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Summary

Organoids have become valuable tools for disease modeling. The extracellular matrix (ECM) guides cell fate during organoid generation, and using a system that resembles the native tissue can improve model accuracy. This study compares the generation of induced pluripotent stem cells-derived human intestinal organoids in animal-derived ECM and xeno-free hydrogels.

Abstract

Extracellular matrix (ECM) plays a critical role in cell behavior and development. Organoids generated from human induced pluripotent stem cells (hiPSCs) are in the spotlight of many research areas. However, the lack of physiological cues in classical cell culture materials hinders efficient iPSC differentiation. Incorporating commercially available ECM into stem cell culture provides physical and chemical cues beneficial for cell maintenance. Animal-derived commercially available basement membrane products are composed of ECM proteins and growth factors that support cell maintenance. Since the ECM holds tissue-specific properties that can modulate cell fate, xeno-free matrices are used to stream up translation to clinical studies. While commercially available matrices are widely used in hiPSC and organoid work, the equivalency of these matrices has not been evaluated yet. Here, a comparative study of hiPSC maintenance and human intestinal organoids (hIO) generation in four different matrices: Matrigel (Matrix 1-AB), Geltrex (Matrix 2-AB), Cultrex (Matrix 3-AB), and VitroGel (Matrix 4-XF) was conducted. Although the colonies lacked a perfectly round shape, there was minimal spontaneous differentiation, with over 85% of the cells expressing the stem cell marker SSEA-4. Matrix 4-XF led to the formation of 3D round clumps. Also, increasing the concentration of supplement and growth factors in the media used to make the Matrix 4-XF hydrogel solution improved hiPSC expression of SSEA-4 by 1.3-fold. Differentiation of Matrix 2-AB -maintained hiPSC led to fewer spheroid releases during the mid-/hindgut stage compared to the other animal-derived basement membranes. Compared to others, the xeno-free organoid matrix (Matrix 4-O3) leads to larger and more mature hIO, suggesting that the physical properties of xeno-free hydrogels can be harnessed to optimize organoid generation. Altogether, the results suggest that variations in the composition of different matrices affect stages of IO differentiation. This study raises awareness about the differences in commercially available matrices and provides a guide for matrix optimization during iPSC and IO work.

Introduction

The extracellular matrix (ECM) is a dynamic and multifunctional component of tissues that plays a central role in regulating cell behavior and development. As a complex network, it provides structural support, cell adhesive ligands¹, and storage of growth factors and cytokines that regulate cell signaling. For example, during wound healing, the ECM serves as a scaffold for migrating cells and as a reservoir of growth factors involved in tissue repair². Similarly, dysregulation in the ECM can lead to an increase in the severity of various diseases such as fibrosis and cancer^3,4. During embryonic development, the ECM guides tissue morphogenesis. For example, in the development of the heart, ECM components play a role in creating the correct architecture and function of the heart tissue⁵. Over a decade of research has shown that the stiffness of the microenvironment alone^6,7 can control stem cell lineage specification. Therefore, it is not surprising that during in vitro cell differentiation, ECM influences stem cell fate by providing signals for differentiation.

Organoids can be generated from induced pluripotent stem cells (iPSCs). Starting with a properly characterized iPSC line is required to generate organoids successfully. However, the lack of physiological cues in classical cell culture materials hinders efficient iPSC differentiation and organoid generation. Moreover, recent research has emphasized the significance of the composition of the extracellular matrix (ECM), interactions between cells and the ECM⁸, as well as mechanical and geometrical cues^9,10,11 in the context of organoid expansion and differentiation¹². Advancing organoid technology by improving reproducibility will involve incorporating tissue-specific physical and chemical cues.

Organoids aim to recapitulate the native tissue within a physiologically similar microenvironment. Choosing an ECM system that closely mimics the native tissue ECM is crucial for achieving physiological relevance regarding cell behavior, function, and response to stimuli¹³. The choice of ECM components can influence the differentiation of stem cells into specific cell types within the organoid. Different ECM proteins and their combinations can provide cues that guide cell fate¹⁴. For example, studies have shown that using specific ECM components can promote the differentiation of intestinal stem cells into mature intestinal cell types, resulting in physiologically relevant intestinal organoids¹⁵. While organoids are a valuable tool during disease modeling and drug testing, selecting an appropriate ECM system is pivotal to this application. An appropriate ECM system can enhance the accuracy of disease modeling by creating a microenvironment that resembles the affected tissue¹⁶. Furthermore, tissue-specific ECM can help generate organoids that better recapitulate disease-associated phenotypes and drug responses¹⁷. Optimizing the ECM system used in organoid differentiation is critical for achieving desired differentiation outcomes.

Commercially available basement membrane systems derived from animal ECM sources (e.g., Matrigel, Cultrex) and xeno-free hydrogel (e.g., VitroGel) are widely used in iPSC and organoid research. Companies that commercialize them and researchers that use them have laid out many instructions for their specific products and applications over the years. Many of these instructions served as a guide for the generation of this protocol. Furthermore, the benefits and setbacks associated with their intrinsic properties have been individually noted by many^18,19,20,21. However, there is no systematic workflow to guide the selection of optimal systems for iPSC and organoid work. Here, a workflow to systematically evaluate the equivalency of ECM systems from various sources for iPSC and organoid work is provided. This is a comparative study of the maintenance of two different human iPSC lines (hiPSC) and human intestinal organoids (hIO) generation in four different matrices: Matrigel (Matrix 1-AB), Geltrex (Matrix 2-AB), Cultrex (Matrix 3-AB), and VitroGel (Matrix 4-XF). For organoid culture, four versions of the xeno-free matrix VitroGel that were previously optimized for organoid culture were used: ORGANOID 1 (Matrix 4-O1), ORGANOID 2 (Matrix 4-O2), ORGANOID 3 (Matrix 4-O3), ORGANOID 4 (Matrix 4-O4). Also, animal-derived matrices optimized for organoids were used: Matrigel High Concentration (Matrix 1-ABO) and Cultrex Type 2 (Matrix 3-ABO). Commercially available stem cell culture media (mTeSR Plus) and organoid differentiation kit (STEMdiff intestinal organoid kit) were used. This protocol combines the individual instructions from the products' manufacturers with lab experiences to guide the reader toward a successful optimization of ECM for their specific iPSC and organoid work. Altogether, this protocol and representative results emphasize the importance of selecting the optimal microenvironment for stem cell work and organoid differentiation.

Protocol

1. hiPSC maintenance

CAUTION: All work is done in a Biosafety Cabinet (BSC) following standard aseptic techniques. Must follow OSHA safety standards for laboratories, including proper use of personal protective equipment such as lab coats, gloves, and goggles.

1. Preparation of matrices aliquots and cell culture media
   1. For commercially available animal-derived basement membranes (BMs; Matrix 1-AB, Matrix 2-AB, Matrix 3-AB), prepare aliquots of working volumes following the manufacturer's recommendation summarized in Table 1 and store them at -20 °C or -80 °C for long term storage. Avoid the formation of bubbles. If some bubbles are formed, before storing, centrifuge samples at 4 °C, at <200 x g, for ~1-2 min to force the bubble to come to the surface.
      NOTE: Making single-use aliquots helps avoid repeated freeze-thaw cycles that disrupt ECM architecture. Because BM concentration varies with lot number, make sure to follow the manufacturer's recommendation to prepare single-use aliquots and make coating solutions. High-concentrated BMs are viscous and difficult to pipette; use cold tips previously stored at -20 °C.
   2. Prepare stem cell culture medium following manufacturer's recommendation. To prepare the specific commercially available complete media used in this protocol, add 100 mL of 5x supplement cocktail to 400 mL of basal medium. Then, mix this media thoroughly and aliquot into 40 mL volumes, stored at 20 °C. For use, thaw each aliquot of complete media, use the aliquoted complete medium immediately, or store at 2 – 8 °C for up to 2 weeks. Do not re-freeze it.
   3. Prepare aliquots of the stem cell complete medium containing 3x the concentration of the supplement cocktail to prepare the Matrix 4-XE Hydrogel, i.e., dilute 5x supplement to 3x.
2. Coating tissue culture plastic with animal-derived BMs (Matrix 1-3AB)
   1. Prepare the following before starting: always keep the matrices in ice when thawing and handling to prevent them from gelling. Use cold media to prepare the diluted matrices. Prepare enough replicate wells per condition to assay cells before starting and at every stage of the hiPSC differentiation process and hIO organoid generation.
      NOTE: Here, 24-well plates were used for this study. Refer to Table 2 for recommended coating volumes for other plate sizes.
   2. For each type of matrix, prepare 25 mL of cold advance DMEM/F-12 containing 15 mM HEPES in a 50 mL conical tube. Keep these on ice.
   3. Add the thawed matrices to their respective cold advance DMEM/F-12 and mix well. Keep the media cold during the mixing process. Visually check for homogenous mixing by ensuring there are no clumps.
   4. Immediately use the diluted matrices solutions to coat each well selected for use (250 µL/ well if using 24 well plates).
   5. Gently tilt the culture ware to spread the coating solution evenly across the surface. Incubate at room temperature (15 – 25 °C) for at least 1 h before use.
      NOTE: If not used immediately, the plate can be stored at 2 – 8 °C for up to 1 week after coating, but it must be sealed with a transparent film to prevent evaporation. When using stored plates, let them reach room temperature (15 – 25 °C) for 30 min before proceeding to the next step.
   6. Gently remove the excess solution using a serological pipette or by aspiration. Ensure that the coated surface is not scratched.
   7. Add warm complete stem cell medium immediately (50% of total volume needed for specific well, e.g., 250 µL/well if using a 24-well plate).
3. Coating tissue culture plastic with Matrix 4-XE hydrogel
   1. Prepare the following before starting: take the Matrix 4-XE hydrogel out of the refrigerator and let it warm to room temperature (25 °C). Prepare enough replicate wells per condition to assay cells before starting and at every stage of the hiPSC differentiation process and hIO organoid generation.
      NOTE: Here, 24-well plates were used for this study. Refer to Table 3 for recommended coating volumes for other plate sizes.
   2. Prepare a complete stem cell culture media containing 3x growth factor concentration compared to the standard formulation for stem cell culture (3x stem cell medium).
   3. Mix Matrix 4-XE hydrogel and 3x stem cell medium at a 2:1 v/v mixing ratio and gently pipette up and down 5x-10x to mix thoroughly.
   4. Transfer the hydrogel mixture to a well plate and carefully tilt the culture ware to spread the mixture evenly across the surface. Use 250 µL/well if using a 24-well plate. See Table 3 for recommended volumes.
   5. Wait 10-15 min at room temperature for soft gel formation. During the hydrogel-formation process, do not disrupt the hydrogel by tilting or shaking the plate.
4. hiPSC enzyme-free clump passaging and seeding
   1. Prepare the following before starting: at least 1 h before passaging, coat desired plasticware with matrices. Aliquot sufficient complete stem cell media and warm to room temperature (15 – 25 °C). Avoid multiple heating cycles for the complete media.
      NOTE: The steps below describe the passage of an already established and >90% confluent culture of iPSCs on a 6-well plate and seeding them in 24-well plates for differentiation of intestinal organoids.
   2. Rinse cells with 1 mL of D-PBS (without Ca++ and Mg++) and aspirate. Add 1 mL of enzyme-free human pluripotent stem cell selection reagent, carefully swirl to spread evenly, and aspirate within 1 min. Colonies only need to be exposed to a thin film of liquid.
   3. Incubate at 37 °C until colonies start looking less compacted, which will take ~ 3-8 min.
      NOTE: Optimal incubation time may vary depending on the cell line used. The optimal incubation time must be determined when passaging each cell line with the enzyme-free human pluripotent stem cell selection reagent for the first time.
   4. Add 1 mL of complete stem cell media. Detach the colonies by carefully tapping firmly on the side of the plate. Make sure to hold the plate with the other hand.
   5. With a 1 mL or larger pipette, transfer the cell clumps suspension to a 15 mL conical tube. Evaluate the size of the clumps using a brightfield microscope and make sure the cell clump's size is between 50-200 µm; if they are larger, gently shake the conical tube to break them down.
   6. Draw an x-y grid at the bottom of each well of a flat-bottom 96-well plate that will be used for clump counting and add 50 µL of D-PBS to each of the wells. It is recommended to average the clumps counted from 3 wells.
   7. Ensure cell clumps are evenly distributed by gently shaking the tube and then transfer 5 µL of clump suspension into each of the wells of the flat-bottom 96-well plate.
      NOTE: It is not recommended to use an automatic cell counter to count clumps. A manual counting is recommended.
   8. Count the total number of clumps in each well that are 50 – 200 µm in diameter. If most cell clumps are > 200 µm in diameter (see Figure 1), repeat steps 1.4.5 – 1.4.7.
   9. Calculate the volume (in µL) of clump suspension required to seed 6,000 clumps as follows:
      
      NOTE: Optimal seeding density must be optimized with every cell line. To determine optimal seeding density, it is recommended to perform an initial seeding in a range of clump densities (e.g., 4,000, 5,000, and 6,000 clumps per well) and check 24-48 h after seeding if 85%-90% confluency is reached.
   10. As a quality control, separate a sample of cells for flow cytometry examination of stem cell markers. Refer to Table 4 for a list of common markers.
   11. Separate the required volume of iPSC clump suspension to seed on the wells coated with different ECM into separate 15 mL tubes.
   12. Centrifuge the tube containing the iPSC clump solution at 200 x g for 5 min to remove media containing enzyme -free human pluripotent stem cell selection reagent. While waiting for centrifugation, add 50% of the desired volume per well to each well if it was not done already.
5. iPSC culture in plates containing animal-derived matrices
   1. Gently resuspend iPSCs clumps in the 50% calculated volume of complete stem cell media to get the desired density and plate the cell aggregate mixture onto coated wells containing 50% of calculated complete stem cell media.
      NOTE: Adding ROCK-1 inhibitor might be necessary for some cell lines or when clumps are 50 µm or smaller. It is recommended to add <10 µm to avoid early gastrulation.
   2. Carefully add medium with cells on top of the matrices, 250 µL/well if using a 24-well plate. For the recommended volume of cell medium for other size well plates, check Table 2.
   3. Tilt the plate in several short back-and-forth and side-to-side motions to distribute the cell clumps evenly.
      CAUTION: Uneven distribution of aggregates results in increased spontaneous differentiation of human iPSCs. Circular spreading of the clumps' suspension results in clumps agglomerating on the border of the wells and lower density at the center.
   4. Incubate the plate at 37 °C and perform medium changes using complete stem cell media daily or every other day. When performing medium changes, visually assess cultures to monitor growth and determine whether the cells require passaging time or are ready for differentiation. To skip two consecutive days of feeding, add 2x the volume of the medium needed for a single day.
6. iPSC culture in plates containing Matrix 4-XE
   1. Gently resuspend iPSCs clumps into the total calculated volume of 3x complete stem cell media to get the desired density.
   2. Carefully add medium with cells on top of the hydrogel 250 µL/well if using a 24-well plate. For the recommended volume of cell medium for other size well plates, check Table 3.
      CAUTION: The hydrogel will swell and occupy a larger volume than freshly made. iPSC colonies will be partially embedded in the hydrogel at the bottom of the plate, so it is recommended to change 50%-80% of the top medium without disturbing the hydrogel.
   3. Move the plate in several quick, short, back-and-forth, and side-to-side motions to distribute the cell clumps evenly.
   4. Place the plate in a 37 °C incubator with 5% CO₂ and 95% humidity. Change the cell medium using complete stem cell media. Perform medium changes daily or every other day; for the latter one, add 2x the volume of the medium.
   5. Visually inspect the cultures to track their growth as they reach the stage suitable for differentiation.

Figure 1: Optimal clump size. Images of clumps of iPSC cell line SCTi003A depicting an example of optimal clump size. Scale bar = 200 µm. Please click here to view a larger version of this figure.

2. hiPSC differentiation and intestinal organoid generation

CAUTION: All work is done in a Biosafety Cabinet (BSC) following standard aseptic techniques. Must follow OSHA safety standards for laboratories, including proper use of personal protective equipment such as lab coats, gloves, and goggles.

1. Prepare the following before starting: prepare aliquots of differentiation medium required for each stage. After thawing the aliquots, use them immediately. Do not re-freeze. Follow Table 5 for a general guide of each stage media type and volumes needed for a 24-well plate.
   NOTE: Optimal time at each differentiation stage might vary per cell line. Checking the expression of each stage cell marker is recommended to improve differentiation efficacy. Periodical assessment of gene expression on the iPSCS cultures was not included in this research since we leveraged vendor-qualified batches at low passages, but it is recommended as a quality control step for new/not yet qualified iPSC lines during subsequent differentiation steps.
2. Stage 1: Definitive endoderm (DE)
   1. To assess readiness to start differentiation, check for the following criteria.
      1. Using a microscope, evaluate the confluency of the stem cell colonies and the amount of spontaneous differentiation in the culture. Do not rely only on morphological evaluation alone. Optimal confluency should be between 85%-90% (above ~10⁵ cells/cm²), and minimal spontaneous differentiation observed should be <5% differentiation.
      2. Collect cells from a sacrificial well for flow cytometry characterization of hiPSC markers before proceeding with the next step. A forward and side scatter gating strategy was used for the representative flow cytometry characterization. If cells meet the optimal criteria, start differentiation. Cell passing criteria is >85% expression of ideally 3 markers (Table 4), especially in case of new/not yet qualified iPSC lines in the lab.
         NOTE: iPSC cultures grown on different matrices undergo the same overall process with only iPSC growth on Matrix 4-XE, requiring more gentle media changes not to disturb the semi-embedded cells. Significant cell death is expected during the induction of definitive endoderm, as cells are particularly sensitive at this stage. Exercise caution during medium changes, aiming to minimize the duration that cells are exposed outside the 37 °C incubation environment as much as possible.
   2. Start differentiation of cell colonies as described below.
      1. Day 0: Warm an aliquot of DE medium to 37 °C. To avoid denaturing of media from warming-cooling cycles, ensure to warm only the volume required for Day 0 (0.7 mL/well).
      2. Aspirate the media from hiPSCs. Carefully add 0.7 mL of DE Medium per well down the side of the well. Avoid harsh pipetting that could detach or damage the colonies. Incubate the plate at 37 °C with 5% CO₂ and 95% humidity for 24 h.
      3. Day 1: Warm an aliquot of DE medium containing only the volume needed for Day 1 (0.5 mL/well) to 37 °C. Aspirate the DE medium from cells and carefully add 0.5 mL of DE medium per well down the side of the well. Incubate at 37 °C with 5% CO₂ and 95% humidity for 24 h.
      4. Day 3: Cells are ready to be assayed for definitive endoderm formation. Refer to Table 4 for recommended markers. Before proceeding with the next phase, sacrifice a well to perform a flow cytometry characterization of specific markers associated with DE. A forward and side scatter gating strategy was used for the representative flow cytometry characterization. It is recommended that the expression of more than 1 marker be checked, especially in the case of new/not yet qualified iPSC lines in the lab.
3. Stage 2: Midgut/Hindgut (MH)
   1. Day 3: Warm only the volume of MH medium needed for day 3 (0.5 mL/well) to 15 – 25 °C. Aspirate DE medium from cells and replace with 0.5 mL of MH Medium. Incubate at 37 °C with 5% CO₂ and 95% humidity for 24 h.
   2. Days 4 – 9: Before daily medium changes, use the microscope at 2x or 4x magnification to assess spheroids formation by checking for visible 3D structures (might happen from day 4); free-floating spheroids (might occur from day 5).
   3. Once spheroids start detaching, use a 1 mL pipettor to gently transfer the medium from cells to a sterile 15 mL conical tube to assess the number of spheroids detached from cells.
      NOTE: Avoid any shear force that might detach 3D structures from the monolayer; once ready, the spheroids will detach on their own.
   4. If less than 50 spheroids are detached, centrifuge at 200 x g for 5 min, remove the old medium, and resuspend spheroids on 0.5 mL/well of medium and transfer to respective well until enough spheroids have matured.
      NOTE: The duration of exposure to the midgut/hindgut medium plays a crucial role in determining the regional identity of developing small intestinal organoids, such as duodenum (shorter exposure) or ileum (longer exposure). To achieve more uniform cultures with the same identity, aim to collect spheroids exposed to midgut/hindgut medium simultaneously.
   5. Repeat step 2.3.3 until enough spheroids are detached to embed and initiate human intestinal organoid (hIO) culture.
   6. Before proceeding with the next phase, separate a sample of spheroids and perform a flow cytometry characterization of specific markers associated with Midgut/Hindgut (MH). A forward and side scatter gating strategy was used for the representative flow cytometry characterization. Refer to Table 4 for a list of commonly used markers. It is recommended that the expression of more than 1 marker be checked, especially in the case of new/not yet qualified iPSC lines in the lab.
4. Stage 3: Spheroid embedding
   1. Prepare the following before starting the procedure.
      1. Thaw aliquots of Matrix 1-ABO and Matrix 3-ABO on ice. Consider the number of spheroids collected to determine the total amount of matrix needed if 30-40 µL of matrix will be required per dome. 
      2. Prepare 25 mL of cold Advance DMEM/F-12 containing 15 mM HEPES. High-concentrated ECMs are viscous and difficult to pipette. Place a box of sterile 100 µL pipette tips at -20 °C for preparing cold tips that can help the process. Prepare aliquots of Intestinal Organoid Growth Medium (OGM), which requires 4 feeds (0.5 mL/well per feed). See Table 5 for preparation.
      3. Thaw an aliquot of the OGM supplement on ice. After thawing the aliquots, use them immediately. Do not re-freeze. Store complete OGM at 2-8 °C for up to 2 weeks.
      4. Prepare OGM containing 3x of the supplement compared to the standard formulation for (3x OGM). Take the Matrix 4-XFO1 thought Matrix 4-XFO4 out of the refrigerator and let them warm to room temperature (25 °C).
   2. For the animal-derived system, perform the steps described below.
      1. Place a sterile 24-well tissue culture plate in the incubator to warm to 37 °C while preparing the spheroids and matrices. Let the collected spheroids settle to the bottom of a 15 mL conical tube. Carefully aspirate and discard the supernatant.
         NOTE: To make a more accurate comparison of the effect of the matrices, make sure to keep the spheroids collected from the hiPSC differentiated on each matrix in separate tubes.
      2. Add 1 mL of DMEM/F-12 with 15 mM HEPES to the spheroids. Centrifuge at 300 x g for 5 min at room temperature (15 – 25 °C).
      3. Carefully remove as much of the supernatant as possible. It is recommended to start removing with a 1 mL pipettor and switch to 100 µL and even 10 µL pipettes to remove as much as possible without disturbing the pellet of spheroids.
         NOTE: The larger the media volume left, the more diluted the matrices will be. Diluted matrices can lead to gelling issues or softer domes. Softer domes lead to higher chances of collapsing with minimal disturbances.
      4. Using a pipettor with a cold 100 µL pipette tip, add 40 µL/50 spheroids of cold (2 – 8 °C) Matrix 1-ABO or Matrix 3-ABO to their respective tube. Gently distribute spheroids into the matrix by pipetting up and down ~5x. Do not completely empty the pipette tip, which may introduce bubbles.
      5. Bring the plate from the incubator, and using a cold pipette tip, gently transfer embedded spheroids into the center of the plate following steps in Figure 2. Avoid dispensing Matrix 1-ABO and Matrix 3-ABO too quickly into the culture dish, as this will flatten the dome.
      6. Carefully transfer the plate into a 37 °C incubator and incubate for 30 min to ensure dome gelation. Be especially careful of any shaking or harsh movement during transport and in the incubator.
      7. While incubating the dome for gelling, warm at 37 °C a sufficient volume of intestinal OGM (0.5 mL/well) for the number of wells to be used.
         NOTE: Warm only the volume needed. Avoid warming-cooling cycles. Make sure the media is warm before adding it to the dome; cold media can lead to the dome collapsing.
      8. After 30 min, carefully add 0.5 mL of intestinal OGM to the side of the well to not disturb the dome. Incubate at 37 °C with 5% CO₂ and 95% humidity. Perform a full-medium change every 3 – 4 days by removing the medium and then add fresh medium.
   3. For the Matrix 4-XF organoid systems follow the steps described.
      NOTE: Matrix 4-XF organoid system includes four different options (O1-O4); the formulation of each type varies in bio-functional ligands, stiffness, and degradability. Consequently, performing initial experiments to determine the optimal type for the specific application is recommended. This protocol describes using the four options to find an optimal formulation for hiPSC-derived intestinal organoids. Additionally, while there are multiple culture protocols that can be used (e.g., 3D encapsulation, dome encapsulation), this protocol involves using the dome method such that it is possible to compare directly with the common methods used when working with animal-derived basement membrane systems.
      1. Add 1 mL/well of cold advance DMEM/F-12 containing 16 mM HEPES to the spheroids generated on the Matrix 4-XFO4 system. Centrifuge at 300 x g for 5 min at room temperature (15 – 25 °C).
         NOTE: To find the optimal formulation for the application, make sure to distribute spheroids into 4 separate tubes before centrifugation. Spheroids generated on animal-derived matrices can also be matured into intestinal organoids using this system.
      2. Using a 1 mL pipettor, remove the supernatant without disturbing the spheroids. Add 50 µL of 3x OGM to the spheroids pellet (~100 spheroids). Add 100 µL of the selected Matrix 4-XFO to the 50 µL spheroid suspension and gently mix 5x-10x times. Keep the Matrix 4-XFO to 3x OGM at a 2:1 v/v mixing ratio to have a final concentration of 1x.
      3. Add 40 µL of the hydrogel-spheroid mixture to the center of a 24-well tissue culture plate. Carefully transfer the plates into a 37 °C incubator and incubate for 30 min.
         NOTE: Matrix 4-XFO does not require 37 °C incubation for gelation; however, it is recommended to keep organoids exposed to similar conditions when compared to Matrix 1-ABO and Matrix 3-ABO systems that require 37 °C for better gelation of the domes.
      4. After 30 min, carefully add 0.5 mL of intestinal OGM to the side of the well so as not to disturb the dome. Incubate at 37 °C with 5% CO₂ and 95% humidity. Perform a full-medium change every 3 – 4 days by removing the medium and then adding fresh medium.
5. hIO passaging and maturation
   1. Prepare the same chemicals, solutions, and reagents as done in step 2.4.1.
   2. Add 1-2 mL of anti-adherence rinsing solution to a 15 mL conical tube (1 per condition) and swirl to coat the tube.
   3. Remove the anti-adherence solution and rinse the tubes with 5 mL of D-PBS (without Ca++ and Mg++). Cap all coated tubes and keep them at room temperature (15 – 25 °C) until needed.
   4. Aspirate the medium from the domes. Using a 1 mL pipette, add 1 mL of cold DMEM/F-12 to the dome directly. The goal is to detach the domes from the plate.
   5. Add an additional 1 mL of cold DMEM/F-12 to the well and pipette up and down to harvest any remaining organoids. Transfer to the coated 15 mL conical tubes.
      NOTE: Verify the successful harvest of organoids by visually examining the well under a microscope. If any residual organoids are observed, repeat step 2.5.5.
   6. With a 1 mL pipettor, perform up-and-down pipetting of the suspension to disintegrate organoids until a uniform fragment suspension is achieved with the desired organoid size (e.g., 100 – 500 µm).
      NOTE: Utilize a 200 µL pipettor to verify that the organoids conform to the recommended size. Employing the 200 µL pipettor facilitates additional fragmentation as needed, ensuring that the fragments can smoothly pass through a 200 µL pipette tip.
      CAUTION: Avoid breaking up fragments into single cells by harsh or prolonged pipetting.
   7. Establish the desired organoid density by either counting fragments or employing the split ratio. Separate an aliquot and perform counting using the same procedure as in steps 1.4.6-1.4.9 for clump counting.
      NOTE: Optimal organoid density should be optimized per line; generally, a density of 40 – 80 intestinal organoids per dome is recommended.
   8. Ensure that the tube is placed on ice during the organoid counting process. After approximately 5 min, organoid fragments will have settled to the bottom of the tube due to gravity.
      NOTE: The larger the volume of the organoid solution, the longer it will take to settle to the bottom.
   9. Gently eliminate as much supernatant and the cloudy layer formed on top of the organoids. In the early passages when organoids are maturing, this cloudy phase encompasses a matrix and individual cells.
   10. Add 2 mL of cold DMEM/F-12 by directly pipetting onto the pellet. Centrifuge at 200 x g for 5 min at room temperature (15 – 25 °C).
   11. For animal-derived system, carefully remove and discard supernatant following the same steps described in steps 2.4.2.4-2.4.2.9.
   12. For the Matrix 4-XFO system, carefully remove and discard the supernatant following the same steps described in steps 2.4.3.2-2.4.3.4.
       NOTE: When working with Matrix 4-XFO system, it is recommended to use a xeno-free hydrogel organoid recovery solution for optimal removal of hydrogel residual. This recovery solution is especially recommended when switching from animal-based to xeno-free hydrogel systems to ensure the elimination of any xenogeneic material.

Figure 2. Schematic of technique recommended for dome formation. The schematic describes the step-by-step process recommended for successful dome formation for all systems. Please click here to view a larger version of this figure.

3. IO size characterization

NOTE: The size of the organoids was characterized by brightfield images taken at 4x and 10x. The image processing analysis was automated using MATLAB. The overall steps of the process are described below, and a sample of the code is included in Supplementary File 1.

1. Define the directory containing the images and list all the image files in the directory. Initialize the table to store the results. Read the brightfield images in a file.
2. Define pixel to µm conversion and set the scale. If the image does not contain a scale bar, request the user the pixel-to-µm conversion factor.
3. Convert the image to grayscale. Apply a Gaussian filter to reduce noise in the image. Threshold the filtered image to separate organoids from the background.
4. Fill small holes and remove small objects to clean up the binary image. Perform connected component analysis to identify individual organoids and calculate their properties such as area, centroid, major axis length, and minor axis length.
5. Calculate organoid size based on the major and minor axis lengths. Display the segmented organoids on the original image and label them with their size. Print the organoid sizes and store results in .cvs file.
   NOTE: The first couple of images were analyzed manually to determine the optimal parameters for the Gaussian filter standard deviation and the area to the threshold. The code in Supplementary File 1 provides an example of the basic framework for organoid size analysis from brightfield images; however, further refinement is needed to suit the specific requirements for each image type and quality. The same process can be done using open-source software like FIJI from Image J.
6. Follow the steps below to use the FIJI software.
   1. Define pixel to µm conversion and set the scale. If the image does not contain a scale bar, request the user the pixel-to-µm conversion factor by clicking Analyze > Set Scale.
   2. Convert the image to grayscale by clicking Image > Type > 8bit. Apply a Gaussian filter to reduce noise in the image by clicking Process > Filter > Gaussian blurb > Sigma (radius) used: 2.
   3. Threshold the filtered image to separate organoids from the background by clicking Image > Threshold > MaxEntropy > Apply.
   4. Fill small holes and remove small objects to clean up the binary image by clicking Process > Noise > Remove Outliers > 20 pixels.
   5. Perform connected component analysis to identify individual organoids and calculate their properties such as area, centroid, major axis length, and minor axis length by clicking Analyze > Set measurement > Make sure Area, Perimeter, and Diameter are included > Analyze > Analyze Particles > Show Outlines.
7. Statistical analysis
   1. Assess the data distribution for normality through the Saphiro-Wilk test using JMP (SAS) software. To examine statistical differences between groups, perform a two-way ANOVA, and conduct post-hoc testing using the nonparametric Wilcoxon method in JMP(SAS) software. Significance was established at an alpha level of p ≤ 0.05.

Representative Results

Following this protocol, commercially available basement membranes and a xeno-free hydrogel system were successfully utilized to cultivate hiPSC cells and differentiate them into hIO. The main objective of these experiments was to systematically evaluate the equivalency of matrices from various sources for hiPSC and hIO work. The first section of this protocol focused on the maintenance and characterization of a healthy iPSC culture that yields an efficient intestinal organoid generation. The process of coating the culture ware with animal-derived matrices Matrix 1-AB, Matrix 2-AB, and Matrix 3-AB was described and compared with a xeno-free hydrogel system Matrix 4-XF.

As shown in Figure 3A, the hiPSCs BXS 0116 (CD34+ derived) show comparable morphology when cultured on the three animal-derived matrices. Nevertheless, comparing the cell line BXS 0116 with SCTi003-A (PBMC derived) cultured on Matrix 1-AB-coated wells emphasizes that intrinsic differences between cell lines (shown in Table 6) can lead to variation regarding cell proliferation, differentiation, and morphology. For example, while it took 3 days to generate DE for the SCTi003-A cell lines, it took 5 days for the BXS0116 line. Moreover, as shown in Figure 3B-C, culturing hiPSCs in Matrix 4-XF hydrogel systems lead to colonies forming in a partially embedded 3D colony that is drastically different from the flat morphology of hiPSC culture on plates coated with Matrix 1-AB or equivalent systems. Also, one of the key aspects to note from these experiments is that making a xeno-free hydrogel system using 3x growth factors can considerably improve cell viability and expression of stem cell markers. Altogether, hiPSC cultures on animal-derived matrices and xeno-free Matrix 4-XF made with 3x growth factors concentration led to an equivalent expression of SSEA-4 (stem cell marker)²²; however, the hydrogel system promotes the formation of partially embedded 3D colonies.

The second section of this protocol focuses on the differentiation of hiPSCs to generate hIO. Here, a commercially available kit developed based on Spence et al.²³ was used to differentiate into definite endoderm (DE), then midgut/hindgut (MH), and finally collect spheroid to mature into IO (Figure 4C). For an efficient generation of IO, examining that the hiPSC culture has minimal spontaneous differentiation before starting the differentiation protocol is crucial. As shown in Figure 4A, the lower the expression of stem cell markers such as SSEA-4 when starting the DE differentiation, the less efficient the process will be at every stage (i.e., lower expression of MH markers) and subsequent poor generation of hIO. Utilizing a compact benchtop flow cytometry device facilitated quick access to flow cytometry results and deciding about the quality of differentiation before moving to the next stage. A peculiar observation was that Matrix 2-AB-maintained hiPSC resulted in fewer spheroid releases during midgut/hindgut stage compared to the other animal-derived systems. Another key aspect to note is that because colonies of hiPSC maintained in Matrix 4-XF already had a 3D structure at the beginning of the differentiation, these systems led to larger spheroids.

Spheroids were embedded into Matrix 1-ABO, Matrix 3-ABO, and Matrix 4-XFO1- Matrix 4-XFO4 and were matured into hIO using a kit. After maturing spheroids for 7 days, they were passaged and embedded into their respective matrices systems. The growth of the organoid on each system was tracked for 7 days from brightfield images. While the culture started with relatively similar-sized organoids, after 5 days, hIO size varied as a function of the matrix in which they were embedded (Figure 5). Of the four Matrix 4-XFO, the formulation that resulted in larger organoids was Matrix 4-XFO3. The Matrix 4-XFO systems are formulated with different ligands and mechanical properties to fit a variety of applications²⁴. Specifically, Matrix 4-XFO3 stiffness²⁵ is closer to human gut stiffness²⁶ than the other Matrix 4-XFO formulations, which supports recent findings about the role of ECM mechanical properties on organoid expansion^12,27,28. Our representative results indicate that Matrix 1-ABO and Matrix 3-ABO equally facilitate the growth and expansion of intestinal organoids.

Figure 3: Selecting matrix system for iPSC maintenance. (A) Representative images of two cell lines grown on culture ware coating with the three different animal-derived matrix systems. (B) Representative images comparing the use of Matrix 4-XF with 1x vs. 3x growth factor supplementation show that increasing growth factor supplementations improves the viability of hiPSC on Matrix 4-XF hydrogel. (C) Flow cytometry comparison of SSEA-4 expression of Matrix 1-AB, Matrix 3-AB, Matrix 4-XF (3X) shows equivalent expression while Matrix 4-XF (1X) led to lower expression of SSEA-4. Bars represent mean ± SD of n = 5; two-way ANOVA, and post-hoc testing with nonparametric Wilcoxon method was used for significance testing. Scale bar = 200 µm. Please click here to view a larger version of this figure.

Figure 4: Differentiating iPSC into definite endoderm (DE) and midgut/hindgut (MH). (A) Representative images of iPSC differentiation into DE starting with different levels of SSEA4 marker highlight the importance of starting differentiation with a hiPSC population with high expression of stem cell markers. The red arrows point to areas of spontaneous differentiation prior to starting DE. (B) Example of flow cytometry analysis of the stem cell marker SSEA4 expression before starting differentiation and DE marker FOXA2 after 3 days of DE differentiation for a poor DE differentiation (left) and a good differentiation (right). Each graph represents the analysis of cells extracted from 1; for statistical analysis, it is recommended to average the results from at least 3 flow cytometry runs. (C) Representative images of iPSC differentiation into hIO on all matrices studied. Scale bar = 200 µm. Please click here to view a larger version of this figure.

Figure 5: Selecting matrix system for spheroid embedding and hIO maturation. (A) Sample images of maturation of hIO embedded in Matrix 4-XFO 1-4. The elastic modulus of the Matrix 4-XFO systems is around 50-300 Pa, with the highest for Matrix 4-XFO3 > Matrix 4-XFO4 > Matrix 4-XFO2 > Matrix 4-XFO1. Matrix 1-ABO and Matrix 3-ABO stiffness range from 440-800 Pa depending on the batch. (B) Sample images of the maturation of hIO were embedded in Matrix 1-ABO, Matrix 3-ABO, and Matrix 4-XFO3 on the day of embedding (Day 0) and after 1 week of culture (Day 7). (C) Data showing size comparison of hIO matured on the different matrix systems. Bars represent mean ± SD of n = 7; two-way ANOVA, and post-hoc testing with nonparametric Wilcoxon method was used for significance testing. Scale bar = 200 µm. Please click here to view a larger version of this figure.

Figure 6: Representative immunofluorescence images of intestinal organoids grown in the different matrices confirm the epithelial cell population. Whole intestinal organoids grown in the different matrices were fixed and stained with epithelial cell adhesion molecule (EpCAM) (green) and counterstained for cell nuclei DAPI (blue). All the organoids show EpCAM-positive cells, confirming the epithelial cell population localized on the exterior surface of the organoids. Scale bar = 200 µm. Please click here to view a larger version of this figure.

Table 1: Summary of aliquot preparation, storage, and dilution of matrices used to coat culture ware used for iPSC. * For accurate dilution factor, check the Dilution factor recommended and/or protein concentrations in the Certificate of Analysis of each lot. Please click here to download this Table.

Table 2: Reference table of the volume of coating solutions per surface area of culture ware and mTeSR Plus Medium. Please click here to download this Table.

Table 3: Reference table of the volume of Matrix 4-XF precursor solutions per surface area of culture ware and complete stem cell medium. Please click here to download this Table.

Table 4: List of common markers recommended to characterize hiPSC and hIO during the process. Please click here to download this Table.

Table 5: Summary of media needed during hiPSC differentiation into Intestinal Organoids. Please click here to download this Table.

Table 6: Specific details about the two hiPSC lines used in this study. Please click here to download this Table.

Supplementary File 1: General MATLAB Code to modify according to specific image type. Please click here to download this File.

Discussion

Selecting the optimal microenvironment for stem cell and organoid work is a pivotal early step when using these platforms for a wide range of applications. Our representative results show that Matrix 4-XFO3, in combination with a higher concentration of growth factors, leads to larger organoids, suggesting that the physical properties of xeno-free hydrogels can be harnessed to optimize organoid generation using these systems. It has been previously shown that the unique characteristics of the extracellular matrix (ECM) are closely intertwined with the identity of the cells within the organ¹². Because cells sense external forces via integrins and other receptors, the mechanical characteristics of the ECM profoundly influence stem cells, and in response, stem cells can generate internal forces¹². Interactions between the stem cells and their microenvironment are key in initiating diverse signaling pathways¹². Additionally, it has been shown that the ECM can serve as a reservoir for storing and releasing growth factors and other signaling molecules that impact cell behavior. Hence, our representative results and decades of research emphasize the importance of considering biochemical and physical aspects of the native cell microenvironment and tissue properties when selecting matrix systems for in vitro investigations^29,30.

Several critical steps and recommendations were explained through the protocol to advise the users of potential issues and ways to avoid them. Nevertheless, two critical steps are presented in these methods that are pivotal for successfully implementing this protocol. First, checking for specific markers associated with the stem cell population is crucial to determining the optimal coating system to maintain the stem cell population ready for efficient controlled differentiation. Similarly, confirming the expression of markers associated with each differentiation step is essential to deciding when to proceed to the next stage. This protocol provides a general guideline regarding the minimum time required at each stage. Nevertheless, the optimal time at each stage will vary for different cell lines and experimental conditions (e.g., genotype, stiffness). Therefore, characterizing multiple stage-specific markers is the best approach to improve the efficiency of hIO generation.

The second critical step was incorporating a higher concentration of growth factors when using the xeno-free Matrix 4-XF system, which is of utmost importance. One frequently highlighted limitation of xeno-free biomaterial systems is the lack of growth factors³¹. However, this weakness can become a strength: full control of growth factors in the systems to improve reproducibility. While this protocol describes using a commercially available media and growth factor supplementation kit, the same rationale can be used when making the complete media in-house with individual components. The specific ratio hydrogel/media guides how much higher needs to be the concentration of growth factors; for example, a 2:1 ratio might require 3x concentration of growth factors to end up with a concentration representative of the standard complete media used. The full control of growth factors present in the system is beneficial to answering mechanistic questions and avoiding regulatory issues for many applications.

The image processing steps described in step 3 are limited by the quality of the images and the type of images taken. It might require refinement to suit the specific requirements of each image type and quality. However, the overall steps of the process and a sample of MATLAB code of the basic framework for organoid size analysis from brightfield images help to define a workflow that can be further optimized and expanded. Furthermore, access to a brightfield microscope is more common than other types of microscopes. While lack of access and familiarity with MATLAB might limit the use of this code, the general steps of the process are compatible with open-source software such as Fiji by Image J. Organoid size alone is not a recommended metric to assess maturation; it is encouraged to follow with an evaluation of specific markers representative of the target organ; some of these markers for intestinal tissue are included in Table 4. Figure 6 shows an example of immunofluorescence staining of whole organoids with the epithelial cell adhesion molecule (EpCAM), which is localized to the exterior surface (basolateral) of the organoids. While EpCAM was visualized in the organoids cultured in all the ECM systems, there were differences in the size of these organoids. Thus, information about organoid size helps to optimize the culturing conditions. Furthermore, in drug response studies, organoid size serves as a metric of the response to the tested compound; changes in size can reflect the effectiveness or toxicity of the tested compounds. Importantly, generating multiple batches of organoids with standardized size is a current challenge in the field, but it is an important metric of experimental reproducibility so results can be compared across different experiments or laboratories. Altogether, an automated workflow that allows the extraction of important information about organoids from brightfield images is a beneficial tool for many researchers.

Both systems, animal-derived matrices, and xeno-free hydrogels, have their advantages and disadvantages. Matrix 2-AB leading to fewer spheroid releases during the MH stage than the other animal-derived matrices could result from lot-to-lot variations, a widely known characteristic of commercially available animal-derived matrices. It is important to note the concentration of the commercially available ECM systems varies; for example, the dilution factors in the product specification sheets were used to prepare the coating solutions which might lead to slightly different concentrations in the case of the animal-derived matrices. These differences in protein concentration can significantly impact differentiation leading to variation in differentiation efficiency when using the same product from different lots. To select between these systems, consider factors such as specific experimental goals, ethical considerations, and regulatory requirements. For example, in cases of comparing in vitro experimental results with a mouse animal model to answer tumor and cancer biology questions, using matrices derived from the Engelbreth-Holm Swarm EHS mouse can be a great choice. Also, because products such as Matrigel have been used for decades, there are many protocols, data, and references. However, using well-defined xeno-free matrices that allow the customization of physical and chemical properties might be a more suitable option when the research question is associated with human diseases or therapies. Also, most xeno-free hydrogel systems are designed to meet regulatory standards, which makes them a suitable choice for research that requires compliance with good manufacturing practices (GMP) or other regulatory guidelines.

In summary, while animal-derived matrices have been widely used for iPSC maintenance and organoid generation due to their biocompatibility, xeno-free hydrogel systems offer advantages in terms of reproducibility, cost-effectiveness, regulatory compliance, and ethical considerations. The choice between the two depends on the specific needs of the research or clinical application. Xeno-free hydrogel systems are gaining popularity as a more versatile and sustainable alternative that can support the utilization of organoids and microphysiological systems for the pre-clinical shortlist of therapeutic candidates. This application can significantly improve early drug development and help tackle current clinical translation challenges in the pharmaceutical industry.

Materials

24-Well Plate (Culture treated, sterile)	Falcon	353504
37 °C water bath	VWR
96-well plate	Fisher Scientific	FB012931
Advanced DMEM/F12	Life Technologies	12634
Anti-adherence Rinsing Solutio	STEMCELL Technologies	7010
Biological safety cabinet (BSC)	Labconco	Logic
Brightfield Microscope	Echo Rebel	REB-01-E2
BXS0116	ATCC	ACS-1030
Centrifuge with temperature control (4 °C capabilities)	ThermoScientific	75002441
Conical tubes, 15 mL, sterile	Thermo Fisher Scientific	339650
Conical tubes, 50 mL, sterile	Thermo Fisher Scientific	339652
Cultrex RGF BME, Type 2	Bio-techne	3533-005-02
Cultrex Stem Cell Qualified RGF BME	Bio-techne	3434-010-02
D-PBS (Without Ca++ and Mg++)	Thermo Fisher Scientific	14190144
GeltrexLDEV-Free, hESC-Qualified Reduce Growth Factor	Gibco	A14133-02
GlutaMAX Supplement	Thermo Fischer Scientific	35050-061
Guava Muse Cell Analyzer or another flow cytometry equipment (optional)	Luminex	0500-3115
HEPES buffer solution	Thermo Fischer Scientific	15630-056
Heralcell Vios Cell culture incubator (37 °C, 5% CO2)	Thermo Scientific	51033775
JMP Software	SAS Institute	JMP 16
MATLAB	MathWorks, Inc	R2022b
Matrigel Growth Factor Reduced (GFR) Basement Membrane Matrix LDEV free	Corning	356231
Matrigel Matrix High Concentration (HC), Growth Factor Reduced (GFR) LDEV-free	Corning	354263
mTeSR Plus Medium	STEMCELL Technologies	100-0276
Nunclon Delta surface treated 24-well plate	Thermo Scientific	144530
PE Mouse Anti-human CD326 (EpCAM)	BD Pharmingen	566841
PE Mouse Anti-human CDX2	BD Pharmingen	563428
PE Mouse Anti-human FOXA2	BD Pharmingen	561589
PerCP-Cy 5.5 Mouse Anti-human SSEA4	BD Pharmingen	561565
ReLeSR	STEMCELL	5872
SCTi003-A	STEMCELL Technologies	200-0510
Serological pipettes (10 mL)	Fisher Scientific	13-678-11E
Serological pipettes (5 mL)	Fisher Scientific	13-678-11D
STEMdiff Intestinal Organoid Growth Medium	STEMCELL Technologies	5145
STEMdiff Intestinal Organoid Kit	STEMCELL Technologies	5140
Vitrogel Hydrogel Matrix	TheWell Bioscience	VHM01
VitroGel ORGANOID Discovery Kit	TheWell Bioscience	VHM04-K