A Protocol for co-Injecting Cells with Pulverized Fibers for Improved Cell Survival and Engraftment

Materials and reagents

Reagents

1. Polylactide-co-glycolide acid (PLGA) 82:18 lactide:glycolide (Corbion Purac, catalog number: PLG8218)

2. Hexafluoroisopropanol (Oakwood Chemical, catalog number: 3409)

3. 0.9% sodium chloride solution (saline) (Henry Shein, catalog number: 1047098)

4. Ethanol, 200 Proof (100%) (Decon Labs, catalog number: 2701)

5. Betadine (Henry Schein, catalog number: 67618–150-09)

6. Isopropyl alcohol (Fisher Scientific, catalog number: A416P-4)

7. Isoflurane (Primal Health Care, catalog number: NDC 66794–017-10)

8. Artificial tear ointment (Henry Schein, catalog number: 1338333)

9. Nairing cream (Nair, catalog number: 339823)

Solutions

1. 70% ethanol solution (see Recipes)

Recipes

1. 70% ethanol solution

   Mix 35 mL of 100% ethanol with 15 mL of sterile water, to create 50 mL of 70% ethanol solution.

Laboratory supplies

1. 0.5 mm sieve (IKA, model: 2939000)

2. 0.25 mm sieve (IKA, model: 2938900)

3. 2 mL Eppendorf Tubes (VWR, catalog number: 87003-298)

4. 50 mL Falcon tubes (VWR, catalog number: 89039-660)

5. Alcohol swabs (Henry Shein, catalog number: HS1007)

6. Cotton tip applicator (McKesson, Q-Tip, catalog number: 785468)

7. Liquid glue (3M Vetbond, catalog number: B07Q39FL9M)

8. 6–0, C-22 black braided silk sutures (Henry Schein, catalog number: 101–2636)

9. Surgical tape (3M, catalog number: 1527–0)

10. 18-gauge needle (Central Infusion Alliance, catalog number: BD 305196)

11. 3 mL syringe (Fisherbrand, catalog number: 14–955-457)

12. Parafilm (Bemis, catalog number: PM999)

Equipment

1. Electrospinner (Glassman High Voltage, model: PS/FJS0R02.4)

2. Micro-miller (IKA Mini-Mill, model: 2836001)

3. Cold centrifuge (VWR, Eppendorf, model: 5430 R, catalog number: 76458-526)

4. Weigh scale (VWR, model: VWR-6001E, catalog number: 10204-996)

5. Analytical balance (VWR, model: VWR-310AC/CAL, catalog number: 89422-676)

6. Slide warmer with temperature control (4MD Medical, catalog number: CASCXH-2001)

7. Somnosuite low-flow anesthesia system (Kent Scientific, catalog number: 13-005-111)

8. Induction chamber (Kent Scientific, catalog number: Somno-0705)

9. Warming pad (Kent Scientific, catalog number: RT-0501)

10. Electric razor with vacuum (Remmington, catalog number: VPG-6530)

11. Tenotomy scissors (Fine Science Tools, catalog number: 14066–11)

12. Ridge-less forceps (Fine Science Tools, catalog number: 11016-17)

13. Germinator 500 (Braintree Scientific, catalog number: GER5287120V)

Procedure

1. Pulverized electrospun PLGA fiber fabrication

   For fiber fabrication, electrospinning was used to create ultra-fine fibers from poly(lactic-co-glycolic acid) (PLGA) polymer. The process begins with the preparation of a solution where PLGA is dissolved in an organic solvent (hexafluoroisopropanol), resulting in a homogeneous solution. This solution is then loaded into a syringe fitted with a needle, and a high-intensity electric field is applied between the needle tip and a grounded collector plate at a constant flow rate. The electric field induces the formation of a liquid droplet at the needle's tip, which elongates into a jet and is ejected toward the collector. As the jet travels, the solvent evaporates, and the polymer solidifies into very thin fibers, forming a non-woven mat. This process is driven by the interaction between electrostatic forces and the surface tension of the polymer solution, and it depends on factors such as the solution's viscosity, the strength of the electric field, and the distance between the needle and the collector. Pulverization of PLGA is then performed by breaking down the electrospun fiber mat into fine fragments. This is achieved through mechanical grinding, where the mat is subjected to shear and impact forces, reducing it to smaller fragments. Milled and un-milled fibers were obtained from Nanofibers Solutions (Columbus, OH) as outlined below:

   1. Dissolve PLGA, at a ratio of 82:18 lactide to glycolide, in hexafluoroisopropanol at 6% w/w (Figure 1).

      
      

      
      Figure 1. Pulverized electrospun poly(lactic-co-glycolic acid) (PLGA) fibers. A. PLGA at a lactide to glycolide ratio of 82:18. B. Electrospinning to get an electrospun fiber mat. C. The resulting PLGA injectable fiber fragments. Scanning electron microscopy (SEM) micrographs of the PLGA before and after the milling process.

      
      

   2. Mix at room temperature for at least 24 h.

   3. Electrospin the polymer solution at +12 kV/-3 kV at a flow rate of 5 mL/h until reaching a 0.1–0.3 mm thick nanofiber sheet at room temperature and humidity with a tip-to-collector distance of 20 cm using an 18-gauge needle.

   4. Evaporate the solvent from the nanofiber sheet for ~16 h at room temperature.

   5. Cut the electrospun mat into 1 cm wide strips and feed into the mini mill. This will grind the fibers into smaller fragments.

   6. Perform milling with the 60-mesh sieve for the first pass. Then, re-run the fibers that passed through that sieve through the 40-mesh sieve mill. Then, run the fibers that passed through that sieve through the 20-mesh sieve to achieve the final desired particle size.




2. Co-injectable nanofiber and adipose tissue preparation

   Adipose tissue was sourced from Cosmetic & Plastic Surgery of Columbus via standard minimally invasive procedures such as single port cannula liposuction; no further characterization of the tissue was done (See General Note 1).

   Note: Preparation should be done in a sterile environment for best results.

   1. Transport the tissue on ice and place it in appropriate containers (See General Note 2).

   2. Centrifuge at 195× g for 1 min at 4 °C (Figure 2).

      Note: This step will separate the tissue from oils and anesthetics that can permeate the tissue during liposuction surgery.

      
      

      
      Figure 2. Co-injectable nanofiber and adipose tissue. A. Adipose tissue collected from patients using standard liposuction procedures. B. The tissue is centrifuged to separate fat oils and anesthesia from viable cells. C. Pulverized fibers (50 mg) are combined with (D) adipose tissue (1 mL) to form an injectable solution (50 mg/mL).

   
   

   3. Discard excess oils and debris.

   4. Place 50 mg of fibers into a 2 mL tube.

   5. Add 1 mL of tissue to the fiber-containing tube (See General Note 3).

   6. Mix tissue and fibers into a 3 mL syringe and cover the end with parafilm for surgical use.




3. Fat engraftment surgery preparation

   1. Sterilize all surgical tools, surgical trays, and materials before surgery.

      Note: Between same-day animal surgeries, clean instruments with 70% ethanol solution and sterilize them using the germinator prior to the next surgery.

   2. Set and maintain the surgical warming pads at 37 °C during surgery.

   3. Place mouse cages onto a slide warmer set to 37 °C.

      Note: Each mouse should be placed in individual cages to prevent agitation of the wound site.

   4. Weigh each mouse and record baseline weights.

      Note: Baseline weight will be compared with post-surgery values to evaluate mouse recovery in the form of post-implantation weight loss.

   5. Completely remove hair from the incision area 24 h prior to surgery. Shave hair with the razor, apply Nairing cream directly to the skin with a cotton tip applicator, and leave it for 10 s. Remove the cream entirely with warm water and cotton balls.

      Note: It is crucial to remove the Nairing cream thoroughly to prevent serious skin burns.




4. In vivo fat engraftment surgery

   1. Place the mouse into the induction chamber (see General Note 4).

   2. Set the Somnosuite low-flow anesthesia system at 5% isoflurane in room air at ~500 mL/min and induce flow into the induction chamber to anesthetize the mouse (see General Note 5).

   3. At full anesthetic depth, move the mouse to the nose cone on the warming pad.

      Note: Confirm the anesthetic depth of the mouse by testing the toe-pinch reflex.

   4. Quickly set the anesthesia system at 1.5%–2% isoflurane at ~250 mL/min and induce flow into the nose cone for the duration of the procedure.

   5. Apply artificial tear ointment to the mouse’s eyes to maintain moisture.

   6. Apply betadine followed by isopropyl to the incision area using cotton tip applicators (repeat this step three times).

   7. Create a 1-inch incision horizontally on the upper and lower dorsal area using ridge-less forceps to gently pull the skin taut, allowing for an easier cut (Figure 3).

      
      

      
      Figure 3. Fat engraftment surgery. Prepare the back of the athymic nude mice for injection. Create a 1-inch incision horizontally on the upper and lower dorsal area for the injection of adipose + fibers and their internal control with only adipose tissue, respectively.

   
   

   8. Form a cavity under the mouse’s skin by detaching the surrounding fascia using tenotomy scissors.

   9. Push approximately 100–120 μL of warm saline into each side of the incision.

   10. Connect a 3 mL syringe loaded with adipose tissue and fibers to a 4 mm cannula.

       Note: Inject adipose tissue alone as a control, as opposed to the adipose tissue–fiber combination treatment.

   11. Push 1 mL from the syringe load into the cavity (see General Note 6).

       Note: Larger syringes (>3 mL) may be required to push the adipose tissue and fibers through the 4 mm cannula, as this can facilitate the displacement of larger volumes while generating less pressure, minimizing the risk of clogging. The syringe may have to be pumped back and forth to ensure that any remaining contents are injected, and it may require readjusting the cannula and extension of the syringe to push in all of the load.

   12. Close the cavity by suturing the skin with 6–0 braided sutures along the incision in a single interrupted suture pattern.

   13. Clean the skin of the mouse using an alcohol pad.

   14. Seal the incision line using liquid glue/bandage.

   15. Remove the mouse from the nose cone, weigh the mouse, and record post-surgery weight.

   16. Place the mouse into a new, clean cage and allow the mouse to recover.

   17. Monitor the status of the mouse until fully awake (1–5 min).

   18. Keep mice on the slide warmer at 37 °C throughout the duration of the study.

   19. Place mash (i.e., wetted food) and hydrogel in Petri dishes into each mouse cage to facilitate fluid/food intake.

Data analysis

Scanning electron microscopy (SEM) is widely used to characterize the physical properties of electrospun fiber scaffolds before and after milling, evaluating the effects on porosity, pore size, and fiber diameter, which impact cellular responses.

To assess the biocompatibility of the milled fibers with adipose tissue, in vitro cultures of adipocytes with the fibers can be used to run viability assays (e.g., live/dead kit) and fluorescence microscopy for quantifying live cells. Similarly, in vivo biocompatibility can be evaluated through subcutaneous implantation in immunocompetent mice, followed by histological analyses (e.g., hematoxylin and eosin staining) to detect cytoarchitecture differences and immunohistochemistry to analyze immune cell infiltration and expression of inflammatory mediators, as previously described [19].

For evaluating volume retention after adipose tissue implantation, width and length measurements are commonly used to assess graft volume [20–22] (Figure 4). Additional methods include water displacement, magnetic resonance imaging (MRI), computed tomography (CT), and ultrasound. Immunostaining is employed to verify graft vascularity using endothelial markers (e.g., CD31, VEGF, bFGF, lectin) [23,24]. Laser speckle imaging (LSI) can be used to evaluate graft perfusion, while flow cytometry, MRI, ultrasound, and micro-CT provide further insights into vascularization by measuring endothelial marker expression at the single-cell level (see General Note 7). When using this model, sample sizes should be estimated based on a power analysis. It is recommended to include at least four biological and technical replicates per group in each experiment. Additionally, to minimize bias, blinding of the animals or experimental groups is advised.

Figure 4. Fat engraftment outcomes. Monitor the volume retention regularly between the day of engraftment and the final day post-engraftment. Remove the graft and evaluate volume retention, displacement, vascularization, and perfusion.

Validation of protocol

This protocol has been used and validated in the following research article:

1. Das, et al. [19]. Injectable pulverized electrospun poly(lactic-co-glycolic acid) fibers improve human adipose tissue engraftment and volume retention. Journal of Biomedical Materials Research Part A (Figures 1 and 3).

General notes and troubleshooting

General notes

General note 1: Engraftment outcomes can be affected by the source of adipose tissue, with decreased cell proliferation being linked to factors such as increasing donor age, higher body mass index, diabetes mellitus, and other comorbidities [25].

General note 2: To ensure maximum cell viability, keep tissue samples on ice unless otherwise specified.

General note 3: While this study focused on the application of PLGA fibers, the protocol can be adapted to substitute or incorporate other materials, allowing for modified properties.

General note 4: Consider mouse characteristics such as age, sex, and housing status (single vs. group) as these factors may influence outcomes. This procedure is most commonly performed on mice aged 6–9 weeks. It is important to evaluate the immunoreactivity profile using immunocompetent mice. In this study, we used wildtype C57/BL6 (Jackson Laboratories, stock #000664) mice to test the immunoreactivity of PLGA fibers via subcutaneous implantation. Once the immunoreactivity was evaluated, male athymic nude mice (Jackson Laboratories, strain #002019) were used due to their immunodeficient nature, which allows for the study of human tissue engraftment without immune response interference [26]. Evaluating the immune response is crucial since it could lead to cell death, inflammation, and calcification of the engrafted fat tissue, complicating the study of fat engraftment.

General note 5: While the duration of the procedure and recovery period may vary, we recommend anticipating approximately 21 min from the initial induction of the mouse with isoflurane (step D2) until the completion of the procedure (step D16). Post-procedure, the mouse should be awake and alert within 3–5 min of the postoperative recovery period (step D18).

General note 6: In this study, human adipose tissue was engrafted into a mouse model. It is important to note that outcomes may vary as different adipose tissue sources or engraftment models are used, as the results are specific to human adipose tissue engraftment in mouse models.

General note 7: One limitation of this study is the evaluation period of adipose tissue engraftments. In this study, evaluations were conducted over 21 days, but a longer period can fully assess the host's response.

Acknowledgments

Schematics were created with BioRender.com. Funding for this work was partly provided by NIH grants (DP1DK126199, DP2 EB028110–01 to D.G.P).

Competing interests

Jed Johnson is a co-founder of Nanofiber Solutions, LLC. (Columbus, OH).

Ethical considerations

All animal experiments were performed in accordance with protocols approved by the Laboratory Animal Care and Use Committee at The Ohio State University (IACUC # 2016A00000074-R1 and IACUC # 2009A0006-R3).

All studies involving human adipose tissue were completed in accordance with a protocol that was approved by the Institutional Review Board (IRB) no. 180822–2. Informed consent was obtained by all human subjects prior to study.

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