**3. Discussion**

The fast and efficient establishment of blood perfusion in engineered constructs after transplantation represents one of the major challenges for the incorporation of TE products into the clinical practice. Some early vascularization approaches focused on vascular ingrowth stimulation in tissue constructs by optimizing scaffolds' material properties [28] or by incorporating growth factor delivery systems [29,30]. Both strategies proved to be inefficient, however, since vascular ingrowth is a slow process [2]. To overcome this, TERM strategies started to incorporate the in vitro creation of a prevascular network. ECs are seeded in scaffolds and supplemented with angiogenic growth factors that induce a vasculogenic-like process, ultimately forming a prevascular network [31]. However, the native vasculogenic process requires different growth factors produced by different cell types that interact with target receptors in a coordinated manner to ultimately yield a mature vascular network [9]. This is why early prevascularization strategies using only ECs, typically HUVECs, were often found to yield non-mature prevascular that is prone to regression after some period of time [32]. However, to recapitulate, the in vitro complexity of native vasculogenesis is a formidable challenge, both technically and in terms of costs due to the diverse cell types and different culture media and growth factors required. These facts urge the development of a streamlined and more cost-effective strategy to promote the vascularization of constructs.

In recent years, the SVF of adipose tissue became a focus of attention mainly due to its intrinsic angiogenic potential. However, studies demonstrating the angiogenic potential of SVF commonly use freshly isolated samples, which may limit its clinical applicability to specialized centers. The development of cell-banking strategies for SVF while maintaining its angiogenic potential would boost its clinical potential. The development of cell-banking strategies for SVF while maintaining its angiogenic potential would boost its clinical potential. Some studies describe the use of cryopreserved SVF for the production of vascularized adipose tissue, showing its ability to create capillary-like structures after preservation [33]. However, such studies use supplementation with angiogenic growth factors to induce capillary-like structure formation, which introduces a significant degree of complexity to the system. A previous study from our laboratory has shown that fresh SVF can spontaneously produce capillary-like structures as early as 5 days of culture, without the addition of angiogenic growth factors, representing a cost reduction and a more organic vasculogenic process since it is orchestrated by the cells themselves [13]. In this sense, we explored if cryopreserved SVF, in the absence of extrinsic angiogenic growth factors, retained this ability to spontaneously create a prevascular network in 3D conditions. SVF consists of a heterogeneous population of cells with an intrinsic capacity to secrete several angiogenic-associated growth factors, creating the perfect angiogenic microenvironment capable of promoting the formation of in vitro capillary-like networks in the absence of extrinsic angiogenic growth factors [13].

Vasculogenesis and angiogenesis are complex processes involving the coordinated action of several families of growth factors such as the vascular endothelial growth factor (VEGF), platelet-derived growth factor (PDGF), tissue inhibitor of metalloproteinases (TIMP), fibroblast growth factor (FGF), angiopoietin (ANG), interleukins (ILs), and matrix metalloproteinases (MMPs). All these players contribute not only ro capillary formation but, critically, to their stabilization and maturation [13]. In the 3D SVF cultures reported herein, we verified the protein expression of several angiogenic modulators over the culture time. Factors involved in ECM remodeling, namely urokinase (uPA), plasminogen activator inhibitor (PAI-1), and TIMP-1, increased from day 5 to day 7. Plasminogen activators such as (uPA) are key mediators of the ECM degradation process by converting inactive plasminogen to active plasmin, and it is in turn capable of degrading specific matrix constituents and of activating matrix-degrading metalloproteases such as MMP-9 [34]. Concurrently, it has been demonstrated that the increased production of uPA by microvascular ECs in response to angiogenic stimuli is accompanied by an increase in the production of the PAI-1 [35], which is in line with what we observed. This is most likely related with

the need of a proteolytic balance that allows ECM degradation for cell migration but in a controlled fashion to keep the three-dimensional matrix intact, into which ECs form capillary-like networks [36]. The secretion of MMPs inhibitors is also influenced by this synergistic effect. Increased TIMP-1 expression is important for vessel stabilization by limiting matrix degradation and allowing matrix depositions that could explain its increase over culture times [37]. The decrease in IL-8 and MCP-1 expression also suggests that the angiogenic process is more directed to capillaries stabilization and maturation. In an initial phase of angiogenesis, MCP-1 recruits macrophages [38] that in turn secrete a variety of angiogenic-related factors such as thrombospondin-1 and IL-8 [39]. The latter is in fact essential for EC proliferation and migration [40,41], allowing capillary formation and elongation. The role of MCP-1 in angiogenesis promotion is not exclusively for macrophage recruitment. It up-regulates VEGF expression [42], increases vascular permeability [43], and is involved in pericytes recruitment [44]. The incorporation of pericytes initiates the stabilization and maturation of new vessels [45]. They are capable of modulating the ECM remodeling capacity of ECs by inducing the upregulation of PAI-1 in ECs and, in this way, limiting its migration and branching [46]. We verified the presence and incorporation of pericytes in the pre-vascular nerwork after 7 days, together with an increase over culture time of the levels of thrombospondin-1 and a decrease in levels of IL-8 and MCP-1 and stagnation in VEGF release. This strongly suggests a move towards the stage of capillary stabilization and maturation. This is further reinforced by decreased levels of ANG-2, involved in angiogenesis initiation [47]. The synergy between all cells present in the SVF and the secretion of all of these factors ultimately led to the formation of a prevascular network, as demonstrated by CD31's expression pattern, and it is anchored by perivascular cells identified by the expression of CD146 and the lack of expression of CD31. Although the present study did not directly compare the angiogenic capacity of cryo-preserved and fresh SVF, our results are in agreemen<sup>t</sup> with what was reported by Costa et al. for fresh SVF. That study showed the spontaneous formation of a network of capillaries, with increased secretions of several angiogenic modulators from 5 to 8 days of culture. All of this, of course, occurred in the absence of extrinsic angiogenic factors [13]. Collagen scaffolds also provided structural and mechanical support and permitted significant capillary formation within its structure in vitro. The ability of collagen, as a three-dimensional matrix, to support the adherence and proliferation of endothelial cells in vitro has been observed for several years [48]. While mammalian collagen is the standard, new collagen sources have emerged. The skin and bones of several marine organisms are abundant in collagen, presenting very similar characteristics to that of mammalian origin. Blue shark skin collagen demonstrated a comparable chondrogenic differentiation of human adipose stem cells compared to the commercial alternative comprising bovine collagen [27]. In the case of a direct comparison between collagen from marine tilapia skin and bovine skin collagen, both showed a similar performance in a wound-healing scenario, allowing fibroblasts infiltration, vascularization, reduced inflammation, and collagen deposition [49]. In fact, the existing differences are only noticeable in proline and hydroxyproline contents [27]. The lower content of this amino acids in comparison with mammalian collagen, affects the denaturation temperature and, consequently, its thermal stability, resulting in a faster degradation of scaffolds [50]. Despite presenting a faster degradation rate due to its lower denaturation temperature, the use of crosslinkers allowed the stabilization of sponges. The large pores created (averaging 250–300 μm) [27] with highly interconnective microporosities positively influenced endothelial cell migration, rearrangement, and vessel density. Microporosity is considered critical for the new vessel's size and number [51,52]. High pore interconnectivity results in significantly higher blood vessel density in vitro and increased blood vessel density and average invasion depth after implantation [53]. These architectural features present in the sponges used in this study allowed for the cell migration, proliferation, and organization of vascular networks, validating marine-derived collagen as a suitable raw material for TE of vascularized tissues. This alternative source of collagen for TE products not only surpasses disease-transmission concerns, such as bovine spongiform encephalopathy (BSE) [22], but

also religious constraints, making it suitable for broader applications. Furthermore, new applications of marine by-products are of extreme importance, especially those that are environmentally friendly [54]. Its isolation represents low costs, creating value from products that are considered wasteful for the fish transformation industry.

To assess the in ovo functionality of the created prevascular network, a CAM assay was performed. The membrane of chick embryo provides a non-innervated and rapidly growing vascular bed, which can serve as a blood supply for engineered tissues and, therefore, be a useful model for testing prevascularization strategies [55]. In particular, its use has been reported in approaches ranging from cell sheets [56] to spheroids [57]. Herein, prevascularized collagen sponges were implanted onto the CAM and collected after 4 days. Empty sponges were implanted as controls. Both prevascularized and empty sponges were able to recruit blood vessels from the host in the implant region. Nonetheless, this effect was significantly improved for prevascularized constructs, suggesting that the presence of SVF can accelerate graft's vascularization in ovo. Furthermore, the rapid infiltration of the implanted construct by host cells, without visible inflammatory response, revealed a superior integration with CAM when SVF cells were present. It is known that the cell complexity present in SVF can have an effect on neovascularization in ischemic tissues [58,59]. By injecting SVF in a hind limb ischemic mouse model, previous studies demonstrated that SVF cells not only improved blood flow [58,59] but were also able to integrate blood vessel linings [58]. It is thought that this improvement is mainly mediated by angiogenic cytokines secreted from implanted cells [60]. Since the latter two studies were conducted by injecting cells, the residence time for implanted cells at the injured site was probably reduced; thus, the release of angiogenic stimulators is limited in time. While the encapsulation of SVF can improve residency times, the creation of a 3D prevascular network prior to implantation [61] may further accelerate the vascularization of the graft in vivo through a rapid connection to recipient blood vessels [61,62]. In fact, the maturation of the created prevascular network also influences the angiogenic potential. Cerino et al. demonstrated that faster inosculation and the enhanced survival of transplanted cells in full-thickness rat wounds is associated to pericytes, for which its number was significantly higher in more mature constructs [62]. In the present study, it was clearly demonstrated that implanted human cells, namely CD31-positive cells, were able to integrate newly formed blood vessels at the interface between the CAM and implanted sponges. While this was not specifically tested, no evidence of prevascular network inosculation with the host's circulation was found. Nevertheless, ISH and CD31 results, together with the demonstrated increase in vessel recruitment from the host, show that cryopreserved SVF had a positive effect on vascularization after implantation, underscoring the potential of this fraction for TE applications.

### **4. Materials and Methods**

### *4.1. Collagen Acid Extraction*

Collagen was extracted from blue shark (Prionace glauca) skin at Instituto de Investigaciones Marinas (CSCI, Vigo, Spain), according to a previously described protocol [27]. Blue shark skin was stirred with 0.1M NaOH in a cold room (3–5 ◦C) for 24 h to remove non-collagenous proteins and pigments. After a centrifugation step, the remaining pellet was washed with distilled water and incubated overnight with 0.5 M acetic acid under agitation to start the acid extraction process. The obtained extract was centrifuged at 10 ◦C, and the supernatant dialyzed against distilled water for 2 days in a cold room (3–5 ◦C). Finally, the obtained collagen extract was freeze-dried.

### *4.2. Collagen Sponge Fabrication*

The production of highly interconnective microporous collagen sponges was performed as previously described [27]. Briefly, the previously extracted collagen was solubilized at 1% (*w/v*) concentration in a 10 mM hydrochloric acid (HCl, Sigma-Aldrich, St. Louis, MO, USA) solution. Cryogelation and crosslinking reactions were carried out at

−20 ◦C for 4 h by the addition of 1-[3-(dimethylamino) propyl]-3-ethylcarbodiimide hydrochloride (EDC) (Mw: 191,70 g.mol−1, Sigma-Aldrich, St. Louis, MO, USA) at 60 mM of final concentration. To remove the residual crosslinker, cryogels were rinsed with distilled water before freeze drying.

### *4.3. Isolation and Cryopreservation of Human Adipose-Derived SVF Cells*

Human subcutaneous adipose tissues were obtained from surgical procedures performed at Hospital de S. João (Porto), after obtaining the patient's written informed consent, and within the scope of a collaboration protocol approved by the ethical committees of both institutions for this work (Comissão de Ética do Hospital de S. João/University of Minho: 217/19; CEICVS 008/2019). SVF was obtained as previously described [13]. Briefly, adipose tissue was digested with a collagenase type II (Sigma Aldrich, St. Louis, MO, USA) solution of 0.05% (*w/v*), for 45 min at 37 ◦C under agitation. After centrifugation, the obtained SVF was incubated with red blood lysis buffer and centrifuged, and the supernatant was resuspended in Minimum Essential Medium alpha-modification (α-MEM) (Life Technologies, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (FBS) and 1% antibiotic/antimycotic. Cell nuclei were stained using a solution of 3% (*v/v*) acetic acid (VWR, Lutterworth, UK) and 0.05 wt % methylene blue (Sigma Aldrich, St. Louis, MO, USA) in water to count nucleated cells. Finally, cells were cryopreserved in 10% (*v/v*) dimethyl sulfoxide (DMSO) in FBS with a controlled freeze rate of 1 ◦C/min for at least 7 days.

### *4.4. Cell Seeding*

After thawing, a pool of five different donors was made. SVF was seeded on collagen sponges by dispensing 25 μL on top of the dried sponges and another 25 μL at the bottom of a 50 μL cell suspension comprising 1.5 × 10<sup>6</sup> cells. Constructs were incubated for 1 h at 37 ◦C, 5% CO2, to allow maximum cell entrapment within the structures, and then fresh medium was added to a total volume of 1 mL. Constructs were cultured for 7 days in α-MEM to allow the formation of capillary-like structures.
