SLAS Technology

Three-dimensional (3D) printing is increasingly used in cell biology to prototype custom cell culture devices, microscopy chambers, and other experimental hardware, but practical guidance on sterilization and material compatibility remains limited. Here, we develop and validate a low-temperature paraformaldehyde vapor sterilization method for fused filament fabrication (FFF) components that avoids heat-induced deformation. Using bacterial challenge assays with Escherichia coli and Geobacillus stearothermophilus, we confirm effective sterilization of 3D printed components. We also assess the effects of common filaments and adhesives on HL-60 human cell growth to identify materials suitable for cell culture workflows. Most untreated plastics were well tolerated over 48 hours, whereas some formaldehyde-treated materials required post-treatment with ammonia vapor to restore compatibility. Together, these results provide a practical framework for sterilizing and deploying 3D-printed materials in cell culture and biological research. Multidisciplinary Abstract3D printers allow laboratories to quickly make custom experimental equipment, but many printed materials cannot be sterilized using standard methods. We developed a low-temperature sterilization procedure using paraformaldehyde vapor and tested whether common printing materials and adhesives are compatible with cultured human cells. We identified several plastics, silicone-based adhesives and acrylic tapes that can be safely used after sterilization, while others require additional ammonia treatment. These results provide guidance for researchers using 3D printing to build custom laboratory and cell culture devices. Methods SummaryA low-temperature paraformaldehyde vapor sterilization method for 3D-printed FFF components was developed and validated using bacterial challenge assays. Compatibility of common printing filaments, adhesives, and tapes with cell culture workflows was assessed by exposing HL-60 cells to untreated, sterilized, and ammonia- neutralized materials and quantifying cell growth by flow cytometry.

Three-dimensional (3D) cell culture models, particularly multicellular tumor spheroids, have become essential tools for studying cancer biology, drug screening, and preclinical testing due to their ability to mimic physiological tumor microenvironments. However, traditional invasion assays, such as Boyden-chamber- or Transwell-based systems, often suffer from variability introduced by spheroid handling and transfer, compromising data reproducibility. Here, we present a novel, 3D-printed migration and invasion platform --the MQm-sert-- designed to standardize and streamline spheroid-based invasion assays while maintaining spheroid integrity. Fabricated via fused filament fabrication using biobased polylactic acid, the MQm-sert integrates a hanging-drop spheroid culture system (MQm-sert) with a membrane-based invasion chamber (M-sert), eliminating the need for disruptive spheroid transfer steps. Using synthetic tumor environment mimics (STEMs) composed of breast cancer cells (MDA-MB-231 and MCF7), mesenchymal stromal cells (MSCs), and human pulmonary microvascular endothelial cells (HPMECs), we quantified invasion dynamics and cellular interactions. This innovation significantly reduces experimental variability, as demonstrated by lower variance in invasive cell mass dimensions and cell density compared to conventional workflows. Beyond biological insights, the platform aligns with sustainability goals by leveraging cost-effective, open-source 3D printing, reducing reliance on commercial labware, and addressing key challenges in 3D cell culture standardization.

Organ-on-chip (OoC) systems are increasingly used to generate human-relevant evidence in drug discovery and preclinical development, yet interpretation can be limited by the interaction of biological variability, device-to-device variability, and technical assay noise. Here, we describe the UstarFlowAI barrier-confidence framework, an in silico assay-readiness approach that quantifies whether a predefined permeability-related effect is resolvable under specified uncertainty. The proposed context of use is decision support for laminar, low-Reynolds-number airway epithelial monolayer barrier OoC assays in which device fabrication, pump performance, flow, wall shear stress, and biological fluctuation may affect permeability-related readouts. Monte Carlo simulations propagate defined manufacturing, biological, systemic, and well-level uncertainty and estimate the minimum detectable shift (MDS) under a prespecified decision rule. In the present proof-of-concept design, the reference workflow was parameterized at 9.4% technical coefficient of variation (CV), whereas the standardized workflow was parameterized at 5.0% CV, corresponding to an approximately 46.8% reduction in the modeled technical-noise component. At n = 48 replicate wells per group and a nominal total assay-noise operating point of 17% CV, a predefined 10% permeability shift exceeded the upper MDS uncertainty bound for the standardized workflow in both best- and worst-case scenarios, whereas it overlapped the 95% MDS interval for the reference workflow and therefore did not meet the prespecified PASS criterion. These findings demonstrate the behavior of the computational framework under the stated assumptions; they do not constitute experimental validation, regulatory qualification, or evidence that a 10% shift is universally meaningful across barrier OoC systems. We therefore propose a prospective wet-lab validation strategy in which model predictions are tested against airway barrier measurements generated under controlled sources of technical and biological variability. The framework is intended to complement, rather than replace, the underlying biological model by making assay uncertainty and decision thresholds explicit and auditable.

Point-of-care (POC) molecular detection has enabled rapid and accessible testing at or near the site of need, facilitating timely decision-making in clinical, field, and industrial settings. Despite these advancements, molecular detection platforms remain limited in their ability to prepare DNA from complex tissue matrices, restricting their applicability to a narrow range of sample types. To address sample preparation inefficiencies, focused ultrasound extraction (FUSE) has been developed for the rapid release of DNA from robust tissue matrices. FUSE disintegrates target tissue and releases DNA by delivering high pressure ultrasound pulses that generate a cavitation bubble cloud. In prior work, FUSE was performed using large, laboratory-based equipment. Here, a miniaturized FUSE device was designed and tested to enable POC DNA sample preparation. Design constraints were defined based on prior FUSE experiments, and based on these metrics, a 750 kHz cylindrical transducer was fabricated. The pressure output was measured, and high-speed optical imaging demonstrated that the transducer generated sustained cavitation within the defined focal region. The feasibility of DNA extraction from piscine and timber tissues was evaluated, and results show that the device can release DNA with quality suitable for PCR-based detection from both tissue types. These results demonstrate the potential of this device to rapidly extract DNA from complex samples and expand the applicability of molecular detection platforms.

At-home sampling can address obstacles to participation in transcriptomics research. We previously developed the homeRNA and homeRNAmax platforms; these custom-designed tubes containing RNA stabilizer interface with a blood tube compatible with upper-arm blood collection devices, allowing for blood collection and RNA stabilization to occur from a participant's home. In this work we introduce homeRNAdrop, an insert for a commercially available blood tube (BD Microtainer) that divides the tube into two compartments: the top compartment holds 200 L blood and the bottom compartment holds ~590 L liquid stabilizer (RNAlater). By combining blood collection and RNA stabilization into the same tube, homeRNAdrop streamlines sample preparation for participants and allows for more compact sample storage for the lab. In a remote pilot study, n=25 participants from across the United States tested the homeRNAdrop kit. The n=23 samples successfully received by the lab show that samples collected using homeRNAdrop are of sufficient RNA quality (all RINs > 6, a common cutoff for analysis) and yield (mean yield=1.44 g) for downstream gene expression analysis. Almost all participants who successfully collected blood using the Tasso Mini device found the homeRNAdrop device easy to use (n=23/24). Overall, this work adds a simplified, user-friendly platform to the growing suite of tools for remote blood RNA stabilization.

Background: Point-of-care manufacturing of CAR-T cells could reduce cost and improve patient access, but few closed, automated systems support both flexible process development and GMP production. Methods and results: We describe LimCORE, a single-use consumable for cell incubation and processing, implemented in LimGROW (manual system) and LimONE (closed and automated system). In LimGROW, T cells expanded 246 +/-18 fold over 10 days at optimized seeding density, versus 89 +/-9 fold in a G-Rex control, with similar viability (>90%) and phenotype. Process parameters directly transferred to LimONE, where automated, closed buoyancy-based CD3+ selection achieved 90.3 +/-3.5% purity and 59.2 +/-17.0% recovery in under two hours. Using this workflow, we manufactured CD19 CAR-T cells in a fully closed, automated 7-day LimONE process requiring 155 minutes of operator intervention. LimONE manufactured CAR-T cells reached 42.6 +/-2.4 fold expansion (532 +/-29x10e6; cells at harvest), with 43.6 +/-6.0% transduction efficiency and viability similar to G-Rex controls. CD4+/CD8+ ratios, CD8+ differentiation subsets, and cytotoxicity were comparable between platforms, while LimONE-manufactured cells showed increased spare respiratory capacity and oxygen consumption. Across four manufacturing runs on the LimONE, no hardware or software failures occurred, and automated liquid transfers and volume concentration stayed within +/-5% accuracy. Conclusions: These data show that LimCORE supports T cell expansion, selection, and complete CD19 CAR-T manufacturing with performance similar to existing platforms while reducing operator time, across R&D and GMP scale closed-system formats.

Recombinase polymerase amplification (RPA) enables rapid nucleic acid testing in low-resource environments, but poorly characterized byproducts can compromise assay specificity and cause false-positive results. Here, we amplified the thirteen original CODIS core loci and Amelogenin to characterize recurrent RPA artefacts and establish conditions that reduce their formation. First, RPA products were analyzed for two reference samples by Oxford Nanopore Technologies sequencing. This revealed two distinct classes of multimeric products: primer multimers and amplicon multimers, consisting of repeated primer or amplicon sequences, respectively. Individual artefacts contained up to 281 primer copies or 22 amplicon copies, demonstrating the extensive range of these products. Next, we performed an optimization study to evaluate the effects of reaction temperature and reagent concentrations at two representative loci, D3S1358 and D5S818. Among the conditions tested, temperature had the most pronounced effect. Reducing the temperature from 42{degrees}C to 34{degrees}C increased the relative target amplicon fraction from 15% to 83% for D3S1358 and from 84% to 98% for D5S818, while maintaining or increasing absolute target concentration. Lower primer concentrations and higher T4 UvsX concentrations also reduced multimer formation, although lower primer concentrations reduced target yield and caused allelic dropout. Finally, amplification at 34{degrees}C was evaluated across all fourteen loci by sequencing. Relative to 42{degrees}C, the target read fraction increased by more than 5 percentage points for 7/14 loci in one reference sample and 9/14 loci in the other, with the largest improvements at multimer-prone loci. These findings identify multimers as an important class of RPA artefacts and establish reaction temperature and T4 UvsX concentration as promising conditions to improve RPA specificity.

Fermentation is a process through which microorganisms are used to transform basic molecules into high value substances in various applications such as biopharma, food production, and biofuel generation. In such processes, monitoring the health of microbial cells is essential, as it is a key factor in the final product yield. Traditional methods of assessing the health of fermenter populations have involved manually extracting samples from the fermenter for benchtop testing, which lacked the accuracy and efficacy of real-time continuous measurements. This has opened the door for inline sensors to be integrated into fermenters for continuous measurement; including microfluidic based Electrochemical Impedance Spectroscopy (EIS). EIS utilizes a range of frequencies to gain insight into the characteristics of microbial populations, such as cell size and membrane capacitance. EIS cytometry, in particular, has emerged as a relevant tool, as it enables single-cell level detection in real-time. However, current EIS cytometry devices cannot be deployed directly within fermenters due to the high cell density conditions. This has limited EIS analysis in fermenters to bulk measurements averaged across a population. Here, we present a microfluidic chip that uses passive cell-flux-partitioning geometry to direct cells from high-density culture through a single-cell-constrained flow path containing integrated impedance-sensing electrodes. The proposed chip combines geometric channel design and embedded electrodes to enable continuous impedance measurements from cell-containing flow while operating at input cell densities on the scale of 10^9 CFU/mL. This work demonstrates a first step toward integrating single-cell-constrained microfluidic handling with impedance sensing for inline monitoring in industrial biomanufacturing.

Suspension culture is the gold standard for ex vivo hematopoietic stem cell (HSC) expansion; however, high media requirement results in expensive cell therapy products. Hydrogel-based HSC culture represents a more realistic in vitro representation of human in vivo HSC microenvironments and may support a more efficient use of media during ex vivo HSC expansion. However, media conditions used for hydrogel-supported cell culture often simply mimic that which are currently used in suspension culture approaches, potentially resulting in an oversupply of cell growth proteins. In this work, we present a microfluidic culture system capable of supporting hydrogel-based cell culture and generating spatial distributions of independent growth factor combinations. Using this device, we image, in real-time, the behaviour of human umbilical cord blood-derived CD34+ cells, within fibrin-based hydrogels, in response to spatial gradients of the human recombinant growth factors stem cell factor, thrombopoietin, angiopoietin-2 and insulin-like growth factor-II. Using live microscopy and image-based single-cell segmentation and quantification, local changes in cell density over time in relation to spatial concentration of factors were identified. This novel microfluidic device has the potential to screen combinations of growth factors required for hydrogel-supported CD34+ cell expansion and inform culture media formulations of future hydrogel-based cell manufacturing processes.

Direct experimental analysis of the mammalian oviduct is constrained by limited tissue access and the short lifespan of ex vivo preparations. Extracellular matrix-embedded three-dimensional epithelial organoids provide longer-term in vitro models. However, their inward-facing apical surface and the absence of supporting stromal cells limit physiological studies of the oviduct, including ciliary activity and maternal-embryonic interactions. Here, we provide a step-wise protocol detailing the generation of mouse and human oviductal assembloids in which epithelial cells form an outward-facing (apical-out) layer around a stromal core. Epithelial and stromal cells from adult mouse oviducts or human Fallopian tubes are isolated, expanded separately, and subsequently aggregated in a rotational culture system. The protocol also outlines morphological and immunostaining criteria for confirming cellular organization, whole-mount detection of external cilia, measurement of ciliary beat frequency, and co-culture of mouse assembloids with preimplantation embryos. Mouse and human assembloids retained epithelial and stromal identity and displayed cilia at the accessible outer surface. In a proof-of-concept experiment, embryos co-cultured with the assembloids developed to blastocysts at a rate similar to that of in vivo-derived blastocysts. This reductionist system provides a straightforward and tractable model to investigate oviduct physiology and embryo-maternal communication while allowing direct manipulation and observation of the epithelial interface. Graphical Abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=148 SRC="FIGDIR/small/743297v1_ufig1.gif" ALT="Figure 1"> View larger version (50K): org.highwire.dtl.DTLVardef@1917a7borg.highwire.dtl.DTLVardef@41d7org.highwire.dtl.DTLVardef@e2bf98org.highwire.dtl.DTLVardef@90c9f3_HPS_FORMAT_FIGEXP M_FIG C_FIG SummaryThe protocol for generating mouse and human oviductal assembloids by combining epithelial and stromal cells for studying oviductal function in an in vitro setting.

Organoids-on-chip combine the 3D complex microenvironment and cellular composition of organoids with microfluidic flow, increasing nutrient-waste exchange and mimicking the contributions of in vivo interstitial and vascular flow. However, the widespread adoption of organoid-on-chip platforms is limited by the lack of incubator-friendly flow control systems. Existing approaches often rely on commercially available syringe or peristaltic pumps, but these are bulky, lack scalability, and present a significant barrier for clinical translation. To circumvent these issues, we present the Compact Active Perfusion Standalone Organoid-on-Chip (CAPS-OC) platform, a fully integrated and battery-powered microfluidic system capable of culturing organoids in active media flow. To achieve this, we introduce a novel low-power mechanism of pressure pulse generation using an off-the-shelf compact rotary actuator (CRA), and package it into a compact electromechanical assembly. This assembly provides timed pneumatic inputs to achieve programmable control of membrane-based peristaltic pumps, with [~]100 {micro}L/hr dynamic range achieved on a custom microfluidic organoid chip. We biologically validated this system by culturing pancreatic cancer organoids derived from a KrasLSL-G12D/+; Trp53LSL-R172H/WT; Pdx1-Cre (KPC) genetically engineered mouse model. We found that our chip enhances proliferation and helps sustain a population of larger (>150 {micro}m) organoids compared to standard dome based static culture. Additionally, through immunostaining, we observe that KPC organoids cultured in the chip show more aggressive PDAC phenotype with reduced nuclear expression of GATA6, whereas organoids in static culture retain less aggressive classical- like subtype. Finally, testing of a RAS inhibitor drug, daraxonrasib, on the KPC organoids on chip showed size-based sensitivity elucidating the impact of active perfusion on the drug diffusion kinetics. Altogether, we establish CAPS-OC as a valuable tool for the bioengineering community and for clinically translating organoid model systems.

Electrical stimulation (ES) and transepithelial/transendothelial electrical resistance (TEER) measurements are essential techniques in cell biology and tissue engineering, yet commercial devices for these applications cost between USD 2,500-9,000 and typically offer only one functionality. We present LATEER (Low-cost Arduino-based TEER and Electrical stimulation device), an open-source hardware platform that combines both ES and TEER measurement capabilities at a total cost below USD 100. The device features four independent channels, configurable pulsatile signals (amplitude up to 8.2 V, frequency 0.1-500 Hz, pulse width [&ge;]0.1 ms), and a resistance measurement range of 300 {Omega} to 1 M{Omega}, with <5% error for R {gtrsim} 4.7 k{Omega}. LATEER uses commercially available graphite pencil leads as electrodes ([~]USD 2 vs. USD 350 for commercial Ag/AgCl electrodes), which demonstrated excellent biocompatibility in cell culture. The system includes 3D-printed electrode holders compatible with standard 12-well and 24-well plates, allowing microscope visualization without electrode removal, and a Python-based graphical user interface for parameter configuration and real-time data acquisition. Because the electrodes remain fixed in the plate lid and only a single cable enters the incubator, both stimulation and resistance measurement can run continuously under standard culture conditions (37 {degrees}C, 5% CO2) without removing the plate or repositioning the electrodes, avoiding the temperature excursions and placement variability inherent to manual chopstick measurements. Validation with human pluripotent stem cell-derived cardiomyocytes demonstrated reliable frequency capture (electrical pacing) of the contracting monolayer, with a capture threshold between 250 and 400 mV/mm and controlled pacing across the 0.5-5 Hz range. TEER functionality was verified with mesenchymal stem cells, where the device resolved cell-density-dependent differences in electrical resistance in real time. All design files, firmware, and software are freely available under the CERN-OHL-S v2 license, enabling replication and customization by research laboratories worldwide. HighlightsO_LIAn open-source device combines electrical stimulation and TEER measurement under $100 C_LIO_LIGraphite electrodes offer biocompatibility at 0.6% cost of commercial alternatives C_LIO_LIFour independent channels with configurable parameters and real-time data logging. C_LIO_LIContinuous run setup in-incubator; no electrode repositioning needed C_LIO_LIValidated with stem cell-derived cardiomyocytes, achieving frequency capture (threshold 250-400 mV/mm) C_LIO_LI3D-printed holders enable microscope visualization without electrode removal C_LI Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=78 SRC="FIGDIR/small/743263v1_ufig1.gif" ALT="Figure 1"> View larger version (28K): org.highwire.dtl.DTLVardef@98f9deorg.highwire.dtl.DTLVardef@13c73aborg.highwire.dtl.DTLVardef@1cdf099org.highwire.dtl.DTLVardef@16ed0cf_HPS_FORMAT_FIGEXP M_FIG C_FIG Specifications Table O_TBL View this table: org.highwire.dtl.DTLVardef@4ef802org.highwire.dtl.DTLVardef@7c651borg.highwire.dtl.DTLVardef@d20013org.highwire.dtl.DTLVardef@102fc89org.highwire.dtl.DTLVardef@111b927_HPS_FORMAT_FIGEXP M_TBL C_TBL

Three-dimensional (3D) spheroid models have become essential in cancer biology, drug screening, and tissue engineering. However, their small size, fragile structure, and tendency to disintegrate during routine histoprocessing present persistent technical challenges. Conventional paraffin embedding often results in tissue fragmentation, loss of spatial orientation, and poor section quality, whereas cryosectioning often compromises cellular morphology. Here, we present a robust, cost-effective protocol for preserving and sectioning fragile 3D spheroids, resulting in high-quality histological sections with intact architecture and excellent cellular detail. The method involves optimized handling and embedding procedures that stabilize spheroids during standard formalin fixation, paraffin infiltration, and microtomy, eliminating mechanical distortion and preserving spherical integrity for consistent sectioning. We demonstrate successful application across different cell line spheroids, with subsequent compatibility with hematoxylin and eosin (H&E) staining protocols. Compared to conventional methods, our approach significantly reduces sample loss, improves inter-section reproducibility, and preserves fine structural features such as necrotic cores, proliferative zones, and extracellular matrix components. This protocol provides a reliable, accessible solution for routine histological analysis of fragile 3D spheroids, facilitating more accurate morphological and molecular assessment in translational research settings. Key featuresO_LIMaintains spheroid integrity: Prevents mechanical distortion, fragmentation, and loss of spatial orientation during processing. C_LIO_LISignificantly reduces sample loss: Decreases failure rate compared to traditional methods, conserving valuable samples. C_LIO_LIBroad spheroid compatibility: Works effectively with primary tumor-derived, stem cell-derived, and co-culture spheroid models. C_LIO_LIEnables high-quality sectioning and staining: Delivers consistent, reproducible sections that are fully compatible with H&E, IHC, and IF. C_LI Graphical overview O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=140 SRC="FIGDIR/small/743094v1_ufig1.gif" ALT="Figure 1"> View larger version (44K): org.highwire.dtl.DTLVardef@1670c4org.highwire.dtl.DTLVardef@145810aorg.highwire.dtl.DTLVardef@1accb1org.highwire.dtl.DTLVardef@17481c0_HPS_FORMAT_FIGEXP M_FIG C_FIG

Functional protein micro/nanofibers integrate high specific surface areas with bioactive architectures but remain hampered low production throughput and severe processing instability. Focused rotary jet spinning (FRJS) shows promising to break these throughput constraints while enabling direct, conformal deposition onto complex, irregular substrates. However, navigating FRJS's narrow processing windows in proteins remains failure-prone without closed-loop experimental guidance. Here, we report FiberPro 1.0, a large language model driven multi agent framework that unites high-throughput spinning with real-time experimental feedback for autonomous design, execution, and optimization. Across three protein systems, FiberPro 1.0 converged on spinnable formulations within an average of two iterations. To demonstrate high-throughput conformal deposition in a translational setting, FiberPro designed a zein-based active packaging system applied directly onto diverse food matrices. The resulting conformal coating combined potent antibacterial activity with real-time freshness monitoring, markedly suppressing Escherichia coli and extending shelf life. This work connects autonomous AI reasoning with high-throughput processing, establishing a verifiable route for scalable manufacturing and conformal coating of functional protein micro/nanofibers.

The modeling of aroma release in food science is a complex, multidisciplinary field integrating mass transfer theory and principles of physical chemistry. While mechanistic models have long been used to understand volatile release, their practical adoption has been hindered because existing implementations are often private, "in-house" tools that lack reproducibility and extensibility. This paper introduces digimouth, an open-source Python framework designed to provide a unified, modular, and reproducible environment for simulating and fitting aroma release kinetics. A key innovation of digimouth is its handling of events and stages. Events represent discrete, instantaneous changes in the system (such as opening a bottle or swallowing), while stages allow for multi-phase simulations where experimental conditions or physical dynamics change over time. To demonstrate the tool's utility, the authors implemented established in vitro convection models and performed sensitivity analyses on parameters such as air-water partition coefficients and mass transfer coefficients. The package also includes an optimization module based on the least-squares method to fit parameters to experimental data. Validation against real-time data from Proton Transfer Reaction-Time of Flight- Mass Spectrometry (PTR-ToF-MS) shows a satisfactory fit, with an average error of approximately 2%. Ultimately, digimouth facilitates robust model development and hypothesis testing, offering the food science community a collaborative tool to better understand the mechanisms of flavor release.

High-throughput enzymology enables quantitative characterization of enzyme function across hundreds to thousands of sequence variants and experimental conditions. Nevertheless, the scale and complexity of these datasets create substantial challenges for analysis and quality control. High-Throughput Microfluidic Enzyme Kinetics (HT-MEK), for example, generates large microscopy datasets that must pass through multiple analytical stages, including image processing, initial-rate fitting, and kinetic modeling. Choices or errors made at any stage can propagate into the final kinetic parameters without being evident from fit statistics alone. Here, we provide a broadly applicable guide for analyzing high-throughput enzymology data from the HT-MEK platform, using Michaelis-Menten kinetics as a representative example. We first outline the conceptual workflow from raw fluorescence measurements to estimates of kcat, KM, and kcat/KM. We then discuss common experimental and analytical failure modes and present a framework for deciding when data should be refitted, filtered, qualified, or repeated. Finally, we provide a step-by-step workflow using Mercury, an open-source Python framework that integrates scalable HT-MEK data processing with traceable quality control and diagnostic visualization. This workflow preserves the connection between reported parameters and their underlying measurements and can be adapted to other kinetic models and biochemical assays.

Primary fetal human epithelial organoids are canonically derived from tissue samples obtained after termination of pregnancy. Recently, we have demonstrated that these organoids can be consistently generated from amniotic fluid cells isolated prenatally during diagnostic and interventional procedures. Amniotic Fluid-derived Organoids (AFOs) are promising fetal epithelial lung, kidney and small intestine tissue models that bypass some of the ethical and legal constraints associated with obtaining primary fetal tissue. Despite this, the widespread adoption of this technology is limited by the lack of standardised and accessible pipelines for the biobanking, culturing and distribution of organoid-forming amniotic fluid cells. To enable broader AFO use for research and clinical purposes, this work investigates two cryopreservation strategies to optimise AF cell recovery for organoid derivation. In Strategy 1, viable AF cells were sorted before freezing, while in Strategy 2 unsorted AF was frozen, and viable AF cells were sorted for viability after thawing. We present here a comprehensive evaluation of 5 commercially available GMP-compliant freezing media (FM1-5) alongside a standard lab-grade control (FM CT). AFOs were assessed for formation efficiency, morphological characteristics, proliferation capacity, epithelial identity, and tissue type. Our results demonstrate that organoid-forming AF cells can be successfully cryopreserved both pre- and post-sorting. Strategy 1 yielded higher organoid formation efficiency, with AFOs derived from cryopreserved and fresh cells exhibiting comparable expansion potential. Finally, FM4 provided minimal to no decline in survival rate, making it the most effective GMP-grade freezing medium tested. In conclusion, we present two viable cryopreservation strategies adaptable to different laboratory settings and identify optimal GMP-compliant freezing media, to support safe distribution and centralised processing, thereby facilitating scalability and collaborative work on the AFO technology.

We present an active mixing toolhead for extrusion bioprinting. The tool enables the programmable fabrication of tumor microenvironment gradient arrays, through controlled deposition of mixed hydrogel precursor formulations into 384-well plates, pre-seeded with tumor cells. It operates on an open-source bioprinter and can actively mix arbitrary ratios of two hydrogel precursors prior to extrusion. These concentration gradient arrays are compatible with quantitative image analysis of cell viability and morphological responses to hydrogels conditioned with drug or extracellular matrix (ECM) proteins. The tools capacity to mix and print hydrogel precursor gradients was demonstrated using alginate and highly concentrated mCherry-conjugated mini-spidroin solutions. Hydrogel precursor stocks contained fluorescent reporters to facilitate quantifications of mixing efficiency, and as proxies for drug and ECM protein concentrations. The tool was applied to generate hydrogel-based gradients of the apoptosis-inducer staurosporine, from which concentration-dependent MDA-MB-231 breast cancer cell death responses were quantified. Gradient arrays of the ECM protein laminin-511, implicated in breast cancer tumorigenesis, were generated and revealed that increasing laminin-511 concentrations potentiated staurosporine-induced cell death. The study demonstrates the utility of this active mixing toolhead for producing hydrogel gradient arrays, and demonstrates the relevance of studying drug-responses in tumor microenvironment models that account for disease-specific ECM components.

mRNA therapeutics have demonstrated transformative potential in infectious disease, oncology and genetic disorders. Parallel to the rapid clinical advancement of mRNA therapies, research into in vitro transcription (IVT), the pivotal process of mRNA manufacturing, has intensified significantly. However, the majority of these studies omitted cap analog, an essential substrate for co-transcriptional capped IVT (co-IVT), likely to simplify experimental systems. Nevertheless, co-IVT remains the dominant industrial standard for the production of mRNA therapeutics. Here, we report an optimized co-IVT process that achieves a yield of more than 26 g/L while maintaining capping efficiency of over 99.5%, and achieving a 30-fold reduction in double-stranded RNA (dsRNA), the most critical process-related impurity. These performance gains substantially improve mRNA production efficiency and enhance mRNA quality, thereby accelerating the transition to commercial-scale manufacturing of mRNA therapeutics. Moreover, the performance was attained using either of two mechanistically distinct engineered T7 RNA polymerase (RNAP) mutants. Both mutants enabled robust mRNA synthesis under process conditions that have not been previously reported. These observations provide mechanistic insights into IVT and advance our fundamental understanding of this critical bio-manufacturing step.

Influenza A virus (IAV) defective interfering particles (DIPs) are regarded as promising broad-spectrum antivirals due to their high efficacy demonstrated in animal trials. Previously, an OP7 chimera DIP was generated that showed markedly enhanced antiviral efficacy compared to conventional IAV DIPs. In addition, a high-yield cell culture-based production process for OP7 chimera DIPs was established in perfusion mode. However, due to the technical complexity of this technology, we aimed to combine the advantages of perfusion in terms of yield with that of a simple and less complex batch process. In the present work, we established a fed-batch production process for OP7 chimera DIPs that allows a more straightforward implementation into existing production facilities. Using a feeding regime that was able to overcome the cell density effect, we achieved productive virus infection in stirred tank reactors until a cell density of 10E+6 cells/mL. Furthermore, we obtained a high virus titer of 3.8 log10(HAU/100 uL), corresponding to a 3.4-fold increase compared to a reference batch production. Moreover, the produced material demonstrated a significantly increased interfering bioactivity shown by an in vitro potency assay. Interestingly, this efficacy was similarly high as compared to OP7 chimera DIP material previously produced in perfusion mode, despite twofold lower virus titers. Finally, we demonstrate robustness of OP7 chimera DIP seed virus trains, which is significant step towards later pharmaceutical production. Further, corresponding GMP MDCK-PB2(sus) cell and OP7 chimera virus banks are now available for later biomanufacturing further use. In conclusion, the established fed-batch process offers a valid n additional option for pharmaceutical grade OP7 chimera DIP GMP manufacturing production under GMP conditions required for subsequent clinical testing and for later large-scale manufacturing.