Perfusion-Limited Efficacy of Platelet-Rich Plasma in Adipose Tissue Grafts

TL;DR

Adding high-dose platelet-rich plasma to fat tissue grafts improves blood flow, survival, and reduces scarring, offering better results for reconstructive surgery.

Contribution

This study establishes a dose-response relationship for PRP in adipose tissue grafts, identifying optimal concentrations for improved clinical outcomes.

Findings

High-dose PRP increased highly perfused regions in grafts by 8-fold.

PRP reduced fibrosis by 1.67-fold and improved adipocyte survival by 3.8-fold.

Graft mass retention was doubled with high-dose PRP compared to native adipose tissue.

Abstract

Autologous adipose tissue (AT) grafting is often compromised by insufficient early vascularization, leading to ischemia, fibrosis, and inconsistent long-term volume retention. Incorporating platelet-rich plasma (PRP) into AT bioinks offers a clinically accessible means to enhance vascular recruitment, but the in vivo impact of PRP dosage remains unclear. Here, we investigated how PRP concentration, uniformly integrated into a previously reported clinically relevant AT bioink, regulates vascular infiltration, tissue remodeling, and overall graft survival. High-dose PRP markedly improved graft performance, including an 8-fold increase in highly perfused regions, a 3.8-fold enhancement in adipocyte survival, a 1.67-fold reduction in fibrosis, and a 2.51-fold increase in collagen III deposition compared with PRP-free AT grafts. Histological analysis further demonstrated that PRP mitigates the adverse effects of poor perfusion, reducing regional disparities in survival and extracellular matrix (ECM) remodeling. High-dose PRP also maximized graft retention, preserving 103% of graft mass relative to 50.6% in native AT. Together, these results establish a clear in vivo dose–response relationship for PRP-enhanced AT grafts and highlight platelet concentration as a key design parameter for soft-tissue reconstruction. This work provides a translational framework for optimizing PRP-functionalized bioinks to improve clinical outcomes in reconstructive surgery.

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1. Introduction

Autologous adipose tissue (AT) grafting is widely used for soft-tissue reconstruction but continues to suffer from unpredictable long-term retention due to early ischemia, inadequate vascular integration, and progressive fibrotic remodeling. Following implantation, adipocytes experience a hypoxic interval before host vessels can infiltrate the graft. During this period, limited nutrient and oxygen delivery lead to cell death, fibrosis, and variable volumetric outcomes [1,2,3,4,5]. Numerous biomaterial strategies including scaffold porosity optimization, angiogenic factor delivery, and pre-vascularized constructs have sought to accelerate vascular ingress, however, achieving a rapid and spatially uniform perfusion throughout volumetric AT grafts remains a fundamental challenge [6,7,8].

Platelet-rich plasma (PRP) has emerged as a clinically accessible, autologous approach for enhancing soft-tissue regeneration [9,10]. Platelet activation triggers the release of pro-angiogenic and pro-regenerative cytokines [11], including vascular endothelial growth factor (VEGF), platelet-derived growth factor-BB (PDGF-BB), and transforming growth factor, beta 1 (TGF-β1). These support endothelial recruitment, stromal remodeling, and adipocyte survival [12,13,14,15,16]. However, PRP therapies show variable outcomes across clinical and preclinical studies, as the concentration impact on graft-level biological responses remains poorly defined [17,18]. Prior work indicates that early PRP degranulation produces a rapid burst release of cytokines largely governed by activation rather than platelet quantity. This leaves uncertainty regarding how the PRP dose affects longer-term vascular integration, extracellular matrix (ECM) organization, and tissue viability after implantation [19,20,21].

Three-dimensional bioprinting provides an opportunity to incorporate PRP directly into adipose-mimetic matrices with precise spatial control, enabling controlled evaluation of platelet dose within a well-defined graft architecture [22,23]. We previously reported a clinically relevant AT bioink capable of supporting high-fidelity extrusion printing and maintaining adipocyte viability [24]. Building upon this work, we sought to determine whether increasing platelet concentration within this bioink would produce measurable, dose-dependent improvements in vascular infiltration, adipocyte preservation, and ECM remodeling in vivo.

In this work, we investigated two biologically validated PRP concentrations, 0.1 × 10^6^ and 1.0 × 10^6^ platelets/μL, chosen to reflect the minimum dose known to elicit cellular responses and the widely adopted therapeutic range used in regenerative medicine [12,25,26,27]. First, we evaluated whether PRP incorporation and activation affected adipocyte viability or acute cytokine release. Additionally, we implanted 3D-printed constructs subcutaneously to examine how PRP dose influences graft retention, spatial patterns of vascular integration, adipocyte survival, fibrosis, and collagen remodeling. Importantly, because vascular supply in implanted grafts is inherently heterogeneous, we analyzed both whole-construct outcomes and region-specific responses across high- and low-infiltration zones.

By integrating quantitative vascular mapping, histological analyses, and ECM profiling, this work defines how PRP dose and vascular accessibility interact to regulate adipose graft healing. These insights provide mechanistic clarity to long-standing questions surrounding PRP efficacy and establish a foundation for rationally engineering PRP-functionalized bioinks for translational soft-tissue reconstruction.

2. Results

2.1. Early Biological Response of PRP-Integrated AT Bioink: Viability and Growth Factor Release

To verify that PRP incorporation and activation do not adversely affect adipocyte viability or platelet activation-dependent cytokine release, PRP was isolated using a two-step centrifugation protocol and uniformly mixed into the previously reported AT bioink prior to CaCl_2_/thrombin activation [24]. Photographs of the step-by-step isolation procedure are shown in Figure 1A, and a schematic illustration is provided in Figure S1. Two PRP doses were selected: 0.1 × 10^6^ platelets/μL, representing a minimally enriched control, and 1.0 × 10^6^ platelets/μL, corresponding to the widely reported optimal range for regenerative PRP applications [25,26,27]. Live/dead imaging demonstrated that adipocyte viability remained unchanged following PRP incorporation and activation, regardless of PRP concentration (Figure 1B,C). A detergent-treated AT construct was included as a negative control to confirm complete loss of viability and assay specificity (Figure S2). ELISA measurement revealed minimal cytokine release in non-activated controls, whereas activated PRP induced marked early secretion of VEGF, PDGF-BB, and TGF-β1 (Figure 1D–F), with increases of ~9.5-fold, ~2.2-fold, and ~3.8-fold, respectively. Notably, cytokine levels were nearly identical between low- and high-dose PRP, indicating that platelet activation state, rather than platelet quantity, dominates early release kinetics at the 1-h timescale.

Furthermore, to assess whether PRP incorporation alters the physicochemical behavior of the AT bioink, rheological characterization was performed before and after crosslinking. Prior to crosslinking, shear-rate-dependent viscosity measurements demonstrated pronounced shear-thinning behavior across all groups, with comparable viscosity profiles for PRP-free, low-dose PRP, and high-dose PRP bioinks (Figure S3), indicating preserved flow behavior during extrusion. Following crosslinking, dynamic frequency sweep analysis showed that the storage modulus (G′) exceeded the loss modulus (G″) for all formulations (Figure S4), confirming the maintenance of solid-like viscoelastic properties after PRP incorporation. Together, these results indicate that PRP addition within the tested range does not adversely affect either the pre-gelation flow behavior or post-crosslinking mechanical integrity of the AT bioink.

2.2. PRP Dose-Dependent Enhancement of Graft Retention and Spatial Vascular Integration

To assess the effects of PRP concentration on graft preservation and spatial vascular integration, 3D-printed constructs were implanted subcutaneously and evaluated post-harvest (Figure 2). The implantation workflow is outlined in Figure S5, and construct orientation and dimensions are shown in Figure S6. Grafts were positioned so that one surface directly contacted the overlying skin, while the opposite surface interfaced with the underlying muscle. Accordingly, the “front” view represents vascular ingress from the skin side, whereas the “back” view reflects infiltration originating from the muscle interface. Across both orientations, gross morphology, backside inspection, and blood-infiltration heatmaps revealed a pronounced PRP dose-dependent increase in vascular penetration, with high-dose PRP constructs showing the deepest and most spatially uniform perfusion throughout the 5 × 5 printed lattice (Figure 2A). PRP also markedly improved graft preservation. Weight retention increased from 50.6% in native AT to 95.5% in the PRP-free AT construct, 98.7% with low-dose PRP, and 103.0% in the high-dose group, a 2.04-fold improvement over native tissue (Figure 2B). Spatial infiltration maps demonstrated a parallel dose-dependent enhancement in perfusion. Native AT was dominated by poorly infiltrated regions (84% of the construct at 0–25% infiltration), whereas PRP-free constructs showed only modest improvement. PRP addition further shifted this distribution, culminating in the high-dose group where highly infiltrated regions (≥75%) increased to 32%, an 8-fold rise relative to native AT and a 2.7-fold improvement compared with PRP-free constructs (Figure 2C). Despite these substantial gains, central regions remained less perfused than pore-adjacent areas, indicating that although PRP significantly enhances vascular integration, it does not fully homogenize perfusion across the entire construct thickness.

2.3. Dose-Dependent PRP Effects on Adipocyte Preservation Modulated by Local Vascular Infiltration Gradients

To evaluate how PRP influences adipocyte preservation and microvascular formation within the implanted constructs, perilipin^+^ adipocytes and CD31^+^ vessels were first visualized across the full graft thickness (Figure 3A), which revealed substantial regional heterogeneity in vascular penetration. The constructs displayed distinct high- and low-infiltration zones, prompting a spatially resolved quantitative analysis. Correspondingly, whole-construct measurements (Figure 3B,C) were complemented by regional quantification in Figures S7 and S8. In high-infiltration zones, perilipin^+^ adipocyte density increased from 63.2 cells/mm^2^ in native AT to 122.7 cells/mm^2^ in PRP-free constructs, 118.5 cells/mm^2^ with low-dose PRP, and 240.1 cells/mm^2^ with high-dose PRP—a 3.8-fold improvement relative to native AT. Low-infiltration regions showed the same directional trend but markedly lower absolute values, rising from 21.0 to 100.7 cells/mm^2^ under high-dose PRP (~4.8-fold increase), confirming the strong dependence of adipocyte survival on local perfusion capacity (Figure S7). CD31^+^ vessel density followed a similar pattern: in high-infiltration areas, vessel area increased from 0.026 to 0.0547 mm^2^/mm^2^ with high-dose PRP (~2.1-fold), while low-infiltration areas showed an increase from 0.0122 to 0.0281 mm^2^/mm^2^ (~2.3-fold; Figure S8). Together, these data demonstrate that PRP produces a clear dose-dependent enhancement in adipocyte survival and neovascularization, but also revealed a persistent infiltration-dependent disparity, wherein high-infiltration zones consistently outperformed low-infiltration zones across all groups. Although PRP substantially improves outcomes even in poorly perfused regions, it does not fully equalize survival between infiltration states, indicating that PRP enhances, but cannot completely overcome the perfusion-limited constraints imposed on adipocytes within low-infiltration domains.

2.4. Influence of PRP Dose and Local Infiltration Level on Fibrotic Remodeling Within Implanted Constructs

To determine whether the spatial disparity in perfusion also translated to differences in fibrotic remodeling, fibrosis was quantified across high- and low-infiltration regions in all groups (Figure 4A). In high-infiltration areas, fibrosis declined from 24.9% in native AT to 14.9% with high-dose PRP, representing a 1.67-fold reduction. Low-infiltration areas—consistently more fibrotic across all groups—showed a similar PRP-dependent trend, decreasing from 31.5% in native tissue to 20.5% with high-dose PRP (1.54-fold reduction; Figure 4B,C). Despite these improvements, low-infiltration zones remained more collagen-dense than high-infiltration zones in all conditions, underscoring the intrinsic remodeling disadvantage imposed by limited vascular access. Nevertheless, high-dose PRP markedly narrowed this disparity, reducing fibrosis in low-infiltration regions by ~35% compared with native AT while simultaneously producing the lowest fibrosis values in well-perfused regions. Together, these findings demonstrate that fibrotic severity is governed jointly by infiltration status and PRP dose: high-dose PRP attenuates collagen accumulation across the graft but cannot fully eliminate the remodeling disadvantage imposed by poor vascular access.

2.5. Synergistic Effects of PRP Dose and Infiltration Level on Collagen Remodeling in Implanted Constructs

To determine whether the infiltration-dependent differences observed in adipocyte survival and fibrosis also extended to extracellular matrix composition, collagen I and III deposition were quantified across groups using Picrosirius Red polarization imaging (Figure 5A). Collagen I, which dominated the matrix of native AT (15.34%), was reduced in all implanted constructs, with levels decreasing to 5.58% in the PRP-free AT construct, 5.26% with low-dose PRP, and 4.27% with high-dose PRP. High-dose PRP therefore achieved a 3.6-fold reduction relative to native AT and a 1.3-fold decrease compared with PRP-free constructs (Figure 5B). In contrast, collagen III, associated with early compliant regenerative matrices, showed the opposite trend. Native AT showed low baseline collagen III content (0.0122 mm^2^/mm^2^), which increased to 0.0191 mm^2^/mm^2^ in PRP-free constructs and 0.0231 mm^2^/mm^2^ with low-dose PRP, reaching 0.0303 mm^2^/mm^2^ under high-dose PRP. This represents a 2.51-fold increase over native AT and a 1.6-fold increase relative to PRP-free constructs (Figure 5C). The findings demonstrate that collagen remodeling mirrors previously observed infiltration-dependent disparities: while all groups exhibited higher collagen III content in better-perfused zones, PRP, particularly at the high dose, shifted the matrix composition toward a less fibrotic, more regenerative ECM phenotype. However, consistent with fibrosis patterns, low-infiltration regions retain comparatively higher collagen I and lower collagen III levels, indicating that limited vascular access inherently constrains ECM normalization, even when PRP-driven remodeling is present.

3. Discussion

The central objective of this work was to determine whether the increase in the concentration of platelets within biologically validated ranges generated measurable, dose-dependent improvements in cytokine release, vascular infiltration, and adipocyte survival in vivo. Two PRP doses were tested based on established biological response thresholds: the low dose (0.1 × 10^6^ platelets/μL), which reflects the minimum concentration shown to elicit cellular responses, and the high dose (1.0 × 10^6^ platelets/μL), which corresponds to the widely used therapeutic concentration that has been demonstrated to improve AT graft outcomes [12,26,28,29,30].

The two PRP formulations differed by tenfold in platelet content, however, their 1-h VEGF, PDGF-BB, and TGF-β1 release profiles were nearly identical. This finding is consistent with the well-described activation-dependent burst release of platelet-stored growth factors, during which a substantial fraction of available cytokines is rapidly released upon activation, potentially masking dose-dependent differences at early time points. In addition, cytokine levels were normalized to platelet input (pg per 10^6^ platelets), which reflects the early release on a per-platelet basis rather than total cytokine availability within the construct. Together, these factors likely contribute to the similar cytokine profiles observed at this early time point despite substantial differences in platelet concentration [19,27]. Given this rapid release kinetics, our goal was to implant the constructs immediately following activation. Hence, subsequent analyses focused on in vivo outcomes rather than extended in vitro release profiling. This strategy allowed us to evaluate whether platelet dose exerts its primary influence during later phases of integration through sustained paracrine signaling, matrix remodeling, and vascular guidance rather than during the initial activation window. Notably, the PRP incorporation did not induce cytotoxic effects, as adipocyte viability remained unchanged across all groups, confirming that both activation and dose escalation were well-tolerated within the AT bioink. This is consistent with the current literature [18,31,32].

In contrast to in vitro early release kinetics, in vivo results revealed clear dose-dependent differences, consistent with the reported literature [33]. High-dose PRP improved graft preservation and increased the proportion of highly infiltrated regions by more than eightfold relative to native tissue, and supported deeper and more uniform vascular penetration through the printed pore network [34,35]. These findings suggest that platelet concentration influences biological processes that act beyond the initial activation burst, potentially including sustained paracrine signaling, modulation of early inflammatory activity [36,37], interactions with the evolving fibrin matrix [38] and infiltrating host vessels [39], ultimately enhancing fat survival [12,18,40,41].

The presence of pronounced regional heterogeneity in perfusion needs to be discussed. As shown in the vascular infiltration maps (Figure 2), the constructs did not achieve uniform vascularization but instead developed distinct high- and low-infiltration zones [42]. It is acknowledged that high PRP doses may transiently increase vascular permeability during early angiogenic remodeling, raising the possibility of localized blood extravasation. However, in the present study, vessel infiltration mapping was restricted to discrete, morphologically identifiable vascular structures, and diffuse blood-filled regions without clearly distinguishable vessels were excluded from analysis. In addition, because implantation was performed in immunodeficient nude mice, this model is not suited to fully assess immune-driven inflammatory responses; nevertheless, preserved adipocyte viability at 2 weeks supports productive vascular integration rather than overt inflammatory tissue damage under the conditions tested. This spatial disparity influenced nearly every biological outcome quantified in this study. Adipocyte density, capillary formation, fibrosis severity, and collagen subtype composition were all strongly dependent on the local infiltration state [43]. Even with optimal PRP supplementation, low-infiltration regions, particularly those located near the construct’s geometric center retained reduced adipocyte density and elevated fibrosis compared with better-perfused regions [42,44]. This consistent pattern suggests that diffusion-limited nutrient and oxygen transport impose a central perfusion barrier that PRP alone cannot fully overcome [17].

Despite this challenge, high-dose PRP still improved outcomes within poorly perfused zones, increasing adipocyte density nearly 5-fold and reducing fibrosis by ~35% compared with native AT. These improvements indicate that PRP can partially mitigate the adverse biological conditions created by limited vascular access [18,30]. However, the persistent gap between high- and low-infiltration zones across all groups, together with the consistently poor integration observed in the geometric center of the construct, suggests that PRP dose alone cannot fully overcome spatial perfusion constraints. This is visible across vascular infiltration maps and all subsequent tissue-level measurements, implying that the distribution of pro-angiogenic cues within the graft may be just as important as their total amount. Accordingly, strategies that incorporate the spatial control of PRP or angiogenic factors, particularly targeting regions prone to poor perfusion, may further enhance uniform vessel ingrowth and improve overall graft viability beyond what dose optimization alone can achieve [45].

ECM remodeling analyses further supported these relationships. High-dose PRP shifted collagen composition toward a more compliant, regenerative phenotype, reducing collagen I and increasing collagen III [46]. However, low-infiltration domains retained a more fibrotic matrix, indicating that sufficient vascular access remains essential for complete ECM normalization regardless of biochemical supplementation.

The findings establish a clear in vivo dose–response relationship for PRP-functionalized AT bioinks and help explain the variable outcomes reported across the PRP literature. Thus, it demonstrates that (i) platelet activation dictates immediate cytokine release, (ii) platelet dose governs long-term remodeling and tissue rescue, and (iii) the therapeutic effect of PRP is fundamentally modulated by local perfusion gradients. This framework unifies prior inconsistencies in PRP-based soft-tissue regeneration studies and provides mechanistic guidance for rational PRP design.

From a translational perspective, this study suggests several key design considerations. First, PRP concentrations within the upper therapeutic range may be required to maximize long-term vascular integration and tissue preservation. Second, because spatial perfusion heterogeneity limits the full impact of PRP, integrating strategies that enhance early vessel ingress, such as pore-guided architectures, spatial patterning of PRP, localized angiogenic factors, or pre-vascularized microvascular networks, may synergize with PRP to overcome central perfusion barriers. Finally, spatially controlled delivery approaches capable of directing vessel entry toward poorly perfused regions may yield more uniform adipocyte survival and reduced fibrosis across the entire graft volume.

4. Conclusions

This study demonstrates that the integration of PRP into a clinically relevant adipose tissue bioink enhances graft performance through a clear in vivo dose–response relationship. While early cytokine release was governed primarily by platelet activation rather than platelet quantity, high-dose PRP significantly improved graft retention, vascular infiltration, adipocyte survival, and ECM remodeling. The identification of persistent high- and low-infiltration zones across all groups highlights that vascular access remains a dominant regulator of regenerative outcomes, and that PRP, while beneficial, cannot fully overcome perfusion-limited constraints within poorly vascularized regions. Our findings indicate that platelet concentration and vascular accessibility synergistically shape tissue repair, and that optimizing both biochemical cues and spatial perfusion will be essential for uniformly restoring adipose graft viability. We establish the use of PRP-functionalized AT bioinks as a promising and tunable platform for soft-tissue reconstruction and highlights the need for future strategies that pair PRP dose optimization with spatially guided vascularization.

5. Methods and Materials

Isolation and Preparation of Biological Components. Subcutaneous AT and whole blood were collected from male Sprague–Dawley rats (8–10 weeks old; ORIENT BIO Inc., Gyeong-gi, Republic of Korea). Posterior and thigh fat pads were excised and manually minced using surgical scissors. AT micro-fragments were generated using a custom four-blade tissue micronizer, as previously described [24]. The minced tissue was combined with PBS (Catalog #PR2004-100-72, Biosesang) supplemented with 1% (v/v) penicillin/streptomycin (Catalog #PS-B, Capricorn Scientific, Ebsdorfergrund, Germany) in a Luer-lock syringe (Thermo Fisher Scientific, Waltham, MA, USA) and mechanically fragmented by repeatedly passing the suspension through the micronizer (approximately 20 back-and-forth passes). The resulting AT micro-fragment suspension was used immediately after preparation to maintain tissue viability. For PRP preparation, whole blood was collected from the inferior vena cava into anticoagulant-treated tubes and processed by sequential centrifugation at 1300 rpm for 5 min followed by 2000 rpm for 10 min to isolate PRP, as previously reported [47]. Isolated PRP was used immediately after preparation. All animal procedures were conducted in accordance with the IACUC guidelines at UNIST (Ulsan, Republic of Korea) under the approved protocol UNISTIACUC-22-48.

Bioink Preparation. AT bioinks were formulated by incorporating freshly prepared AT micro-fragments into a gelatin–fibrinogen hydrogel matrix, following an adapted protocol based on previously reported methods [24]. Gelatin (75 mg mL^−1^; Catalog #G6144, Sigma-Aldrich, St. Louis, MO, USA), fibrinogen (20 mg mL^−1^; Catalog #F8630, Sigma–Aldrich), and HA (6 mg mL^−1^) were dissolved in minimum essential medium (MEM; Catalog #M8167, Sigma–Aldrich) to generate the base hydrogel. To achieve the final bioink composition, the hydrogel phase was prepared at twice the target concentration and subsequently mixed 1:1 (v/v) with AT micro-fragments, resulting in final concentrations of 37.5 mg mL^−1^ gelatin, 10 mg mL^−1^ fibrinogen, 3 mg mL^−1^ HA, and 50% (v/v) AT. For PRP-containing formulations, isolated PRP was incorporated into the AT-based bioink at the desired final platelet concentrations of 0.1 × 10^6^ platelets μL^−1^ (low-dose PRP-AT) or 1.0 × 10^6^ platelets μL^−1^ (high-dose PRP-AT) immediately prior to printing. To enable structural stability during printing, the bioink was incubated at 4 °C for 10 min to induce partial gelatin gelation. Following printing, fibrin polymerization and platelet activation were initiated by treating the constructs with thrombin (10 U mL^−1^; Catalog #T7326, Sigma–Aldrich) supplemented with calcium chloride (22.8 mM CaCl_2_; Catalog #10035-04-8, Junsei Chemical Co., Tokyo, Japan) in PBS for 30 min at room temperature.

3D Bioprinting. All constructs were fabricated using an in-house-built multi-material extrusion bioprinter operated under sterile conditions. Polycaprolactone (PCL) scaffolds (12 × 12 × 1.8 mm^3^, 25-pore design) were printed by extruding PCL through a 200-µm metal nozzle at 200 kPa using a pneumatic dispenser (ML-808, Musashi Engineering, Tokyo, Japan). The AT bioink was printed through a 400-µm nozzle at a flow rate of 0.34552 µL/s and a printing speed of 75 mm/min using a mechanical dispenser (SMP III, Musashi Engineering). Following fabrication and platelet activation, constructs were implanted subcutaneously.

ELISA-Based Cytokine Quantification. Cytokine release from PRP-integrated bioinks was evaluated after a 1 h static incubation at 37 °C. PRP was adjusted to predefined platelet concentrations prior to incorporation into AT-based bioink formulations. Bioink samples were incubated in culture medium, after which the conditioned medium was collected for analysis. Cytokine levels were quantified using ELISA kits (R&D Systems, Minneapolis, MN, USA) according to the manufacturers’ instructions: VEGF (Rat VEGF Quantikine ELISA, Catalog #RRVOO), PDGF-BB (Quantikine ELISA, Catalog #MBB00), and TGF-β1 (Human/Mouse/Rat/Porcine/Canine Quantikine ELISA, Catalog #DB100C). ELISA measurements were used to calculate the total amount of cytokine released into the conditioned medium. These values were normalized to the predefined platelet input for each PRP-bioink condition by dividing the total cytokine amount by the corresponding platelet number and scaling to 10^6^ platelets. Normalized cytokine release is reported as picograms per 10^6^ platelets, enabling a direct comparison across formulations with different platelet concentrations.

Live/Dead Viability Assessment. Cytocompatibility was evaluated using the LIVE/DEAD Viability/Cytotoxicity Kit (Invitrogen, Catalog #L3224, Waltham, MA, USA). Fresh AT fragments were incubated in high-glucose DMEM with or without PRP for 1 h to match the pre-printing incubation conditions. Printed AT constructs and native AT controls were subsequently assessed. To verify staining specificity and confirm assay performance, a negative control was included to induce complete cell death (Figure S2). AT samples designated as negative controls were incubated in detergent-containing solution for 1 h, then the samples were stained using the LIVE/DEAD Kit under the same conditions as experimental groups. A Leica DM2500 microscope (Leica Microsystems, Wetzlar, Germany) was used during imaging. Live and dead cells were identified based on calcein-AM (green, live) and ethidium homodimer-1 (red, dead) signals, respectively. Quantitative analysis of live/dead ratios was performed using ImageJ, version 1.53a.

In Vivo Experimental Design, Control Groups, and Species Selection. To enable clear interpretation of PRP-mediated effects, control groups were defined to distinguish between unprocessed AT and biofabricated AT constructs while controlling for graft mass. Native AT controls consisted of freshly harvested AT implanted directly without micronization, hydrogel encapsulation, or printing. These grafts retained native tissue architecture and irregular geometry. The wet weight of native AT grafts was matched to that of implanted AT bioink constructs (without PCL) to ensure equivalent tissue mass across groups. In contrast, AT bioinks without PRP controls were generated by processing AT into micro-fragments, incorporating them into a base hydrogel at the same final concentrations described above and fabricating constructs with defined dimensions using identical bioprinting parameters as PRP-containing groups. This design isolates the effects of biofabrication and scaffold-mediated structural support from those of PRP incorporation while maintaining equivalent AT mass. A rat-to-mouse xenogeneic implantation model was employed to facilitate the controlled in vivo evaluation of PRP-mediated effects. This approach was selected because the preparation of PRP requires relatively large blood volumes to achieve precise and reproducible platelet concentrations. To obtain sufficient PRP without necessitating the sacrifice of multiple mice per experimental group, PRP was prepared from rats, while implantation was performed in immunodeficient nude mice to minimize immune-mediated responses. This strategy enabled ethical animal use while maintaining experimental consistency and physiological relevance.

In Vivo Implantation. Eight-week-old male nude mice (ORIENT BIO Inc.) were used for subcutaneous implantation. All procedures were performed under isoflurane inhalation anesthesia. A 2 cm dorsal incision was made, and a subcutaneous pocket was created for implantation. Constructs were implanted subcutaneously (n = 3 per group) according to the experimental groups defined above, including native AT, AT bioink without PRP, and PRP-integrated AT bioinks containing low or high PRP doses. Implants were retrieved 2 weeks post-implantation. Gross images were obtained at the time of harvest, and graft retention was quantified as the percentage of mass remaining at retrieval relative to the initial implanted mass.

Vessel Infiltration Heatmap. Spatial vessel infiltration heatmaps were generated from gross photographs of explanted constructs based on the presence of macroscopic, blood-filled vascular structures visible within the grafts. Vessel infiltration was defined as the spatial distribution and density of visually identifiable vessels penetrating the construct, representing functional vascular ingrowth at the tissue level. High-resolution images were imported into ImageJ (NIH), where color channel separation was used to enhance the contrast between blood-containing vessels and the surrounding scaffold matrix. Each construct was subdivided into a uniform 5 × 5 grid. Within each grid cell, visible vessels were manually identified and counted, and the degree of vessel infiltration was quantitatively classified into five predefined categories (0%, 1–25%, 26–50%, 51–75%, and 76–100%) based on the proportion of the grid area occupied by vascular structures. The resulting values were recorded as a spatial intensity matrix for each construct. Spatial intensity matrices were subsequently visualized as color-coded heatmaps to illustrate regional vessel distribution across constructs. Heatmap visualization was performed by converting the manually generated numerical matrices into graphical representations using a Python-based (Python, version 3.14.3) rendering workflow implemented via a generative AI computational tool (ChatGPT, OpenAI, San Francisco, CA, USA, GPT-4).

Rheological Characterization. The rheological properties of the bioinks were characterized using a rotational rheometer (Anton Paar, Graz, Austria). Viscosity measurements were performed using a shear-rate sweep following thermal gelation of the gelatin component by incubation at 4 °C for 10 min. Measurements were conducted using a 20 mm parallel-plate geometry over a shear-rate range of 0.1–100 s^−1^ at 4 °C. After crosslinking, the storage modulus (G′) and loss modulus (G″) were measured by dynamic frequency sweep analysis over a frequency range of 0.1–100 rad/s at 1% strain using the same rheometer and geometry.

Histological and Immunostaining Analysis. Following retrieval, samples were immersion-fixed in 4% paraformaldehyde at 4 °C for 24 h, processed through sequential ethanol and xylene dehydration steps, and subsequently paraffin-embedded. Sections (8 µm) were stained with Masson’s trichrome (Catalog #25088-1, Polysciences, Warrington, PA, USA) and Picrosirius Red (Catalog #ab150681, Abcam, Cambridge, United Kingdom). Masson’s trichrome-stained sections were imaged using a dotSlide digital virtual microscope (BX-51/22; Olympus, Tokyo, Japan), while Picrosirius Red-stained sections were imaged under polarized light using a THUNDER imaging system (Leica Microsystems). Fibrosis was quantified in ImageJ as the percentage of Masson’s trichrome-positive area within regions of high and low vascular infiltration, identified directly from histological sections based on the local density of visible blood vessels, whereas collagen content was quantified as the percentage of Picrosirius Red-positive area across the entire construct. For immunostaining, paraffin sections were deparaffinized, subjected to antigen retrieval in citrate buffer (Sigma–Aldrich), and blocked with 5% bovine serum albumin (BSA; Sigma–Aldrich). Sections were incubated overnight at 4 °C with primary antibodies against CD31 (AF3628, 1:40; R&D Systems) and perilipin-1 (PA1-1051, 1:200; Thermo Fisher Scientific). After washing with PBS, sections were incubated with Alexa Fluor-conjugated secondary antibodies (Abcam), counterstained with DAPI, and imaged using a confocal microscope (LSM700; ZEISS, Oberkochen, Germany). CD31 density was quantified as the CD31-positive vessel area normalized to total tissue area (mm^2^/mm^2^), and adipocyte retention was quantified as the number of perilipin-positive cells per mm^2^. Both parameters were analyzed separately in regions of high and low vascular infiltration within each section to capture spatial differences in vascularization and adipose tissue preservation.

Statistical Analysis. All results are shown as the mean ± standard deviation (SD). Statistical tests were conducted using GraphPad Prism 9 (GraphPad Software, San Diego, CA, USA). Two-tailed Student’s t-tests were used to compare two groups, and one-way ANOVA with Tukey’s test was used when comparing more than two groups. Differences were considered significant at p < 0.05 (* p < 0.05, ** p < 0.01, *** p < 0.005).