MEW Bio 3D Printing Cultivates Muscle
Strong Adhesion, Neat Alignment, Striations Formed:
MEW Bio 3D Printing Cultivates Muscle
bioRxiv 2025 · University of Oregon
Once muscle loss exceeds 20% of its original volume, it essentially cannot regrow — trauma and tumor resection can both cause this “volumetric muscle loss,” and existing treatments struggle to rebuild muscle’s most valuable asset: its neatly aligned fibrous architecture.
A team at the University of Oregon posted a combination approach on bioRxiv: melt electrowriting (MEW) first weaves PCL microfiber “tracks” in three diameters, then wraps the fibers in a hyaluronic acid (HA) hydrogel coating, with the “cell-gripping” RGD peptide optionally tethered to the coating. Once muscle cells move in, they adhere more firmly and align more neatly, differentiating into myotubes oriented along the fibers within 7 days — the HA-coated group even showed sarcomeric striations.
Highlights
- MEW weaves “tracks”: full printing parameters for three microfiber diameters
- The chemistry of the “sugar coating”: dynamic covalent hydrazone crosslinking, RGD photografted in one minute
- Adhesion showdown: HA-RGD wins decisively, 20 µm is the “golden diameter” for muscle cells
- 7-day differentiation: oriented myotubes form, HA-coated group shows sarcomeric striations
01 Why Does 20% Muscle Loss Not Grow Back?
Skeletal muscle is highly ordered: the epimysium wraps the muscle bundle, the perimysium encloses fiber bundles inside, and each fiber is further wrapped by the endomysium — all fibers are aligned in the same direction, so contractile forces can sum into useful movement. This anisotropic arrangement is precisely the hardest part of muscle regeneration to replicate.
Volumetric muscle loss (VML) — over 20% volume loss caused by trauma, chronic disease, or surgical resection — has limited regenerative capacity; without intervention, it progresses to fibrosis and fatty infiltration. Existing options are either autologous muscle transfer (scarce tissue, donor-site damage) or rehabilitation plus long-term complication management — none can rebuild the ordered tissue architecture.
Tissue engineering scaffolds are a popular alternative route, but the two traditional paths each have weaknesses:
Synthetic fiber scaffolds (e.g., electrospinning, 3D-printed PCL): precise alignment, but lacking biochemical cues — cells adhere poorly and integrate badly;
Hydrogel scaffolds (collagen, PEG, GelMA, etc.): friendly microenvironment, but unable to support oriented muscle architecture, and with poor long-term stability.
The University of Oregon team’s approach fuses the two: MEW-printed microfibers provide “physical alignment cues,” then an ECM-derived HA hydrogel coating provides a “biochemical comfort zone” — physical and biochemical cues each handle one aspect, complementing each other’s weaknesses.
The experimental design is a 3×3 systematic comparison: three fiber diameters × three surface coatings, plus tissue culture plates as baseline — the effects of thickness and coating on muscle cell behavior, resolved in one go.
02 Weave the “Tracks” First: MEW Prints Three Microfiber Diameters
The tracks are woven by melt electrowriting (MEW) — one of the highest-precision members of the bio 3D printing family. The principle can be imagined as “charged winding”: PCL is melted, and when extruded from a fine nozzle, is drawn by a high-voltage electric field into stable, straight micron-scale filaments, deposited in pre-defined paths onto the collector. Compared to the tangled mess of electrospinning, MEW fibers are individually obedient, with controllable position — exactly the capability needed to make “alignment cues.”
The scaffold design is highly targeted: square 7×7 mm, two-layer structure — myoblast-alignment fiber spacing 150 µm (responsible for guiding alignment), perpendicular reinforcement fiber spacing 500 µm (responsible for holding the structure), with overall height controlled at about 200 µm. The three diameters (10/20/30 µm) are tuned by printing parameters, not by changing materials.
Material and nozzle: medical-grade PCL; melted at 75 °C; 22G grounded metal nozzle (protruding 1 mm from printhead); nozzle–collector distance 3.5 mm; printed on 1 mm glass slide (paper Methods)
Three parameter sets: 10 µm group 5.5 kV / 0.17 bar / 500 mm/min (6 aligned + 14 alternating layers); 20 µm group 5.5 kV / 0.65 bar / 345 mm/min (3 + 7 layers); 30 µm group 6.2 kV / 1.40 bar / 345 mm/min (2 + 5 layers) (paper Table 1)
Scaffold design: 7×7 mm; aligned fiber spacing 150 µm, perpendicular reinforcement fiber spacing 500 µm; total height about 200 µm; 10 µm group additionally has anti-buckling edge reinforcement (printed at 1 mm/min) (paper Methods)
Pre-treatment: 0.1 M NaOH etching 30 min → sterile ethanol → 10% serum-containing medium at 4 °C overnight equilibration (paper Methods)
03 Then Put On the “Sugar Coating”: How HA Hydrogel Wraps the Fibers
Choosing hyaluronic acid (HA) for the coating is deliberate: it is a natural component of the muscle extracellular matrix, capable of directly “talking” to muscle cells via cell-surface CD44 and RHAMM receptors, promoting migration, adhesion, and proliferation. But HA hydrogel has an inherent contradiction — the crosslinked polyanionic surface is actually unfavorable for cell adhesion. The team’s solution is to add “grips” to the coating: covalently graft fibronectin-derived RGD peptides.
The full chemical route is in three steps: first, HA is oxidized to generate aldehyde groups (HA-Ox, modification degree 51.0±10.5%); another pathway uses ion exchange to attach norbornene groups (HA-Nor, 24.0±2.3%), then via amide reaction attaches adipic acid dihydrazide (HA-Nor-ADH, 40.5±2.3%); finally, photoinitiated thiol-ene click reaction (365 nm UV for 1 min) attaches the cysteine-terminated RGD peptide — actual grafting reached 55.4±29.3 nmol/mg HA, below the 2.5 mM design target but still within the concentration range reported to support cell adhesion.
The coating itself is a simple “mix-and-set” operation: place the scaffold in an 8-chamber culture slide, first add 25 µL of 2% HA-Nor-ADH (or HA-RGD version), then drip 25 µL of 2% HA-Ox — aldehyde and hydrazide spontaneously undergo a Schiff base reaction to form hydrazone bonds, gelling in place into a thin layer of hydrogel within 30 minutes, fully wrapping the fiber network. This dynamic covalent crosslinked gel has low swelling, can remain stable for over 28 days, and is a mature system previously characterized by the team.
Base polymer: 100 kDa sodium hyaluronate; HA-Ox oxidized with 0.6 equivalents sodium periodate, protected from light, 4 h, modification degree 51.0±10.5% (paper Methods/Results)
Functionalization: norbornene modification degree 24.0±2.3%; ADH modification degree 40.5±2.3% (¹H-NMR quantification) (paper Results)
RGD grafting: CGRGDSG peptide, Irgacure 2959 photoinitiator, 365 nm irradiation 1 min; Ellman’s reagent measured 55.4±29.3 nmol/mg HA, about 1.1 mM / 25 µL (target 2.5 mM) (paper Methods/Results)
Coating: 25 µL 2% HA-Nor(-ADH) + 25 µL 2% HA-Ox, in-situ gelation over 30 min; SEM observation with freeze-dried samples, 7 nm titanium sputter coating (paper Methods)
04 Adhesion Showdown: HA-RGD Wins Decisively, 20 µm Is the Golden Diameter
The first-round test is adhesion: C2C12 mouse myoblasts seeded at about 57,000 cells/cm², high-serum proliferation for 24 h then switched to low-serum medium for 3 days to initiate early differentiation, then fixed and stained — F-actin labels the cytoskeleton, DAPI labels nuclei.
Quantitative results nail the conclusion: HA coating significantly improves adhesion at every diameter, and HA-RGD is significantly higher than HA. Taking the 20 µm group as example, no coating gives about 8–150 nuclei, HA coating 122–412, and HA-RGD directly to 815–1273 — an order of magnitude difference. Fiber diameter also matters: on HA-RGD coating, the 20 µm group has the highest nucleus count, exceeding both 10 µm and 30 µm groups.
Even more interesting is alignment. Nucleus orientation analysis showed: uncoated and HA-coated groups had randomly oriented nuclei (uniform distribution from −90° to 90°); while HA-RGD-coated 10 µm and 20 µm groups showed nuclei clearly concentrated at 0° — exactly the direction of aligned fibers. RGD “grips” not only make cells adhere in larger numbers, but also line them up along the tracks.
05 From Cells to Myotubes: 7-Day Differentiation, Striations Appear
Adhesion is only the first step — the ultimate task of muscle cells is fusion into myotubes: multinucleated tubular structures formed by fusion of multiple cells, the precursor to muscle fibers. The differentiation experiment switched to low-density seeding (about 14,000 cells/cm²), switched to 2% horse serum differentiation medium after 24 h, cultured for 7 days, then stained for myosin heavy chain (MHC, a late differentiation marker).
Fluorescence images continue the same trend: HA and HA-RGD coated groups had more cells and more obvious myotube formation; except for the uncoated 10 µm group, myotubes on all scaffolds were oriented along 0° — the physical “tracks” guided throughout the differentiation process.
Quantification added detail: nucleus count still highest in HA-RGD, optimal in the 20 µm group; on myotube length, the HA and HA-RGD coatings in the 10 µm group were significantly longer than uncoated (about 22–23 µm vs 14 µm), while in the 20 and 30 µm groups there was no difference between coatings — when the track is too fine, the “nourishing” effect of the sugar coating becomes prominent.
Myotube “maturity” is judged by two objective indicators. First is creatine kinase (CK) activity — enzyme activity peaks when muscle cells fuse into tubes. Measured at days 1/5/7: all groups collectively reached a peak at day 5, then declined at day 7, matching the myotube formation timeline.
Second is sarcomeric striations — the periodic arrangement of α-actinin within myotubes, the morphological gold standard for myotube maturation. Day-7 α-actinin staining gave an intriguing result: only the HA-coated group showed distinct striations, while uncoated and HA-RGD groups had not yet developed them.
At the gene level, differentiation progress is also evident: day-5 RT-qPCR showed early differentiation genes Pax7, MyoD, Myf5 and the late gene MyoG were all upregulated relative to undifferentiated cells, with MyoD significantly higher in the HA-coated group than HA-RGD, and MyoG significantly higher in the HA-coated group than uncoated — echoing the staining result that “the HA group has the most mature striations.”
Cells: C2C12 mouse skeletal myoblasts (ATCC), passages 7–9; adhesion experiments seeded at ~57,000 cells/cm², differentiation experiments at ~14,000 cells/cm² (paper Methods)
Culture timeline: adhesion group high serum 24 h → low serum 3 days; differentiation group high serum 24 h → 2% horse serum differentiation medium 7 days, medium changed every 3 days (paper Methods)
Readouts: MHC/α-actinin immunofluorescence + DAPI; spinning-disk confocal z-stack (slice thickness 0.4/2.5 µm, average depth 68–129 µm); IMARIS for nucleus counting and orientation analysis; CK activity kit colorimetric method (450 nm), BCA normalization (paper Methods)
Gene detection: day 5 TRIzol extraction, RT-qPCR for Pax3/Pax7/MyoD/Myf5/MyoG, GAPDH internal reference (paper Methods)
Not reported in paper: myotube diameter distribution, sarcomere number density statistics, gene expression fold-change values of coated groups vs culture-plate controls
06 Why 20 µm, and What Comes Next
The victory of 20 µm fibers is likely not coincidental: C2C12-formed myotubes have diameters between 5–25 µm — 20 µm fibers match the size of myotubes themselves, so cells can “wrap around” and adhere firmly; 10 µm is too fine to grip fully, and 30 µm too thick to reach. Mechanically, the tensile stiffness of the uncoated 20 µm scaffold is 0.843±0.219 MPa, in the same order of magnitude as the ~0.44 MPa reported for human parallel muscle fibers in the literature, providing appropriately scaled support for myotubes.
What’s intriguing is the division of labor between HA and HA-RGD: RGD maximizes “adhesion and alignment,” but striations and higher MyoD/MyoG expression appear in the HA group — the authors speculate this may relate to HA’s own signaling via CD44/RHAMM receptors, and to the hydrogel’s viscoelastic properties leaving room for cell migration and fusion. More adhesion does not equal faster maturation; the “formulation” of biochemical signals still needs further tuning.
The authors also candidly list remaining items: the system lacks exogenous growth factors (such as IGF-1, bFGF), myotubes are not fully mature; there is a gap between C2C12 cell line and primary muscle cells. Next steps include in vivo validation of a muscle repair patch in a rat VML model, and exploring conductive materials such as PPy/PEDOT with additional peptide signals grafted onto the HA scaffold.
Tensile testing: uncoated 20 µm scaffold along the aligned fiber direction, 0.005 mm/s constant-rate stretching, slope taken from 1–5% strain segment; stiffness 0.843±0.219 MPa (paper Results; universal solid mechanics analyzer)
Native reference: human skeletal muscle parallel fibers about 0.44 MPa (cited value); scaffold same order of magnitude as native tissue (paper Discussion)
Limitations (self-described): myotubes not fully mature (missing striations / only HA group has them); no growth factors added; gap between C2C12 and primary cells; reinforcement fibers occupy a share of aligned fiber content (paper Discussion/Conclusion)
MEW handles aligning the cells neatly, HA sugar coating handles keeping the cells comfortable
— rather than letting either side fight alone.
The systematic comparison of three diameters also yielded a practical conclusion:
fiber thickness must match the “body size” of the target cells,
and 20 µm happens to hit the exact size of myotubes.
📄 Paper Information
Title: Galindo AN, Chi AK, Liashenko I, et al. Hyaluronic Acid-Coated Melt Electrowritten Scaffolds Promote Myoblast Attachment, Alignment, and Differentiation. bioRxiv preprint, 2025.
DOI: 10.1101/2025.03.06.641880
Copyright: CC BY-NC-ND 4.0; this article is a preprint that has not undergone peer review
Author affiliation: University of Oregon Knight Campus et al.
This article is a research literature interpretation; all content is quoted from the published paper, for scientific knowledge dissemination only, and does not constitute any diagnostic or treatment recommendation, nor any product performance promise. The technologies and materials described have not entered clinical application; experimental data come from in vitro studies; the original paper is a preprint that has not undergone peer review, and conclusions should defer to the subsequent peer-reviewed published version.
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