Mechanical Processing of SVF-Enriched Microfat for Reconstruction of Traumatic Soft Tissue Defects

Summary

This protocol describes a reproducible method for preparing mechanically processed SVF-enriched microfat from autologous adipose tissue and injecting it into cavity-type traumatic soft tissue defects for clinical reconstruction.

Abstract

Traumatic soft tissue defects pose significant challenges for reconstruction due to tissue loss, impaired vascularity, and difficulties achieving durable coverage. Adipose tissue offers a practical autologous tissue source, and mechanically processed stromal vascular fraction (SVF)-enriched microfat can be prepared intraoperatively without enzymatic digestion.
This study presents a standardized clinical protocol for harvesting autologous adipose tissue and processing it into SVF-enriched microfat for injection into cavity-type traumatic soft tissue defects. Fat is harvested manually from the thigh or abdomen under low negative pressure, mechanically fragmented by cutting and syringe-to-syringe emulsification, filtered to achieve uniform microfat consistency, and centrifuged to isolate the SVF-containing fraction. The processed microfat is injected throughout the wound cavity in a multilayered pattern. Postoperative assessment includes serial clinical evaluation, photographic documentation, and measurement of wound-area reduction until epithelialization.
In a small cohort, the method was associated with progressive wound contraction and complete epithelialization within approximately 4-8 weeks, with no major complications. Although cellular composition and viability were not quantified, the technique provided a feasible intraoperative approach suitable for settings without access to enzymatic processing or laboratory facilities. This protocol offers a practical, minimally manipulated method for delivering SVF-enriched microfat in managing traumatic cavity-type defects and may serve as a foundation for further controlled studies.

Introduction

Traumatic cavity-type soft tissue defects remain a major reconstructive challenge because they combine tissue loss, impaired local perfusion, and a high risk of infection and dead-space formation. Conventional coverage techniques, such as split-thickness skin grafts or flap transfers, provide durable coverage in many cases. However, they are frequently limited by donor-site morbidity, technical complexity, and variable long-term outcomes, especially in contaminated or scarred beds^1,2,3.

Adipose tissue is an abundant, easily accessible source of a heterogeneous cell population referred to as the stromal vascular fraction (SVF). SVF includes mesenchymal stromal cells, endothelial progenitors, pericytes, and supporting stromal elements. When retained within fat tissue particles, the native extracellular context of SVF is preserved during handling and clinical delivery⁴. Clinically, fat grafts enriched for SVF (SVF-enriched microfat) have been reported to improve graft retention and to be associated with accelerated wound epithelialization across a variety of indications^5,6,7.

Two principal strategies exist for obtaining SVF from lipoaspirate. Enzymatic digestion typically yields higher nucleated cell counts per unit volume but requires dedicated reagents, laboratory infrastructure, longer processing times, and is subject to regulatory restrictions in many jurisdictions^4,8. In contrast, mechanical processing methods include cutting, syringe-to-syringe emulsification, filtration, and the use of closed-system mechanical devices. These approaches permit rapid intraoperative preparation of SVF-enriched microfat with minimal manipulation and shorter turnaround times^6,9,10,11. Recent mechanical systems have reported processing times <15 min and yields approaching those of enzymatic methods in some series, though measured cell counts and viability can vary by device and operator⁹.

Despite the expanding literature on mechanically processed SVF and on fat grafting for chronic ulcers and diabetic foot disease, standardized, reproducible protocols specifically targeted to cavity-type traumatic soft tissue defects are scarce^7,12. Published clinical reports commonly address chronic ulcers, diabetic wounds, or aesthetic fat grafting. However, only a few provide an intraoperative stepwise protocol that specifies harvest parameters, emulsification endpoints, filtration pore sizes, centrifugation force (× g), dosing per wound area, and wound-readiness criteria for injection in trauma settings^7,13.

Practical applicability guidance is therefore important for surgical teams considering this approach. Based on the available literature and our operative experience, typical intraoperative aspirate volumes for a single cavity are ~20-40 mL. This volume generally yields sufficient processed microfat to fill small to moderate cavities. In contrast, large-volume reconstructions are likely beyond the scope of point-of-care mechanical processing and may require staged procedures or alternative strategies^9,12. Mechanical SVF approaches are also less suitable for grossly contaminated wounds until infection is controlled; in such cases, adjunctive debridement and infection management (including targeted antibiotics and, where appropriate, negative pressure wound therapy) should precede grafting⁷.

The present work aims to provide a detailed, reproducible intraoperative protocol for preparing SVF-enriched microfat by mechanical processing and for injecting this product into cavity-type traumatic soft tissue defects. The protocol emphasizes explicit operational parameters (harvest, mechanical fragmentation and emulsification, filtration, centrifugation expressed as × g, injection technique, and objective wound-area measurement) so that other surgical teams can adopt and validate the method in their own settings.

Protocol

All procedures were approved by the Institutional Ethics Committee (Approval No. KL-2025062) and conducted in accordance with the Declaration of Helsinki. Written informed consent was obtained from all patients prior to participation.

1. Patient selection and preoperative assessment

1. Include adult patients presenting with cavity-type traumatic soft tissue defects requiring reconstructive intervention after completion of adequate surgical debridement.
   ​NOTE: Defects should demonstrate a well-defined cavity with surrounding viable tissue and no ongoing necrosis at the time of reconstruction.
2. Exclude patients with active systemic infection or uncontrolled local wound infection, poorly controlled diabetes mellitus (defined as HbA1c persistently >8% despite treatment), significant peripheral vascular disease affecting the involved limb, known coagulation disorders or current anticoagulation that cannot be safely interrupted, malignancy involving the defect site, or contraindications to liposuction or anesthesia as determined by preoperative assessment.
3. Perform Baseline data collection and wound assessment
   1. Record baseline patient characteristics, including age, sex, wound location, injury mechanism, and time from injury to reconstruction.
   2. Perform wound assessment after standard wound preparation. Measure wound length and width using a sterile ruler at the widest points. Acquire standard digital photographs with a fixed distance and orientation. Calculate wound area (cm²) using planimetric analysis based on calibrated photographs (see Step 6.6).
4. Provide preoperative counselling. Explain the procedural steps, anticipated benefits, potential risks (including infection, fat resorption, and need for additional procedures), postoperative care requirements, and follow-up schedule to the patient.
5. Confirm understanding and obtain written informed consent prior to surgery

2. Preoperative preparation

1. Preparation of sterile equipment
   1. Prepare the required sterile equipment, including: liposuction cannulas (2-3 mm diameter, blunt tip), Luer-lock syringes (10-20 mL capacity), sterile Luer-lock connectors for syringe-to-syringe transfer, surgical scissors, blunt-tip injection cannulas (22G × 50 mm).
   2. Confirm the integrity and sterility of all devices before use.
2. Preparation and infiltration of tumescent solution
   1. Draw 1,000 mL of 0.9% normal saline into a sterile container
   2. Add 2 mL of 1:1,000 epinephrine to achieve 1:500,000 final concentration. Mix thoroughly under sterile conditions.
   3. Connect the solution to a Luer-lock syringe with a 2-3m blunt infiltration cannula
      ​NOTE: Lidocaine is intentionally omitted to allow separate, dose-controlled anesthesia administration.
   4. Introduce the solution into the subcutaneous donor site (e.g., abdomen or thigh) using slow, fan-shaped passes, from deep to superficial planes.
   5. Adjust the total volume according to donor-site surface area and anticipated lipoaspirate volume.
   6. Infiltrate until tissue demonstrates uniform tumescence and vasoconstriction.
   7. Wait 10-15 min after infiltration to allow maximal vasoconstriction before harvesting.NOTE: Epinephrine reduces intraoperative bleeding and facilitates adipose harvest. Omitting lidocaine prevents exceeding safe anesthetic doses and allows separate local or regional anesthesia.
3. Administration of anesthesia
   NOTE: Anesthesia is administered separately from the tumescent solution, selected based on defect size, donor site, and patient tolerance. One of the following anesthesia approaches is selected based on defect size, donor site, and patient tolerance.
   1. Local infiltration (field block): Infiltrate 0.5-1% lidocaine with epinephrine 1:200,000 using a blunt tip (22G × 50 mm) cannula. Spread radially around sensory nerves supplying the donor site. Allow 5-10 min for full anesthesia effect before donor-site manipulation or tumescent infiltration.
      ​NOTE: Maximum dose: 7 mg/kg lidocaine with epinephrine, adjusted for patient weight and comorbidities.
   2. Regional anesthesia (optional): Use ultrasound-guided peripheral nerve blocks based on the donor site. Use bupivacaine (0.25-0.5%) or ropivacaine (0.5% ), dosed per standard guidelines. Confirm block onset (10-20 min) before harvesting.
   3. General anesthesia: Reserved for large defects or combined procedures. Administer per institutional protocols with standard monitoring.
      CAUTION: Ensure that the total systemic epinephrine dose from all infiltrated solutions remains within accepted clinical safety limits.

3. Fat harvesting

1. Select the donor site (abdomen and/or lateral thigh) based on the following criteria
   1. Availability of sufficient subcutaneous adipose tissue to allow adequate harvest without contour deformity
   2. Absence of local scarring, infection, or previous surgery that may compromise tissue quality.
   3. Patient accessibility and positioning on the operating table to permit sterile infiltration and aspiration.
      ​NOTE: Selecting a site with adequate tissue and minimal prior trauma facilitates consistent adipose harvest and reduces procedural complications.
2. Make a 2-3 mm skin incision using a No. 11 scalpel, under sterile conditions. Incise through the epidermis and dermis down to the subcutaneous layer without penetrating deeper fascia or muscle.
3. Maintain a small, controlled incision to minimize scarring.
4. Attach a 2-3 mm liposuction cannula to a 10-20 mL Luer-lock syringe.
5. Insert the cannula through the skin incision into the subcutaneous adipose layer, remaining superficial to the underlying fascia.
6. Use gentle, multidirectional movements to distribute the cannula evenly throughout the target subcutaneous plane.
7. Apply manual low negative pressure by gently withdrawing the syringe plunger to aspirate adipose tissue.
8. Avoid aggressive suction to minimize mechanical trauma to adipose-derived cells.
9. Continue aspiration in small, controlled aliquots, repositioning the cannula within the subcutaneous layer to maximize harvest efficiency.
   NOTE: Maintain the syringe at room temperature (20-25 °C) to preserve cell viability.
10. Harvest approximately 20-40 mL of lipoaspirate, adjusted to the defect size.
11. After aspiration, remove the cannula and close the skin incision with a single 3-0 or 4-0 nylon suture, or use sterile adhesive strips for smaller stab incisions. Apply a sterile dressing to the donor site.
12. The aspirated lipoaspirate is initially in the Luer-lock syringe used for harvesting. Gently detach the harvesting cannula and transfer the lipoaspirate into a new sterile 10-20 mL Luer-lock syringe for further processing.
13. Avoid exposure to air, high temperatures, or excessive mechanical force. Maintain the aspirate at room temperature (20-25 °C) until processing (e.g., purification, centrifugation, or injection).
    NOTE: Using a fresh sterile syringe prevents contamination and allows standardized processing. Gentle handling preserves adipocyte and stromal vascular fraction viability

4. Mechanical processing of SVF-enriched microfat

1. Allow the harvested lipoaspirate to stand upright in sterile 10-20 mL Luer-lock syringes for 5 min at room temperature (20-25 °C) to permit gravity-based separation.
2. After standing, three distinct layers (upper oil layer [free lipid fraction], middle adipose layer, lower aqueous/blood layer) become visible.
3. Using sterile technique, hold the syringe vertically with the nozzle pointing upward and gently advance the plunger to expel the upper oil layer until only the adipose fraction remains. Invert the syringe and carefully remove the lower aqueous/blood layer by slow plunger advancement or aspiration using a sterile syringe. Retain only the middle adipose fraction for further processing.
   NOTE: Removal of free oil and aqueous components reduces inflammatory by-products and improves graft consistency and cell viability.
4. Transfer the retained adipose fraction into a sterile stainless-steel or glass dish. Using sterile surgical scissors, gently mince the adipose tissue into fragments approximately 1-2 mm in size, avoiding excessive compression or shear forces.
5. Load the minced adipose tissue into a 10 mL Luer-lock syringe. Connect this syringe to a second 10 mL Luer-lock syringe using a sterile female-to-female Luer-lock connector.
6. Mechanically emulsify the adipose tissue by transferring it back and forth between the two syringes for 20-30 passes at a steady, moderate rate. Continue until a uniform, injectable microfat consistency is achieved.
   NOTE: The processed fat should appear homogeneously emulsified with minimal visible oil separation.
7. Detach the Luer-lock connector and transfer the emulsified microfat from the syringe into a sterile centrifuge tube compatible with the centrifuge rotor. Ensure tubes are balanced by volume before centrifugation.
8. Centrifuge the emulsified fat at approximately 400 × g for 3 min at room temperature.
9. After centrifugation, three layers can be seen: Upper oil layer, Middle SVF-enriched microfat fraction, Lower aqueous/blood layer (Figure 1).
10. Using a sterile syringe, carefully aspirate only the middle SVF-enriched microfat layer, avoiding contamination from the adjacent layers.
11. Pass the collected fraction through a 500-1,000 µm sterile stainless-steel mesh filter to remove fibrous debris and large particles. This step facilitates smooth injection and minimizes cannula clogging during graft delivery.
12. Load the filtered SVF-enriched microfat into 1-5 mL sterile syringes for immediate injection.

5. Injection into the defect site

1. Perform final wound debridement followed by thorough irrigation with sterile saline until all necrotic tissue is removed.
2. Confirm wound readiness by the presence of healthy granulation tissue and/or punctate bleeding, indicating adequate perfusion and tissue viability.
3. Using sterile technique, introduce a blunt-tip cannula (1.2-2.0 mm diameter) into the defect through the wound edge or adjacent intact skin, avoiding direct entry through the central wound base when possible.
   1. Depth and tissue plane: Advance the cannula into the subcutaneous tissue plane immediately superficial to the wound bed; intramuscular placement is avoided unless specifically indicated by defect depth.
   2. Angle of insertion: Insert the cannula at a low oblique angle (approximately 10-30°) relative to the wound surface to facilitate controlled, layered deposition.
   3. Cannula positioning: Maintain the cannula tip within well-vascularized tissue planes to optimize graft survival and minimize extrusion.
4. Inject the SVF-enriched microfat using a multilayered, retrograde fanning technique, depositing small aliquots during slow cannula withdrawal.
5. Distribute the graft evenly across the wound base, margins, and surrounding subcutaneous tissue to achieve uniform filling and maximize contact with vascularized recipient tissue.
6. Adjust injection volume according to the size and depth of the cavity, typically ranging from 5-12 mL.
7. Injection is discontinued once the defect is adequately filled and tissue contours are restored, ensuring no over-correction or excessive tissue tension.
   NOTE: Avoid overfilling to reduce the risk of fat necrosis, impaired perfusion, or graft extrusion.
8. Cover the wound with Vaseline (petrolatum) gauze, placed in non-compressive contact with the wound surface.
9. Apply secondary dressing to protect the site while allowing passive drainage and minimizing shear forces on the injected microfat.

6. Postoperative care and follow-up

1. Administer prophylactic antibiotics according to institutional protocol, taking into account wound size, contamination status, and patient specific risk factors.
2. Instruct patients to avoid pressure, shear, or friction at both donor and graft sites for 1-2 weeks postoperatively to minimize graft displacement and optimize integration.
3. Inspect the wound dressing at each follow-up visit and replace Vaseline (petrolatum) gauze as needed. Do not forcibly remove adherent gauze; allow spontaneous detachment during epithelialization to prevent disruption of the regenerating tissue.
4. Schedule follow-up visits at 1, 2, 4, and 12 weeks post-procedure.
5. Acquire standardized digital photographs at each visit using a fixed camera at the wound distance, consistent lighting conditions, and a scale reference (e.g., sterile ruler) placed in the same plane as the wound. This ensures consistency for longitudinal assessment.
6. Measure wound area using ImageJ software (National Institutes of Health (NIH), USA) via planimetric analysis.
   1. Import standardized wound photographs into ImageJ
   2. Calibrate image scale using the reference ruler
   3. Manually trace the wound margin using the polygon selection tool.
   4. Calculate wound area automatically using the software's measurement function.
7. For outcome assessment, define complete epithelialization as full wound surface coverage without exudate, dressing requirement, or need for secondary intervention.
8. Record all postoperative complications, including infection, fat necrosis, hematoma, seroma, or delayed wound healing.
   CAUTION: Dispose of all biological waste in accordance with institutional bio-safety regulations.

Results

A total of eight patients with cavity-type traumatic soft tissue defects were treated using the described protocol.

Cohort characteristics

The cohort included five males and three females, with a mean age of 51.5 ± 11.7 years (range, 38-74 years). Defects were located on the lower limb (n = 5), upper limb (n = 2), and trunk (n = 1). The mean maximal wo...

Discussion

This study describes a clinically applicable and reproducible protocol for the mechanical processing and transplantation of SVF-enriched microfat in managing traumatic cavity-type soft tissue defects. The protocol is intended for point-of-care implementation in a standard operating room setting and prioritizes procedural simplicity, safety, and feasibility over biological characterization. In this small clinical series, all treated defects demonstrated progressive wound closure and achieved complete epithelialization wit...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.9% Normal saline	Baxter Healthcare (or equivalent)	Various	Used as a base solution for tumescent solution
Blunt-tip injection cannula (22G × 50 mm)	CONPUVON (;), China	DZ 22×50-C5	Used for multilayered injection of SVF-enriched microfat
Centrifuge	Longtime Biotechnology (), China	LTA-1600	Clinical centrifuge capable of generating approximately 400 × g
Digital camera / Smartphone	Any	N/A	Standardized wound photography during follow-up
Epinephrine	Local hospital pharmacy	Various	Added to saline to achieve a final concentration of 1:500,000
ImageJ software (Version 1.53 or later)	National Institutes of Health (USA)	Free software	Used for planimetric wound area measurement
Liposuction cannula (2–3 mm)	Standard medical supplier	N/A	Used for harvesting adipose tissue from donor site
Luer-lock connector (female-to-female)	Becton Dickinson (or equivalent)	Various	Used for syringe-to-syringe mechanical emulsification
Luer-lock syringe (1, 5, 10, 20 mL)	Hongda Medical Devices (), China	Not specified (institutional supply)	Used for aspiration, mechanical processing, and injection
Sterile dressing / bandage	Hospital Pharmacy	N/A	For postoperative wound coverage
Surgical scissors (sterile)	Guangzhou Baitang Medical Devices Co., Ltd.	BT00301 (or similar representative model)	Used for mechanical fragmentation of adipose tissue
Vaseline gauze (10 cm × 10 cm)	Huaxi Medical Dressing Co., Ltd. (), China	Not specified (institutional supply)	Non-adherent dressing used for postoperative wound care