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. Author manuscript; available in PMC: 2023 Jan 12.
Published in final edited form as: Methods Cell Biol. 2022 Jan 12;168:235–247. doi: 10.1016/bs.mcb.2021.12.017

Methods for Induction and Assessment of Intestinal Permeability in Rodent Models of Radiation Injury

Laura E Ewing *, Prabath G Biju †, Rupak Pathak ǂ, Stepan Melnyk §, Martin Hauer-Jensen ǂ, Igor Koturbash ¶,‡,1
PMCID: PMC9808921  NIHMSID: NIHMS1855684  PMID: 35366985

Abstract

Ionizing radiation (IR) is a significant contributor to the contemporary market of energy production and an important diagnostic and treatment modality. Besides having numerous useful applications, it is also a ubiquitous environmental stressor and a potent genotoxic and epigenotoxic agent, capable of causing substantial damage to organs and tissues of living organisms. The gastrointestinal (GI) tract is highly sensitive to IR. This problem is further compounded by the fact that there is no FDA-approved medication to mitigate acute radiation-induced GI syndrome. Therefore, establishing the animal model for studying IR-induced GI-injury is crucially important to understand the harmful consequences of intestinal radiation damage. Here, we discuss two different animal models of IR-induced acute gastrointestinal syndrome and two separate methods for measuring the magnitude of intestinal radiation damage.

Keywords: abdominal irradiation, intestinal permeability, leaky gut, radiation injury, rodent model

1. Introduction

Ionizing radiation (IR), a potent genotoxic and epigenotoxic agent, is a ubiquitous environmental stressor, a significant contributor to the contemporary market of energy production, and an important diagnostic and treatment modality (Miousse et al., 2017). Exposure to high doses of IR, either as a result of accidental exposures (i.e., catastrophic events or terroristic attacks) or during cancer therapy may result in a number of negative health effects. Among the latter, the effects to the gastrointestinal (GI) tract present the gravest concerns because of its high sensitivity to IR (Hauer-Jensen et al., 2014). This concern is further compounded by the fact that there are currently no effective countermeasures or mitigators to target radiation-induced gastrointestinal toxicity.

In these regards, it is imperative to understand the mechanisms underlying GI radiation-induced toxicity, as understanding the exact mechanisms will expedite the development of effective countermeasures. Intestinal permeability is an important physiological aspect of the GI function that often exerts whole organismal effects. Indeed, impaired intestinal permeability has been documented in numerous diseases, such as Crohn’s disease, inflammatory bowel disease, irritable bowel syndrome, a number of autoimmune diseases, and even diabetes and asthma (Turner, 2009). Exposure to IR is known to negatively regulate intestinal permeability, often leading to the development of a “leaky gut syndrome.” The latter is characterized by bacterial, bacterial remnant, and lipopolysaccharide (LPS) translocation into the blood stream and the loss of fluids and electrolytes into the gut lumen.

Mechanistically, radiation-induced disruption of intestinal permeability is a multifactorial process associated with direct damage to intestinal mucosa (with particular emphasis on the mucosal stem cell compartment that is critical to intestinal wall renewal) and disruption of dynamic tight junction structures that exist and function between intestinal epithelial cells. Radiation-induced alterations in the gut ecology, with increases in abundance of pathogenic bacterial species at the expense of commensal bacteria, can often trigger pro-inflammatory signaling in the gut and further exacerbate the manifestation of “leaky gut syndrome.”

Here, we will discuss two mouse models of radiation-induced gastrointestinal toxicity that we use in our laboratories, as well as two approaches to evaluate post-exposure intestinal permeability.

2. Models of Radiation-Induced Gastrointestinal Toxicity

Two most popular approaches of radiation-induced intestinal toxicity are: 1) total-body irradiation with hind-leg shielding and 2) targeted abdominal irradiation with an image-guided irradiator. Both approaches inflict significant intestinal damage without causing substantial injury to the bone marrow. At UAMS, we use a J. L. Shepherd Mark I model 25 137Cs irradiator (J. L. Shepherd & Associates, San Fernando, CA) for the total-body irradiation with hind-leg shielding, while the Small Animal Radiation Research Platform or SARRP (Xstrahl, Camberly, Surrey, GB) is used for targeted abdominal irradiation. The first approach allows to irradiate multiple (4 mice) unanesthetized mice, while only one anesthetized mouse can be irradiated with SARRP.

2.1. Total-body irradiation with hind-leg shielding

Hind-limb protection by medical-grade tungsten blocks can effectively spare ~5% to 10% of the mouse bone marrow, thus preventing the development of acute radiation-induced hematopoietic syndrome. Therefore, exposures at doses below those that cause acute neurotoxic effects would allow for the study of acute radiation toxicity with prevalence of possible GI syndromes. For instance, exposure of CBA/CaJ mice at a dose of 8.5 Gy under certain conditions (i. e., dietary modulation) is ~LD100/30 (100% mortality that occurs within 30 days of irradiation) for this strain; however, hind-limb protection decreases the mortality to levels of less than 10% (Miousse et al., 2020) (Notes 1 and 2).

  1. Irradiation procedures are performed in a special custom-made radiation jig with un-anesthetized mice in a Plexiglas restrainer with the animal’s hind limbs shielded by a tungsten block (Figure 1) (Note 3).

  2. Both hind limbs of the mouse are carefully pulled out through the holes located at the rear part of Plexiglas restrainer using blunt tip tweezers. To keep the hind limbs in place, a medical grade adhesive tape should be used, which will allow air to reach the skin during irradiation.

  3. A groove at the base of the tungsten block is placed carefully over the taped hind limbs, ensuring that it would not exert any pressure on the hind limbs, but will simultaneously effectively attenuate radiation exposure. The tungsten block is secured using rubber bands.

  4. Usually, four animals can be exposed in the irradiation chamber at a given time. It is important that the direction of the radiation beam be perpendicular to the orientation of the animals, so that uniform doses can be delivered over the exposed body parts.

  5. After exposure to the desired dose, the animals are returned to their respective cages (initial cages where they were housed prior to irradiation). This is particularly important in regards to male mice that can exhibit strong territorial instincts and, if not housed together previously, will result in aggressive behavior. IR is known to further exacerbate aggression in male mice.

  6. !NB! It is critical to perform systemic and well-documented monitoring of animals after irradiation. If an animal is found in the moribund condition, or exhibiting any signs of distress, including but not limited to loss of appetite, lethargy, huddling, shivering, severe diarrhea, vocalization, or loss of 20% body weight, an attending veterinarian should be notified immediately to take appropriate actions. For further details, please, refer to Note 4.

Figure 1.

Figure 1.

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The method of partial body irradiation: bone marrow shielding approach. The mice are placed in custom-made Plexiglas restrainers, the hind-legs are covered by tungsten blocks, and the tungsten blocks are secured with rubber bands.

2.2. Targeted abdominal irradiation with an image-guided irradiator

As mentioned above, targeted delivery can be achieved with SARRP, thus avoiding radiation exposure to the parts of the body not in the radiation field. The steps involved in irradiation with SARPP are as follows.

  1. Mouse is anesthetized with 2–3% isoflurane by inhalation and is immediately placed onto a platform that connects to a robotic arm inside the irradiator.

  2. The imaging is performed at 60-kV and 0.4-mA filtered with aluminum for 2 min. Occasionally, these scans may take up to 5 minutes. This procedure will provide visualization of internal organs, allowing radiation to be targeted to a specific region (in this case, to selected abdominal area). During the scan, an animal will receive a negligible dose of IR.

  3. For abdominal irradiation, the imaging is performed by Muriplan (Xstrahl), the preclinical treatment planning system. It is used to specifically target the beam using the last rib as a cranial limit marker and the spinal cord as a dorsal limit marker to ensure all mice will be irradiated in approximately the same location.

  4. For the development of acute radiation-induced gastrointestinal syndrome, irradiation procedures are performed with the gantry at 90° using a 10×10 mm collimator, delivering 220-kV and 13 mA X-ray with a single dose of between 12.5 and 15 Gy (depending on the mouse strain) (Please, consult Note 2). The control mice are usually anesthetized and placed inside the SARRP, but not exposed to radiation.

  5. The entire procedure takes ~15 minutes of total anesthesia time.

  6. After irradiation, the animals are first placed in a recovery cage until they have recovered from the anesthesia. Mice are then returned to their original cages and should be monitored daily for health and well-being (please refer to points 5 and 6 in Section 2.1. and Note 4).

3. Evaluation of Intestinal Permeability

Here, we will provide the details on two distinct approaches that can be used for the analysis of intestinal permeability in both of the above-discussed animal models. The lactulose/mannitol assay is a non-invasive and generally atraumatic approach that allows for continuous analysis of the gut integrity from the same mouse in dynamics. Although it can be used in the same settings as the lactulose/mannitol assay, the so-called fluorescein isothiocyanate (FITC)/dextran 4000 assay, when used in conjunction with a non-survival surgery, can serve as a more sensitive approach for the analysis of intestinal permeability after radiation exposure.

3.1. Lactulose/Mannitol Assay

The lactulose/mannitol assay is the most frequently utilized assay for the analysis of intestinal permeability in both human and experimental animals, including mice (Grootjans et al., 2010; Volynets et al., 2016).This assays considers collection of mouse urine after administration of a lactulose/mannitol cocktail with further processing of urine samples using liquid chromatography – mass spectrometry (LC-MS). It is considered the most recommended assay for the analysis of small-intestinal permeability in both humans and experimental rodent models (Anderson et al., 2004; Meddings and Gibbons, 1998; Volynets et al., 2016).

3.1.1. Urine Sample Collection

  1. Mice are individually housed in metabolic cages. Metabolic cages allow for the collection of urine prior to and after lactulose/mannitol administration.

  2. The urine collection cups contain 10% thymol (Millipore-Sigma, St. Louis, MO, USA) in isopropyl alcohol as a preservative, and paraffin oil (Millipore-Sigma, St. Louis, MO, USA) to prevent evaporation (Arrieta et al., 2009; Kish et al., 2013).

  3. Mice are fasted for five to twelve hours in the metabolic cages, but allowed unlimited access to water, prior to lactulose/mannitol administration.

  4. Pre-gavage urine samples are collected from the urine cups, pipetted into 1.5 mL microcentrifuge tubes, and then centrifuged at 2000 × g at 4°C for 15 min. The urine layer is then pipetted into new 1.5 mL microcentrifuge tubes and flash frozen in liquid nitrogen prior to storage at −80°C.

  5. New urine collection cups, also containing 10% thymol and paraffin oil, are placed under the metabolic cages just prior to gavage.

  6. Lactulose and mannitol are orally gavaged simultaneously in a single solution containing 20 mg/mL of lactulose (Millipore-Sigma, St. Louis, MO, USA) and 50 mg/mL of mannitol (MP Biomedicals, Los Angeles, CA, USA) in 250 μL of sterile water (5 mg and 12.5 mg of lactulose and mannitol, respectively, per mouse).

  7. Fasting continues for one hour after gavage, and then the standard chow is supplied ad libitum.

  8. Mice are kept in metabolic cages overnight (approximately 12 to 16 hours). The overnight urine samples are then processed as described above with the addition of calculating total urine output in order to determine total mannitol and lactulose that was excreted.

3.1.2. Liquid Chromatography-Mass Spectrometry (LC-MS) Procedure

  1. Prepare standard solution (1mg/ml) of mannitol and lactulose (both Sigma-Aldrich, St. Louis, MO, USA) to perform calibration and test solutions.

  2. Mix the thawed urine samples with acetonitrile at a 1:1 ratio for protein precipitation, vortex for 20 sec following 10 min centrifugation at 3000g.

  3. Filter the supernatant through 0.22 μm Nylon Centrifuge Tube Filters (Costar, Corning, NJ) and inject 10 μl for analysis.

  4. Mobile phase (A) acetonitrile (Sigma-Aldrich) and (B) water (LC-MS grate) are used at 85:15/100:0 (A:B), flow rate 1.6 ml/min, Accucore HILIC column 100 mm, I.D. 2.1 mm, 2.6 μm particle size (Thermo-Fisher, Waltham, MA) at +300C column heater set point and dual wavelength UV 190/260 nm detector setting.

  5. The samples are then eluted with an acetonitrile and water mixture (85:15) at a flow rate of 0.6 mL/min.

  6. UHPLC Ultimate 3000 and LC-MS detectors (Thermo-Fisher) equipped with mobile phase degasser, automated sampler and column heater are used.

  7. Calibration curves solutions of appropriate concentration were prepared in LC-MS grate water. Urinary concentration of mannitol and lactulose are calculated from the calibration curves by peak-height analysis. The limits of detection are the following: for lactulose – 0.1–25 μmol/L and for mannitol – 0.2–20 μmol/L.

  8. Recovery rates are calculated as the percent of the gavaged dose, based on total urine output. Permeability is assessed using the ratio of the recovered lactulose to recovered mannitol.

3.2. Fluorescein Isothiocyanate (FITC)/Dextran 4000 Assay

This approach has been used extensively in numerous rodent models, with some modifications, including models of radiation-induced GI toxicity (Biju et al., 2012; Cani et al., 2008). Furthermore, the FITC/dextran methodology has previously been recommended for global assessment of intestinal permeability in mice (Volynets et al., 2016).

  1. First, the mouse is placed in the chamber of isofluorane vaporizer and anesthetized with mixture of isoflurane (2%) and oxygen (2 LPM).

  2. After achieving the initial anesthesia, the mouse is taken out of the chamber and placed in supine position and kept anesthetized with isoflurane (2%) with a mask connected to the isofluorane vaporizer (Note 5).

  3. Midline laparotomy is performed.

  4. Renal artery and vein are isolated and ligated on both kidneys with 5–0 thread.

  5. A 10 cm segment of small intestine is isolated beginning at 5 cm distal to the Treitz ligament without damaging intestinal and mesenteric structures and ligated at both ends with 5–0 thread (as described in Figure 2).

  6. 0.1 ml of 4-kDa fluorescein isothiocyanate-conjugated dextran (FITC-dextran 25 mg/ml in phosphate–buffered saline; Sigma, catalog # FD4) is injected into the isolated intestine from the proximal end through a 30 gauge needle connected with a 1ml syringe.

  7. The injection site is sealed with tissue glue (3M Vetbond, catalog # 1469SB). Abdominal incision will be closed with 4–0 suture.

  8. The mouse is placed back into the chamber of isofluorane (2%) vaporizer with an isothermal pad on the bottom of the chamber to maintain animal body temperature.

  9. After 90 min of continuous isofluorane anesthesia, blood is collected from the portal vein. If unsuccessful, the mouse is anesthetized with Ketamine/Xylazine (50 mg/10mg/kg body weight, I.P.), and blood is collected by retro-orbital bleeding using heparanized microcapillary tube (Fisher Scientific, catalog # 22-362-566).

  10. The mouse is euthanized by cervical dislocation. Death is further ensured by dissection of thoracic section of aorta.

  11. Plasma is separated by centrifuging at 4°C, 8000 rpm for 10 min.

  12. The concentration of FITC-dextran is determined with a fluorescence spectrophotometer (Synergy HT, Bio-Tek Instruments, Winooski, VT) at an excitation wavelength of 480 nm and an emission wavelength of 520 nm.

  13. Standard curves to calculate FITC-dextran concentration in the plasma samples are prepared from dilutions of FITC-dextran in PBS.

Figure 2.

Figure 2.

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Schematic representation of the small intestine ligation surgery.

Notes

  1. Mouse strain and animal’s age are two important determinants in manifestation of acute radiation toxicity in mice. A number of strains, such as CBA/CaJ, are known to be considerably radiosensitive and may develop severe acute radiation syndrome and succumb at lower doses compared to more radioresistent strains, such as C57BL6/J (Miousse et al., 2020). Furthermore, besides the genotype, phenotypic peculiarities must be taken into consideration. For instance, obese NZO/HlLtJ mice exhibit a high degree of radioresistance at doses 7.4 Gy and below; exposure to higher doses, however, is characterized by high sensitivity with manifested GI syndrome (Ewing et al., 2020a).

    Age is another important determinant of sensitivity to IR exposure. It is generally accepted that juvenile animals are considerably more sensitive to IR due to higher rates of proliferation and higher number of proliferative cells; conversely, older animals (6 months and up) are more sensitive due to impaired immune function and repair capacity (Hudson et al., 2011; Kovalchuk et al., 2014). Per our observations, even 10–15 days in age difference may play a significant role, as 45 days-old animals were more sensitive to IR than 60 days-old animals (unpublished data). It is also important to highlight the differences in rodents’ sensitivity to radiation exposure associated with conditions in animal husbandries, circadian rhythms (early morning exposures usually result in more severe manifestations of acute radiation syndrome, possibly due to circadian rhythms in stem cells physiology) and even time of the year (mice exposed in the middle of summer usually manifest higher sensitivity to IR than those exposed in winter).

  2. Selection of appropriate doses and dose regimen are important aspects in the design of studies devoted to intestinal permeability. Higher doses delivered to selective areas of the body allow for more robust organ/tissue response and to avoid the whole-organismal response and increased mortality. Our partial irradiation model with bone marrow shielding approach (described in Section 2.1) allows us to deliver doses as high as 8.5–9 Gy without evidence of animal mortality, but with substantial damage to the gut and development of a “leaky gut syndrome.” Utilization of SARRP (described in Section 2.2.), allows to further decrease the surface of area of exposure and minimize the systemic damage. According to our experience, the doses between 12.5 and 15 Gy can be safely considered in this case, with about 50% and 75% loss in intestinal crypts respectively on day 4 after irradiation (Ewing et al., 2020b). However, mouse strain peculiarities and other factors that can influence radiosensitivity (discussed Note 1) have to be taken into consideration. Furthermore, even little changes in the diet may significantly alter the course of radiation-induced gastrointestinal syndrome. For instance, supraphysiological concentrations of methionine in the diet can cause mortality after total body irradiation at doses as low as 3 Gy and at 7.6 Gy after partial body irradiation with bone marrow shielding (Miousse et al., 2020).

  3. For the irradiation procedures described in Section 2.1., we use un-anesthetized animals. The reason for this is that anesthetics modulate various target proteins and may work as weak protectors of living organisms against IR (Merriam et al., 1968). Therefore, in order to avoid the potential influence of anesthetics in modulating the biological effects of IR, we suggest using un-anesthetized animals where possible. However, for successful irradiation procedures using SARRP, animals will need to be anesthetized.

  4. Development of radiation syndrome in mice may be associated with morbidity or mortality. Radiation syndrome is characterized by immune system impairment with bacteremia and, at higher doses, diarrhea with fluid loss through the gastrointestinal tract. In some cases, hemorrhage may develop because of thrombocytopenia. The time during which pain and distress from the radiation syndromes develop after the radiation doses discussed here is from around day 6 until around day 24. Therefore, special care has to be given during this window. Animals should be inspected diligently at least twice daily and animals that are moribund (more than 20% weight loss, lethargy, huddling and/or shivering, loss of appetite, hunched posture, severe diarrhea, and vocalization) must be euthanized without delay. Written records should be maintained for all the monitoring sessions. It must be kept in mind that pain relieving drugs or any other form of interventions may significantly alter the course of radiation syndrome.

  5. Surgical anesthesia is defined by the following: 1) reduced but regular respiratory rate and depth; 2) discontinuation of eye movement; 3) absence of lid and corneal reflexes; and 4) elimination of pain reflexes.

Table 1.

Ranges of lactulose and mannitol in the urine of control and exposed mice fed either methionine-adequate or methionine-supplemented diets.

Lactulose and Mannitol Urine Concentrations by Diet, Radiation, and Time after Radiation
Diet Radiation Days after Radiation Lactulose (μmol/mL) Mannitol (μmol/mL)
Adequate Methionine Sham 7 0.135–0.217 12.3–19.8
Adequate Methionine Sham 10 0.103–0.192 11.4–20.4
Adequate Methionine Local Abdominal 7 0.151–0.294 16.2–28.6
Adequate Methionine Local Abdominal 10 0.183–0.327 16.1–27.5
Supplemented Methionine Sham 7 0.181–0.267 12.8–25.3
Supplemented Methionine Sham 10 0.158–0.279 14.8–27.9
Supplemented Methionine Local Abdominal 7 0.245–0.396 18.5–31.4
Supplemented Methionine Local Abdominal 10 0.226–0.344 20.5–30.5
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Local abdominal radiation was performed with the Small Animal Radiation Research Platform. Mice were exposed to 12.5 Gy in a one cm transverse section just below the last rib and just ventral to the spine. Values given are ranges (n = 6 per diet, radiation, and days after radiation).

Table 2.

Ranges of lactulose and mannitol in the urine of control and exposed mice of different ages.

Lactulose and Mannitol Urine Concentrations by Age at Time of Radiation, Radiation, and Time after Radiation
Age (days) Radiation Days after Radiation Lactulose (μmol/mL) Mannitol (μmol/mL)
62 Sham 7 0.135–0.164 12.3–15.6
62 Local Abdominal 7 0.205–0.294 17.9–28.6
62 Sham 10 0.104–0.176 12.6–17.4
62 Local Abdominal 10 0.286–0.327 21.3–26.8
70 Sham 7 0.168–0.206 15.1–17.9
70 Local Abdominal 7 0.189–0.266 16.2–20.8
70 Sham 10 0.103–0.192 11.4–15.2
70 Local Abdominal 10 0.244–0.265 20.2–27.5
78 Sham 7 0.182–0.217 17.5–19.5
78 Local Abdominal 7 0.151–0.218 17.6–18.3
78 Sham 10 0.177–0.192 16.5–20.4
78 Local Abdominal 10 0.183–0.302 16.1–19.4
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Local abdominal radiation was performed with the Small Animal Radiation Research Platform. Mice were exposed to 12.5 Gy in a one cm transverse section just below the last rib and just ventral to the spine. Values given are in ranges (n = 4 per age, radiation, day after radiation).

Acknowledgements

Research reported in this publication was supported by an Institutional Development Award (IDeA) from the National Institute of General Medical Sciences of the National Institutes of Health under grant number 1P20GM109005; Clinical and Translational Science Awards UL1TR000039 and KL2TR000063; and the Arkansas Biosciences Institute. The authors are thankful to Christopher Fettes for editing this manuscript.

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