Oral Gavage in Neonatal Mouse Pups and Functional Assessment of Gut Barrier Integrity Using Ussing Chambers

Gut barrier integrity plays a fundamental role in maintaining overall health by coordinating mucosal immune response and regulating the selective permeability of the intestinal epithelium. This barrier system controls the passage of nutrients while preventing the translocation of pathogens, antigens, and toxins into systemic circulation¹. Chronic disruption of this interface, associated with increased intestinal permeability, commonly referred to as "leaky gut", is now identified not only as a consequence but also as a driver of chronic inflammatory states and a range of extra-intestinal pathologies².

For instance, compromised gut barrier function has been shown to be a key component of autoimmune and metabolic disorders such as inflammatory bowel disease (IBD), juvenile idiopathic arthritis (JIA), and type 1 diabetes (T1D). This highlights the gut barrier state as a key biomarker, but also as a therapeutic target to prevent the initiation of pathophysiological processes as early as possible and to prevent the progression of acute inflammation to a chronic state^3,4,5,6.

In this context, the neonatal period represents a unique and highly sensitive window of development during which the establishment and maturation of the gut-microbiota interface profoundly influence lifelong intestinal and systemic health⁷. At this early stage, the intestinal epithelium and immune system display heightened sensitivity to both endogenous and exogenous factors, undergoing rapid growth and differentiation while simultaneously adapting to microbial colonization and dietary changes. Disruptions impairing gut barrier integrity during this critical developmental window may play a pivotal role in the onset of a wide spectrum of acute and chronic disorders later in life. These conditions include immune-mediated diseases such as food allergy, and more severe chronic illnesses, such as IBD, JIA, and T1D^7,8,9. These diseases underscore how pathophysiological processes often originate early, as the primary onset generally occurs between childhood and early adulthood^10,11. Therefore, understanding the mechanisms that govern gut barrier function in neonates is essential for developing preventive and therapeutic strategies aimed at reducing the burden of many chronic diseases.

Given this context, there is a pressing need for reliable, physiologically relevant tools to study gut barrier integrity, chronic disease mechanisms, and inflammation specifically within the neonatal setting. Such tools must allow the replication of experimental assays already proven in the adult context without sacrificing precision and efficacy. Additionally, it is becoming increasingly urgent to investigate both short- and long-term effects of many molecular candidates of interest, which are expected to have beneficial or harmful effects on gut homeostasis in the neonatal context. Among the tools available, oral gavage administration in neonatal mouse pups represents a more than reliable method of delivering molecules of interest at a defined developmental stage. It allows for controlled investigations into the effects of various bioactive compounds, drugs, or microbial metabolites on gut maturation and barrier function.

The first part of this article describes a method for gavage of mouse pups as early as Day Of Life (DOL) 6, allowing the study of interventions before, during and after the massive expansion of the gut microbiota species numbers induced by food diversification. Thus, this technique enables longitudinal analyses of treatment effects on intestinal health as pups grow and develop.

In the second part of this article and complementing this approach, ex vivo assessment of colonic epithelial barrier function using Ussing chambers provides a sensitive and quantitative measure of permeability characteristics. Evaluation of both paracellular and transcellular permeability in colon biopsies from neonatal mice (e.g., DOL 10 pups) allows for precise determination of the impact of several key candidates on epithelial barrier integrity and function. Paracellular transport is an essentially passive mechanism occurring through the intercellular space of the epithelial layer. It allows for the passage of small solutes driven by electrochemical and osmotic gradients from the lumen to the basolateral side. This mechanism is mainly regulated by proteins of the tight junction family, such as claudins and occludin. Transcellular transport, in contrast, is an active mechanism, allowing for the absorption of bigger solutes and occurring through the epithelial cells via specific channels, carriers, and pumps.

Fluorescein isothiocyanate (FITC)-dextran (4 kDa) predominantly undergoes paracellular transport due to its size, whereas horseradish peroxidase (HRP; 44 kDa) is selectively conveyed to the basolateral side via transcellular routes. Quantification of these non-endogenous markers thus enables distinct assessment of paracellular and transcellular permeability, respectively. The Ussing chamber assay with FITC and HRP preserves tissue architecture and allows real-time measurement of respectively paracellular and transcellular permeability of any segment of the intestinal tract. This combination of neonatal gavage and Ussing chamber analysis offers a powerful strategy to study how specific molecules impact gut barrier function both immediately and over longer timeframes, informing strategies to mitigate inflammation and prevent chronic disease from the earliest stages of life.

In summary, this protocol describes two complementary experimental procedures allowing for: (1) administration of molecules of interest in the early neonatal period to mouse pups (step 1); (2) assessment of both para and transcellular permeability in intestinal tract segments from neonatal to weaning pups using Ussing chambers (steps 2-6).

All animal procedures were conducted in accordance with the applicable national regulations and approved by the French Ministry of Higher Education, Research and Innovation via the APAFIS platform (Authorization number: APAFIS#23855) and grants from the Agence Nationale pour la Recherche (ANR): 11-IDEX-0005-02, ANR-24-CE15-7152, and 18-IDEX-0001. The reagents and the equipment used are listed in the Table of Materials.

1. Gavage of neonatal mouse pups

NOTE: The following method is adapted from Francis et al.¹². While the original protocol permits gavage of mouse pups as early as DOL 2, gavage was not performed before DOL 6 here, as the technical demands at earlier ages make the procedure considerably more challenging. In addition, using younger pups complicates studies as it increases the risk of animal loss during the first days of treatment, thus requiring larger cohorts.

1. Gavage solutions prep and setup
   1. Weigh each pup individually.
   2. Adjust the concentration and volume of the gavage solution so that the total volume administered does not exceed 20 µL per gram of body weight.
      NOTE: Under standard conditions, mouse pups rarely weigh less than 2.5 g at DOL 6.
   3. Wash a 24-G round-tip feeding needle by flushing it thoroughly with deionized water, followed by 70% ethanol using a syringe, then autoclave it.
      NOTE: It is recommended to use a separate feeding needle for each treatment group to prevent cross-contamination.
   4. Place clean paper towels on a disinfected heating pad set to ~38 °C.
   5. Wash gloves with 70% ethanol prior to opening the animal cages to limit odor transfer to the pups and reduce the risk of cannibalization.
   6. Transfer the dam to a separate cage in another room to limit stress caused by the pups' possible vocalizations.
   7. Rub cleaned gloves with nesting materials to limit the risk of transfer of external odors to the pups during gavage.
   8. Measure the length between the xiphoid process (lower end of the sternum) and the snout of a pup to mark the maximum insertion length on the feeding needle (Figure 1A,B).
2. Intra-esophageal gavage of mouse pups
   1. Attach the head of the needle to a syringe in a sterile manner and draw more than the desired volume of gavage solution to administer the entirety of the required volume without any risk of administering air bubbles, as they dramatically increase the risk of aspiration.
   2. Using the thumb and index of the non-dominant hand, gently pinch the skin between the scapulas to lift the pup.
      NOTE: Do not draw too much skin from the pup's back, as this may induce suffocation.
   3. Hold the pup in a near-horizontal position and gently insert the feeding needle into the oral cavity, advancing it perpendicular to the pharynx until it reaches the back of the throat (Figure 1C).
   4. While keeping the same angle and depth, slide the needle to the right or left part of the oral cavity to avoid the tongue.
   5. Slowly adjust the angle of the needle while gently tilting the pup's head backward, until the syringe, head, and back are aligned, with the head slightly inclined toward the back (Figure 1D).
   6. Without applying any pressure, allow the syringe to descend slowly under its own weight until the maximum insertion mark is close to or reaches the snout level (Figure 1E).
      NOTE: If resistance is encountered, gently rotate the syringe by rolling it back and forth between the fingers to facilitate descent. If the syringe still does not advance, try adjusting the insertion angle or restart the procedure using a feeding needle lubricated with the gavage solution. The needle shouldn't remain inserted for more than 20 s to avoid asphyxiation.
   7. Once the maximum insertion mark is reached or close, quickly dispense the desired volume. If resistance is met whilst dispensing the volume, check the syringe-head-back alignment and slightly adjust the depth of insertion.
      NOTE: Quick dispensing of the volume is recommended, as prolonged needle insertion significantly increases the risk of suffocation, struggling, and potential injury to the animal.
   8. Once the correct volume is administered, gently withdraw the needle along the same angle used for insertion.
   9. Place the pup on the paper towel previously placed on the heating pad and monitor recovery. A normal breathing pattern should come back in less than 20 s.
      NOTE: Apparition of bubbles from the snoot or mouth of the animal indicates aspiration and requires immediate euthanasia in pups younger than DOL 14. The presence of a very small amount of blood in the mouth of the animal caused by a lesion between the oropharynx and esophagus requires monitoring of the animal, but not systematic euthanasia. If pup recovery is uncertain, place it on its side. If it does not attempt to right itself, euthanize immediately.
   10. Once recovery of the animal is confirmed, place the pup back in the nest and regroup it with its dam.
       NOTE: Up to DOL 10, pups have low mobility and can remain on the heating pad until gavage of every pup in the cage is completed. Older pups must not be left unattended on the pad, as they may try to escape.

Figure 1: Gavage of neonate mouse pup at DOL 10. The distance between the snout and xyphoid process is measured (A), allowing for marking of maximal insertion length on the feeding needle (B). The feeding needle is inserted into the left or right part of the oral cavity (C). Once the back of the throat is reached, the angle of the needle is adjusted until the syringe, head, and back of the animal are aligned (D). The needle is allowed to gently slide downwards without resistance until the maximum insertion mark is nearly reached or reached (E). Please click here to view a larger version of this figure.

2. Neonatal mouse pup euthanasia and colon retrieval

1. Perform euthanasia using the appropriate method mandated by national legislation.
   NOTE: Decapitation of pups up to DOL 10 and cervical dislocation of pups of age superior to DOL 10 is recommended. Euthanasia with CO₂ or anesthetics is not advised, as pups display a large tolerance to these methods.
2. Place the euthanized animal under a binocular microscope.
3. Apply 70% ethanol to the abdomen of the animal.
4. Using forceps, pinch the abdominal skin and underlying peritoneum and make a large transverse abdominal incision using fine scissors (perpendicular to the rostral-caudal axis) without cutting the intestine.
5. Gently insert the fine scissors into the incision and extend the cut laterally through the peritoneum on both sides.
6. Using forceps and fine scissors, cut both superior and inferior parts of the peritoneum open, from the thoracic region to the lower pelvic area (Figure 2A).
7. Identify the distal part of the colon and make an incision parallel to its axis, towards the rectum.
8. Make an incision in the terminal/rectal portion of the colon to mobilize it.
9. Gently pull the colon and progress towards the cecum, removing potential pieces of connective or fat tissue.
10. Free the colon from the abdomen by cutting at the ileocecal junction, above the cecum (Figure 2B).
11. Discard the cecum and the initial few millimeters of the proximal colon, then collect one to two samples from the proximal colon that are larger than the aperture size of the Ussing slider (Figure 2C).
    NOTE: Securing two over-sized biopsies is advised, as the neonatal colon is extremely fragile and can easily be torn, lacerated, or pierced during cleaning and mounting.
12. With the scissors closed, gently remove the fecal content from the flattened biopsy by applying light pressure and sliding from the proximal to the distal end.
13. Place the emptied colon piece in ice-cold KRB buffer.

Figure 2: Dissection of the colon of DOL 10 pup. The epidermis and peritoneum of the pup are cut and opened in two steps to avoid unvoluntary incision of the colon, until the totality of the abdominal cavity is exposed (A). Colon is freed from the abdomen through a first cut of the distal/rectal section, followed by a second cut above the cecum (B). The initial millimeters of proximal colon are discarded, and one to two colonic samples larger than the aperture size of the Ussing sliders are collected (C). Please click here to view a larger version of this figure.

3. Colonic permeability assay using mouse biopsies in Ussing chambers

1. Preparation of the assay
   1. On the day prior to the day of the Ussing experiment. To ensure optimal assay conditions, preparation of several solutions on the day before the experiment is recommended. These solutions are prepared as instructed in Table 1.
      NOTE: the following tables indicate the required amounts for 12 Ussing chambers, with KRB 10x in large excess to allow aliquoting.
      NOTE: The KRB 10x solution can be stored at 4 °C for up to 6-12 months.
   2. On experiment day, 30 min prior to animals' euthanasia:
      1. Prepare 400 mL of KRB buffer, as instructed in Table 2.
         NOTE: The KRB 10x, deionized water, and bicarbonate must be pre-mixed and bubbled using carbogen gas (95% O₂; 5% CO₂) for 15 min before adding CaCl₂ to avoid precipitation.
      2. Using a carbogen system (95% O₂; 5% CO₂), bubble the fresh KRB buffer for 15 min.
      3. Split the 400 mL of KRB buffer into two and prepare 200 mL of Krebs-Glucose, 200 mL of Krebs-Mannitol, and the HRP solution as described in Table 3.
   3. Ussing system setup
      1. Pre-heat the Ussing System to 37 °C.
      2. Pre-assemble the chambers as follows:
         1. Lift the latches of each slot.
         2. Place the chambers in their slots without tightening the system.
         3. Insert the two gas-supplying tubes into each part of the chambers.
         4. Introduce the four electrodes in the front openings (if available - not covered here) or seal the four openings using 200 µL cones filled with glue ahead of the experiment.
         5. Place trays under the chambers in anticipation of potential leaks.
      3. Tissue mounting
         1. Lightly dry the previously cleaned colon sample with a paper towel to remove excess moisture.
         2. Place the piece of colon under a magnifying glass and very gently insert one of the blades of fine scissors in the luminal part to cut it open (Figure 3A).
            NOTE: The cut can be made in one movement once the scissors are fully inserted. The colon section used must be as straight as possible while avoiding torsion.
         3. Using fine forceps, mount the tissue on the white part of the slider while keeping the luminal part upwards, toward the operator. The tissue must be devoid of traces of fat, feces, wrinkles, perforations, and lacerations, and must widely cover the aperture (Figure 3B).
         4. Close the slider under the binocular microscope and check for potential displacement of the tissue. A slight deformation of the tissue must be visible when pressure is applied to the slider, which indicates that the slider is tightly closed.
            NOTE: Install the sliders in the Ussing chamber (Figure 4) and start the equilibration step individually as soon as it is ready, to prevent sample dehydration.
         5. While maintaining light pressure on the slider to keep it closed, insert the slider into the Ussing chamber, ensuring that the lumen (transparent part of the slider) faces to the left.
         6. Close both latches and tighten the screw located on the right until the chamber is airtight and the slider cannot move.
      4. Final setup and balancing
         1. Add 4 mL of Krebs-Mannitol to the left (apical) compartment of the chamber.
         2. Add 4 mL of Krebs-Glucose to the right (basolateral) compartment of the chamber.
         3. Place a small piece of parafilm between both compartments to avoid cross-contamination.
         4. Open the gas intake (95% O₂ / 5% CO₂) and set the output between 1-2 bubbles/s.
         5. Eliminate any large bubbles located on the tissue or in the system using a Pasteur pipette.
         6. Let the system balance for up to 20-30 min.
         7. Prepare the FITC 4kDa + HRP + Mannitol solution as described in Table 4.
2. Starting the colonic permeability assay
   1. Once equilibrium of the system is reached, replace 200 µL of Krebs-Mannitol from the apical compartment (left) with 200 µL of FITC 4Kd + HRP + Mannitol solution and immediately start a timer. Wash the tip by doing gentle ups and downs in the compartment and discard the tip between each chamber.
   2. For each chamber, collect 2 x 100 µL (duplicates) from the basal (right) compartment and deposit the volumes in a black 96-well plate. Cover the plate with aluminum foil to avoid photodegradation of the FITC present in the plate.
   3. Immediately replace the 200 µL previously collected with 200 µL of fresh Krebs-Glucose.
   4. Repeat the steps (2) and (3) every 30 min until the 2 h time-point is reached, to obtain the time-points: T0, T30, T60, T90 and T120.
   5. Seal the plate with aluminum foil to avoid photodegradation of the FITC and proceed to FITC quantification.

Table 1: Solution preparation on the day prior to the Ussing experiment.

Table 2: KRB buffer preparation on the day of the Ussing experiment.

Table 3: Preparation of Krebs-glucose/mannitol and HRP solutions on the day of the Ussing experiment.

Table 4: Preparation of FITC 4 kDa + HRP + mannitol solution on the day of the Ussing experiment.

Figure 3: Mounting of a colon sample from a DOL 9 pup. One blade of a fine pair of scissors is inserted in the luminal segment of the sample to cut it in one movement (A). The opened sample is mounted on the Ussing slider aperture with the luminal side facing towards the operator (B). Please click here to view a larger version of this figure.

Figure 4: Picture of fully prepared Ussing chambers. Please click here to view a larger version of this figure.

4. Colonic para-cellular permeability assessment through FITC quantification

1. FITC Standards preparation
   1. Dilute the FITC 4Kd + HRP + Mannitol solution in Krebs-Glucose to prepare the standard dilutions as described in Table 5 and Figure 5.
   2. Deposit 2 x 100 µL (duplicate) of each standard dilution from n°1 through n°7 in the plate. Seal the plate with aluminum foil to protect from light until plate reading begins.
2. Data acquisition: Using a plate reader, set the excitation wavelength to 485 nm and emission to 535 nm, and read the plate as soon as possible.
   NOTE: Reading the plate at both optimal gain and with a manually set gain is advised to facilitate inter-plate values comparison and allow for interpolation of data if a mistake was made during standards dilutions.
3. Plate storage: Once FITC quantification data has been stored, store the plates sealed with aluminum foil at 4 °C overnight if HRP quantification is to be conducted in the following days, or frozen at -80 °C for longer storage.

Figure 5: Visual representation of FITC standard dilutions preparation. Please click here to view a larger version of this figure.

Table 5: Preparation of FITC standard dilutions on the day of the Ussing experiment.

5. Colonic trans-cellular permeability assessment through HRP quantification

1. On the day prior to the day of the HRP quantification assay:
   1. Ensure optimal conditions on the day of the assay; preparation of several solutions is required on the day prior to the experiment. These solutions are prepared as described in Table 6.
      ​NOTE: The following tables indicate the required amounts for one 96-well plate.HRP Standards can be aliquoted and stored at -20 °C or -80 °C for up to several months. Due to instability, use only fresh carbonate/bicarbonate buffers.
   2. Upon Anti-HRP antibody solution preparation, immediately coat a Clear 96-well Clear Flat Bottom High Binding microplate with 100 µL per well. Wrap it in aluminum foil and incubate overnight at 4 °C.
2. On the day of the HRP quantification assay:
   1. Prepare the 5% BSA Blocking buffer as described in Table 7.
   2. Discard the contents of the plate and wash with 280 µL of PBS Tween 4 times with 1 min of shaking at 350 rpm between each wash.
   3. Add 200 µL of BSA blocking buffer to each well.
   4. Seal the plate with aluminum foil and incubate on a shaker at 500 rpm for a minimum of 1 h. In the meantime, thaw and dilute samples (1:10 or 1:50) and prepare the standards dilutions as described in Table 8.
   5. Discard the contents of the plate and wash with 280 µL of PBS Tween 4 times with 1 min of shaking at 350 rpm between each wash.
   6. Add 50 µL of standard or diluted samples, seal the plate, and incubate at 300 rpm for 1 h at RT.
   7. During incubation, bring the QuantaBlu reagents to RT and prepare the working solution as described in Table 9.
   8. Discard the contents of the plate and wash with 280 µL of PBS Tween 4 times with 1 min of shaking at 350 rpm between each wash.
   9. Add 100 µL of QuantaBlu Working Solution to each well at RT or 37 °C (1.5-90 min, 30 min on average).
      NOTE: Assay progress can be followed by the appearance of a strong blue fluorescence under a UV light source.
   10. Add 100 µL of QuantaBlu Stop Solution to each well to stop the peroxidase activity. Seal plate and incubate for 10 min at RT under agitation at 350 rpm.
   11. Using a plate reader, set the excitation wavelength to 315 nm and emission to 470 nm. Read the plate as soon as possible.
       ​NOTE: Reading the plate at both optimal gain and with a manually set gain is advised to facilitate inter-plate values comparison and allow for interpolation of data if a mistake was made during standards dilutions. Alternatively to the QuantaBlu Fluorogenic Peroxidase Substrate kit, other peroxidase substrates such as Tetramethylbenzidine can be used (measured at 450 or 650 nm with or without the use of stop solution, respectively), but use of this alternative comes at the cost of lower sensitivity.

Table 6: Solution preparation on the day prior to the HRP experiment.

Table 7: BSA blocking buffer preparation on the day of the HRP experiment.

Table 8: HRP standard preparation on the day of the HRP experiment.

Table 9: QuantaBlu working solution preparation on the day of the HRP experiment.

Standard curve and well concentration quantification
The average value of blanks was subtracted from every well of the plate. With absorbance values on the Y axis and concentrations on the X axis, a standard curve was created using the "Interpolate a standard curve" function from GraphPad Prism. If all standard points are aligned, the "Line" model provides the best results. Alternatively, the "Sigmoidal, 4 PL, X is concentration" model offers great flexibility and accuracy with non-linear standard curve interpolation (Figure 6). The absorbance values of samples on the Y axis were pasted below the standard's and recovered the concentrations from the "Interpolated X mean values" from the results tab (Figure 6).

Figure 6: FITC Standard curve and samples concentration interpolation. Please click here to view a larger version of this figure.

Data analysis
Starting from T0+1, each previous time-point(T-1) was subtracted from the current time-point(T) to calculate Δ-FITC or Δ-HRP for each interval: T30-T0, T60-T30, T90-T60, and T120-T90. Using the following equation, the flux for each sample was calculated at each time point, where J is the flux, ΔC is the change in concentration between the two time points, V is the volume of the basal compartment, ΔT is the time interval between measurements, and A is the exposed surface area.

Graph each individual value per ΔT to identify potential anomalies or outliers.

The results are displayed as the average evolution of Colonic Flux of FITC-Dx or HRP of each group for each time window (Figure 7).

Figure 7: Colonic FITC-Dx 4kDa Flux (nmol.min-1.cm-2) values across different time-windows of the Ussing chamber experiment conducted on 24 colonic samples from pups. 8 pups per group were treated for several days through oral gavage with either PBS, miR-A or miR-B. Please click here to view a larger version of this figure.

However, it is worth noting that several factors may vary between samples and can impact the precision of the assay, including the time between euthanasia and tissue mounting, the duration spent for system equilibrium, the presence or movement of air bubbles on or close to the biopsy, and cell death. Checking for anomalies in the evolution of concentrations or flux remains essential. Due to varying incubation times between the first and last mounted sliders, plus notable cell death toward the assay's end, T0 and T120 values are sometimes discarded. Sudden drops or increases in flux values also warrant attention, as flux increases over time-points should stay stable. For instance, prior testing showed that adding 10 mM EDTA to one colonic sample at T0 caused a flux rise from 97% to 64% across time-points compared to controls, yet the overall flux evolution mirrored that of untreated samples (Figure 8).

Figure 8: Colonic FITC-Dx 4kDa Flux (nmol.min-1.cm-2) values across different time-windows of the Ussing chamber experiment conducted on 6 colonic samples from pups. Samples #1 through #5 serve as controls, with sample #6 receiving 10 mM EDTA at T0. Please click here to view a larger version of this figure.

Oral gavage enables repeated administration of a precise, defined luminal dose to individual neonatal mice. In contrast, voluntary feeding, although a viable alternative, requires extended administration time per animal to deliver a lower compound quantity with reduced dosing precision¹². Moreover, oral gavage accommodates larger volumes than those feasible via injectable routes, facilitating higher total doses at lower concentrations and thereby minimizing potential toxicity upon administration¹³. Transit through the digestive tract further restricts the compound's distribution volume, which is advantageous for studies targeting intestinal permeability and barrier function. However, toxicity varies substantially with developmental window and molecular class; therefore, caution must be applied when administering any treatment to neonatal subjects for the first time. The gavage of neonatal pups is a delicate procedure that requires training, particularly for pups under DOL 10. Training with PBS containing dark food coloring is advised, with successful gavage inducing a visible darkening of the pups' stomach. Apparition of coloring outside of the stomach, especially around the throat or thoracic region, is a sign of failed gavage and should result in immediate euthanasia of the pup. Additionally, gavage should not be conducted under anesthesia, as swallowing of the feeding needle is required for correct administration of the solution and limits the risks of damage to the upper esophageal sphincter and aspiration¹².

The ex vivo paracellular and transcellular permeability assays with Ussing chambers are very sensitive methods, and the successful completion of the assay is reliant on many parameters^14,15,16. Amongst them, dissection and mounting of the tissues remain the most common causes of failure. Despite showing great elasticity, colon dissection should be conducted quickly while minimizing physical stress to the tissue, as excessive extension can induce micro-tears. Removal of connective tissue and fat from the colon should be performed with care, as excessive traction on adipose and connective tissues can compromise the structural integrity of the sample¹⁵. Upon collection of the colonic biopsy, immediate removal of the feces is recommended, as fully hydrated stools are more prone to disruption, making sample cleaning more difficult. Prompt removal of feces also minimizes friction on the luminal surface and improves preservation of the epithelium.

Aside from removing fecal material and avoiding excessive disruption of the mucus layer, excision of the serosal-muscle layer (SML) from biopsies is not recommended in this protocol. The necessity of removing the muscle layer, as described by Hempstock, Ishizuka, and Hayashi (2021)¹⁷ is still debated among the scientific community. Conventionally, the SML is considered to impair oxygenation and nutrient delivery to the epithelium and to favor the accumulation of toxic metabolites during the assay¹⁵. While SML removal has been reported to lower transepithelial electrical resistance (TER) and prevent tissue shortening due to muscle contraction, Sjögren et al.¹⁸ found no significant effect on drug permeability in rat jejunum when comparing stripped and unstripped preparations. Moreover, several of their stripped samples failed to satisfy quality criteria, whereas all native tissue segments were deemed suitable for experimentation, suggesting that the stripping procedure itself may negatively impact tissue integrity. For these reasons and given the particularly fragile nature of mouse pup biopsies, SML removal is not advised here, as it proves impossible on samples from young pups, and would substantially increase the number of biopsies required and consequently the number of older animals used.

Mounting the tissue is another critical step requiring finesse. Different types of sliders are available to accommodate varying tissue sizes. Larger sliders are typically equipped with pins, which prevent tissue retraction and associated wrinkling while ensuring that the entire aperture is covered. In contrast, smaller sliders are usually pinless, as the use of pins in this context would greatly increase the risk of tearing thin tissues. The choice of the slider model also influences assay precision, as biopsies that match the aperture width too closely are more prone to leakage. Colonic biopsies from the proximal colon of DOL 10 pups were large enough for sliders with an aperture size of 2.8 x 1.5 mm (0.04 cm²), while samples from DOL 7 pups were too small to fit the aperture. However, more distal colon samples from DOL 7 and younger pups had a greater width and were compatible with the aperture size. Small intestine samples from DOL 10 pups also proved significantly smaller than colon samples and thus too small to fit the sliders. Sliders with smaller apertures are available for use with pups under DOL 10 and smaller segments of the digestive tract. However, the minimal age required to obtain reliable permeability measurements remains unclear. Limited data are available on gut permeability in mouse pups at or below postnatal day 10 (DOL 10). Some studies suggest that intestinal permeability may be similar to, or even higher than, that observed at DOL 10^19,20,21, which would make detecting modest effects particularly challenging. Furthermore, it remains uncertain what minimal colonic surface area is required to reliably detect differences under treatments with moderate effects. Previous experiments yielded satisfactory and significant results using aperture surfaces of 0.04 cm² down to 0.031 cm². However, smaller surfaces can substantially diminish assay sensitivity and amplify biases from minor variations in tissue handling, particularly during slider mounting and dissection. Assigning a single operator for all dissections and mounting within each experiment helps eliminate inter-operator technique variations. Employing specialized equipment, such as user-friendly sliders and the dissection instruments outlined in this article, enhances reproducibility, shortens handling time, and reduces tissue damage risk.

Using the same colonic segment within and across experiments is also recommended, as it is essential for obtaining comparable permeability results. Note that biopsies taken from different regions of the same animal can yield markedly different values. The time required to mount all chambers is also a critical parameter. Although this issue is minor in small setups with only a few chambers, the interval between euthanasia and assay initiation, as well as the overall incubation time, can become significantly prolonged if the operator waits to start until all chambers are prepared. To mitigate this, one option is to mount the first sliders and hold them in the Krebs buffer at 4 °C, then insert all sliders simultaneously to begin the assay. Complementarily, TER assessment is a reliable method to determine viability of the tissue, as TER tends to decrease with cell death, ~120-150 min after colon sampling¹⁶.

Alternatively to the Ussing technique, the in vivo FITC-dextran assay also enables evaluation of global intestinal permeability through dosage of serum FITC after gavage, it neither discriminates between paracellular and transcellular pathways nor localizes changes to specific gut segments, making the Ussing chamber approach more appropriate for this study¹⁴. Complementarily, contrary to the in vivo-FITC, which generally requires fasting, the latter is not necessary for colonic permeability assays. Intestinal permeability in the small intestine is strongly influenced by fasting status and the interval since the last meal, but such effects have not been demonstrated in the colon. Moreover, fasting pups would require maternal separation, and severe stressors such as neonatal maternal deprivation are known to exert long-lasting effects on paracellular permeability and mucosal immunity. Although the minimum duration required to induce these effects has not been clearly defined, it is preferable to avoid stressing the pups prior to euthanasia. It is also important to note that the gavaged-FITC plasma method is incompatible with this protocol, as residual FITC from previous experiments could confound permeability measurements obtained with the Ussing chambers.

Finally, the presence or absence of colonic permeability modulation may be confirmed using an independent method, such as qPCR-based quantification of tight junction-related transcripts, including ZO-1, occludin, and members of the claudin family. Overall, the methods described in this article offer complementary tools for investigating the neonatal period, particularly the maturation of the early-life intestinal barrier and its susceptibility to various interventions. They can generate insights that are highly relevant for studies on the physiological and pathophysiological mechanisms underlying early-life infections such as necrotizing enterocolitis and complications of prematurity. In addition, they are well-suited to address more fundamental questions, including the impact of human milk oligosaccharides, microRNAs, and probiotics on gut development and function.

In summary, oral gavage of molecules of interest during the neonatal period, combined with assessment of both paracellular and transcellular colonic permeability using Ussing chambers, provides an efficient approach to investigating the short-term impact of potentially harmful substances, as well as their short- and long-term effects, particularly in the context of chronic inflammatory diseases.