The implementation of additive manufacturing in the field of biomedicine is known as 3D bioprinting. This technique has become a powerful and cost-effective method in tissue engineering and regenerative medicine, particularly when compared to traditional manual manufacturing methods. Many 3D bioprinters offer high precision and resolution, allowing for the creation of complex and finely detailed tissue structures in a relatively short time making the technology easily accessible for research institutions and medical facilities. Therefore, 3D bioprinting can significantly reduce the waiting time for tissue and organ transplants, potentially saving lives by providing timely replacements for damaged organs. As the technology becomes widely adopted, it has the potential to become more affordable, making it accessible to a broader range of research and healthcare facilities [71]. Four categories of 3D bioprinting can be distinguished based on the processes of fabrication: extrusion, inkjet printing, laser-induced forward transfer, and vat polymerization. 3D bioprinting is a cutting-edge technology that has gained significant attention in the field of tissue engineering. It involves the precise deposition of biological materials, such as cells and bioinks, layer by layer, to create complex three-dimensional structures.

Extrusion-based bioprinting (EBB) represents the most widely used bioprinting technique. In particular, coaxial bioprinting, a type of EBB, enables the fabrication of concentric biomaterial layers with cells and thus, can mimic crucial aspects of native tissues [72]. EBB is performed by loading specific bioinks into cartridges, which are then subsequently extruded onto a surface through a nozzle via either pneumatic pressure or mechanical forces. In general, three distinct methods are used for EBB: pneumatic-based extrusion, screw-based extrusion, and piston-based extrusion [72].

In the last decade, coaxial bioprinting contributed significantly to the further development of tissue-engineered constructs with vascular networks. This type of bioprinting can be applied to control concentric multi-material deposition or improve resolution through inline crosslinking. Distinct combinations of hydrogels, cell-laden materials, or crosslinkers can be applied for the creation of vascular tubular structures, composite 3D structures, and complex multilayered structures. For example, perfusion of decellularized ECM-based proteins, followed by a low-viscosity cell-laden hydrogel was designed to increase cell attachment and migration. The first bioinks to be utilized in coaxial bioprinting were alginate and collagen, a mixture of which resulted in constructs that had the benefits of both collagen and alginate, namely soft material allowing cell interaction and aggregation and strong mechanical properties, respectively. In particular, the creation of vasculature is one of the major applications for coaxial bioprinting (Fig. 7). Importantly, coaxially-bioprinted constructs can mimic the key characteristics of the circulatory system. Currently, vascular structures including arteries and veins, which are 10–300 mm in diameter, as well as arterioles can be generated by coaxial printing [72].

Applications of coaxial printing in vasculature engineering: A Vessels were fabricated by extruding sodium alginate onto a rotating rod (upper), followed by assembly into multiscale vasculature (lower). B Coiled-rope structures (upper right) were formed within a glass tube by crosslinking and adjusting the viscosity of the core and sheath fluids (left), providing space for endothelial cell lumen formation (lower right). C Artificial bio-blood vessels (BBV) were designed using a sacrificial core fluid (left), showcasing enhanced limb salvage in a mouse model when loaded with cells (EBBV) and a statin drug (EABBV) (center right). D Coaxially printed vessels (left) demonstrated cell differentiation in vitro (center) compared to single fluid printing (right) in a rat model. Scale bars: A 5 mm; B 200 μm; D 200 μm [Modified after 72]

However, one of the main challenges of coaxial printing remains the inclusion of cells since small nozzle sizes result in higher shear stress and damage to cells. In contrast, large nozzle sizes present an opposite as the bioink yield stress may be overcome by gravity, making printing actuation impossible. Thus, careful choice of nozzle dimensions and flow may enable a physiologically relevant scaffold organization.

Microfluidics can facilitate the formation of a perfusable and functional 3D microvasculature in different organ-on-a-chip systems. Therefore, microfluidic technologies have emerged as useful tools, which can offer precise control over various aspects of the cellular microenvironment such as a different profile of fluid flow, gradient of various GFs, and mechanical properties of versatile biomaterials (e.g. stiffness, orientation, etc.). Furthermore, microfluidic technologies hold great potential to model pathological conditions to study vascular-related diseases [73].

Importantly, microfluidics can mimic physiological 3D microstructure of blood vessels (i.e., circular cross-section) and blood vessel polarization from the apical (luminal) to basal (abluminal) axis, which is important for directed secretion of proteins involved in the cell shape to form a lumen [73]. Besides that, microfluidics also enables the continuous perfusion of cell culture medium to supply oxygen and nutrients as well as the removal of waste products, which is crucial to maintaining the long-term survival of vascularized microtissues. With the combination of various biomaterials and tissue engineering techniques, three main types of 3D in vitro microfluidic vascularization strategies have been developed: (1) EC lining-based methods, (2) vasculogenesis and angiogenesis-based methods, and (3) hybrid methods.

The EC-lining method represents one of the first established techniques, which has been broadly used. It is achieved bycreating a microfluidic channel and lining it with EC to form a monolayer on the inner walls of the microchanneles (Fig. 8).

Vascularization strategies using ECs in vitro: A Microneedle-based Removable Method: (i) A needle-molding technique to form fluidic channels within hydrogels. (ii) Endothelialized microchannels under the influence of fluid flow. B Micropatterned Planar Hydrogel Slab Bonding Method: (i) Fluidic hydrogels created via micromolding from PDMS or silicon molds, followed by bonding. (ii) Micrographs exhibit the shape of microvessel networks within collagen gel constructed from a micropatterned silicon stamp, along with confocal sections of endothelialized microfluidic vessels immunostained with CD31. Scale bar, 100 μm. C Dissolvable Materials-Based Sacrificial Micromolding Method: (i) A schematic showcases a 3D interconnected microvessel network formed by casting a carbohydrate glass lattice as the sacrificial element with a 3D printer. (ii) Micrographs reveal HUVECs expressing mCherry attached to the hydrogel wall for generating the microvessel network and an endothelial monolayer-lined vascular lumen surrounded by 10T1/2 cells after 9 days in culture. Scale bars, 1 mm and 200 μm. D EC Lining Inside a PDMS-Based Microfluidic Channel: (i) A schematic depicts a PDMS-based microfluidic channel, alongside a confocal image of endothelial cells within the channel. (ii) Confocal reconstruction images show the complete lumen formed by HUVECs inside the PDMS tube, along with fluorescence micrographs of cross-sectional views of an endothelialized PDMS tube stained with CD31/nuclei. Scale bars, 100 μm (top) or 200 μm (bottom) [modified after 73]

The major advantage of the EC-lining method is that the vascular geometry and dimensions can be easily controlled. Besides, shear stress can be precisely calculated based on the channel dimensions and the applied flow rate. The hollow single channel or a channel network can be made of either hydrogel or polydimethylsiloxane (PDMS). For hydrogel-based microchannel, various micro-molding methods reported in the literature so far can be grouped into three main types: (1) microneedle-based removable method for a single microchannel, (2) micropatterned, planar hydrogel method for single-layer microchannel network, and (3) dissolvable material-based sacrificial micro-molding method for multi-layer microchannel network. Since cells might not adhere tightly to the PDMS surface inside a PDMS-based microfluidic channel, a thin layer of basement membrane ECM proteins (e.g., laminin, fibronectin, collagen IV, etc.) need to be coated onto the microchannel inner walls to enhance cell adherence. However, the main drawback of EC-lining methods is the fact that they cannot mimic physiological vascular formation in vivo [73]. Indeed, this can be only reached using vasculogenesis and angiogenesis-based methods.

Several studies highlight the diverse approaches employed to enhance vascularization in skin tissue engineering. Skardal et al. demonstrated an innovative approach to enhance the vascularization of skin defects in a mouse wound model. They utilized bioprinted amniotic fluid-derived stem cells embedded within fibrin-collagen hydrogels [74]. This combination effectively promoted the formation of new blood vessels in the injured skin tissue [74]. In another study, Michael et al. employed laser-assisted bioprinting to create artificial skin by implanting keratinocytes and fibroblastsin Matriderm®, a dermal template [75]. Upon implantation of these constructs into mouse dorsal skinfold chambers, the authors observed the development of new blood vessels originating from the surrounding host tissue. This vascularization process was mainly stimulated by the secretion of vascular endothelial GF (VEGF) by the co-implanted keratinocytes [75]. Additionally, researchers utilized a microfluidic water-in-oil emulsion technique to produce microporous annealed particle (MAP) gels, which are injectable materials for in situ skin regeneration (Fig. 9A). These scaffolds facilitated accelerated skin regeneration and the ingrowth of microvasculature in vivo [76]. Importantly, the vascularization process was characterized by early pericyte stabilization of the newly formed microvessels within the first seven days (Fig. 9B, 9C), indicating a crucial role in vascular maturation and stability [76].

Development of MAP gel using microfluidics. A Application of MAP gel to produce any kind of 3D shape through 25-gauge syringe. B–C Detection of vasculature by staining for endothelial cell marker PECAM-1 in MAP scaffolds, 5 days after transplantation onto mice injured skin [modified after 76]

One important concept in vessel network formation is the establishment of a vascular hierarchy, where vessels of different sizes and functions are organized hierarchically. This hierarchical organization is critical for efficient blood flow distribution and tissue perfusion. Arterioles, capillaries, and venules have distinct structural and functional characteristics, and their formation involves spatially and temporally coordinated interactions between ECs, mural cells, and perivascular cells. Studies have elucidated various mechanisms involved in the formation of hierarchical vessel networks. For example, recent research has highlighted the role of EC heterogeneity, differential expression of angiogenic factors, and dynamic cell–cell interactions in shaping vessel hierarchy. Additionally, emerging evidence suggests that mechanical forces, such as blood flow patterns and tissue stiffness, contribute to the spatial organization and remodeling of vessel networks.

In human skin, the capillary network consists of both blood and lymphatic capillaries present through the entire dermis. Interestingly, the upper papillary and lower reticular dermal parts differ with respect to vascularization pattern. There are two primary horizontal plexuses composed of arterioles and veins in the dermal compartment, namely the superficial sub-papillary plexus (SSP) and the deeper cutaneous plexus (DCP). Arterioles that supply blood to the muscles and the hypodermal layer form the DCP, which reside between the subcutaneous and the cutaneous compartment. The arterioles and veins are arrange in a vertical vascular pattern to connect to the SSP, which is located between the epidermis and the papillary dermal compartment. The vessels in the DCP differ in morphology from those in the SSP. DCP vessels have a wider diameter (10–35 µm in the SSP and 40–50 µm in the DCP) and thicker walls. Individual arterioles in the dermal papilla form separate capillary vessel loops that have an ascending limb, an intra-papillary loop, and a descending limb that fuses with post-capillary venules. The capillary loop supplies blood to a skin area that is between 0.04 and 0.27 mm2 in size. Numerous anastomoses in the skin vasculature through which blood flows play an important role in temperature regulation. So far the complex vascular plexus of human skin could not be reproduced in vitro [77].

Recently, researchers have presented a novel approach for constructing a hierarchical vessel network using induced spontaneous anastomosis in a tumor model-on-a-chip. They developed a microfluidic platform that enables the formation of hierarchical vessel networks to support tumor growth. Fabrication of a microfluidic device, consisting of multiple interconnected channels, was designed to mimic the hierarchical organization of blood vessels in vivo. Next, ECs were seeded within the channels and spontaneous anastomosis, a process where adjacent vessels fuse to form interconnected networks, was induced. This approach allowed the generation of capillary-sized vessels, arteriole-sized vessels, and venule-sized vessels within the microfluidic device, resembling the complexity of native vascular networks [78].

There are a few important issues to think about before building a new in vitro microvasculature or modifying an existing one depending on size and flow characteristics. First, any microfluidic system containing cultivated cells is severely damaged by air bubbles, but this can be avoided with the right approach (priming tubing and connecting fluidic ports in a bubble of liquid). Second, it is difficult to evenly distribute a high density of ECs inside the microfluidic channels with small diameters due to the blockage, thus EC lining methods are only suitable to construct large blood vessels with diameter greater than 50 μm. The time required for device design and development is another factor to take into account; it can take anywhere from days for PDMS devices to weeks for certain gel devices, however, a cutting-edge platelet-rich plasma-culturing method may significantly accelerate this procedure. The choice of assays that can be successfully carried out using in vitro microvascular systems is the final important factor to take into account. Many of these assays may require live-cell microscopy imaging, keeping in mind that microvascular failure might manifest as bleeding, thrombosis, remodeling, or altered perfusion. Whereas immunofluorescent labeling of specific antigens and adhesion molecules represents an easy method, single-cell-based approaches linked to proteomics and lipidomics may require the pooling of many devices to obtain the necessary numbers of cells for analysis [79].

In conclusion, 3D bioprinting is an innovative technology with immense potential to revolutionize medicine and healthcare. While it offers numerous advantages, including tissue customization and complex structure creation, it also faces technical, cost, and regulatory challenges that must be addressed to unlock its full potential and ensure its safe and effective use in clinical practice. Continued research and development in the field will likely lead to further advancements and overcome many of the current limitations.

GelMA, a gelatin derivative with a significant number of methacrylamide groups and fewer methacrylate groups, is another type of hydrogel with several biomedical applications due to its versatile physical attributes and compatible biological properties. Several authors used multiple terms for GelMA, such as gelatin methacrylamide, methacrylated gelatin, methacrylamide modified gelatin, or gelatin methacrylate (GelMA). Because GelMA hydrogels contain peptide motifs that bind to cells and activate matrix metalloproteinase (MMP), they bear a striking resemblance to some of the fundamental characteristics of native ECM. These peptides facilitate the growth and division of cells within GelMA-based scaffolds [80].

To create covalently crosslinked hydrogels, GelMA is exposed to UV light in the presence of a photoinitiator, a process known as photoinitiated radical polymerization. Gelatin is a byproduct of the hydrolysis of collagen, which is the main component of ECM in most tissues. It contains numerous arginine-glycine-aspartic acid (RGD) sequences that facilitate cell attachment, as well as matrix MMP target sequences that are appropriate for cell remodeling. Since its first synthesis, GelMA hydrogels have been thoroughly studied in terms of physical and biological properties. GelMA applications range from tissue engineering to medication and gene delivery. Various forms of GelMA have been utilized so far, including 3D hydrogel, electrospun fibrous membranes, and 3D printed scaffolds. These scaffolds may fulfill the needs of skin engineering repairs, tendon, bone, cartilage, vasculature, etc. when combined with other polymers, GFs, and small molecule drugs [80]. Moreover, GelMA is a potential raw material for organ-on-chip due to its high-performance biofabrication and biocompatibility. Moreover, modified GelMA can be used for the fabrication of food analysis tools [80].

GelMA scaffolds have been demonstrated by Zhao et al. to facilitate the multi-layered epidermis development with high keratinocyte proliferation [81]. To further enhance wound healing, they also constructed a 3D, completely cellularized scaffold that mimics the natural dermal ECM using GelMA hydrogels. In addition, Zhao et al. applied GelMA in a rat full-thickness skin wound healing model to increase vascularization for the treatment of random skin flap distal necrosis [81]. Furthermore, Zhou et al. developed GelMA-based intelligent, responsive wound dressing vesicle systems that give fluorometric/colorimetric response (green) to bacterial infection on the wound site and release an encapsulated anti-microbial agent of vesicles to either block or kill pathogenic bacteria, including P. aeruginosa and S. aureus, while providing a visual infection alert [82]. This approach aims to mitigate antibiotic resistance and the excessive use of antimicrobials while extending the efficacy and stability of the encapsulated antimicrobial agents. This is achieved by ensuring the controlled release of antimicrobials specifically triggered in response to pathogenic bacteria. A recent contribution by Jahan et al. demonstrated that Ag-nanoparticle entrapped GelMA scaffolds improve wound healing, particularly for deep skin wounds83 . When employed as a wound dressing, these scaffolds trigger fibroblast migration and reduce microbial infections [83].

Moreover, GelMA has a wide range of applications to facilitate 3D vascular network formation because of its adaptable mechanical characteristics. For example, Chen et al. demonstrated that GelMA hydrogels may promote the development of human progenitor cells-based vascular networks [84]. Additionally, they demonstrated that the degree of methacrylation affects the malleable physical properties of GelMA modulating the extent of vascular development [84]. In particular, GelMA with 49.8% methacrylation creates a  softer hydrogel than GelMA with 73.2% methacrylation [84], with the latter showing enhanced vascular development. GelMA’s vascular formation performance can be further improved by combining it with drug release systems. For instance, Chen et al. created a GelMA hydrogel to deliver desferrioxamine through a continuous and steady release [85]. This hydrogel network promotes the expression of HIF-1α, an important activator of vessel formation, and thus creates a favourable environment for EC proliferation and migration [85]. Further, using an extrusion technique, Wang et al. created a living photosynthetic scaffold made of microalgae, alginate, and GelMA that resembles the properties of tissues or organs [86]. As the inner and outer phases, respectively, they employed a solution of microalgae, alginate, and GelMA and gelatin fluid containing calcium chloride. The hollow fibers were created by the cross-linking of alginate and Ca2+, and the GelMA component was then photopolymerized by UV light. Moreover, it was shown that the resulting hydrogel fibers may be piled on tissue in situ to create a 3D scaffold, suggesting that this technique could be a novel and interesting approach for 3D bioprinting [86].

Despite great advancements in 3D bioprinting, printing multi-layered tubular tissues, like arteries, is still challenging. In this respect, GelMA represents a promising biomaterial for vascular regeneration. GelMA has been synthesized in various forms including hydrogels, 3D bioprinted scaffolds, and electrospun fibers, thanks to its controllable physical properties. Pi et al. demonstrated that multi-layered tubular tissues may be bioprinted in a single step using the multichannel coaxial extrusion system using GelMA, alginate, and 8-arm polyacrylate as the bioink [87]. Meanwhile, a favorable environment for cell adhesion and proliferation may be provided by GelMA hydrogel, which shows strong hydrophilicity and naturally occurring cell-binding motifs. Apart from 3D printing, there are other uses for the combination of GelMA and electrospinning in vascular tissue engineering. For example, Hassanzadeh et al. used a chitin nanofibrous system to create a self-assembly GelMA hydrogel, resulting in the formation of hybrid films with varying chitin nanofiber content [88]. The straightforward self-assembly procedure of these hydrogels is compatible with soft lithography techniques for microstructure fabrication. Importantly, HUVECs co-cultured with human MSCs demonstrated high proliferation and alignment on the micropatterned hydrogels. Furthermore, those matrices showed expression of vasculogenic markers, confirming cell differentiation and the establishment of stable vasculature in these substrates. [88].

Altogether, hydrogels like GelMA, silicone, and ECM like collagen type I are some of the most frequently applied biomaterials that are used in 3D bioprinting techniques. However, these biomaterials have also some drawbacks, including the need for the material to be liquid before printing, the requirement for quick molding after printing, insufficient physical-mechanical properties, channel collapse, inconsistent biological drop and tissue regeneration, high oil-liquid interfacial tension, and other restrictions. The process of extruding bioink or cell-laden materials through a nozzle or printhead can subject the printed cells to shear stress. Excessive shear stress can damage or disrupt delicate biological structures and compromise the viability and functionality of the printed cells. Printhead clogging is a common issue in 3D bioprinting systems. The bioinks used in the process can contain particulate matter, cells, or biomaterials that may clog the printhead or nozzle. This can lead to interruptions in the printing process, affecting the quality of the printed tissue or organ. The diameter of the printing nozzle or printhead can impose limitations on the size and resolution of the printed structures. Smaller nozzle diameters may offer higher resolution in coaxial bioprinting but can limit the speed of printing, while larger nozzles may enable faster printing but with reduced precision. Balancing these factors can be challenging, especially for complex tissue engineering applications [89].

Biomaterial models are designed and manufactured layer by layer using techniques like photopolymerization and microfluidics to manipulate the bioink. As a result, the bioinks used to produce the biomaterials mostly define their biocompatibility. The development of highly organized microvascular networks with distinct branching and hierarchical patterns is made possible by two fast-evolving technologies: microfluidics and 3D bioprinting. On the one hand, phase-changing hydrogels, soluble factors, and endothelial and parenchymal cells can be co-assembled using 3D bioprinting in a high-throughput and reliable manner. Implementation of the proper bioreactor perfusion methods enables ECs to be uniformly seeded into the microfluidic channels of 3D hydrogels. Skin tissue engineering is recently beginning to employ these approaches, although these techniques have been extensively studied in the fundamental angiogenesis and vascular research field.

Conventional skin substitutes often fall short of recapitulating the intricate architecture and multifaceted functionality of native dermal tissues. Therefore the emergence of decellularized extracellular matrix (dECM) generated after the removal of cellular components, holds promise in circumventing these challenges. dECM, derived from endogenous ECM, closely mimics its structural and compositional properties, thereby fostering a conducive microenvironment for cellular regeneration. Additionally, dECM exhibits favorable bioactivity, minimal immunogenicity, and abundant availability, rendering it a promising biomaterial for the restoration and rejuvenation of cutaneous tissue. Recent research has highlighted the pivotal role of dECM in the intricate process of skin wound repair. Characterized by its abundance of biomolecules and unique 3D structure conducive to cellular behavior, dECM stands as an optimal bioactive scaffold for skin repair and regeneration. The fibrous network architecture of dECM fosters an appropriate microenvironment for cellular proliferation, while its bioactive constituents modulate crucial cellular functions such as adhesion, migration, proliferation, and immune regulation during wound healing.

The inception of decellularization dates back to 1948, and since then, the advancement of decellularization technologies has been substantial. For example, in the 1970s, the isolation of basement membrane was achieved, while the production of acellular small intestinal submucosa matrices began in 1995. Subsequently, decellularized whole heart, lung, and kidney scaffolds were developed in 2008, 2010, and 2013, respectively. Simultaneously, evaluation methodologies were further developed; the establishment of ECM’s matrisome occurred in 2012, followed by proteomic, and glycosaminoglycanomic analyses in 2020 and 2021, respectively. The development of a commercial acellular dermal matrix in 1995 allowedthe firsthuman application in 1996. Since then, a plethora of dECM and dECM-based materials have been synthesized and utilized for skin regeneration. A recent study by Jiang et al. revealed that dECM derived from skin tissue exhibits superior physical stability and biological composition compared to conventional collagen bioink [90].

The ECM serves as a bioactive substrate rich in biochemical cues essential for cellular events crucial in wound healing. Nonetheless, the presence of cellular and nuclear remnants within the ECM may induce cytotoxic effects in vitro and trigger immune responses afterwards in vivo, potentially compromising the wound repair and thus, therapeutic outcomes. Consequently, a meticulous decellularization protocol for ECM-derived materials becomes imperative before application, with the primary objective of minimizing immunogenic components. Physical techniques, such as freeze-thaw cycles, mechanical agitation, pressure application, sonication, and supercritical CO2 treatment, represent straightforward approaches to ECM decellularization (Fig. 10). Notably, their avoidance of chemical reagents renders them particularly favored among researchers. Deng et al. [91] employed freeze-thaw cycling methods to produce ECM-PLGA materials, with subsequent DNA electrophoresis and quantitative analysis indicating significant removal of DNA components. Moreover, various chemical agents, encompassing acids, bases, detergents, and hypo/hypertonic solutions, have demonstrated notable efficacy in removing nuclear and cytoplasmic components. In investigation performed by Ng et al., Triton X-100 and NH4OH solution were utilized to decellularize human umbilical cord-derived mesenchymal stem cells, yielding favorable results [92, 93]. Unlike physical decellularization methods, theis method using chemicals offer efficient removal of potentially immunogenic ECM components within a short timeframe, alongside with inherent sterilizing properties. However, excessive detergent concentrations and prolonged exposure to chemicals may lead to the depletion of vital GFs and disruption of ECM ultrastructure. Consequently, optimizing the duration and concentration of chemical agents is of paramount importance for achieving satisfactory decellularization outcomes. Furthermore, process-related impurities and chemical residuals within dECM poss a significant challenge, and may cause host inflammatory responses. In addition to physical and chemical methodologies, biological techniques are also available for the removal of cellular components. For example, nucleases exhibit notable efficiency in eliminating DNA and RNA contents, while lipases are utilized for specific lipid removal. However, akin to chemical approaches, biological methods often result in incomplete decellularization, residual reagents, disruption of ECM ultrastructure, and potential inflammatory reactions. For instance, prolonged exposure to trypsin may induce irreversible damage to ECM collagen components. Conversely, when employed in conjunction with the chelating agent ethylenediaminetetraacetic acid (EDTA), the risk of immune response activation can be mitigated.

Schematic diagram for preparation and application of dECM (prepared with BioRender)

dECM can be categorized into two distinct groups according to the source of the ECM: organ-/tissue-derived and cell-derived dECM (Fig. 10). The dECM sourced from organs/tissues and cells each possesses distinct attributes. Organ/tissue-derived dECM boasts an optimal yield and preserves its native 3D structure along with multidirectional active components, yet it carries inherent risks such as immunogenicity, cell permeability, and potential disease transmission. In contrast, cell-derived dECM inherits natural bioactive factors and proteins from donor cells, mitigating the risk of pathogens and immunogenic macromolecules compared to organ/tissue-derived dECM. However, cell-derived dECM lacks a three-dimensional structure, necessitating the incorporation of additional molecules or the combination with other scaffold materials to bolster mechanical strength in practical applications [94]. Concurrently, the conventional two-dimensional (2D) culture system falls short in meeting the demands of large-scale preparation, prompting the need for the development of three-dimensional (3D) culture systems to rapidly upscale cell quantity and enhance ECM production efficiency.

Thus far, a diverse array of organs/tissues has been employed in skin tissue engineering endeavors, encompassing skin tissues, perinatal-related tissues, adipose tissues, small intestinal submucosa, fish skin, heart, lung tissues, and beyond. Pigs represent the primary source among animal-derived dECM biomaterials due to their widespread availability and ample supply. Numerous investigations have highlighted the close resemblance in composition and functionality between porcine skin ECM and its human counterpart as compared to ECM from other animal sources. In a recent study by Liu et al. [95] proteomic analysis of porcine skin-derived dECM revealed elevated levels of angiogenesis-associated proteins, pointing towards its potential role in wound healing by regulating angiogenesis. Currently, porcine skin and small intestine submucosa rank as the most extensively utilized dECM tissues, both of which have transitioned into  commercial products for clinical applications. Human skin, adipose, and perinatal tissues represent the primary sources of dECM utilized in skin regeneration. Perinatal tissues, encompassing the placenta, umbilical cord, and amniotic membrane, fulfill various functions during fetal development but are typically discarded postpartum. Extensive research has underscored the rich trove of diverse GFs and cytokines present in perinatal-related tissues, including EGF, TGF-β, FGF, PDGF, and VEGF. These bioactive agents play pivotal roles in fibroblast migration, MSCs homing, re-epithelialization, and neovascularization, rendering them indispensable for wound healing and early-stage tissue repair and regeneration. In a recent study [92], the dECM extracted from human, pig, and rat skin underwent comprehensive proteomic and bioinformatics analyses. In comparison to human-derived dECM, both pig and rat skin-derived dECM showed a deficiency in proteins linked to inflammatory modulation and the innate immune response. Additionally, they exhibited markedly lower levels of most proteases and protease inhibitors. These findings suggest that human-derived dECM may establish a more favorable microenvironment for enhancing wound healing. Compared to organ/tissue-derived sources, cell-derived dECM presents a reduced presence of immunogenic components and a diminished potential for pathogen transmission. Furthermore, dECM can be obtained through the cultivation of the patient’s own cells, facilitating “autologous tissue engineering”. Notably, during in vitro ECM deposition, specific modifications of dECM can be achieved by altering culture conditions or introducing particular stimuli, a feat challenging to replicate in organ-/tissue-derived dECM. For instance, Dong et al. [96] successfully generated dECM with immunomodulatory properties by supplementing IFN-γ during culture. Similarly, the bioactivity of cell-derived dECM can be augmented through the co-culture of two or more cell types. Carvalho et al. [97] observed that dECM produced via the co-culture of bone marrow-derived MSCs (BMSCs) and HUVECs significantly enhanced the osteogenic differentiation and angiogenic potential of BMSCs in vitro. Moreover, cell-derived dECM can be tailored into various formats depending on the intended applications, including 2D layers and 3D structures. Cell-derived dECM has found widespread application in skin, nerve, bone regeneration, and other domains, yielding favorable tissue regeneration outcomes. In the context of skin regeneration, diverse fibroblast types serve as primary sources of dECM. As the predominant cell type in the dermis, fibroblasts possess robust ECM secretion capabilities, resulting in dECM primarily comprised of collagen, closely resembling native skin tissue in comparison to traditional biopolymers. Furthermore, various fibroblast subtypes exhibit distinct phenotypes, protein compositions, secretory profiles, and mechanical properties. Researchers have highlighted the morphological, functional, and compositional differences observed in ECM derived from three distinct fibroblast subtypes within the human skin dermis. These findings underscore the versatility of maximizing ECM bioactivity through diverse methodologies, suggesting that cells originating from the same source are not invariably essential.

To sum up, dECM encompasses a plethora of GFs and bioactive molecules crucial for regulating cell maturation, migration, proliferation, and other vital biological functions essential for skin repair processes. Following decellularization, dECM can be generated in various forms such as powders, gels, foam, and sheets. Moreover, it can be synergistically combined with cells, GFs, and other synthetic peptides to create advanced bioactive scaffolds. Over time, a myriad of dECM-based materials have been developed for skin repair and regeneration, spanning from porous scaffolds and fibrous scaffolds to hydrogels and bioinks.

Porous scaffolds, organ-/tissue-derived dECM, sourced from both prenatal and sk