Compression bioreactor for cultured meat

Abstract

Cultured meat is emerging as a sustainable and ethical alternative source of dietary protein. However, the commercial viability of cultured meat is fundamentally constrained by prolonged tissue maturation timelines and the high energy demands of conventional, motor-driven electronic bioreactors. To address these bottlenecks, this study introduces a novel, scalable, magnetic-valve controlled pneumatic compression bioreactor, which utilizes a mechanically tunable magnetic circuit to deliver precise cyclic compression without internal electronic sensors or digital feedback loops. We evaluated the system's biological efficacy by applying cyclic compressive strain (10% strain, 0.27 Hz, 1 h/day) to murine C2C12 myoblasts cultivated on scaffolds. Optical motion tracking validated the device's kinematic stability and stroke-to-stroke repeatability during continuous operation. Biological assays confirmed that passive pneumatic actuation dampened the impact forces, maintaining high cell viability comparable to static controls. Mechanical stimulation profoundly accelerated early myogenesis, inducing a highly significant 2.93-fold upregulation of the master transcriptional regulator MyoD within just 3 days. These findings demonstrate that precise mechanotransduction can replace prolonged static culture and reduce the industry's reliance on exogenous biochemical growth factors. Ultimately, this novel bioprocessing paradigm offers a sustainable, highly scalable framework to accelerate tissue organization and reduce operational costs in industrial cultured meat manufacturing.

Graphical abstract

Highlights

• •A pneumatic driven bioreactor without electrical control for cultured meat production.
• •Bioreactor mechanical loading significantly promotes myogenesis.
• •Stable cyclic compression maintains high cell viable (>90%).
• •In situ tissue maturation combined with minimized downstream support rapid and scalable biomanufacturing.

1. Introduction

Cultured meat has emerged as a promising alternative to conventional animal agriculture, driven by an increasing demand for sustainable, ethical, and resource-efficient protein sources (Jara et al., 2023). However, despite significant biological advances, large-scale production remains constrained by prolonged manufacturing timelines, substantial material consumption, and energy-intensive bioprocessing conditions. A key impediment is the long time needed for the formation of structured muscle tissue, which requires extended culture phases to achieve functional maturation (Allan et al., 2019).

The conventional production workflow, which encompasses cell seeding, expansion, myogenic differentiation, and tissue assembly, frequently takes several weeks per batch (Reiss et al., 2021). This increases operational expenses and limits throughput. The acceleration of these stages has primarily relied on exogenous biochemical growth factors to promote proliferation and differentiation (Ahmad et al., 2023). Despite the efficacy of this approach, concerns have been raised regarding its cost, scalability, and the potential ethical implications (Santos et al., 2023).

To address these biochemical limitations, mechanical stimulation has gained attention as a non-chemical, biomimetic strategy to modulate cell behavior and tissue development (Hu et al., 2023; Chiu et al., 2023; Sunadome et al., 2023; Shou et al., 2023). In contrast to exogenous growth factors, which depend exclusively on molecular signaling, mechanical cues engage cell-intrinsic mechanosensitive pathways (Haroon et al., 2021). These pathways emulate the physical microenvironment of native muscle. When applied with precision, these stimuli enhance not only cell proliferation and alignment but also accelerate myogenic differentiation and tissue maturation, which are needed to produce structured muscle in vitro.

Mechanotransduction is defined as the process by which mechanical forces are converted into cellular signals, and it governs how muscle cells interpret and respond to external mechanical loading (Sunadome et al., 2023; Mueller et al., 2021). Three principal modes of mechanical force are typically considered: tensile strain, which elongates cells and guides alignment (Chiu et al., 2023); shear stress, often induced by fluid flow, which influences membrane dynamics and metabolic activity (Hong et al., 2022); and compressive loading, a modality that has received less attention but shows considerable promise in musculoskeletal tissue modeling (Nordgaard et al., 2022).

Compression force exerts a unique and multifaceted influence on muscle tissue formation (Tao et al., 2023). Although skeletal muscle is not typically subjected to high compressive loads under normal physiological conditions, controlled compression in vitro simulates the mechanical resistance experienced during development, exercise, or load-bearing activity (Jorgenson et al., 2020). At the cellular level, cyclic compression promotes actomyosin contractility, reinforces cytoskeletal tension, and enhances cell–matrix interactions by activating integrin-mediated signaling pathways (Muntz et al., 2022). These effects are associated with the increased expression of myogenic regulatory factors, including myoblast determination protein (MyoD), myogenin (MyoG), and myosin heavy chain (MyHC), which are needed for muscle cell fusion and sarcomere organization (Luo et al., 2024). Furthermore, compressive forces modulate extracellular matrix (ECM) deposition and remodeling, thereby contributing to the mechanical robustness of engineered tissues (Mwase et al., 2022). Increased matrix density and collagen cross-linking under compressive conditions significantly improve tissue stiffness and resilience. These factors are necessary to mimic the texture and tensile properties of natural meat. Notably, compression exerts a significant influence on metabolic activity and nutrient transport, particularly within densely packed muscle constructs, where diffusion limitations may compromise cell viability (Johnson et al., 2023).

From a bioprocessing perspective, the incorporation of compressive stimulation into the fabrication of cultured meat is advantageous because it enables the transition from passive, time-consuming static cultures toward dynamic, feedback-driven systems that more closely resemble physiological muscle growth. This not only accelerates differentiation, thereby reducing the duration of cultivation, but also makes the process less reliant on expensive biochemicals sourced from animals or recombinant production supplies.

Despite the well-documented biological advantages of mechanical loading, an engineering gap persists in translating these techniques to industrial bioprocessing. Current dynamic bioreactors predominantly rely on complex, motor-driven electronic actuation systems (Mitra and Murthy, 2022; Ghanim et al., 2025; Dinara et al., 2025). When deployed within standard high-humidity (95% RH) cell culture incubators, these electronic components are highly susceptible to corrosion, short-circuiting, and failure. Furthermore, motor-driven systems generate undesirable localized heat, require sophisticated digital feedback loops, and incur high energy overheads. This reliance on energy-intensive, failure-prone equipment contradicts the fundamental sustainability and scalability objectives of cellular agriculture, presenting a major barrier to commercial viability.

To overcome these constraints, we previously designed and patented a customized pneumatic compression bioreactor to harness the biological benefits of mechanical loading without the drawbacks of electronic actuation (Vogel, 2021). This system utilizes compressed air regulated by a tunable magnetic circuit to deliver precise, physiologically relevant compressive stress. Given the robust and magnetic-valve composition of its components at the device level, we recently published a comprehensive review describing its potential for scalable cultured meat production (Ngwa et al., 2025). The review highlighted several distinct biomanufacturing advantages of the system, including its modular capacity for horizontal scale-out via parallelization, its ability to produce discrete, finished cuts of meat within individual vessels, and the elimination of fragile tissue transfer steps between distinct expansion and differentiation bioreactors, which mitigates cell damage and significantly reduces contamination risks.

Building upon that theoretical framework, here we describe the design architecture and operational principles of our pneumatic bioreactor. In addition, we evaluate its biological performance by applying a dynamic compression force onto murine myoblasts embedded in a plant-based scaffold, which we have already developed for cultured meat applications (Chiu et al., 2025). We hypothesize that this mechanically driven will effectively stimulate myogenesis, significantly reduce differentiation timeframes, and enhance the morphological integrity of the engineered tissue. Ultimately, these findings aim to position pneumatic mechanical stimulation as a scalable, and industrially relevant technology for next-generation cultured meat manufacturing.

2. Materials and methods

2.1. Bioreactor fabrication and system assembly

The compression bioreactor was custom-fabricated to provide a sterile, pneumatically driven mechanical stimulation environment (Fig. 1). The culture chamber was machined from food-grade 316L stainless steel to ensure corrosion resistance and autoclavability. The actuation system consists of a pneumatic piston driven by a compressed air supply, regulated by a tunable magnetic circuit that controls the pressurization frequency without the use of electronic controllers. Unlike motor-driven devices, the cyclic strain and operational frequency of our device are tuned via manual structural configurations. Compressive strain amplitude is governed by the physical positioning of the magnetic valve assembly, which sets the stroke limit and driving range of the pneumatic piston. Cyclic frequency is modulated by adjusting the tightness of the adjustment screw on the pressure escape/exhaust valve; tightening the screw restricts the rate of pressure exhaust, thereby lengthening each decompression phase and lowering the compression frequency. While the system does not support automated digital parameter adjustment, manual physical calibration enables the device to run with exceptional stroke-to-stroke precision and operational stability once the target thresholds are established. To maintain sterility and dampen mechanical impact, the culture chamber is sealed with a flexible, biocompatible silicone rubber membrane (thickness: 0.3 mm). The piston applies force externally to this membrane, transmitting compressive strain to the underlying scaffolds while maintaining a closed system. The compressing piston facilitates the delivery of nutrients by forcing waste out of the scaffold during compression and drawing in fresh medium during retraction.

Fig. 1.

Design and operational characteristics of the pneumatically driven compression bioreactor. (A) Schematic of the overall system architecture, highlighting the pneumatic actuation unit regulated by a tunable magnetic circuit. (B) Detailed view of the modular culture chamber.

2.2. Mechanical characterization via optical motion analysis

To validate the precision of the pneumatic actuation system without introducing invasive sensors into the sterile culture environment, we used a non-contact optical tracking method. A marker (black circular indicator, Ø = 1 mm) was affixed to the moving piston shaft to serve as a tracking point. To ensure measurement accuracy, a rigid calibration standard (precision rule) was positioned directly adjacent to the piston within the same focal plane to minimize parallax error. Video footage of the bioreactor during operation was acquired using a high-resolution CMOS camera (iPhone 16 Pro, Apple Inc., USA) mounted on a stabilized tripod. Recordings at 60 frames per second (fps) ensured sufficient temporal resolution to capture the peak displacement points of the compression cycle.

The acquired video data were analyzed using open-source physics modeling software (Tracker Video Analysis and Modeling Tool v6.1, Open-Source Physics). The video coordinate system was calibrated by defining the pixel-to-millimeter ratio based on the visible calibration standard. The vertical displacement of the fiducial marker was tracked over a continuous duration of 108 cycles using the software's auto-tracker algorithm. Mechanical stability was quantified by calculating the coefficient of variation (CV) of the peak amplitudes across replicate cycles as follows:

$$CV(\%)=\left(\sigma /\mu \right)\times 100\%$$

where σ represents the standard deviation of peak amplitudes, and μ represents the mean of peak amplitudes (Stibbs et al., 2024).

2.3. Scaffold preparation and cell cultivation

Plant-based composite scaffolds composed of alginate/guar gum/konjac glucomannan (AGK) were fabricated as previously described (cylinder scaffold with a diameter and height of 10 mm) (Chiu et al., 2025). The scaffolds were sterilized by autoclaving at 121 °C for 20 min. Murine C2C12 myoblasts were expanded in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS). For seeding, cells were harvested and resuspended at a density of 5 × 10⁷ cells/mL. Each scaffold was seeded with 100 μL of the cell suspension and allowed the cell attachment for 1 day before transfer to a new uncoated 24-well plate to remove the non-adhered cells. The same culture medium was used for entire experiment to minimize variation. The cells were then cultured in static conditions for 3 days adaptation before the dynamic compression experiments.

2.4. Dynamic compression culture

After 4 days overall adaptation, cell-laden scaffolds were divided into two experimental groups: a static control and a dynamic compression group. Both groups were maintained in identical culture medium within 24-well plates under standard incubator conditions (37 °C, 5% CO₂) to ensure identical baseline growth environments. During the mechanical stimulation over a period of three consecutive days (D4-D6), samples from both groups were placed in vessels which possesses a maximum capacity of 250 mL. The static control group samples were just kept in the vessel without compression to decouple the effects from potential handling and environmental confounding variables. The dynamic compression group's samples were placed on the scaffold stand in the bioreactor inner vessel and subjected to a cyclic compression regimen mechanically tuned to deliver a frequency of 0.27 Hz and a strain amplitude of ∼10%. This specific parameter combination was selected based on prior literature and preliminary testing, which demonstrated that higher frequencies and amplitudes not only exacerbate cell death but also compromise the structural integrity of the scaffold matrix (Yao and Mak, 2016). Following the daily 1 h compression sequence, both groups' samples were immediately transferred back to their original 24-well plates for a further 23 h of cultivation before the next daily compression cycle.

2.5. Cell viability analysis

Cell viability in the compression and static groups was compared using a Cell Viability Imaging kit (Roche Diagnostics, Germany; 06432379001). After stimulated with and without compression, scaffolds were stained for 1 h with 4 μM calcein-AM (labeling live cells) and 4 μM propidium iodide (PI) (BioLegend, USA) (labeling dead cells). The cells were then washed three times with PBS and stained with Hoechst 33342 for 30 min (labeling nuclei). Cells were visualized by fluorescence microscopy (excitation/emission 495/515 nm for calcein-AM, 540/615 nm for PI, and 361/497 nm for Hoechst 33342). Digital images were captured from three random positions in at least three individual samples. Images were analyzed using ImageJ. All images were converted to 8-bit grayscale, and the threshold was automatically adjusted to segment fluorescence signals. The live cell ratio was calculated using the formula:

$$LiveCellRatio=\left(LiveCells/\left(LiveCells+DeadCells\right)\right)\times 100\%$$

where “Live Cells” represents the number of spots detected in the green channel and “Dead Cells” represents the number of spots detected in the red channel. Data from individual samples were pooled to calculate mean values and standard deviations.

2.6. Analysis of cell differentiation

Cells embedded in the scaffold were fixed with 4% paraformaldehyde overnight at 4 °C for immunofluorescence staining. The samples were washed three times with PBS, permeabilized with 0.2% v/v Triton X-100 (Fluka, Germany) for 1 h, and non-specific binding sites were blocked by incubation in 2% w/v bovine serum albumin (BSA; Sigma-Aldrich), shaking gently for 2 h at room temperature (RT). MyoD was labeled overnight at 4 °C with a rabbit anti-mouse primary antibody (ABclonal, Germany; A0671) diluted 1:400 in 2% w/v BSA/PBS. After another three washes as above, the samples were incubated with a goat anti-rabbit secondary antibody labeled with Alexa Fluor 594 (Abcam, UK) diluted 1:1000, and Alexa Fluor 488 Phalloidin (Thermo Fisher Scientific) diluted 1:250, in 2% w/v BSA/PBS for 2 h at RT. After another three washes as above, nuclei were stained with DAPI (Carl Roth, Germany; 6335.1) diluted 1:3000 in PBS for 2 h at RT. After another three washes as above, the signal was visualized using a BZ-X800 fluorescence microscope (Keyence, Germany). To evaluate myogenic differentiation inside the 3D construct and prevent superficial surface-level bias, optical z-stack imaging was conducted to observe the cellular structures not only at the superficial surface but deep within the interior of the porous plant-based texturized carrier matrix. The automated ‘full-focus' function of the Keyence microscope software was subsequently employed to compress the compiled optical sections of the z-stack (∼800 μm) into a single, fully resolved, and in-focus 2D projection image. Differentiation was quantified by calculating the intensity of MyoD expression normalized to DAPI, and evaluating the co-localization area, analyzing at least three random fields per sample.

2.7. Statistical analysis

All experiments were conducted in triplicate, with at least three biological replicates (n ≥ 3) per independent condition. Data are expressed as means ± standard deviation (SD). The statistical significance of differences between the static control and dynamic compression groups was determined using a one-tailed, unpaired Student's t-test in GraphPad Prism v10 (GraphPad Software, USA). A p-value <0.05 was considered statistically significant. Otherwise, results were labeled “ns” (not significant). Significance levels were indicated as follows: p < 0.05 (∗), p < 0.01 (∗∗), p < 0.005 (∗∗∗), and p < 0.001 (∗∗∗∗).

3. Results and discussion

3.1. Design and operational principles of the compression bioreactor

The transition of cultured meat production from laboratory-scale tissue engineering to commercially viable biomanufacturing requires robust, scalable equipment. Conventional dynamic bioreactors rely extensively on motorized electronic actuation and digital feedback loops to deliver mechanical stimulation. However, continuous operation within standard mammalian cell incubators (typically at 37 °C and 95% RH) eventually leads to sensor corrosion, electronic short-circuiting, and high equipment failure rates (Li et al., 2024). To overcome these drawbacks, we engineered a pneumatic compression bioreactor lacking internal electronic components (Fig. 1A). The system architecture consists of three primary subsystems: a pneumatically driven actuation unit, a modular culture chamber, and a tunable magnetic circuit control mechanism. By regulating compressed air delivery using the tunable magnetic circuit, the system translates pneumatic pressure into controlled, mechanical piston strokes. This approach effectively isolates all electronic control units outside the humidified incubation environment, thereby avoiding the mechanical failure that can result from long-term operation.

The core of the bioreactor is the customized culture chamber (Fig. 1B), which is designed to ensure sterility, optimal mass transport, and effective force transmission. The chamber was fabricated from food-grade stainless steel, ensuring high durability and ease of sterilization, consistent with food industry standards. This apparatus secures the cell-laden scaffolds on a fixed base. To maintain a sterile environment while enabling dynamic loading, the chamber was sealed with a flexible, sterilized elastomeric membrane (silicone rubber diaphragm). The pneumatic piston therefore exerts its force on the external rubber interface rather than making direct contact with the culture medium. This configuration offers a significant biomechanical advantage, as demonstrated by the rubber membrane's function as a mechanical damper, allowing it to absorb the high peak-impact forces generated by the pneumatic piston. The design ensures that the compressive load transmitted to the underlying scaffolds is uniformly distributed, thus avoiding potential shear damage or abrupt mechanical shock while achieving effective stimulation of the cells. The internal scaffold stand was engineered to facilitate unrestricted nutrient diffusion and gas exchange during the compression phase, thereby preventing the onset of hypoxia in the continuous compressed state.

3.2. Magnetic control of distance and frequency

Passive pneumatic systems offer significant advantages in terms of sustainability but they must still achieve the precise mechanical fidelity required to control biological outcomes. Deviations in strain amplitude or loading frequency can severely compromise tissue development, and excessive compression forces can induce localized cell necrosis or damage the structural integrity of hydrogel-based scaffolds (Chiu et al., 2023; Roberts et al., 2023; Wu et al., 2016).

The compression frequency and displacement distance are critical operational parameters that control the pneumatic pressure compressor, and they are controlled via magnetic valve adjustments rather than software (Fig. 2). The displacement distance (and consequently the strain applied to the tissue) is determined by the vertical position of the magnetic circuit components relative to the piston stroke. Adjusting this position sets the physical limit of the driving distance. Concurrently, the compression frequency is modulated by adjusting the tightness of the circuit screw. This modifies the pneumatic resistance and the pressure threshold required to trigger the magnetic switch, thereby accelerating or decelerating the pressurization cycle.

Fig. 2.

Schematic diagram of the compression bioreactor control system. The illustration details the mechanically tuned actuation mechanism by the magnetic valve that operates without electronic feedback loops.

The mechanical microenvironment plays an essential role in directing the behavior and lineage progression of 3D cultured skeletal muscle cells. To systematically optimize the physical operating parameters of our pneumatic bioreactor, we investigated the structural and biological impacts of different cyclic compressive strain amplitudes. From previous study, we found that 10% compression strain might be the proper condition from literature review, while low strain magnitudes at or below 5% are generally insufficient to elicit a robust physiological response (Chiu et al., 2023). Conversely, raising the dynamic mechanical strain to 20% or higher compromises the structural integrity of the plant-based scaffold, inducing localized matrix tearing, cell-matrix detachment, and active cellular death (Gawlitta et al., 2007). The compressing frequency and duration are also a crucial factor that could damage the cells and scaffold with overloading. Mechanical force stimulation seems limited to the frequency around 0.1 Hz to 0.5 Hz with 1 h/day which was known for not injuring the cells. Since our bioreactor are adjusting the frequency based on the screwing valve of pressure releasing, we test different setting which within the range and not able to break the scaffold. After the setup, we calculate the cycle number per minute to get the compression frequency at 0.27 Hz.

To assess the kinematic precision of the mechanically tuned actuation, we applied non-contact optical motion tracking to monitor the piston stroke (Fig. 3). The resulting displacement–time profile demonstrated a highly uniform and stable cyclic loading pattern. The system accurately and consistently delivered the target frequency of 0.27 Hz, characterized by highly reproducible peak-to-peak intervals. This analysis was conducted in triplicate, ensuring the reliability of the findings. We observed a consistent displacement amplitude, with a negligible drift of less than 2% (∼1.77%), thereby substantiating the hypothesis that high precision can be achieved without sophisticated electronic controllers. Crucially, the vertical displacement amplitude, governed by the magnetic circuit's positioning, remained constant throughout the continuous recording period of more than 100 cycles, with no observable baseline drift or mechanical attenuation.

Fig. 3.

Kinematic validation and mechanical stability of the bioreactor. Optical motion tracking illustrates the displacement–time profile of the piston stroke during operation. The piston's drive path was tracked, indicating the horizontal and vertical displacement stability during repetitive cycles. The videos were taken from three independent testing and evaluate the stability.

3.3. Cell viability and morphology under compression

To ensure that the mechanically tuned compression did not induce cytotoxicity or physically damage the cultured tissue, we assessed post-stimulation cell viability with the procedures shown in Fig. 4A. Myoblasts embedded within AGK composite scaffolds were subjected to the optimized regimen of 0.27 Hz and 10% strain for 1 h/day over a 3-day dynamic phase. Live/dead fluorescence microscopy revealed a high density of living cells (calcein-AM positive, green) and a negligible number of dead cells (PI positive, red) across both the static control and the dynamically compressed scaffolds (Fig. 4B). Quantitative imaging confirmed that cell viability remained remarkably high, exceeding 85% in the compression group, with no statistically significant difference compared to the static controls (Fig. 4C).

Fig. 4.

Cell viability under mechanical compression. (A) Schematic timeline indicating the cultivation process. (B) Representative live/dead fluorescence microscopy of C2C12 myoblasts embedded within plant-based AGK composite scaffolds following 3 days of static control (above) or dynamic compression (below; 0.27 Hz, 10% strain, 1 h/day). Live cells are green (calcein-AM), dead cells are red (propidium iodide), and nuclei are blue (Hoechst). Scale bar: 200 μm. (C) Quantitative analysis of the live cell ratio. Data are means ± SD (n ≥ 3), and statistical significance are determined using a one-tailed, unpaired Student's t-test (ns: not statistically significant).

High cell viability during mechanical stimulation is necessary for bioprocessing. In mammalian cell cultures, the inappropriate application of macroscopic forces can induce apoptosis or mechanically rupture cell membranes, particularly in hydrated environments (Wu et al., 2016). Our results confirmed that the bioreactor's pneumatic actuation delivers uniform strain without lethal shear or compressive shock. The silicone membrane interface and compression piston head effectively dampened the impact forces and maintained continuous nutrient diffusion, thereby preventing hypoxia and mechanical necrosis. Furthermore, we also found that the plant-based AGK scaffold has sufficient mechanical resilience to endure repeated cyclic loading while protecting the embedded myoblasts. By establishing this biocompatible operational baseline, we have confirmed that the mechanical parameters are physiologically safe, ensuring that any subsequent cellular responses reflect targeted mechanotransduction rather than stress-induced pathological damage.

3.4. Myogenic differentiation and tissue organization

Having confirmed system biocompatibility, we next investigated the efficacy of the pneumatic compression in actively driving the functional maturation of myoblasts. The primary biological bottleneck in cultured meat production is the protracted timeline required for progenitor cells to commit to the myogenic lineage and organize into structured tissue. We therefore used immunofluorescence analysis to assess the expression of MyoD, a master transcriptional regulator of early myogenic commitment. Qualitative visual inspection revealed a striking discrepancy in tissue development between the control and compression groups (Fig. 5A). Whereas baseline levels of MyoD expression (red) were detected in the static control, the dynamically compressed tissue displayed a profound upregulation of this critical marker. The merged fluorescent images demonstrate strong co-localization of MyoD with the DAPI-stained nuclei (blue) in the dynamic group, confirming the active nuclear translocation required for MyoD to initiate the transcription of muscle-specific genes (Fig. 5B). Furthermore, cytoskeletal staining (phalloidin, green) indicated that the mechanically stimulated cells possessed a denser, more organized F-actin network compared to the relatively sparse distribution observed in static culture (Fig. 5A). Quantitative co-localization analysis corroborated positive myogenesis (Fig. 5B). We normalized marker expression against cell density by using the ratio of the MyoD-positive area to the DAPI area. We observed a mean 2.93-fold increase in the MyoD/DAPI area ratio of the dynamic compression group compared to the static control, revealing a statistically significant difference (p = 0.0071).

Fig. 5.

Mechanically promoted myogenic differentiation (A) Representative immunofluorescence images of C2C12 myoblasts cultivated in static versus dynamic compression conditions for 3 days. Samples were heavily stained for the early myogenic transcription factor MyoD (red), cytoskeletal F-actin (Phalloidin, green), and nuclei (DAPI, blue). (B) Quantitative co-localization analysis depicting the MyoD-positive area normalized to the DAPI area. Data are means ± SD (n ≥ 3), and statistical significance are determined using a one-tailed, unpaired Student's t-test (p < 0.01 (∗∗)). Scale bar: 200 μm.

These findings have profound industrial implications. The pronounced upregulation of MyoD within just 3 days of cyclic loading demonstrates the potent effect of mechanotransduction. In conventional static cultures, achieving comparable levels of myogenic commitment often requires extended incubation periods spanning several weeks, in conjunction with continuous replenishment of costly differentiation media (Martins et al., 2024). By directly translating physical forces into biochemical responses, the pneumatic system effectively shortens this developmental timeline. The 10% compressive strain we applied mimics the physiological loads experienced in native musculoskeletal environments, triggering cell-intrinsic mechanosensitive pathways that bypass the need for prolonged static incubation. For the cultured meat industry, our magnetic-valve controlled pneumatic stimulation method could advance the onset of myogenesis using purely physical forces. This mechanically driven acceleration may contribute to reduce overall batch production times and the industry's reliance on expensive, exogenously supplied biochemical growth factors, thus lowering the economic barriers to commercial-scale production.

Our findings present a compelling paradigm for mechanical tissue stimulation, but it is important to acknowledge certain limitations to guide future efforts to scale up the process. First, our study used the murine cell line C2C12 as a robust and standardized biological model to meticulously isolate and validate the physical and mechanical efficacy of the newly engineered bioreactor. In order to realize its full industrial potential, future research must translate these optimized compression parameters to agriculturally relevant cells (e.g. bovine, porcine, or avian myoblasts), which may exhibit distinct mechanosensitive thresholds. Second, although the significant upregulation of MyoD and enhanced F-actin networking confirm early-stage myogenic commitment at the cellular level, subsequent studies must evaluate the macroscopic rheological properties of the fully matured constructs. Rigorous texture analysis and mechanical shear testing of the final tissues is needed to confirm that this mechanically promoted myogenesis accurately replicates the complex texture, bite and mouthfeel of conventional meat. Third, to address scalability barriers, we provide a new strategy to scale out the bioreactor instead of scale-up designs which frequently suffer from nutrient diffusion gradients, unstable fluid shear, and non-uniform pneumatic force fields (Ngwa et al., 2025). Parallelized vessel with highly controlled individual chambers ensures uniform mechanical stimulation and identical microenvironmental conditions across all constructs, thereby enabling better quality control over individual batches and mitigating the risk of total batch failure. To facilitate routine operation and practicalize this scale-out technology, ongoing development must also address the current operational and physical challenges of our bioreactor prototype, such as the labor-intensive sterilization protocols required for its reusable components, handling difficulties associated with heavy glass and steel structural elements, and the risk of pneumatic pressure leakage at connection joints during high-frequency cycles. To overcome these bottlenecks, future research must focus on optimizing vessel design and utilizing alternative fabrication materials to lower material costs and improve operational ease. Finally, while the current pneumatic setup establishes a reliable system for operating within an incubation environment, we can further optimize energy efficiency and sustainability by integrating alternative driving units. By retaining the core magnetic-valve logic but transitioning to systems such as closed-loop hydrostatic actuation or an external electromechanical drive, the bioreactor can achieve more sustainable and cost-efficient bioprocessing.

4. Conclusion

In this study, we developed and validated a novel magnetic-valve controlled pneumatic compression bioreactor, addressing the major bioprocessing bottlenecks of energy consumption, equipment failure and protracted production timelines in cultured meat manufacturing. The device was engineered to function without motorized electronic components when exposed to high humidity, a feat accomplished by using a mechanically tuned magnetic-pneumatic actuation system. This design ensured that operation would be highly scalable while maintaining exceptional kinematic precision. It reliably delivered a cyclic strain without reliance on continuous digital feedback loops. A rigorous biological assessment was conducted to evaluate the biocompatibility and efficacy of the passive mechanical stimulation framework under investigation. Our results confirmed the framework is both robust and effective. Murine myoblasts cultivated on plant-based AGK scaffolds maintained high viability under dynamic compression, demonstrating the efficacy of the system's pneumatic dampening in preventing cytotoxic shear and mechanical necrosis. The mechanotransducive cues applied in this study resulted in a significant 2.93-fold upregulation of the early myogenic marker MyoD within just 3 days of cyclic loading.

In terms of future applications, our findings demonstrate that precise physical forces can actively promote tissue organization and may also support muscle differentiation, thus circumventing the traditional reliance on prolonged static culture and expensive, exogenous growth factors. We used a robust murine model as a biological proof of concept, but future research must focus on translating these mechanical conditioning parameters to agriculturally relevant bovine, porcine or avian cell lines and expanding the reactor chamber volume. This bioprocessing approach could also be further optimized by using closed-loop hydrostatic actuation or external electromechanical drive to reduce the energy consumption from compressed air for pneumatic pressure. This offers a promising outlook for the sustainable, energy-efficient, and economically viable upscaling of structured cultured meat analogs.

Declaration of competing interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: Stefan Schillberg reports financial support was provided by European Union. Simon Vogel, Stefan Schillberg has patent #WO 2021/148663 licensed to Fraunhofer-Gesellschaft. If there are other authors, they declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Handling Editor: Professor Alejandro G.Marangoni