The research is supported by the CALIPSO project supported by the National Research Foundation, Prime Minister&#39;s Office, Singapore, under its Campus for Research Excellence and Technological Enterprise (CREATE) programme. V.V. acknowledges the support of NRF investigator NRF-NRFI2018-07, MOE Tier 3 MOE2016-T3-1-005, MBI seed funding, and ANR ADGastrulo. A.B. and G.G. acknowledge the support from MBI core funding. A.B. acknowledges Andor Technologies for the loan of the BC43 microscope.

NOTE: The first part of this protocol details the microfabrication of the cell culture device. An original primary mold with pyramidal cavities can be produced in-house-if micro-fabrication facilities are available-or outsourced to external companies. The primary mold used in this work is produced in-house, with fabrication process steps described elsewhere11,13. A basic protocol for the microfabrication of the mold is available in Supplementary File 1. CRITICAL: The operations in steps 1 to 6 need to be conducted in a dust-free environment. A laminar flow hood or a clean room, if available, are preferred. All through these steps, personal protective equipment (PPE) needs to be used, such as gloves, a lab coat, and safety glasses.
1. Dicing of PDMS mold
<ol>
	<li>Cut small portions of the PDMS mold to the dimension required for the final device (e.g., 1 cm x 1 cm [Figure 1B]). Cut them parallel to the XY directions of the array. 
	&#8203;CRITICAL: When cutting the PDMS mold in 1 cm x 1 cm dices, execute the cuts in single steps; using a one-sided razorblade, apply pressure and cut through the PDMS in one single step. This is to prevent the formation of small particles of PDMS, which can deposit on the surface of the mold and affect the quality of the replica steps (step 3).</li>
</ol>
2. Preparation of flat PDMS substrates
<ol>
	<li>Weigh ~15-20 g of PDMS base resin and 1.5-2 g of its reticulation agent (i.e., a ratio of 10:1), mix them carefully, and degas in a vacuum jar for ~20 min.</li>
	<li>Slowly pour the degassed PDMS onto a plastic, 15 cm wide (diameter) Petri dish.</li>
	<li>Cure the PDMS by keeping the Petri dish in an oven set at 65&#176; C for 2 h. After waiting for the curing to complete, a flat PDMS film of ~1.5 mm thickness (contained in the Petri dish) is ready for use (Figure 2A).</li>
	<li>Using a sharp blade, cut 2 cm x 2 cm pieces out of the PDMS (Figure 2B-D). 
	&#8203;NOTE: The exact thickness of this PDMS sheet is not critical; ~1-2 mm is a suitable range. The dimension for the cut should be bigger than the textured mold but not much larger, which would result in wasting of material.</li>
</ol>
3. Production of the textured layer made of UV-curable adhesive
<ol>
	<li>Place one PDMS mold dice (as prepared in step 1) face-down on top of the flat PDMS cut (as prepared in step 2) (Figure 3A,B). 
	NOTE: Make sure the pyramidal protrusions in the mold are oriented toward the flat PDMS substrate and that only the top surface of the truncated pyramids is in contact. Assess the correct placement with an optical microscope (Figure 3C,D).</li>
	<li>To one side of the mold, using a pipette, drop a small amount (two drops, approximately 0.1-0.2 mL) of UV-curable adhesive (Figure 4A). 
	NOTE: As the liquid gets in contact with the edge of the mold, capillarity will drive the liquid to fill the cavity between the mold itself and the flat cut of PDMS used as substrate.
	<ol>
		<li>Follow the progression of the liquid inside the cavity, for example, using an inverted optical microscope with a 10x magnification objective (Figure 4B,C).</li>
	</ol>
	</li>
	<li>Expose the UV-curable adhesive to UV light to cure it. Adjust the time of exposure depending on the power density of the UV source used (e.g., a UV-LED box with a power density of 35 mW/cm2; 2 min at 50% power in this protocol to cure the adhesive completely).</li>
	<li>Using the excess amount of adhesive at one edge, hold the cured adhesive on the flat PDMS substrate by gently pressing with a finger (Figure 5A,B). Meanwhile, use tweezers to pinch one corner of the mold next to the same edge being held down and slowly peel it off while making sure that the textured film is not lifted up as well. 
	NOTE: Figure 5C shows the result of a proper mold removal procedure; Figure 5E-G shows the wrong procedure.</li>
	<li>Trim the excess adhesive and the excess PDMS substrate using a razor blade to leave the cured, textured, adhesive layer flat on the PDMS with excess PDMS on one edge only (Figure 5D), which will be needed in step 5.</li>
</ol>
4. Coverslip substrate preparation
NOTE: As a substrate for the final device, standard rounded 1.5H coverslips with 25 mm diameter are used. Before they can be used, they need to be cleaned to remove dust and/or organic residual from their surface.
<ol>
	<li>Coverslip cleaning
	<ol>
		<li>Immerse the coverslips in soapy water for 5 min while the ultrasound is applied (40 kHz, 110 W sonic power).</li>
		<li>Wash the coverslips in clean deionized (DI) water, first by immersion and finally with running DI water from a tap. Dry the coverslips with an N2 gas blow-gun.</li>
		<li>Immerse the coverslips in an acetone bath for 5 min; immediately move to a 2-propanol (IPA) bath for an additional 5 min.</li>
		<li>Rinse the coverslips with clean IPA using a squeeze bottle. Dry the coverslips with the N2 gas blow-gun. 
		NOTE: Coverslips cleaned using this procedure can be stored in a closed container until needed. Keep them in a dry cabinet to avoid humidity depositing on their surface.</li>
	</ol>
	</li>
	<li>When ready to be used, treat a clean coverslip with a short O2 plasma process to improve its hydrophilicity: O2 20 sccm, 3 mbar pressure, 60 W at the RF power generator, and 60 s duration.</li>
	<li>Immediately after the plasma activation, proceed with spin-coating the thin layer of UV-curable adhesive by placing the coverslip on the vacuum chuck of a standard spin-coater and pouring a small drop of adhesive at the center of the coverslip (Figure 6). Run the spin-coating process: spreading for 5 s at 500 rpm, coating at 3,000 rpm for 45 s (with the acceleration and deceleration set at 100 rpm/s).
	<ol>
		<li>If spin-coating is not available, use the following alternative way to produce a thin film of UV adhesive on the coverslips:
		<ol>
			<li>On a clean coverslip, drop ~0.1 mL of UV-curable adhesive using a pipette (Figure 7A).</li>
			<li>Take a second coverslip and place it on top of the first one to make the liquid adhesive spread evenly between the two coverslips (Figure 7B-D).</li>
			<li>Once the spreading adhesive has reached the edges of the coverslips, gently separate them by sliding one over the other. Once separated, both coverslips are fully coated with a thin layer of liquid adhesive (Figure 7E). 
			NOTE: The coating might not be uniform and smooth only if the separation is not done with a smooth and continuous movement.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Precuring of UV-curable adhesive
	<ol>
		<li>After spin-coating, precure the adhesive by exposure to UV. Adjust the time of exposure depending on the power density of the UV source used (here, a UV-LED box with a power density of 35 mW/cm2 was used for 1 min at 50% power). 
		CRITICAL: The adhesive used here is an optical glue. See the discussion for key points related to the energy doses for its curing.</li>
	</ol>
	</li>
</ol>
5. Transfer of the textured film to the final substrate
<ol>
	<li>Take one of the textured films (prepared in step 3) and place it in contact with the adhesive-coated coverslip (prepared in step 4). Make sure the contact between the partially cured adhesive on the coverslip and the textured film is as uniform as possible (Figure 8A-C). 
	CRITICAL: At this stage, the adhesive on the coverslip should be solid enough to avoid reflowing, which would fill the pyramidal cavities of the textured film when placed in contact but also be plastic and adhesive enough that contact can be optimized by gently pressing on the textured film.</li>
	<li>Expose the coverslips to UV light until the adhesive layer coated on the coverslip is fully cured. This will seal the textured film on the coverslip and provide leak-proof isolation between the pyramidal cavities.</li>
	<li>Finally, peel off the PDMS flat substrate (Figure 8D-F). Using tweezers, pinch the PDMS on one corner at the edge where excess material was left after trimming (step 3.5). This way, the textured film layer is left adhesive to the coverslip with open access at the top for cell seeding.CRITICAL: When peeling off the flat PDMS, the textured film should remain well attached to the coverslip. Adhesion failures are easily confirmed if the textured film can be peeled off from the coverslip after final exposure to UV without any effort.</li>
</ol>
6. Long-term passivation of the cell culture coverslip for cell culture
NOTE: Passivation is achieved by generating a conformal and continuous coating of a biomimetic copolymer with a structure similar to the polar group of phospholipids in the cell membrane. This conformal coating prevents cells adhesion to the cell culture device
<ol>
	<li>Prepare a solution with 0.5% (w/v) of the biomimetic copolymer dissolved in pure ethanol. Store the solution at 4 &#176;C for future use.</li>
	<li>Place the cell culture coverslip in a 35 mm Petri dish and fully cover it with the biomimetic copolymer solution.</li>
	<li>After 5 min, remove the cell culture coverslip from the container with the biomimetic copolymer solution and leave it to dry at room temperature inside the final dish in a biosafety hood (&#62;1 h). 
	NOTE: A thicker coating can be produced by increasing the concentration of the biomimetic copolymer in the coating solution; the results of a thicker coating are visible under a brightfield microscope (Figure 9A).</li>
</ol>
7. Cell seeding
<ol>
	<li>Degassing and sterilization
	<ol>
		<li>Just before cell seeding, dispense sterile phosphate-buffered saline (PBS) into the cell culture dishes (typically 1 mL for a 35 mm Petri dish). Degas the dish with the sterile PBS using an ultrasonic device for ~10 min, followed by several rounds of pipetting to remove all the bubbles. 
		CRITICAL: If air is trapped inside the pyramidal cavities, it will prevent the cells from entering them. To make sure there is no air trapped in the pyramid before cell seeding, it is recommended to visually ensure (under a benchtop brightfield microscope at 10x or 20x magnification) the absence of air in these cavities (Figure 10).</li>
		<li>Replace the PBS with sterile culture medium and sterilize the plate with UV light for 30 min under a cell culture hood. 
		NOTE: From this step onward, the dish should be considered as sterile and manipulated using sterile techniques. An alternative way to remove trapped air from the cell culture device is to use a vacuum jar with a vacuum pump.</li>
	</ol>
	</li>
	<li>Cell suspension preparation 
	NOTE: Cells can be seeded as single or small cell aggregates and enter the sample wells through the top aperture. Over time, the inserted cells aggregate and grow inside the sample wells into spheroids of a size greater than the size of the aperture. As validated cell line models, use HCT116 (CCL-247 ATCC) or MCF7 (HTB-22 ATCC) cancer cells maintained in recommended culture medium (ATCC guidelines).
	<ol>
		<li>Prepare a cell suspension (e.g., using a trypsinization process following ATCC guidelines). Follow trypsinization/cell suspension preparation recommendations for the cells of interests.</li>
		<li>Count and adjust the cell concentration to 0.5 &#215; 106 cells/mL in the recommended culture medium.</li>
	</ol>
	</li>
	<li>Cell dispensing
	<ol>
		<li>Remove the PBS buffer from the 35 mm cell culture dish and then dispense 1 mL of the adjusted cell suspension. See Figure 11A for an optical microscope image of a cell seeding procedure with adequate cell density and homogeneity.</li>
		<li>Place the cell culture dish back into the cell incubator (37 &#176;C, 5% CO2, and 100% humidity) for 10 min. Approximately 80-100 cells will enter each pyramidal cavity. 
		NOTE: It is possible to increase the number of cells per cell culture dish by increasing either the cell concentration or the time spent before cell suspension removal. Typically, spheroid formation takes several hours (depending on cell type) after cell seeding and can be followed with a brightfield microscope (4x to 40x objectives; Figure 11). From here, culture medium, extracellular matrices, and differentiation growth factors can be changed or added to the cell culture dish containing the spheroids in accordance with typical differentiation protocols that could last a few days, weeks, or months.</li>
		<li>After the 10 min incubation, recover the cell culture dish from the incubator and gently aspirate the cellular suspension to remove untrapped cells. Add 1 mL of culture medium to a 35 mm dish and place it back into the cell incubator. 
		CRITICAL: At this stage, because spheroids have not formed yet, it is very important to avoid strong aspiration or dispensing that will result in loss of the cells. Visual control using a benchtop brightfield microscope is highly recommended at this step.</li>
	</ol>
	</li>
</ol>
8. Immunostaining and imaging
<ol>
	<li>Fixation and staining 
	NOTE: Different classical procedures of fixation and immunostaining are completely compatible with the cell culture dish. One typical protocol is described here.
	<ol>
		<li>Fix the organoids/spheroids in the cell culture dish for 20 min in 4% paraformaldehyde at room temperature.</li>
		<li>Permeabilize the organoids for 24 h in 1% surfactant solution in sterile PBS at 4 &#176;C on an orbital shaker and incubate for 24 h in blocking buffer (2% bovine serum albumin [BSA] and 1% surfactant in sterile PBS) at 4 &#176;C on an orbital shaker.</li>
		<li>Incubate the samples with primary antibodies of interest at a dilution between 1/50 and 1/200 (or according to the manufacturer&#39;s recommendations) in antibody dilution buffer (2% BSA and 0.2% surfactant in sterile PBS) at 4 &#176;C for 48 h.</li>
		<li>Rinse the samples 3x with washing buffer on an orbital shaker (3% NaCl and 0.2% surfactant in sterile PBS) and incubate with corresponding secondary antibodies in antibody dilution buffer (dilution between 1/100 and 1/300 or according to the manufacturer&#39;s recommendations), 0.5 &#181;g/mL 4&#39;,6-diamidino-2-phenylindole (DAPI), and 0.2 &#181;g/mL Alexa Fluor 647 or 488 Phalloidin at 4 &#176;C for 24 h on an orbital shaker, followed by five rinsing steps with PBS. Optionally, mount the samples using a water-soluble clearing agent prewarmed to 37 &#730;C.</li>
	</ol>
	</li>
	<li>Imaging 
	NOTE: At this stage, the organoids in the microwell plate can be considered as a normal culture dish containing fixed and stained samples for imaging: any standard imaging procedure can be used with no adaptation or modification required. Figure 12 illustrates a representative result of images and 3D reconstruction obtained using a spinning disk confocal microscope, with a 40x air objective (numerical aperture 0.75).
	<ol>
		<li>Use constructor software for automatic image acquisition process with the following settings: exposure time = 50 ms, with a z motorized stage to acquire a z-stack (1 &#181;m Z-step, for a total height of 70 &#181;m).</li>
		<li>Perform 3D reconstruction using image analysis software.</li>
	</ol>
	</li>
</ol>
9. Release and collection of the organoids
NOTE: The textured adhesive layer of the cell culture dish can be detached from the coverslip to release the living spheroids/organoids (before fixation) contained inside the pyramidal cavities for analysis of the cells with other procedures such as RNA sequencing, -omic approaches, in vitro experiments, and in vivo transplantation.
<ol>
	<li>With the sample still in the Petri dish and in a biosafety hood, use a blade such as a scalpel to cut a corner of the textured adhesive layer.</li>
	<li>With tweezers, pinch the textured adhesive layer on the cut edge and gently peel it off from the glass coverslip but keep it immersed in the medium (some organoids might be remaining with the adhesive layer, Figure 13).</li>
	<li>Rinse 3x with culture medium and collect the organoids by pipetting.</li>
</ol>

<ol>
	<li>Kim, J., Koo, B. -. K., Knoblich, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+organoids:+model+systems+for+human+biology+and+medicine.">Human organoids: model systems for human biology and medicine.</a> Nature Reviews Molecular Cell Biology. 21 (10), 571-584 (2020).</li><li>Takebe, T., Wells James, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organoids+by+design.">Organoids by design.</a> Science. 364 (6444), 956-959 (2019).</li><li>Kratochvil, M. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineered+materials+for+organoid+systems.">Engineered materials for organoid systems.</a> Nature Reviews Materials. 4 (9), 606-622 (2019).</li><li>Rossi, G., Manfrin, A., Lutolf, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progress+and+potential+in+organoid+research.">Progress and potential in organoid research.</a> Nature Reviews Genetics. 19 (11), 671-687 (2018).</li><li>O'Connell, L., Winter, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organoids:+past+learning+and+future+directions.">Organoids: past learning and future directions.</a> Stem Cells and Development. 29 (5), 281-289 (2020).</li><li>Vives, J., Batlle-Morera, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+challenge+of+developing+human+3D+organoids+into+medicines.">The challenge of developing human 3D organoids into medicines.</a> Stem Cell Research &amp; Therapy. 11 (1), 72 (2020).</li><li>Busslinger, G. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+potential+and+challenges+of+patient-derived+organoids+in+guiding+the+multimodality+treatment+of+upper+gastrointestinal+malignancies.">The potential and challenges of patient-derived organoids in guiding the multimodality treatment of upper gastrointestinal malignancies.</a> Open Biology. 10 (4), 190274 (2020).</li><li>Lukonin, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenotypic+landscape+of+intestinal+organoid+regeneration.">Phenotypic landscape of intestinal organoid regeneration.</a> Nature. 586 (7828), 275-280 (2020).</li><li>Rios, A. C., Clevers, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+organoids:+a+bright+future+ahead.">Imaging organoids: a bright future ahead.</a> Nature Methods. 15 (1), 24-26 (2018).</li><li>Dekkers, J. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+3D+imaging+of+fixed+and+cleared+organoids.">High-resolution 3D imaging of fixed and cleared organoids.</a> Nature Protocols. 14 (6), 1756-1771 (2019).</li><li>Galland, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+high-+and+super-resolution+imaging+using+single-objective+SPIM.">3D high- and super-resolution imaging using single-objective SPIM.</a> Nature Methods. 12 (7), 641-644 (2015).</li><li>Beghin, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+content+3D+imaging+method+for+quantitative+characterization+of+organoid+development+and+phenotype.">High content 3D imaging method for quantitative characterization of organoid development and phenotype.</a> bioRxiv. , (2021).</li><li>Beghin, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+high-speed+3D+imaging+of+organoid+cultures+with+multi-scale+phenotypic+quantification.">Automated high-speed 3D imaging of organoid cultures with multi-scale phenotypic quantification.</a> Nature Methods. 19 (7), 881-892 (2022).</li></ol>

An international patent application has been published with the Publication Number WO 2021/167535 A1.

The procedure for the fabrication of the microwell culture dish, which allows high-density organoid culture and differentiation, has been described in this paper. Owing to the geometry and arrangement of the microcavities, thousands of spheroids can be cultured and stained in a single plate for long periods of time (several weeks or more) with nearly no loss of material. As a comparison, an area of 4 mm x 2 mm on the cell culture plate can contain as many spheroids as a single 384-well plate with an area of 12 cm x 8 cm....

In organotypic 3D cell cultures, hereafter referred&#160;to as organoids, stem cells differentiate and self-organize into spatial structures that share strong morphological and functional similarities with real organs. Organoids offer valuable models to study human biology and development outside the body1,2,3. A growing number of models are being developed that mimic the liver, brain, kidney, lung, and many other organs2,4,5. Differentiation in organoids is directed by the addition of soluble growth factors and an extracellular matrix in a precise time sequence. However, in marked contrast to organs, the development of organoids is quite heterogeneous.
Beyond numerous biological challenges6,7, organoid cultures also pose technological challenges in terms of cell culture methods, characterization of transcriptomics, and imaging. In vivo organ development occurs in a biological environment that results in a highly stereotypical self-organization of cell arrangements. Any phenotypic alteration can be used as a proxy to diagnose a diseased state. In contrast, organoids develop in vitro in minimally controlled microenvironments compatible with cell culture conditions, resulting in large variability in the development path and shape formation for each individual organoid.
A recent study8 demonstrated that quantitative imaging of organoid shape (phenotype descriptors) coupled to the assessment of a few genetic markers allow the definition of phenotypic development landscapes. Arguably, the ability to relate the diversity of genomic expression in organoids with their phenotypic behavior is a major step toward unleashing the full potential of organotypic cultures. Thus, it begs for the development of dedicated, high-content imaging approaches allowing the characterization of organoid features at subcellular, multicellular, and whole-organoid scales in 3D9,10.
We developed a versatile high-content screening (HCS) platform allowing streamlined organoid culture (from isolated human embryonic stem cells [hESCs], human induced pluripotent stem cells [hIPSCs], or primary cells to 3D, multicellular, differentiated organoids) and fast, non-invasive 3D imaging. It integrates a next-generation, miniaturized, 3D cell culture device, called the JeWells chip (the chip hereafter), that contains thousands of well-arrayed microwells flanked with 45&#176; mirrors that allow fast, 3D, high-resolution imaging by single-objective light-sheet microscopy11. Compatible with any standard, commercial, inverted microscope, this system enables the imaging of 300 organoids in 3D with subcellular resolution in &#60;1 h.
The microfabrication of the cell culture device starts from an existing micro structured mold, which contains hundreds of micropyramids (Figure 1A) with a square base and sidewalls at 45&#176; with respect to the base. Figure 1C shows electron microscope (EM) images of such structures. The mold itself is made of poly(dimethylsiloxane) (PDMS) and can be made as a replica cast of a primary mold (not shown here) with corresponding features (as cavities) using standard soft-lithographic procedures. The primary mold can be produced by different procedures. The one used for this protocol was made using silicon wet etching as reported in Galland et al.11; the procedure for the fabrication of the primary mold is not critical for this protocol. The pyramids are arranged in a squared array, with the same pitch for the X and Y directions (in this case the pitch is 350 &#181;m).
As an illustration, proof-of-concept experiments12 were published to demonstrate that the chip allows long-term culture (months) and differentiation protocols while precisely defining the number of initial cells in the wells. Individual development of a large number of organoids can be automatically monitored live using standard brightfield and 3D light-sheet fluorescence microscopes. Moreover, organoids can be retrieved to perform further biological investigations (e.g., transcriptomic analysis). This paper outlines detailed protocols for the fabrication of the cell culture coverslips, the seeding and staining procedure for fluorescence microscopy, as well as the retrieval of the organoids.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2-Propanol</td>
			<td>Thermofisher scientific</td>
			<td>AA19397K7</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Thermofisher scientific</td>
			<td>AA19392K7</td>
			<td></td>
		</tr>
		<tr>
			<td>BC43 Benchtop Confocal Microscope</td>
			<td>Andor Technology</td>
			<td></td>
			<td>spinning disk confocal microscope</td>
		</tr>
		<tr>
			<td>bovine serum albumin&#160;</td>
			<td>Thermofisher scientific</td>
			<td>37525</td>
			<td></td>
		</tr>
		<tr>
			<td>Buffered oxide etching solution</td>
			<td>Merck</td>
			<td>901621-1L</td>
			<td></td>
		</tr>
		<tr>
			<td>CEE Spin Coater</td>
			<td>Brewer Science</td>
			<td>200X</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Thermofisher scientific</td>
			<td>62248</td>
			<td></td>
		</tr>
		<tr>
			<td>Developer AZ400K</td>
			<td>Merck</td>
			<td>18441223164</td>
			<td></td>
		</tr>
		<tr>
			<td>DI Milliq water</td>
			<td>Millipore</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>Invitrogen</td>
			<td>10082147</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslips</td>
			<td>Marienfled</td>
			<td>117650</td>
			<td>1.5H, round 25 mm diameter</td>
		</tr>
		<tr>
			<td>Hepes</td>
			<td>Invitrogen</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaris software</td>
			<td>BitPlane</td>
			<td></td>
			<td>image analysis software</td>
		</tr>
		<tr>
			<td>Inverted Transmission optical microscope</td>
			<td>Nikon</td>
			<td>TSF100-F</td>
			<td></td>
		</tr>
		<tr>
			<td>Labsonic M</td>
			<td>Sartorius Stedium Biotech</td>
			<td></td>
			<td>Ultrasonic homogenizer</td>
		</tr>
		<tr>
			<td>Lipidure</td>
			<td>NOF America</td>
			<td>CM5206</td>
			<td>bio-mimetic copolymer</td>
		</tr>
		<tr>
			<td>NOA73</td>
			<td>Norland Products</td>
			<td>17-345</td>
			<td>UV curable adhesive</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Invitrogen</td>
			<td>15070063</td>
			<td></td>
		</tr>
		<tr>
			<td>Phalloidin</td>
			<td>Thermofisher scientific&#160;</td>
			<td>A12379</td>
			<td>Alexa Fluor</td>
		</tr>
		<tr>
			<td>Phosphate Buffer Solution</td>
			<td>Thermofisher scientific</td>
			<td>10010023</td>
			<td></td>
		</tr>
		<tr>
			<td>Photo Resist AZ5214E</td>
			<td>Merck</td>
			<td>14744719710</td>
			<td></td>
		</tr>
		<tr>
			<td>Pico Plasma tool</td>
			<td>Diener Electronic GmbH + Co. KG</td>
			<td>Pico Plasma</td>
			<td>For O2 plasma treatment</td>
		</tr>
		<tr>
			<td>RapiClear 1.52</td>
			<td>Sunjin lab</td>
			<td>RC 152001</td>
			<td>water-soluble clearing agent</td>
		</tr>
		<tr>
			<td>RCT Hot Plate/Stirrer</td>
			<td>IKA (MY)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Reactive Ion Etching tool</td>
			<td>Samco Inc. (JPN)</td>
			<td>RIE-10NR</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>Invitrogen</td>
			<td>11875093</td>
			<td>culture medium for HCT116 cells</td>
		</tr>
		<tr>
			<td>Sylgard 184 Silicone Elastomer Kit</td>
			<td>Dow Corning</td>
			<td>4019862</td>
			<td>Polydimethylsiloxane or in short, PDMS</td>
		</tr>
		<tr>
			<td>Trichloro(1H,1H,2H,2H-perfluorooctyl)silane</td>
			<td>Sigma Aldrich</td>
			<td>448931-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma Aldrich</td>
			<td>T9284</td>
			<td>surfactant</td>
		</tr>
		<tr>
			<td>Trypsin EDTA</td>
			<td>Thermofisher scientific</td>
			<td>15400054</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic Cleaner</td>
			<td>Bransonic</td>
			<td>CPX2800</td>
			<td></td>
		</tr>
		<tr>
			<td>UV-KUB 2</td>
			<td>KLOE</td>
			<td></td>
			<td>UV-LED light source, 365 nm wavelength, 35 mW/cm2 power density</td>
		</tr>
		<tr>
			<td>UV mask aligner</td>
			<td>SUSS Microtec Semiconductor (DE)</td>
			<td>MJB4</td>
			<td></td>
		</tr>
	</tbody>
</table>

a high-throughput platform for culture and 3d imaging of organoids

The characterization of a large number of three-dimensional (3D) organotypic cultures (organoids) at different resolution scales is currently limited by standard imaging approaches. This protocol describes a way to prepare microfabricated organoid culture chips, which enable multiscale, 3D live imaging on a user-friendly instrument requiring minimal manipulations and capable of up to 300 organoids/h imaging throughput. These culture chips are compatible with both air and immersion objectives (air, water, oil, and silicone) and a wide range of common microscopes (e.g., spinning disk, point scanner confocal, wide field, and brightfield). Moreover, they can be used with light-sheet modalities such as the single-objective, single-plane illumination microscopy (SPIM) technology (soSPIM). 
The protocol described here gives detailed steps for the preparation of the microfabricated culture chips and the culture and staining of organoids. Only a short length of time is required to become familiar with, and consumables and equipment can be easily found in normal biolabs. Here, the 3D imaging capabilities will be demonstrated only with commercial standard microscopes (e.g., spinning disk for 3D reconstruction and wide field microscopy for routine monitoring).

Figure 8F shows the typical aspect of a cell culture coverslip after successful fabrication. The UV-curable adhesive layer appears flat and adheres well to the coverslip. The transfer of the adhesive layer on the coverslip might fail if the layer on the coverslip is overcured, or if the removal of the flat PDMS substrate is done incorrectly (as shown in Figure 8G,H). In both cases, the failure is evident as no textured film is transferred to the...

Watch this Scientific Journal Video about A High-Throughput Platform for Culture and 3D Imaging of Organoids at JoVE.com

A High-Throughput Platform for Culture and 3D Imaging of Organoids

This paper presents a fabrication protocol for a new type of culture substrate with hundreds of microcontainers per mm2, in which organoids can be cultured and observed using high-resolution microscopy. The cell seeding and immunostaining protocols are also detailed.

The characterization of a large number of three-dimensional (3D) organotypic cultures (organoids) at different resolution scales is currently limited by ...

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Biology

2.4K Views.  National University of Singapore. This paper presents a fabrication protocol for a new type of culture substrate with hundreds of microcontainers per mm2, in which organoids can be cultured and observed using high-resolution microscopy. The cell seeding and immunostaining protocols are also detailed.

Video: A High-Throughput Platform for Culture and 3D Imaging of Organoids

A Multi-compartment CNS Neuron-glia Co-culture Microfluidic Platform

We present a novel multi-compartment neuron co-culture microsystem platform for in vitro CNS axon-glia interaction research, capable of conducting up to six independent experiments in parallel for higher-throughput. We developed a new fabrication method to create microfluidic devices having both micro and macro scale structures within the same device through a single soft-lithography process, enabling mass fabrication with good repeatability.
The multi-compartment microfluidic co-culture platform is composed of one soma compartment for neurons and six axon/glia compartments for oligodendrocytes (OLs). The soma compartment and axon/glia compartments are connected by arrays of axon-guiding microchannels that function as physical barriers to confine neuronal soma in the soma compartment, while allowing axons to grow into axon/glia compartments. OLs loaded into axon/glia compartments can interact only with axons but not with neuronal soma or dendrites, enabling localized axon-glia interaction studies. The microchannels also enabled fluidic isolation between compartments, allowing six independent experiments to be conducted on a single device for higher throughput.
Soft-lithography using poly(dimethylsiloxane) (PDMS) is a commonly used technique in biomedical microdevices. Reservoirs on these devices are commonly defined by manual punching. Although simple, poor alignment and time consuming nature of the process makes this process not suitable when large numbers of reservoirs have to be repeatedly created. The newly developed method did not require manual punching of reservoirs, overcoming such limitations. First, seven reservoirs (depth: 3.5 mm) were made on a poly(methyl methacrylate) (PMMA) block using a micro-milling machine. Then, arrays of ridge microstructures, fabricated on a glass substrate, were hot-embossed against the PMMA block to define microchannels that connect the soma and axon/glia compartments. This process resulted in macro-scale reservoirs (3.5 mm) and micro-scale channels (2.5 &mu;m) to coincide within a single PMMA master. A PDMS replica that served as a mold master was obtained using soft-lithography and the final PDMS device was replicated from this master.
Primary neurons from E16-18 rats were loaded to the soma compartment and cultured for two weeks. After one week of cell culture, axons crossed microchannels and formed axonal only network layer inside axon/glia compartments. Axons grew uniformly throughout six axon/glia compartments and OLs from P1-2 rats were added to axon/glia compartments at 14 days in vitro for co-culture.

We developed a novel multi-compartment neuron co-culture microsystem platform for in vitro CNS axon-glia interaction research. The platform is capable of conducting up to six independent experiments in parallel and was fabricated using a newly developed macro/micro hybrid fabrication method.

We present a novel multi-compartment neuron co-culture microsystem platform for in vitro CNS axon-glia interaction research, capable of conducting up to ...

Lensless On-chip Imaging of Cells Provides a New Tool for High-throughput Cell-Biology and Medical Diagnostics

Conventional optical microscopes image cells by use of objective lenses that work together with other lenses and optical components. While quite effective, this classical approach has certain limitations for miniaturization of the imaging platform to make it compatible with the advanced state of the art in microfluidics. In this report, we introduce experimental details of a lensless on-chip imaging concept termed LUCAS (Lensless Ultra-wide field-of-view Cell monitoring Array platform based on Shadow imaging) that does not require any microscope objectives or other bulky optical components to image a heterogeneous cell solution over an ultra-wide field of view that can span as large as ~18 cm2. Moreover, unlike conventional microscopes, LUCAS can image a heterogeneous cell solution of interest over a depth-of-field of ~5 mm without the need for refocusing which corresponds to up to ~9 mL sample volume. This imaging platform records the shadows (i.e., lensless digital holograms) of each cell of interest within its field of view, and automated digital processing of these cell shadows can determine the type, the count and the relative positions of cells within the solution. Because it does not require any bulky optical components or mechanical scanning stages it offers a significantly miniaturized platform that at the same time reduces the cost, which is quite important for especially point of care diagnostic tools. Furthermore, the imaging throughput of this platform is orders of magnitude better than conventional optical microscopes, which could be exceedingly valuable for high-throughput cell-biology experiments.

Lensfree on-chip imaging and characterization of cells is illustrated. This on-chip cell imaging approach provides a compact and cost-effective tool for medical diagnostics and high-throughput cell biology applications, making it especially suitable for resource poor settings.

Conventional optical microscopes image cells by use of objective lenses that work together with other lenses and optical components. While quite ...

A High Throughput Screen for Biomining Cellulase Activity from Metagenomic Libraries

Cellulose, the most abundant source of organic carbon on the planet, has wide-ranging industrial applications with increasing emphasis on biofuel production 1. Chemical methods to modify or degrade cellulose typically require strong acids and high temperatures. As such, enzymatic methods have become prominent in the bioconversion process. While the identification of active cellulases from bacterial and fungal isolates has been somewhat effective, the vast majority of microbes in nature resist laboratory cultivation. Environmental genomic, also known as metagenomic, screening approaches have great promise in bridging the cultivation gap in the search for novel bioconversion enzymes. Metagenomic screening approaches have successfully recovered novel cellulases from environments as varied as soils 2, buffalo rumen 3 and the termite hind-gut 4 using carboxymethylcellulose (CMC) agar plates stained with congo red dye (based on the method of Teather and Wood 5). However, the CMC method is limited in throughput, is not quantitative and manifests a low signal to noise ratio 6. Other methods have been reported 7,8 but each use an agar plate-based assay, which is undesirable for high-throughput screening of large insert genomic libraries. Here we present a solution-based screen for cellulase activity using a chromogenic dinitrophenol (DNP)-cellobioside substrate 9. Our library was cloned into the pCC1 copy control fosmid to increase assay sensitivity through copy number induction 10. The method uses one-pot chemistry in 384-well microplates with the final readout provided as an absorbance measurement. This readout is quantitative, sensitive and automated with a throughput of up to 100X 384-well plates per day using a liquid handler and plate reader with attached stacking system.

This protocol describes a high throughput screen for cellulolytic activity from a metagenomic library expressed in Escherichia coli. The screen is solution based and highly automated, and uses one-pot chemistry in 384 well microplates with the final readout as an absorbance measurement.

Cellulose, the most abundant source of organic carbon on the planet, has wide-ranging industrial applications with increasing emphasis on biofuel ...

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

The nematode C. elegans has emerged as an important model for the study of conserved genetic pathways regulating fat metabolism as it relates to human obesity and its associated pathologies. Several previous methodologies developed for the visualization of C. elegans triglyceride-rich fat stores have proven to be erroneous, highlighting cellular compartments other than lipid droplets. Other methods require specialized equipment, are time-consuming, or yield inconsistent results. We introduce a rapid, reproducible, fixative-based Nile red staining method for the accurate and rapid detection of neutral lipid droplets in C. elegans. A short fixation step in 40% isopropanol makes animals completely permeable to Nile red, which is then used to stain animals. Spectral properties of this lipophilic dye allow it to strongly and selectively fluoresce in the yellow-green spectrum only when in a lipid-rich environment, but not in more polar environments. Thus, lipid droplets can be visualized on a fluorescent microscope equipped with simple GFP imaging capability after only a brief Nile red staining step in isopropanol. The speed, affordability, and reproducibility of this protocol make it ideally suited for high throughput screens. We also demonstrate a paired method for the biochemical determination of triglycerides and phospholipids using gas chromatography mass-spectrometry. This more rigorous protocol should be used as confirmation of results obtained from the Nile red microscopic lipid determination. We anticipate that these techniques will become new standards in the field of C. elegans metabolic research.

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

We present robust biochemical and microscopic methods for studying Caenorhabditis elegans lipid stores. A rapid, simple, fixing-staining procedure for fluorescent lipid droplet imaging leverages the spectral properties of the lipophilic dye Nile red. We then present biochemical measurement of triglycerides and phospholipids using solid phase extraction and gas chromatography-mass spectrometry.

The nematode C. elegans has emerged as an important  model for the study of conserved genetic pathways regulating fat metabolism as  it relates to human ...

High Throughput Microinjections of Sea Urchin Zygotes

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.

Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of ...

High-throughput Image Analysis of Tumor Spheroids: A User-friendly Software Application to Measure the Size of Spheroids Automatically and Accurately

The increasing number of applications of three-dimensional (3D) tumor spheroids as an in vitro model for drug discovery requires their adaptation to large-scale screening formats in every step of a drug screen, including large-scale image analysis. Currently there is no ready-to-use and free image analysis software to meet this large-scale format. Most existing methods involve manually drawing the length and width of the imaged 3D spheroids, which is a tedious and time-consuming process. This study presents a high-throughput image analysis software application &#8211; SpheroidSizer, which measures the major and minor axial length of the imaged 3D tumor spheroids automatically and accurately; calculates the volume of each individual 3D tumor spheroid; then outputs the results in two different forms in spreadsheets for easy manipulations in the subsequent data analysis. The main advantage of this software is its powerful image analysis application that is adapted for large numbers of images. It provides high-throughput computation and quality-control workflow. The estimated time to process 1,000 images is about 15 min&#160;on a minimally configured laptop, or around 1 min&#160;on a multi-core performance workstation. The graphical user interface (GUI) is also designed for easy quality control, and users can manually override the computer results. The key method used in this software is adapted from the active contour algorithm, also known as Snakes, which is especially suitable for images with uneven illumination and noisy background that often plagues automated imaging processing in high-throughput screens. The complimentary &#8220;Manual Initialize&#8221; and &#8220;Hand Draw&#8221; tools provide the flexibility to SpheroidSizer in dealing with various types of spheroids and diverse quality images. This high-throughput image analysis software remarkably reduces labor and speeds up the analysis process. Implementing this software is beneficial for 3D tumor spheroids to become a routine in vitro model for drug screens in industry and academia.

We present a high-throughput image analysis software application to measure the size of three-dimensional tumor spheroids imaged with bright-field microscopy. This application provides a fast and effective way to examine the effects of therapeutic drugs on spheroids, which is beneficial for researchers who wish to use spheroids in drug screens.

The increasing number of applications of three-dimensional (3D) tumor spheroids as an in vitro model for drug discovery requires their adaptation to ...

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...

High-throughput Screening for Chemical Modulators of Post-transcriptionally Regulated Genes

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays focus on transcriptional regulation, being essentially aimed at the identification of promoter-targeting molecules. As a result, post-transcriptional mechanisms are largely uncovered by gene expression targeted drug development. Here we describe a cell-based assay aimed at investigating the role of the 3&#39; untranslated region (3&#8217; UTR) in the modulation of the fate of its mRNA, and at identifying compounds able to modify it. The assay is based on the use of a luciferase reporter construct containing the 3&#8217; UTR of a gene of interest stably integrated into a disease-relevant cell line. The protocol is divided into two parts, with the initial focus on the primary screening aimed at the identification of molecules affecting luciferase activity after 24 hr of treatment. The second part of the protocol describes the counter-screening necessary to discriminate compounds modulating luciferase activity specifically through the 3&#8217; UTR. In addition to the detailed protocol and representative results, we provide important considerations about the assay development and the validation of the hit(s) on the endogenous target. The described cell-based reporter gene assay will allow scientists to identify molecules modulating protein levels via post-transcriptional mechanisms dependent on a 3&#8217; UTR.

Here we describe a cell-based reporter gene assay as a valuable tool to screen chemical libraries for compounds modulating post-transcriptional control mechanisms exerted through 3&#8217; UTR.

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays ...

qPCRTag Analysis - A High Throughput, Real Time PCR Assay for Sc2.0 Genotyping

The Synthetic Yeast Genome Project (Sc2.0) aims to build 16 designer yeast chromosomes and combine them into a single yeast cell. To date one synthetic chromosome, synIII1, and one synthetic chromosome arm, synIXR2, have been constructed and their in vivo function validated in the absence of the corresponding wild type chromosomes. An important design feature of Sc2.0 chromosomes is the introduction of PCRTags, which are short, re-coded sequences within open reading frames (ORFs) that enable differentiation of synthetic chromosomes from their wild type counterparts. PCRTag primers anneal selectively to either synthetic or wild type chromosomes and the presence/absence of each type of DNA can be tested using a simple PCR assay. The standard readout of the PCRTag assay is to assess presence/absence of amplicons by agarose gel electrophoresis. However, with an average PCRTag amplicon density of one per 1.5 kb and a genome size of ~12 Mb, the completed Sc2.0 genome will encode roughly 8,000 PCRTags. To improve throughput, we have developed a real time PCR-based detection assay for PCRTag genotyping that we call qPCRTag analysis. The workflow specifies 500 nl reactions in a 1,536 multiwell plate, allowing us to test up to 768 PCRTags with both synthetic and wild type primer pairs in a single experiment.

Designer chromosomes of the Synthetic Yeast Genome project, Sc2.0, can be distinguished from their native counterparts using a PCR-based genotyping assay called PCRTagging, which has a presence/absence endpoint. Here we describe a high-throughput real time PCR detection method for PCRTag genotyping.

The Synthetic Yeast Genome Project (Sc2.0) aims to build 16 designer yeast chromosomes and combine them into a single yeast cell. To date one synthetic ...

Automated Modular High Throughput Exopolysaccharide Screening Platform Coupled with Highly Sensitive Carbohydrate Fingerprint Analysis

Many microorganisms are capable of producing and secreting exopolysaccharides (EPS), which have important implications in medical fields, food applications or in the replacement of petro-based chemicals. We describe an analytical platform to be automated on a liquid handling system that allows the fast and reliable analysis of the type and the amount of EPS produced by microorganisms. It enables the user to identify novel natural microbial exopolysaccharide producers and to analyze the carbohydrate fingerprint of the corresponding polymers within one day in high-throughput (HT). Using this platform, strain collections as well as libraries of strain variants that might be obtained in engineering approaches can be screened. The platform has a modular setup, which allows a separation of the protocol into two major parts. First, there is an automated screening system, which combines different polysaccharide detection modules: a semi-quantitative analysis of viscosity formation via a centrifugation step, an analysis of polymer formation via alcohol precipitation and the determination of the total carbohydrate content via a phenol-sulfuric-acid transformation. Here, it is possible to screen up to 384 strains per run. The second part provides a detailed monosaccharide analysis for all the selected EPS producers identified in the first part by combining two essential modules: the analysis of the complete monomer composition via ultra-high performance liquid chromatography coupled with ultra violet and electrospray ionization ion trap detection (UHPLC-UV-ESI-MS) and the determination of pyruvate as a polymer substituent (presence of pyruvate ketal) via enzymatic oxidation that is coupled to a color formation. All the analytical modules of this screening platform can be combined in different ways and adjusted to individual requirements. Additionally, they can all be handled manually or performed with a liquid handling system. Thereby, the screening platform enables a huge flexibility in order to identify various EPS.

We present an automated modular high-throughput-method for the identification and characterization of microbial exopolysaccharides in small scale. This method combines a fast preselection to analyze the total amount of secreted polysaccharides with a detailed carbohydrate fingerprint to enable the fast screening of newly isolated bacterial strains or entire strain collections.