Method Article

Measuring Intestinal Barrier Permeability by Detecting Flux of Fluorescent Tracers Across Human Colonoid Monolayers

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DOI:

10.3791/68857

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October 17th, 2025

 ,  , 
1Department of Pediatrics, Division of Gastroenterology, Hepatology and Nutrition, Children's Hospital Colorado, 2Department of Medicine, Mucosal Inflammation Program, Division of Gastroenterology and Hepatology, University of Colorado, 3Rocky Mountain Veterans Hospital

In This Article

Summary

We describe a technique for measuring intestinal barrier integrity, whereby the rate of translocation of a fluorescently-tagged sugar of known size reflects the permeability of tissue-derived human intestinal colonoids.

Abstract

The intestinal barrier is a critical site of regulation between luminal antigens and the host immune system, and its dysregulation is implicated in multiple gastrointestinal diseases. Intestinal organoids can be a useful tool to understand and model differences in intestinal barrier integrity. Here, we describe a protocol that uses human colonoids to quantify functional intestinal barrier differences by measuring the flux of fluorescent tracers across organoid monolayers. After growing monolayers to confluence, we add a fluorescent tracer, such as FITC-dextran, to the apical chamber and detect its translocation into the basolateral chamber over time. Increased translocation of the fluorescent tracer is reflective of a more permeable, or weaker, epithelial barrier. This protocol is an adaptable platform that allows evaluation of functional intestinal barrier differences across multiple conditions in parallel. Using variations of this protocol, one can assess transepithelial permeability to molecules of various size, shape, and chemical structure, as well as permeability differences in the context of isolated treatments or microenvironmental changes on the apical or basolateral sides of epithelial cells. This allows for a deeper understanding of the mechanism and treatment of barrier disruption in disease states.

Introduction

The intestinal mucosa is a complex structure that plays an important role in regulating the contact between the metabolites, antigens, and microbes in the intestinal lumen and the immune system and enteric circulation. Intestinal epithelial cells are a key component of this system, and in healthy conditions, form a highly selective barrier. Transport across the epithelium can occur through multiple mechanisms: transcellular active or passive transport, paracellular pore or leak pathways regulated by tight junctions and adherens junctions. Whether any of these pathways can be used by a molecule to translocate across the epithelium is determined by the hydrophobicity, size, and chemical structure. Once across the epithelium, these molecules can more readily interact with mucosal immune cells and enter the blood or lymphatic circulation of the intestine1,2 Disruption of the intestinal barrier is a hallmark of several inflammatory gastrointestinal diseases, such as inflammatory bowel disease and celiac disease1,3,4,5,6,7. This weakened intestinal barrier is thought to play a role in the pathophysiology of these disease processes through dysregulated translocation of pathogenic antigens, which can then create a local or systemic inflammatory response.

There are several established protocols for measuring the intestinal barrier. In vitro studies traditionally use cancer cell lines, such as T84 or Caco-2 cells, to measure transepithelial electrical resistance (TEER) and to quantify tight junction and adherens junction components8. Ex vivo, measurements can be taken of transepithelial current using Ussing chambers on biopsy or resected intestine samples9. In vivo studies typically involve giving inert tracers orally, then measuring blood or urine absorption of these tracers10,11. Though these techniques have proved useful in the study of intestinal barrier, they present several limitations: transformed immortal cell lines inherently have metabolic and cytoskeletal changes that affect the intestinal barrier in poorly defined ways; access to patients, patient samples, or model organisms can be limited; in vivo studies are difficult to control for confounding factors or make conclusions at a molecular level. Thus, though evidence of this barrier disruption has been reported in multiple disease models, understanding of the mechanism of these intestinal barrier defects remains limited, and our ability to target and treat this aspect of gastrointestinal disease has remained elusive.

Human intestinal organoids offer a unique and exciting capability to elucidate the cause and treatment of intestinal barrier defects. Tissue-derived human intestinal organoids are primary cell cultures derived from adult stem cells located at the base of intestinal crypts12. These cells can be isolated from biopsy or surgical specimens and cultured to maintain genetic and epigenetic changes seen in the patients from whom they were derived13,14,15. They serve as a more physiologically relevant model than traditional immortal cell lines because they do not have the metabolic changes of cancer cell lines and can be differentiated into the various cell types present in the mature intestinal epithelium16,17,18. In addition, because they can be expanded in vitro, they are more accessible and useful for controlled studies of the cellular effect of various treatments and growth conditions in parallel. Though there are limitations inherent in intestinal organoid models as well, most notably the absence of the other non-epithelial cell types and the context of the whole-body system, it has been shown that human intestinal organoids provide a reliable model for multiple aspects of gastrointestinal disease13.

Among these, intestinal organoids have been shown to closely model intestinal barrier defects, demonstrating expected changes in tight junction organization and TEER in the setting of barrier insults19,20,21. Here, we adapt one technique of assessing barrier integrity in human colonoids, in which flux of fluorescently tagged molecules across organoid monolayers is detected22. In this protocol, organoids are grown and matured into polarized two-dimensional monolayers on semi-permeable membranes. The assay entails the addition of a fluorescent tracer to the apical surface of the epithelial monolayer, and its presence on the opposite side is quantified over time. Increased flux of the tracer is reflective of a leakier or more permeable epithelium. This assay provides a functional barrier readout, which can be additive to evaluations of the structure of tight and adherens junctions. Additionally, though TEER measurements are often used to provide a basic assessment of barrier integrity, the flux assay described here can be adapted to detect permeability of molecules of various classes, charge, and size, thereby characterizing the barrier in much greater detail. Several commercially available fluorescent tracers can be used for this purpose. Further, through the use of non-overlapping fluorescent spectra, permeability to multiple tracers can be assessed simultaneously. Moreover, the monolayer configuration of this assay allows the assessment of barrier response to growth conditions applied specifically to the apical or basolateral surface of cells only. With this, a controlled assessment of the effect of therapeutics, microenvironmental changes, and co-culture conditions is possible.

Intestinal barrier defects are known to play an important pathogenic role in the development of several gastrointestinal diseases, but the term "leaky gut" is increasingly used in popular media without precision23, and efforts to target defective intestinal barrier have been limited in success. This highlights the need for improved models to evaluate and characterize the intestinal barrier. The described system uses physiologic human tissue-derived intestinal organoid monolayers to assess functional barrier integrity under controlled conditions.

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Protocol

All samples were acquired and used in accordance with a protocol approved by the University of Colorado Institutional Review Board (IRB), protocol number 14-2012.

NOTE: All procedures in steps 1-3 should be done in a sterile biosafety cabinet with sterile technique. A schematic of the protocol is presented in Figure 1.

1. Establishment of human colonoids from frozen cultures

NOTE: Ensure all materials are ready in the biosafety cabinet before thawing cells to minimize exposure of thawed organoids to freezing medium. The protocol begins with cryopreserved tissue-derived human colonoids. Protocols for deriving and cryopreserving human colonoids have been described previously24.

  1. Preparation of human intestinal organoid stem medium
    1. Prepare Wnt-3A/Noggin/R-Spondin 3 conditioned media (L-WRN) according to previously described protocols25,26.
    2. Supplement the L-WRN media with the following growth factors: Epidermal growth factor [50 ng/mL], Nicotinamide [10 mM], A83-01 [500 nM], SB202190 [10 µM], CHIR99021 [3 µM], Thiazovivin [4 µM], SB431542 [4 µM], Gastrin I [10 nM].
    3. Mix the supplemented L-WRN media 1:1 with commercial intestinal organoid media.
    4. Just prior to use, supplement with Y-27632 [10 µM].
      NOTE: Optimal intestinal organoid stem medium composition may vary based on lab and colonoid line. Use of supplemented L-WRN media alone or use of commercial intestinal organoid media alone may also be possible. This protocol's use of equal parts L-WRN conditioned media and commercial media is more resilient to batch-to-batch variability compared to the use of conditioned media alone and is more cost-effective compared to the use of commercial media alone.
  2. Preparation of differentiation medium
    1. Supplement DMEM/F12 with 10% fetal bovine serum (FBS), 1% penicillin-streptomycin, and 2 mM L-alanyl-L-glutamine dipeptide.
    2. Mix supplemented DMEM/F12 1:1 with intestinal organoid stem medium from step 1.1
  3. Preparation of extracellular matrix (ECM)
    1. Thaw ECM overnight on ice, then prepare 100 µL aliquots.
      NOTE: Aliquots can be refrozen at -20 °C. See manufacturer instructions for details.
    2. Incubate aliquots on ice until the time of plating cells.
  4. Thawing and plating organoids
    1. Thaw contents of cryovial in a 37 °C water bath, ensuring the vial remains submerged, but the cap is above the water, until just thawed.
    2. Transfer the contents of the vial to a 15 mL conical tube containing 13 mL of ice-cold supplemented DMEM/F12 medium (from step 1.2.1).
    3. Centrifuge at 200 × g for 5 min at 4 °C.
    4. Aspirate the supernatant, leaving approximately 1 mL of residual volume above the cell pellet.
    5. Place the tube on ice. Add 1 mL of ice-cold supplemented DMEM/F12 (from step 1.2.1) and pipette the suspension 10-15 times to resuspend the pellet.
    6. Centrifuge at 200 × g for 5 min at 4 °C.
    7. Remove the supernatant using a pipette, leaving as little residual wash media as possible over the cell pellet.
    8. Add 100 µL of thawed ECM to the pellet. Resuspend thoroughly, ensuring the organoids are evenly dispersed without clumps. Pipette slowly to avoid introducing air bubbles.
    9. Plate ECM by pipetting approximately 10 domes or three-dimensional droplets of ECM, about 10 µL each, into one well of a 6-well plate.
    10. Invert the plate quickly and incubate at 37 °C for 5-10 min or until the domes are solidified.
    11. Once the domes are set, add 2 mL of intestinal organoid stem media (from step 1.1) to the well.
    12. Incubate at 37 °C, in a humidified incubator with 5% CO2. Replace the media every 2 days.
    13. Proceed with step 2 once organoids are either taking up greater than about 75% of the area of the ECM dome or are accumulating central debris.

2. Passaging and expansion of human colonoids

  1. For each well, aspirate the growth media from the wells without disrupting ECM domes.
  2. Wash the domes gently with 1 mL of ice-cold phosphate-buffered saline (PBS), then remove PBS.
  3. Add 1 mL of ice-cold trypsin to each well. Use a pipette to break up the ECM domes by pipetting up and down. Pipette the mixture 10-15 times to fully dissociate the ECM.
  4. Incubate the plate at 37 °C for 3-5 min.
  5. Add 1 mL of ice-cold supplemented DMEM/F12 (from step 1.2.1) to the well to neutralize trypsin. Pipette the mixture vigorously 20-30 times until cells are dissociated into clusters of 2-3 cells each.
  6. Transfer the cell suspension to a 15 mL conical.
  7. Use an additional 1 mL of supplemented DMEM/F12 (from step 1.2.1) to wash the well and add the wash solution to the 15 mL conical.
  8. Add supplemented DMEM/F12 (from step 1.2.1) to reach a total volume of 13 mL in the 15 mL conical.
  9. Proceed with steps 1.4.3-1.4.12 of step 1 using an appropriate amount of ECM to expand into additional wells.
    NOTE: A confluent well of organoids in 100 µL ECM can typically be passaged into 2-3 wells (200-300 µL of ECM). One confluent well of organoids is sufficient for approximately 3-6 monolayers in step 3.

3. Generation of human colonoid monolayers

  1. Collagen coating cell culture inserts
    1. Place polystyrene cell culture inserts (0.4 µm pore size) in a 24-well plate and add 100 µL of collagen coating solution to the apical chamber of each insert.
    2. Seal the plate with sealing film to prevent evaporation.
    3. Incubate at 4 °C for 4-24 h.
      NOTE: There is no difference in results within this incubation time, but a shorter or longer incubation time can be chosen to fit a convenient pause point.
  2. Plating organoid monolayers
    1. Follow steps 2.1-2.5 to dissociate the organoids.
    2. Pass the cell suspension through 70 µm cell strainer into a 50 mL conical tube.
    3. Rinse the wells with 1 mL each of supplemented DMEM/F12 (from step 1.2.1). Collect this wash media and pass it through the cell strainer.
    4. Wash the cell strainer with supplemented DMEM/F12 (from step 1.2.1) until the total volume in the 50 mL conical tube reaches 15 mL.
    5. Transfer the filtrate into a 15 mL conical.
    6. Centrifuge at 200 × g for 5 min at 4 °C.
    7. Aspirate the supernatant, leaving 1 mL above the pellet.
    8. Add an additional 1 mL of supplemented DMEM/F12 medium (from step 1.2.1) and resuspend.
    9. Examine the suspension under a microscope to confirm complete dissociation. If needed, pipette the cells vigorously until they are mostly single cells.
    10. Centrifuge at 200 × g for 5 min at 4 °C. Remove the supernatant using a pipette, leaving as little residual wash media as possible over the cell pellet.
    11. Resuspend in a small volume (200-1000 µL) of intestinal organoid stem medium (from step 1.1). Keep on ice.
    12. Take an aliquot of the cell suspension to use to count viable cells using the preferred method. Use the viable cell count to determine the volume of cell suspension needed to plate 100,000 viable cells per insert.
    13. Aspirate the collagen coating solution from each insert without touching the insert membrane.
    14. Add the appropriate volume of intestinal organoid stem media (from step 1.1) to the apical chamber of each insert to bring the final volume to 200 µL once the cell suspension is added.
      NOTE: If preferred, the cell suspension in step 3.2.12 can be adjusted to a final concentration of 100,000 cells/200 µL, then 200 µL can be plated into the apical chamber. However, if the entire cell suspension is not used for monolayer plating, this may lead to excessive waste of media.
    15. Add the appropriate volume of cell suspension to the apical chamber.
    16. Add 500-1000 µL of intestinal organoid stem media to the basolateral chamber.
      NOTE: 500 µL is sufficient for typical colonoids. If cells are more metabolically active or otherwise sensitive to waste product build-up, use of a larger volume in the basolateral chamber may be helpful to maintain growth.
    17. Repeat for all the inserts/wells.
    18. Incubate at 37 °C, in a humidified incubator with 5% CO2.
    19. Replace the media once per week.
      NOTE: Treatments of interest can be applied to the apical and/or basolateral chambers of inserts at any time before or after plating.

4. Measuring transepithelial electrical resistance (TEER) and monolayer maturation

  1. Measure resistance
    1. Use an epithelial volt/ohm meter on the ohm setting. Clean the probe by wiping it down with a task wipe soaked in 70% ethanol. Place the longer arm of the electrode in the basolateral compartment and the shorter arm in the apical compartment of the insert. Record measurement.
      NOTE: If TEER measurements are also an endpoint of interest, also take resistance measurements of a collagen-coated insert without cells as a background. Then, TEER is calculated as (resistance - background insert resistance) x surface area of insert. The surface area of the insert in this protocol is 0.33 cm2. This calculation will give TEER in ohm·cm2. If using a different epithelial volt/ohm meter than listed in the Table of Materials, use the manufacturer's instructions for use of the meter to measure resistance.
    2. Repeat daily until measurements plateau.
      NOTE: Plateau typically occurs after 2 to 3 weeks with TEER greater than 200 ohm·cm2, but varies by colonoid line and growth conditions.
  2. Once resistance has plateaued, initiate organoid maturation by replacing stem media with differentiation media (from step 1.2).
  3. Continue daily TEER measurements. Expect an increase in TEER and a second plateau.
    NOTE: Second plateau typically occurs around 7 days with TEER of 1500-2500 ohm·cm2, but this varies by colonoid line and growth conditions.
  4. Once TEER measurements have plateaued a second time, proceed with step 5.

5. FITC-dextran flux assay

NOTE: This protocol uses FITC-dextran 4 kDa (Stokes radius 14 Å). However, depending on experimental needs, fluorescent tracers of different sizes, charges, and fluorophores can be used22. Multiple tracers may also be used at the same time, in which case, non-overlapping excitation and emission fluorescence spectra should be confirmed. Steps 5.1-5.3 are suggested to be performed in a warming cabinet (non-humidified, non-CO2 incubator) set to 37 °C. If this is not available, reagents should be warmed to 37 °C prior to use, and cells, buffers, and collected samples should be kept at 37 °C during incubation times.

  1. Reagent preparation
    1. Prepare Hank's buffered saline solution (HBSS) with 10 mM HEPES and adjust pH to 7.4. Warm to 37 °C.
      NOTE: If needed, prepare HBSS+HEPES with treatments of interest. Prepare enough for 80 µL/well in the apical chamber and 1 mL/well in the basolateral chamber.
    2. Pour about 50 mL of the warm HBSS+HEPES into a small beaker for rinsing the inserts. Have a larger beaker nearby for liquid waste.
    3. Completely thaw FITC-dextran 4 kDa. Prepare a 1.25 mg/mL solution in HBSS with HEPES. Prepare an adequate volume of FITC-dextran solution to allot 80 µL per well and 1 µL for the standard curve.
      NOTE: The starting concentration can be adjusted based on the size and expected permeability of the tracer. For example, smaller or more permeant tracers may be added at a lower concentration to conserve reagents. If treatments need to be applied to the apical chamber, prepare FITC-dextran solution in appropriately treated HBSS+HEPES.
  2. Application of FITC-dextran to monolayers
    1. Prepare a fresh 24-well plate by adding 1 mL of HBSS+HEPES (with treatment if desired) into the appropriate wells.
    2. Wash the organoid monolayers by grasping the insert with forceps, lifting it out of the well, dunking it into the beaker containing warm HBSS+HEPES (from step 5.1.2) to fill the apical chamber, then inverting it over the waste beaker to empty the wash solution.
    3. Repeat this wash 2 additional times.
    4. Place the insert into the appropriate well of the fresh plate prepared in step 5.2.1.
    5. Gently pipette 80 µL of FITC-dextran solution into the apical chamber. Pipette onto the side of the insert in order to avoid disturbing the monolayer.
    6. Repeat step 5.2.5 for all monolayers.
    7. Add HBSS or PBS to any empty wells to maintain humidity.
  3. Collecting flux samples
    1. Immediately after step 5.2.7, collect 70 µL from the basolateral chamber of each well into a well of a flat, clear-bottom black 96-well plate and keep protected from light at 37 °C. This will be time point 0.
    2. Incubate the plate, protected from light, at 37 °C.
    3. Every 30 min for 2 h, mix the buffer in the basolateral well, then collect an additional 70 µL into empty wells of the clear-bottom 96-well plate from step 5.3.1.
  4. Preparation of the standard curve
    1. Add 1 µL of the reserved FITC-dextran solution (1.25 mg/mL; step 5.1.3) into 999 µL of HBSS+HEPES. This will be the 1:1000 or 1.25 µg/mL dilution for the standard curve.
    2. Serially dilute this solution 1:1 by taking 150 µL of the solution and adding it to 150 µL of HBSS+HEPES, then mixing well. Repeat this serially ten additional times for a total of a 12-step dilution.
      NOTE: This will create a standard curve of: 1250, 625, 312.5, 156.25, 78.13, 39.06, 19.53, 9.77, 4.88, 2.44, 1.22, and 0.61 ng/mL. Depending on the permeability of the cell lines, the standard curve may need to be more or less concentrated. The first time the experiment is run, reserve an additional FITC-dextran 1.25 mg/mL solution in case the standard curve needs to be remade at a different concentration.
    3. Pipette 70 µL of each dilution of the standard curve into empty wells of the 96-well clear bottom plate from steps 5.3.1 and 5.3.3. Repeat for the second replicate.
  5. Measure fluorescence on a fluorescent plate reader with the following settings: excitation 490 nm, emission to 520 nm, reading from the bottom of the plate, with a gain of 100 and read height of 7 mm.
    NOTE: Adjust excitation/emission wavelengths as appropriate for the fluorophore of choice.
  6. Data analysis
    1. Plot the standard curve with concentration on the x-axis and fluorescence on the y-axis. Use this to determine the linear regression equation.
    2. Use this linear regression equation and the samples' fluorescent readings to calculate the FITC-dextran concentration at each time point for each sample.
    3. Plot the concentration over time for each sample and calculate the slope of a linear regression equation, which represents micrograms/min (µg/min). Divide this by the surface area of the cell culture insert (0.33 cm) to determine the flux rate as microgram per minute per square centimeter (µg/min/cm2).

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Results

Using this protocol, the permeability of healthy human colonoids in the presence and absence of inflammatory signaling was analyzed. Addition of a mixture of inflammatory cytokines, termed cytomix, which includes IFNγ, TNFα, and IL-1β (each at 10 ng/ml), has been previously shown to induce intestinal barrier dysfunction as measured by TEER in intestinal epithelial cell models20,27. We aimed to determine if this barrier dysfunction could be recapitulated using the...

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Discussion

This is a protocol to assess the intestinal barrier in the physiologically representative model of human colonoid monolayers. The fluorescent tracer flux assay described here provides information about the functional integrity of the barrier beyond what is gleaned with other types of barrier assays, such as TEER, Ussing chamber measurements, or evaluation of tight junction protein expression. Additionally, the use of patient-derived intestinal organoid models confers the benefit of recapitulating complex human physiology...

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Disclosures

There are no disclosures or conflicts of interest.

Acknowledgements

Dr. Sean Colgan is funded by NIH grants DK1047893, DK50189, DK095491, DK103639, and VA Merit BX002182.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
15 mL conical centrifuge tubesCelltreat229411
2 mL Internal Threaded Cryogenic Vial Corning430488
50 mL conical centrifuge tubesCelltreat229421
6-well plate, tissue culture treatedCelltreat229106
70 μm cell strainerFalcon352350
96-well black/clear bottom plateThermo Scientific165305
A83-01 R&D2939
BioTek Synergy plate readerAgilentSynergy H1
Cellometer Auto T4 Brightfield cell counterNexcelcomCMT-AT4P
Cellometer cell counting chambersNexcelcomNC1531583
CHIR99021Sigma-AldrichSML1046-5MG
Collagen coating solution Cell Applications125-50
DMEM/F-12Gibco11330057
DMSOSigma-AldrichD2650-100ML
Ethanol 100%Fisher ScientificA4094
EVOM2 Voltohm Meter with Stx2 ElectrodeWorld Precision InstrumentsNC9792051
Fetal Bovine Serum (FBS)CytivaSH30396.03HI
Filter pipet tipsFisher Scientific02-707-000, 02-707-002, 02-707-006, 02-707-008
Fluorescein isothiocyanate (FITC)-dextran, 4kDaSigma-Aldrich46944
Gastrin I (Human)R&D3006
GlutaMAX Fisher Scientific35050061200 mM L-alanyl-L-glutamine dipeptide
Hanks buffered saline solution 10xThermo Scientific14065056
HEPES 1MThermo Scientific15630080
Interferon gamma recombinant humanBioLegend575308
Interleukin 1 beta recombinant humanThermo scientific200-01B-500UG
IntestiCult Intestinal Organoid Growth Medium (Human)STEMCELL Technologies06010 Commercial intestinal organoid media
Live inverted microscopeOlympusIX85
MatrigelCorning354234Extracellular matrix (ECM)
Microbiological incubatorFisher Scientific151030515
Multiwell plate for suspension culture, 24 wellGreiner Bio-one662102
NicotinamideR&D4106
ParafilmMillipore SigmaHS234526BSealing film
Pasteur pipets (borosilicate glass)Fisher Scientific13-678-20B
PBS Gibco10010-023
Penicillin Streptomycin (10,000 U/mL)Gibco15140122
Recombinant moouse epidermal growth factorR&D2028-EG
Refrigerated centrifugeBeckman CoulterBE-AX15R
SB202190 R&D1264
ThiazovivinR&D3845
ThinCert cell culture insert for 24 well plate, 0.4 μmGreiner Bio-one662641
Trypsin-EDTA (0.25%)Fisher Scientific25200114
Tumor necrosis factor alpha recombinant humanR&D10291-TA-100
Y-27632 dihydrochloride TocrisTB1254-GMP

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Intestinal OrganoidsFITC DextranTransepithelial PermeabilityEpithelial BarrierBarrier IntegrityOrganoid Model

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