SLAS Technology

The development of microphysiological cell culture models (MPMs) that align with the throughput demands of drug and chemical testing are needed to help reduce animal testing, aide in the discovery of new drugs, and identify harmful chemical exposures. To address this need, we have developed a process for rapid prototyping MPM devices using computer numerical control (CNC) micromilling of commercially available microplates. Microchannels are cut out of the existing microplate structure and ports are drilled into the bottom of the wells to interface the wells. To test versatility and benchmark to another rapid-prototyping approach, we manufactured common microfluidic features into microplates using four different CNC mills as well as a 3D printer. Cell viability was assessed for polystyrene (PS) well plates and two 3D printed resins (MED610 and VeroClear) with the PS showing >2.5-fold increase in cell growth after three days. Machines were tested on their ability to create common device features including a traditional microfluidic device as well as a custom design incorporating complex geometries. Features were measured by confocal microscopy. We found that features including 1000{micro}m ports, 800{micro}m microchannels, 200{micro}m phase-guides, and 500{micro}m post arrays were machined and the range of CVs for features were 1.02-4.42, 1.32-3.50, 2.34-16.58, 6.25-16.40 respectively, while the 3D printed features exhibited maximal CVs of 20.98, 11.68, 23.60, and 10.01 for the same features. Predictably, more expensive machines generally showed higher accuracy and lower variation, but many features could be created accurately and precisely by inexpensive (<$3000) machines facilitating the broader use of this technology to create a user customizable platform to support the prototyping, development, and testing of human relevant models with broad applications across the life sciences. Graphical abstractMultiple CNC mills are assessed on accuracy and precision of microfluidic features of interest for microphysiological model development creation. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=112 SRC="FIGDIR/small/615399v1_ufig1.gif" ALT="Figure 1"> View larger version (42K): org.highwire.dtl.DTLVardef@1be08eorg.highwire.dtl.DTLVardef@33b0a4org.highwire.dtl.DTLVardef@192167borg.highwire.dtl.DTLVardef@200b7_HPS_FORMAT_FIGEXP M_FIG C_FIG

Biomedicine today is experiencing a shift towards decentralized data collection, which promises enhanced reproducibility and collaboration across diverse laboratory environments. This inter-laboratory study evaluates the performance of biocytometry, a method utilizing engineered bioparticles for enumerating cells based on their surface antigen patterns. In a decentralized framework, spanning 78 assays conducted by 30 users across 12 distinct laboratories, biocytometry consistently demonstrated significant statistical power in discriminating numbers of target cells at varying concentrations as low as 1 cell per 100,000 background cells. User skill levels varied from expert to beginner capturing a range of proficiencies. Measurement was performed in a decentralized environment without any instrument cross-calibration or advanced user training outside of a basic instruction manual. The results affirm biocytometry to be a viable solution for immunophenotyping applications demanding sensitivity as well as scalability and reproducibility and paves the way for decentralized analysis of rare cells in heterogeneous samples.

Liquid handling is a fundamental capability for many scientific experiments. Previously, we introduced the Surface Patterned Omniphobic Tiles (SPOTs) platform, which enables manipulation of hundreds to thousands of independent experiments without costly equipment or excessive consumable expenses. However, the SPOTs platform requires a custom coating formulation and lacks robustness. To overcome these limitations, we introduce EZ-SPOTs. These devices can be created in an hour with common fabrication tools and just three components - glass, a hydrophobic coating, and acrylic. EZ-SPOTs preserve many of the SPOTs platforms strengths - ease of use, ability to handle a wide range of volumes, and scalability - and adopt a durable and abrasion resistant coating that enables multiple reuses of each device. Here, we describe the fabrication of EZ-SPOTs and showcase how its reusability allows antibiotic susceptibility testing of many isolates using a single device. These results quantitatively match current gold standard assays and the increased throughput provides substantially more information than standard approaches.

Live-cell microscopy imaging typically involves the use of high-quality glass-bottom chambers that allow cell culture, gaseous buffer exchange and optical properties suitable for microscopy applications. However, commercial sources of these chambers can add significant annual costs to cell biology laboratories. Consumer products in three-dimensional printing technology, for both Filament Deposition Modeling (FDM) and Masked Stereo Lithography (MSLA), have resulted in more biomedical research labs adopting the use of these devices for prototyping and manufacturing of lab plastic-based items, but rarely consumables. Here we describe a modular, live-cell chamber with multiple design options that can be mixed per experiment. Single reusable carriers and the use of biodegradable plastics, in a hybrid of FDM and MSLA manufacturing methods, reduce plastic waste. The system is easy to adapt to bespoke designs, with concept-to-prototype in a single day, offers significant cost savings to the users over commercial sources, and no loss in dimensional quality or reliability.

Cell culture automation has traditionally been limited to basic tasks at low throughput, which are insufficient for passaging rapidly proliferating cell lines or for generating stable clonal lines. To address unmet needs, this study implemented a Biomek i7 Hybrid automated workstation, integrated with peripheral instruments and coordinated by SAMI EX software, to enable automated, high throughput mammalian cell culture workflows. The workflows support cell density monitoring, arrayed passaging, sample cherry-picking, plate reformatting, cell density normalization, and cryopreservation in 96-well plates. Integration with the CloneSelect imager allows rapid confluency monitoring and monoclonality assessment (<100 sec per plate). Cell passaging and density normalization require 32 minutes for one plate and 61 minutes for two plates. Workflow consistency was demonstrated across multiple cell lines and biological replicates, with wells showing comparable confluency within three standard deviations, lower coefficient of variation, and substantially narrower interquartile ranges after a single cell passage and density normalization. Four automation pipelines, including monoclonality screening, cell passaging and cherry-picking, density normalization, and cryopreservation, collectively enable clonal line establishment. Depending on scale, one to eight 384-well plates were processed in 69 to 355 minutes, yielding an average of 35 clonal lines per plate suitable for downstream genomic DNA sequence confirmation.

Scratch assays are routinely performed for multiple research applications in cell biology (i.e., migration, metastasis, repair mechanisms, etc.). Conventional scratches are usually generated with pipette tips. User inconsistencies and varying pipette tips are major obstacles to reproducibility and quantitative power. Here, we present a novel, scratch/gap generating, cell culture plate-based, platform that addresses these issues, providing consistency and reproducibility across multiple users, significantly reducing variability, and increasing quantitative outcomes.

New in vitro models are an urgent need for to improve both the research data and the preclinical development of new drugs. Current standardized cellular models are mainly based in 2D cell culture, which lacks flow conditions and with complex co-culture settings. In this way, advanced cell culture models, such as organ-on-a-chip (OoC), aim to solve these limitations. OoC systems are composed by a microfluidic chip functionalized with different combinations of extracellular matrixes, coatings and cell cultures to mimic the physiological conditions of human organs. Advantages of OoC include the possibility to add 3D structures, delimited regions for co-culture and dynamic flow conditions to cell cultures. However, to perform reproducible and controlled experiments with OoC, it is necessary to systematically standardize the cell culture conditions in the microfluidic channels. For this is necessary to test both the combination of flow and extracellular matrix (ECM) coating to reliably mimic the human organ physiology. In this work, we standardized both conditions, ECM coating and the flow conditions to functionalize OoC with cell lines from kidney (Caki-1) and from lung (A549) to develop OoC systems beyond the Vessel-on-a-chip setting. In this way, the protocol detailed in this work will allow to standardize cell culture on different optimized OoC types with different cell types from different origins.

In vitro cytotoxicity assessments frequently rely on staining-based methods that indirectly estimate viable cell numbers indirectly. A major limitation of many such techniques is their endpoint nature, requiring cell lysis or irreversible processing that precludes longitudinal monitoring of cellular responses following treatment. An ideal assay for evaluating cell viability and proliferation should be simple, rapid, cost-effective, reproducible, and highly sensitive, while also enabling accurate quantification with minimal interference from test compounds. The resazurin reduction assay satisfies these criteria, offering a sensitive and economical alternative to conventional tetrazolium-based methods. Although both assay types depend on the metabolic reduction of a dye by viable cells, they differ mechanistically. Tetrazolium salts (e.g., MTT) are reduced by cellular dehydrogenases to insoluble formazan crystals that require solubilization before to detection. In contrast, resazurin--a cell-permeable, non-fluorescent blue dye--is reduced to resorufin, a highly fluorescent compound detectable without additional processing steps. This property renders the resazurin assay broadly applicable to viability testing in eukaryotic cells cultured in both 2D and 3D formats, as well as in bacterial systems. Here, we present a streamlined, universal protocol for implementing the resazurin reduction assay across diverse experimental models, emphasizing its practicality, reproducibility, and adaptability for real-time viability monitoring. Key featuresO_LIReal-time, non-destructive monitoring: Enables longitudinal studies by allowing repeated measurements of the same samples over hours without toxicity or disruption. C_LIO_LIStreamlined workflow: A simple "add-incubate-read" protocol eliminates the need for cell lysis, washing, or extraction, saving time and reducing variability. C_LIO_LIBroad sample compatibility: Versatile and reliable for use with 2D monolayers, 3D spheroids, organoids, and bacterial cultures. C_LIO_LIHigh sensitivity: Fluorescent detection of resorufin provides exceptional sensitivity, enabling accurate quantification of even small viable cell populations. C_LIO_LILow background and minimal interference: A clean fluorescent readout reduces the risk of signal artifacts, offering a more reliable alternative to traditional colorimetric assays. C_LIO_LICost-effective and accessible: Utilizes standard laboratory plate readers and commercially available reagents, making it an economical choice for any lab. C_LIO_LIScalable for high-throughput screening: Easily adaptable to various plate formats, supporting both small-scale experiments and large-scale automated screening applications. C_LI Graphical overview O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=141 SRC="FIGDIR/small/718248v1_ufig1.gif" ALT="Figure 1"> View larger version (56K): org.highwire.dtl.DTLVardef@82bcecorg.highwire.dtl.DTLVardef@14164aforg.highwire.dtl.DTLVardef@395118org.highwire.dtl.DTLVardef@fb1349_HPS_FORMAT_FIGEXP M_FIG C_FIG

Stable and productive CHO cell lines are essential for biopharmaceutical manufacturing, yet early expansion steps are often constrained by prolonged period required for suspension adaptation. Single-cell cloning (SCC) ensures monoclonality and regulatory compliance, but cells transitioning from static to suspension culture frequently exhibit variable recovery, which prolongs timelines and increases process variability. To address this challenge, mixing-based microplate culture systems have been developed to improve early expansion efficiency. The C.NEST platform provides controlled pneumatic mixing and environmental monitoring that facilitates earlier adaptation to suspension conditions. At the 96-well and 24-well stages, this system allows cells to establish stable growth under suspension-like environments, thereby shortening the adaptation period following transfer to shaking culture. In this study, we applied C.NEST to the SCC workflow for developing CHO-K1 stable cell lines. Integrating C.NESTs controlled mixing reduced adaptation time, enhanced the consistency of clone expansion, and improved the ability to identify high-yield clones. These findings highlight the potential of C.NEST to streamline cell line development workflows by accelerating early suspension adaptation and improving clone selection reliability. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=165 SRC="FIGDIR/small/693844v1_ufig1.gif" ALT="Figure 1"> View larger version (40K): org.highwire.dtl.DTLVardef@ca76bforg.highwire.dtl.DTLVardef@3a3079org.highwire.dtl.DTLVardef@447e01org.highwire.dtl.DTLVardef@ac8d6f_HPS_FORMAT_FIGEXP M_FIG C_FIG C.NEST mixing shortens suspension adaptation, accelerates clone expansion, and enhances early-stage screening. HighlightO_LIThe C.NEST microplate agitation culture system accelerates early CHO-K1 cell line development. C_LIO_LIControlled pneumatic mixing improved oxygen transfer and medium homogeneity, promoting stable growth during early expansion. C_LIO_LIEarly mixing shortens suspension adaptation by approximately one week. C_LIO_LIMixing cultures enabled more accurate clone performance assessment, revealing high-producing outliers. C_LIO_LIC.NEST provides a scalable and reproducible solution for integrating mixing-based culture into single-cell cloning workflows. C_LI

By combining the radial migration assay with injection molded gaskets and a rigid fixture, we have developed a more reliable and sensitive method for measuring radial cell migration. This method is well adapted for use on high throughput automated imaging systems. The use of injection molded gaskets enables low cost replacement of cell-wetted components. Furthermore, the design enables secondary placement of attractants and co-cultures. This device and high-throughput application permit the use of therapeutic screening to evaluate phenotypic responses e.g. cancer cell migration. This approach is orthogonal to other 2D cell migration applications such as scratch wound assays, although here we offer a non-invasive, high-throughput device which is currently not commercially available. Collectively, we have designed a systematic, reliable, high-throughput application to monitor phenotypic responses to chemotherapeutic screens, genetic alterations (e.g. RNAi; CRISPR; others), supplemental regiments, and other approaches offering a reliable methodology to survey unbiased and non-invasive cell migration.

The expanding scope of laboratory automation increasingly demands systems that can be tailored to specific experimental constraints, including footprint, timing, cost, and control. While open-source software has improved protocol flexibility, liquid-handling hardware itself remains largely closed, limiting the ability of academic and startup laboratories to build instruments around biological requirements rather than vendor defaults. Here, we present a fully open-source, purpose-built liquid-handling robot assembled from commercially available components and developed entirely in a research setting. The platform integrates open hardware, electronics, and a Python-based control stack compatible with PyLabRobot, exposing low-level motion dynamics and liquid-handling behaviors directly to experiment code. We validate the system using a high-throughput turbidostat workflow that requires rapid, closed-loop measurement and actuation to maintain microbial cultures at defined density setpoints. The robot sustains stable steady-state growth across approximately 200 cultures with heterogeneous growth dynamics. A replica build completed by two lab members in approximately one week confirms that the platform can be reproduced from its bill of materials and assembly guide. Its compact footprint and use of off-the-shelf components make it suitable for rapid, parallel deployment in settings such as public health emergencies or by distributed laboratories. Together, these results demonstrate that industry-class liquid handlers can be custom-built for specific experimental goals, establishing a blueprint for open, purpose-driven hardware development across research and industrial automation contexts. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=132 SRC="FIGDIR/small/709168v1_ufig1.gif" ALT="Figure 1"> View larger version (63K): org.highwire.dtl.DTLVardef@1b2cb4eorg.highwire.dtl.DTLVardef@1418e8dorg.highwire.dtl.DTLVardef@f60618org.highwire.dtl.DTLVardef@a3d3b_HPS_FORMAT_FIGEXP M_FIG Open Liquid Handler (OLH) Design Goals. Left: Design goals for a purpose-built platform for time-sensitive, closed-loop biological workflows, emphasizing high-accuracy dosing (low variability liquid handling), rapid integrated measurement (plate deck and isolated workspace), customizable deck and peripheral options, compact footprint with high throughput, containment via an enclosed wet workspace for biosafety and sterility, and a replicable build using off-the-shelf OEM components with open design files. Right: Open Liquid Handler design and physical implementation, with aerial and front views highlighting the enclosed cabinet and the working envelope over a compact deck. C_FIG

O_LIThe escalating threats to biodiversity, public health, and food security posed by emerging infectious diseases and illegal wildlife trafficking requires novel approaches to biosurveillance. This paper introduces two innovations developed to address these multifaceted challenges: the Nucleic Acid Barcode Identification Tool (NABIT) and the Molecular Development Kit (MDK). C_LIO_LIThe NABIT is a handheld, battery-powered device that enables genetic tests to be performed at the point of contact by non-technical users, creating a critical bridge between centralized laboratories and the field by reducing barriers to accessible and routine genetic testing. Verification testing and validation results for the NABIT and the lyophilized assays used with it demonstrate key performance parameters, including sample preparation, detection sensitivity, and stability of field-ready assays after 17 months without refrigeration. C_LIO_LIThe MDK complements the NABIT by providing a framework for third-party development and deployment of field-ready genetic assays. Similar to software development kits (SDKs), the MDK offers documentation, software tools, and NABIT hardware to accelerate the development of new assays, enabling early detection strategies and minimizing future losses. Crucially, the MDK empowers scientists worldwide to contribute to a new ecosystem for wildlife genetics and biosurveillance by developing their own field-ready tests for the NABIT. C_LIO_LIIn summary, the NABIT and MDK present the potential for a paradigm shift in biosurveillance, ecosystem monitoring, and biodiversity conservation, enabling decentralized genetic testing, early disease detection, and rapid response to protect sensitive ecosystems, public health, and food security. C_LI

Cryopreservation depends critically on suppression of ice formation by cryoprotective agents (CPAs), but limited data is available on the CPA concentration required for vitrification (Cv). Here, we introduce a high-throughput 384-well platform that integrates automated liquid handling, randomized plate layouts, and a binary-search strategy to rapidly determine Cv across hundreds of formulations. Relative to conventional methods, this approach increases throughput by [~]50-fold, compressing a year of measurements into one week, while markedly reducing manual labor. Across [~]200 CPA compositions, we demonstrate that environmental boundary conditions strongly influence vitrification behavior: plates sealed with silicone mats exhibited lower Cv than open plates, indicating that sealed configurations promote vitrification. Further, the data reveal a decrease in Cv with increasing CPA molecular weight, consistent with enhanced ice suppression by larger molecules. We also present a simple mixture model that accurately predicts Cv for a broad range of CPA formulations, including mixtures containing up to seven CPAs (R{superscript 2} > 0.94), and use this model to evaluate published CPA toxicity data to identify formulations that operate near their vitrification threshold while maintaining relatively low toxicity. Together, these results establish a framework for rapid Cv determination, predictive modeling of vitrification behavior, and rational design of CPA formulations.

Although numerous cell and gene therapies have received regulatory approval, their adoption has been hampered by high cost and challenges in scaling out manufacturing. Many autologous cell therapies are individually manufactured for each patient using traditional manual methods in high-cost environments. Therefore, robotics and automation offer a potential solution to meet the growing demand for such therapies. To ensure identical biological outcomes, automation must replicate validated manual workflows, a requirement that poses significant engineering challenges, especially for aseptic manipulation and compatibility with manual-centric instruments and consumables. Here we describe the design and performance of a modular robotic cluster consisting of specialized modules containing widely adopted equipment. The robotic arm uses custom end-effectors to handle standard consumables and instruments such as syringes, vials, bags, cell counters, bioreactors, incubators, and closed centrifuges. We compared the performance of skilled human operators against the robotic cluster across multiple tasks: transferring cells between sterile bags, cell counting, drawing volume from a vial to a syringe, and resuspension and sampling from both a bag and G-Rex 100M-CS bioreactor. The robotic system also executed high-complexity operations with industry-standard instruments: cell selection using a CytoSinct 1000 and wash/buffer exchange with a CTS Rotea Counterflow centrifugation system. Experimental results for each unit operation show that the robotic clusters performance is equivalent to manual operations on the selected key metrics. These data demonstrate that the robotic system can efficiently and robustly perform specific unit operations, which can be combined in any order for end-to-end cell therapy manufacturing processes. One Sentence summaryA modular robotic system can perform key unit operations in cell therapy manufacturing with accuracy comparable to human operators while ensuring throughput and flexibility.

Organoid technology offers a powerful platform for modeling human tissues, studying disease mechanisms, and developing personalized therapies. However, widespread clinical application is hindered by challenges in scalability, reproducibility, and the handling of ultra-small tissue samples typical of clinical biopsies. Here, we present a comprehensive and automated protocol for the formulation and characterization of microfluidic droplet-engineered organoids (DEOs) derived from primary tissue samples. This protocol integrates 3 key stages: (1) Extraction and purification of viable primary cells from ultra-small tissue specimens using the small-Tissue Extraction Device (sTED); (2) High-throughput fabrication of uniform cell-laden microspheres using an integrated microfluidic bioprinter (OrgFab), capable of generating over 100 organoids from just 10 L of bioink; and (3) Rapid organoid characterization using lamination-based processing for single-cell analysis while preserving spatial context. The automated workflow minimizes manual intervention, reducing variability and enhancing reproducibility, making it suitable for high-throughput applications such as drug screening and disease modeling. Our method allows for the generation of patient-derived organoids that closely mimic the native tissue microenvironment, including diverse cell types and structural features, within a significantly reduced timeframe. This 3 stage protocol enhances the use of organoids in personalized medicine by enabling the rapid assessment of drug efficacy in patient-specific models. Here, the integration of advanced techniques supports existing organoid protocols, providing a valuable resource for researchers and clinicians seeking to improve patient outcomes through more efficient and precise organoid-based applications.

Data-driven materials development requires large, well-characterized libraries of precisely defined formulations. While microfluidic platforms excel at generating highly controlled materials, their throughput is often limited by the challenge of efficiently interfacing device outputs with standard well plates. This bottleneck frequently necessitates manual transfer or non-microfluidic workflows, constraining both throughput and reproducibility. Here, we present LMNOP-bot (Libraries of Micro- and Nano-materials, OPen-source bot), an open-source robotic platform for the automated generation and collection of micro- and nanomaterial libraries from serial microfluidic outputs. Using synchronized, pressure-driven flow, LMNOP-bot enables continuous formulation and direct deposition into standard well plates. The system is low-cost (<$700, excluding pressure regulators), constructed from readily available or easily fabricated components, and designed for broad accessibility. LMNOP-bot collects [&ge;]30 {micro}L per formulation at a rate of one sample every four seconds, representing an approximately 50x increase in throughput over existing serial microfluidic workflows, and operates robustly for over 10,000 runs without maintenance. We demonstrate compatibility with both PDMS/glass and commercial polycarbonate devices, with seamless interfacing to 96- and 384-well plates. Repeated sampling confirms high precision and reproducibility. By removing a key bottleneck in microfluidic library generation, LMNOP-bot enables rapid, scalable, and accessible exploration of material design spaces.

Precise immobilization of zebrafish larvae is critical for high-resolution neural imaging and behavioral studies, yet conventional methods are often labor-intensive, low-throughput, and can induce stress. To address these challenges, we introduce the Agarose Stamped Device (ASD), a novel platform enabling efficient and gentle immobilization of zebrafish larvae for high-resolution imaging and behavioral assays. By employing a stamping approach to mold an array of consistent agarose wells, the ASD enables rapid, parallel, and reproducible positioning of larvae in a standardized orientation while minimizing stress and preserving viability. We validated ASD performance by comparing larval survival, heart rate, and imaging stability to conventional agarose embedding, finding no adverse effects on larval physiology and significantly enhanced imaging throughput and data consistency. Demonstrations in neuroscience and behavioral experiments underscore the devices versatility for diverse in vivo studies. The ASD offers a simple, cost-effective, and powerful tool for advancing zebrafish-based neuroscience and behavioral research.

Restoring the epidermal barrier after injury requires spatial and temporal orchestration of migration, proliferation, and signaling across many cell types. The mechanisms that coordinate this complex process are incompletely understood. In vitro wound assays are common model systems for examining these mechanisms in wound healing. In the scratch assay, a cell-free gap is created by mechanical removal of cells from a monolayer, followed by monitoring cell migration into the gap over time. While simple and low-cost, manual scratch assays are limited by low reproducibility and low throughput. Here, we have designed a robotics-assisted automated wound healing (AWH) assay that increases reproducibility and throughput while integrating automated live-cell imaging and analysis. Wounds are designed as computer-aided design (CAD) models and recreated in confluent cell layers by the BioAssemblyBot (BAB) 3D-bioprinting platform. The dynamics of migration and proliferation in individual cells are evaluated over the course of wound closure using live-cell fluorescence microscopy and our high-performance image processing pipeline. The AWH assay outperforms the standard scratch assay with enhanced consistency in wound geometry. Our ability to create diverse wound shapes in any multi-well plate with the BAB not only allows for multiple experimental conditions to be analyzed in parallel but also offers versatility in the design of wound healing experiments. Our method emerges as a valuable tool for the automated completion and analysis of high-throughput, reproducible, and adaptable in vitro wound healing assays.

3D spheroids, which closely replicate three-dimensional cell-cell and cell-extracellular matrix interactions, offer superior predictive capabilities compared to conventional 2D monolayer cultures, positioning them as forward-looking platforms in drug testing, cancer biology, and regenerative medicine. However, high-throughput generation of uniform sized spheroids is still a technological challenge. In one hand, the use of conventional ultra-low attachment (ULA) multiwell plates for this purpose is labour intensive and complex. On the other hand, the use of microfabricated facilities demands cutting-edge infrastructure such as clean room, photolithography, and microfluidic setup which are often unavailable for the resource constrained laboratories. In this study, we addressed these problems by developing a low-cost Do-It-Yourself (DIY), polydimethylsiloxane (PDMS) and agarose-based spheroid generation device, capable of producing and maintaining hundreds of spheroids with minimal user intervention. We have demonstrated two variants based on their size, termed here as S1 and S2 devices which fit into 6-well and 12-well plates, and can generate 600 and 1200 uniform-sized spheroids respectively. We validated our device with various cell lines including primary and cancerous cell lines. We further demonstrated the drug testing capabilities of the device by estimating the IC50 value of the anticancer drug Temozolomide on U87-MG. The value was comparable with the same obtained from the spheroids generated using conventional ULA plates. Additional attachment of a perfusion system made the device suitable for long-term spheroid culture without much user intervention. Furthermore, the devices can also be used for the production of spheroids with gradually changing diameters in a controlled manner, resembling a size gradient. This feature is useful for checking the effect of drugs on different-sized spheroids and for co-culturing spheroids with varying cell densities, mimicking the disease architecture. We have co-cultured two types of the placental trophoblast cells, i.e., extravillous trophoblast (HTR-8) and syncytiotrophoblast (BeWo) with varying densities. In summary, this paper demonstrates a unique DIY method for a high-throughput uniform-sized spheroid generation at a fraction of cost which can be deployed to resource-constrained labs.

There is a constant pressure to reduce timelines in mammalian cell line development (CLD) for biotherapeutic protein production. Demonstration of clonal derivation of the generated cell lines is key for health authorities approval. To meet these regulatory and process-oriented demands, single-cell dispensers have become vital instruments for single-cell cloning. We conducted validation experiments with the UP.SIGHT (CYTENA GmbH) to determine this instruments single-cell dispensing efficiency (SCDE) and probability of clonal derivation (p(clonal)). Process optimization to maximize clone recovery with several cell lines was also performed, focusing on cloning media and plate type. With a SCDE >97%, p(clonal) >99.99% and clone recovery values of up to 80%, the data reported here support the notion that the UP.SIGHT covers all steps in the single-cell dispensing process with assurance of clonality and colony tracking, leading to faster and more efficient CLD workflows. This work also serves as a guideline for instrument validation and guidance towards process optimization.