Real‐Time Functional Assay of Volumetric Muscle Loss Injured Mouse Masseter Muscles via Nanomembrane Electronics

Abstract Skeletal muscle has a remarkable regeneration capacity to recover its structure and function after injury, except for the traumatic loss of critical muscle volume, called volumetric muscle loss (VML). Although many extremity VML models have been conducted, craniofacial VML has not been well‐studied due to unavailable in vivo assay tools. Here, this paper reports a wireless, noninvasive nanomembrane system that integrates skin‐wearable printed sensors and electronics for real‐time, continuous monitoring of VML on craniofacial muscles. The craniofacial VML model, using biopsy punch‐induced masseter muscle injury, shows impaired muscle regeneration. To measure the electrophysiology of small and round masseter muscles of active mice during mastication, a wearable nanomembrane system with stretchable graphene sensors that can be laminated to the skin over target muscles is utilized. The noninvasive system provides highly sensitive electromyogram detection on masseter muscles with or without VML injury. Furthermore, it is demonstrated that the wireless sensor can monitor the recovery after transplantation surgery for craniofacial VML. Overall, the presented study shows the enormous potential of the masseter muscle VML injury model and wearable assay tool for the mechanism study and the therapeutic development of craniofacial VML.

Note S1. Fabrication details of the wearable electronics system.

Graphene ink preparation
1. For the electrochemical exfoliation, 10 V was applied between the graphite (Alfa Aesar) and Pt foil in an electrolyte solution of ammonium sulfate ((NH4)2SO4, Sigma-Aldrich). 2. As-exfoliated graphene was purified using deionized water (DI water) and further filtered under vacuum to remove the residuals. 3. The filtered wet powder of graphene was dispersed in DI water and concentrated to 15%. Hard bake at 240 °C for 1 h in a vacuum oven. 8. Spin-coat PR (AZ P4620, Integrated Micro Materials) at 2000 RPM for 30 sec, and soft bake at 90°C for 4 min. Photolithography exposing UV light with intensity of 15 mJ/cm² for 100 s. Develop with a developer (AZ-400K, Integrated Micro Materials) diluted with DI water (AZ-400K: DI water = 1:4). 9. Etch for via hole with reactive ion etcher (RIE). 10. Deposit 2 µm thickness of 2 nd Cu by sputtering. 11. Spin-coat PR (AZ P4620) at 1500 RPM for 30 s, and soft bake at 90°C for 4 min.
Photolithography exposing UV light with intensity of 15 mJ/cm² for 120 s and develop. 12. Etch exposed Cu with Cu etchant. 13. Spin-coat 3 rd PI layer (PI-2610) at 3000 RPM for 60 s. Soft bake at 100°C for 5 min and hard bake at 240°C for 1 h in a vacuum oven. 14. Spin-coat PR (AZ P4620) at 900 RPM for 30 sec, and soft bakes at 90°C for 4 min.
Photolithography exposing UV light and develop. 15. Etch exposed PI with RIE. 16. Peel off the microfabricated circuit with a water-soluble tape from the PDMS/Si wafer. 17. Mount microchip components with screen-print low-temperature solder paste. Figure S1. Printing process of graphene electrodes. A) Schematic illustration of the printing process of electrodes. Atomization begins in the ultrasonic vial, and a carrier gas (N2) flows the graphene ink droplets through the tubing, the diffuser, and the deposition head, where a sheath gas focuses the particles into a narrow stream. B) Details of inks and printing parameters of PI and graphene. Resistivity and skin-contact impedance of the printed graphene were around 2 × 10 -3 Ω cm and 210.5 kΩ, respectively. Figure S2. Biocompatibility study via cell viability measurement. Fluorescence images showing the cultured keratinocyte cells on two types of substrates, including a control (polystyrene petri dish, left) and graphene integrated on an elastomer (right). Tests used human primary keratinocyte cells cultured in an incubator at 37°C with 5% CO2. In the incubator, the material samples were placed in a 24-well plate, and 5000 keratinocytes/cm2 were seeded. After 7 days in the incubator, keratinocyte cells were washed with phosphate-buffered saline (Fisher Chemical) and dyed with 0.1 ml of calcein blue AM (Thermo Fisher) in 0.9 ml of the cultured medium. Keratinocytes and the reagent were additionally stored in the incubator of 37°C for 10 min. The supernatant was then aliquoted in a 96-well plate for further biocompatibility.    Figure S6. Filtered EMG signals of a post-VML-injured mouse. There is no signal variation for chewing behavior due to masseter muscle loss between A) non-eating and B) eating. Chewing EMG signals of 3 mice with C) post-VML-injured after 30 days and D) uninjured masseter muscle.

Figure S7. Dysregulated stem cells in masseter muscles at 3 days post-VML injury. (A)
Scheme of experiments. Tamoxifen was injected Pax7 cre/ERT ;tdTomato mice for 5 days to induce tdTomato fluorescence expression in satellite cells. VML or freeze injury was performed 10 days after tamoxifen injection. Mononucleated cells were isolated for flow cytometry analysis at 3 days after injury. (B) Representative dot plot of satellite cells, which are gated by red fluorescent protein (tdTomato), using flow cytometry. (C) Number of satellite cells from VML-injured masseter muscles is comparable with one of uninjured masseter muscles. Satellite cell numbers are normalized to average satellite cell number of uninjured muscles. Freeze-injured muscles are served as a positive control. Error bars represent standard error of the mean (SEM). (D) Representative histogram of fibroadipose cells (FAPs), which is defined by surface markers (Cd31 -, CD45 -, Sca1 + ) using flow cytometry. (E) Ratio FAPs to satellite cells is increased in VML-injured masseter muscles. Data is analyzed 1-way ANOVA and Kruskal-Wallis method for post-hoc comparison. *p<0.05. Figure S8. Filtered EMG signals of an uninjured mouse while eating. Motion artifacts make a significant and distinguishable amplitude comparing with the chewing signals. Figure S9. (A) Representative images of 3-mm punch on masseter and TA muscles of female mice. (B) Images of the biopsied masseter and TA muscles to compare general mass for transplant. Chewing RMS EMG signals of 3 mice with (C) TA muscle-transplanted and (D) masseter muscletransplanted at craniofacial muscle area. Figure S10. Schematics of the device fabrication process. A) Fabrication process of stretchable graphene membrane electrodes and B) microfabrication of thin film-based flexible circuit. Table S1. Power analysis of animal study Movie S1. Wireless EMG monitoring on the masseter muscle of mouse with miniaturized portable electronics.