regeneration of long-distance sciatic nerve defect in rats, Adv. Healthc. Mater 7 (17) (2018) e1800276. [DOI] [PubMed] [Google Scholar]
- [124].Wakimura Y, et al. , An experimental study to bridge a nerve gap with a decellularized allogeneic nerve, Plast Reconstr. Surg 136 (3) (2015) 319e–327e. [DOI] [PubMed] [Google Scholar]
- [125].Wang H, et al. , Decellularization technology in CNS tissue repair, Expert Rev. Neurother 15 (5) (2015) 493–500. [DOI] [PubMed] [Google Scholar]
- [126].Zilic L, et al. , Decellularisation and histological characterisation of porcine peripheral nerves, Biotechnol. Bioeng 113 (9) (2016) 2041–2053. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [127].Baiguera S, et al. , Electrospun gelatin scaffolds incorporating rat decellularized brain extracellular matrix for neural tissue engineering, Biomaterials 35 (4) (2014) 1205–1214. [DOI] [PubMed] [Google Scholar]
- [128].Crapo PM, et al. , Biologic scaffolds composed of central nervous system extracellular matrix, Biomaterials 33 (13) (2012) 3539–3547. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [129].De Waele J, et al. , 3D culture of murine neural stem cells on decellularized mouse brain sections, Biomaterials 41 (2015) 122–131. [DOI] [PubMed] [Google Scholar]
- [130].Guo SZ, et al. , Preparation of the acellular scaffold of the spinal cord and the study of biocompatibility, Spinal Cord 48 (7) (2010) 576–581. [DOI] [PubMed] [Google Scholar]
- [131].Su LW, Hai-yong He, Wang Hui, Mei-qin Cai, Jin Gong, Ying Guo, Preparation and cytocompatibility of acellular rat brain matrix for tissue engineering scaffolds, J. Sun Yat-sen Univ. (Med. Sci.) 33 (1) (2012). [Google Scholar]
- [132].Lin Q, et al. , Enhanced neuroprotection with decellularized brain extracellular matrix containing bFGF after intracerebral transplantation in Parkinson's disease rat model, Int. J. Pharm 517 (1–2) (2017) 383–394. [DOI] [PubMed] [Google Scholar]
- [133].Liu J, et al. , Acellular spinal cord scaffold seeded with mesenchymal stem cells promotes long-distance axon regeneration and functional recovery in spinal cord injured rats, J. Neurol. Sci 325 (1–2) (2013) 127–136. [DOI] [PubMed] [Google Scholar]
- [134].Medberry CJ, et al. , Hydrogels derived from central nervous system extracellular matrix, Biomaterials 34 (4) (2013) 1033–1040. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [135].Ribatti D, et al. , Angiogenic response induced by acellular brain scaffolds grafted onto the chick embryo chorioallantoic membrane, Brain Res. 989 (1) (2003) 9–15. [DOI] [PubMed] [Google Scholar]
- [136].Sood D, et al. , Fetal brain extracellular matrix boosts neuronal network formation in 3D bioengineered model of cortical brain tissue, ACS Biomater. Sci. Eng 2 (1) (2016) 131–140. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [137].Tukmachev D, et al. , Injectable extracellular matrix hydrogels as scaffolds for spinal cord injury repair, Tissue Eng. Part A 22 (3–4) (2016) 306–317. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [138].Wang Q, et al. , The preparation and comparison of decellularized nerve scaffold of tissue engineering, J. Biomed. Mater. Res. A 102 (12) (2014) 4301–4308. [DOI] [PubMed] [Google Scholar]
- [139].Wu Y, et al. , Implantation of brain-derived extracellular matrix enhances neurological recovery after traumatic brain injury, Cell Transplant 26 (7) (2017) 1224–1234. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [140].Xu HL, et al. , Thermo-sensitive hydrogels combined with decellularised matrix deliver bFGF for the functional recovery of rats after a spinal cord injury, Sci. Rep 6 (2016) 38332. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [141].Yin WH, et al. , Effects of mechanical vibration on the morphology of the acellular scaffold for the spinal cord, Nan fang yi ke da xue xue bao = J. Southern Med. Univ 28 (10) (2008) 1748–1751. [PubMed] [Google Scholar]
- [142].Cai M, et al. , Role of demyelination efficiency within acellular nerve scaffolds during nerve regeneration across peripheral defects, Biomed. Res. Int 2017 (2017) 4606387. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [143].Gao S, et al. , Comparison of morphology and biocompatibility of acellular nerve scaffolds processed by different chemical methods, J. Mater. Sci. Mater. Med 25 (5) (2014) 1283–1291. [DOI] [PubMed] [Google Scholar]
- [144].Pollins AC, et al. , Comparing processed nerve allografts and assessing their capacity to retain and release nerve growth factor, Ann. Plast. Surg 81 (2) (2018) 198–202. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [145].Prest TA, et al. , Nerve-specific, xenogeneic extracellular matrix hydrogel promotes recovery following peripheral nerve injury, J. Biomed. Mater. Res. A 106 (2) (2018) 450–459. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [146].Sondell M, et al. , Regeneration of the rat sciatic nerve into allografts made acellular through chemical extraction, Brain Res. 795 (1–2) (1998) 44–54. [DOI] [PubMed] [Google Scholar]
- [147].Sridharan R, et al. , Decellularized grafts with axially aligned channels for peripheral nerve regeneration, J. Mech. Behav. Biomed. Mater 41 (2015) 124–135. [DOI] [PubMed] [Google Scholar]
- [148].Wen X, et al. , Cauda equina-derived extracellular matrix for fabrication of nanostructured hybrid scaffolds applied to neural tissue engineering, Tissue Eng. Part A 21 (5–6) (2015) 1095–1105. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [149].Giusti G, et al. , Return of motor function after segmental nerve loss in a rat model: comparison of autogenous nerve graft, collagen conduit, and processed allograft (AxoGen), J Bone Joint Surg. Am 94 (5) (2012) 410–417. [DOI] [PubMed] [Google Scholar]
- [150].Hudson TW, et al. , Engineering an improved acellular nerve graft via optimized chemical processing, Tissue Eng. 10 (9–10) (2004) 1346–1358. [DOI] [PubMed] [Google Scholar]
- [151].Hudson TW, et al. , Optimized acellular nerve graft is immunologically tolerated and supports regeneration, Tissue Eng. 10 (11–12) (2004) 1641–1651. [DOI] [PubMed] [Google Scholar]
- [152].Hundepool CA, et al. , Optimizing decellularization techniques to create a new nerve allograft: an in vitro study using rodent nerve segments, Neurosurg. Focus 42 (3) (2017) E4. [DOI] [PubMed] [Google Scholar]
- [153].Isaacs J, Safa B, A preliminary assessment of the utility of large-caliber processed nerve allografts for the repair of upper extremity nerve injuries, Hand (N Y) 12 (1) (2017) 55–59. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [154].Junka R, Yu X, Novel acellular scaffold made from decellularized schwann cell sheets for peripheral nerve regeneration, Regen Eng Transl Med 1 (1) (2015) 22–31. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [155].Moore AM, et al. , Acellular nerve allografts in peripheral nerve regeneration: a comparative study, Muscle Nerve 44 (2) (2011) 221–234. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [156].Rinker B, et al. , Use of processed nerve allografts to repair nerve injuries greater than 25 mm in the hand, Ann. Plast. Surg 78 (6S Suppl 5) (2017) S292–S295. [DOI] [PubMed] [Google Scholar]
- [157].Yampolsky A, et al. , Efficacy of acellular nerve allografts in trigeminal nerve reconstruction, J. Oral Maxillofac. Surg 75 (10) (2017) 2230–2234. [DOI] [PubMed] [Google Scholar]
- [158].Kim JK, et al. , Development of a decellularization method to produce nerve allografts using less invasive detergents and hyper/hypotonic solutions, J. Plast. Reconstr. Aesthet. Surg 69 (12) (2016) 1690–1696. [DOI] [PubMed] [Google Scholar]
- [159].Philips C, et al. , Qualitative and quantitative evaluation of a novel detergent-based method for decellularization of peripheral nerves, Ann. Biomed. Eng 46 (11) (2018) 1921–1937. [DOI] [PubMed] [Google Scholar]
- [160].Philips C, et al. , Evaluation methods as quality control in the generation of decellularized peripheral nerve allografts, J. Neural Eng 15 (2) (2018) 021003. [DOI] [PubMed] [Google Scholar]
- [161].Ren T, et al. , Fetal extracellular matrix nerve wraps locally improve peripheral nerve remodeling after complete transection and direct repair in rat, Sci. Rep 8 (1) (2018) 4474. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [162].Wang W, et al. , Comparative study of the efficacy of decellularization treatment of allogenic and xenogeneic nerves as nerve conduits, J. Biomed. Mater. Res. A 104 (2) (2016) 445–454. [DOI] [PubMed] [Google Scholar]
- [163].Cerqueira SR, et al. , Decellularized peripheral nerve supports Schwann cell transplants and axon growth following spinal cord injury, Biomaterials 177 (2018) 176–185. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [164].Ghuman H, et al. , Long-term retention of ECM hydrogel after implantation into a sub-acute stroke cavity reduces lesion volume, Acta Biomater. 63 (2017) 50–63. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [165].Ghuman H, et al. , ECM hydrogel for the treatment of stroke: characterization of the host cell infiltrate, Biomaterials 91 (2016) 166–181. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [166].Wang JY, et al. , Neurorestorative effect of urinary bladder matrix-mediated neural stem cell transplantation following traumatic brain injury in rats, CNS Neurol. Disord.: Drug Targets 12 (3) (2013) 413–425. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [167].Zhang L, et al. , Effect of an inductive hydrogel composed of urinary bladder matrix upon functional recovery following traumatic brain injury, Tissue Eng Part A 19 (17–18) (2013) 1909–1918. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [168].Badylak SF, et al. , Extracellular matrix as a biological scaffold material: structure and function, Acta Biomater. 5 (1) (2009) 1–13. [DOI] [PubMed] [Google Scholar]
- [169].Kokkalis ZT, et al. , Assessment of processed porcine extracellular matrix as a protective barrier in a rabbit nerve wrap model, J. Reconstr. Microsurg 27 (1) (2011) 19–28. [DOI] [PubMed] [Google Scholar]
- [170].Papatheodorou LK, et al. , Preliminary results of recurrent cubital tunnel syndrome treated with neurolysis and porcine extracellular matrix nerve wrap, J. Hand Surg. Am 40 (5) (2015) 987–992. [DOI] [PubMed] [Google Scholar]
- [171].Cornelison RC, et al. , Development of an apoptosis-assisted decellularization method for maximal preservation of nerve tissue structure, Acta Biomater. 77 (2018) 116–126. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [172].Boyer RB, et al. , Adjuvant neurotrophic factors in peripheral nerve repair with chondroitin sulfate proteoglycan-reduced acellular nerve allografts, J. Surg. Res 193 (2) (2015) 969–977. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [173].Neubauer D, et al. , Chondroitinase treatment increases the effective length of acellular nerve grafts, Exp. Neurol 207 (1) (2007) 163–170. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [174].Seif-Naraghi SB, et al. , Design and characterization of an injectable pericardial matrix gel: a potentially autologous scaffold for cardiac tissue engineering, Tissue Eng. Part A 16 (6) (2010) 2017–2027. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [175].Sullivan DC, et al. , Decellularization methods of porcine kidneys for whole organ engineering using a high-throughput system, Biomaterials 33 (31) (2012) 7756–7764. [DOI] [PubMed] [Google Scholar]
- [176].Piccoli M, et al. , Improvement of diaphragmatic performance through orthotopic application of decellularized extracellular matrix patch, Biomaterials 74 (2016) 245–255. [DOI] [PubMed] [Google Scholar]
- [177].Elder BD, et al. , Developing an articular cartilage decellularization process toward facet joint cartilage replacement, Neurosurgery 66 (4) (2010) pp. 722–7; discussion 727. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [178].Stern MM, et al. , The influence of extracellular matrix derived from skeletal muscle tissue on the proliferation and differentiation of myogenic progenitor cells ex vivo, Biomaterials 30 (12) (2009) 2393–2399. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [179].Uriel S, et al. , Extraction and assembly of tissue-derived gels for cell culture and tissue engineering, Tissue Eng. C Meth 15 (3) (2009) 309–321. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [180].Funamoto S, et al. , The use of high-hydrostatic pressure treatment to decellularize blood vessels, Biomaterials 31 (13) (2010) 3590–3595. [DOI] [PubMed] [Google Scholar]
- [181].Mligiliche N, et al. , Grafting of detergent-denatured skeletal muscles provides effective conduits for extension of regenerating axons in the rat sciatic nerve, Arch. Histol. Cytol 64 (1) (2001) 29–36. [DOI] [PubMed] [Google Scholar]
- [182].Meezan E, et al. , A simple, versatile, nondisruptive method for the isolation of morphologically and chemically pure basement membranes from several tissues, Life Sci. 17 (11) (1975) 1721–1732. [DOI] [PubMed] [Google Scholar]
- [183].Liang HC, et al. , Effects of crosslinking degree of an acellular biological tissue on its tissue regeneration pattern, Biomaterials 25 (17) (2004) 3541–3552. [DOI] [PubMed] [Google Scholar]
- [184].Huang LL, et al. , Biocompatibility study of a biological tissue fixed with a naturally occurring crosslinking reagent, J. Biomed. Mater. Res 42 (4) (1998) 568–576. [DOI] [PubMed] [Google Scholar]
- [185].Speer DP, et al. , Biological effects of residual glutaraldehyde in glutaraldehydetanned collagen biomaterials, J. Biomed. Mater. Res 14 (6) (1980) 753–764. [DOI] [PubMed] [Google Scholar]
- [186].Vasudevan S, et al. , Detergent-free decellularized nerve grafts for long-gap peripheral nerve reconstruction, Plast. Reconstr. Surg. Glob Open 2 (8) (2014) e201. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [187].Roosens A, et al. , Impact of detergent-based decellularization methods on porcine tissues for heart valve engineering, Ann. Biomed. Eng 44 (9) (2016) 2827–2839. [DOI] [PubMed] [Google Scholar]
- [188].Sondell M, et al. , Vascular endothelial growth factor stimulates Schwann cell invasion and neovascularization of acellular nerve grafts, Brain Res. 846 (2) (1999) 219–228. [DOI] [PubMed] [Google Scholar]
- [189].Lin T, et al. , Hydrogel derived from porcine decellularized nerve tissue as a promising biomaterial for repairing peripheral nerve defects, Acta Biomater. 73 (2018) 326–338. [DOI] [PubMed] [Google Scholar]
- [190].IJkeme-Paassen J, et al. , Transection of peripheral nerves, bridging strategies and effect evaluation, Biomaterials 25 (9) (2004) 1583–1592. [DOI] [PubMed] [Google Scholar]
- [191].Marcal H, et al. , A comprehensive protein expression profile of extracellular matrix biomaterial derived from porcine urinary bladder, Regen. Med 7 (2) (2012) 159–166. [DOI] [PubMed] [Google Scholar]
- [192].Nakamura N, et al. , Overview of the development, applications, and future perspectives of decellularized tissues and organs, ACS Biomater. Sci. Eng 3 (7) (2017) 1236–1244. [DOI] [PubMed] [Google Scholar]
- [193].Means KR Jr et al. , A multicenter, prospective, randomized, pilot study of outcomes for digital nerve repair in the hand using hollow conduit compared with processed allograft nerve, Hand (N Y) 11 (2) (2016) 144–151. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [194].Brooks DN, et al. , Processed nerve allografts for peripheral nerve reconstruction: a multicenter study of utilization and outcomes in sensory, mixed, and motor nerve reconstructions, Microsurgery 32 (1) (2012) 1–14. [DOI] [PubMed] [Google Scholar]
- [195].Cho MS, et al. , Functional outcome following nerve repair in the upper extremity using processed nerve allograft, J. Hand Surg. Am 37 (11) (2012) 2340–2349. [DOI] [PubMed] [Google Scholar]
- [196].Rinker BD, et al. , Outcomes of short-gap sensory nerve injuries reconstructed with processed nerve allografts from a multicenter registry study, J. Reconstr. Microsurg 31 (5) (2015) 384–390. [DOI] [PubMed] [Google Scholar]
- [197].Spang MT, Christman KL, Extracellular matrix hydrogel therapies: in vivo applications and development, Acta Biomater. 68 (2018) 1–14. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [198].Stahl EC, et al. , Evaluation of the host immune response to decellularized lung scaffolds derived from alpha-Gal knockout pigs in a non-human primate model, Biomaterials 187 (2018) 93–104. [DOI] [PubMed] [Google Scholar]
- [199].Kumar A, et al. , Understanding the extracellular forces that determine cell fate and maintenance, Development (Cambridge, England) 144 (23) (2017) 4261–4270. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [200].Jensen G, et al. , 3D tissue engineering, an emerging technique for pharmaceutical research, Acta Pharm. Sin. B 8 (5) (2018) 756–766. [DOI] [PMC free article] [PubMed] [Google Scholar]
- [201].Keane TJ, et al. , Methods of tissue decellularization used for preparation of biologic scaffolds and in vivo relevance, Methods 84 (2015) 25–34. [DOI] [PubMed] [Google Scholar]
- [202].Maghsoudlou P, et al. , A decellularization methodology for the production of a natural acellular intestinal matrix, J. Vis. Exp 80 (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]
- [203].Meng F, et al. , Biologic scaffold for CNS repair, Regen. Med 9 (3) (2014) 367–383. [DOI] [PubMed] [Google Scholar]
- [204].David S, Aguayo AJ, Axonal elongation into peripheral nervous-system bridges after central nervous-system injury in adult-rats, Science 214 (4523) (1981) 931–933. [DOI] [PubMed] [Google Scholar]
- [205].Carriel V, et al. , Tissue engineering of the peripheral nervous system, Expert Rev. Neurother 14 (3) (2014) 301–318. [DOI] [PubMed] [Google Scholar]
- [206].Lee SK, Wolfe SW, Peripheral nerve injury and repair, J. Am. Acad. Orthopaedic Surg 8 (4) (2000) 243–252. [DOI] [PubMed] [Google Scholar]
- [207].Johnson PJ, et al. , Tissue engineered constructs for peripheral nerve surgery, Eur. Surg 45 (3) (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]
- [208].Tang P, Chauhan A, Decellular nerve allografts, J. Am. Acad. Orthopaedic Surg 23 (11) (2015) 641–647. [DOI] [PubMed] [Google Scholar]