Murine Fecal Isolation and Microbiota Transplantation

Summary

The goal here is to outline a protocol to investigate the mechanisms of dysbiosis in cardiovascular disease. This paper discusses how to aseptically collect and transplant murine fecal samples, isolate intestines, and use the "Swiss-roll" method, followed by immunostaining techniques to interrogate changes in the gastrointestinal tract.

Abstract

Gut microbiota dysbiosis plays a role in the pathophysiology of cardiovascular and metabolic disorders, but the mechanisms are not well understood. Fecal microbiota transplantation (FMT) is a valuable approach to delineating a direct role of the total microbiota or isolated species in disease pathophysiology. It is a safe treatment option for patients with recurrent Clostridium difficile infection. Preclinical studies demonstrate that manipulating the gut microbiota is a useful tool to study the mechanistic link between dysbiosis and disease. Fecal microbiota transplantation may help elucidate novel gut microbiota-targeted therapeutics for the management and treatment of cardiometabolic disease. Despite a high success rate in rodents, there remains translational changes associated with the transplantation. The goal here is to provide guidance in studying the effects of gut microbiome in experimental cardiovascular disease. In this study, a detailed protocol for the collection, handling, processing, and transplantation of fecal microbiota in murine studies is described. The collection and processing steps are described for both human and rodent donors. Lastly, we describe using a combination of the Swiss-rolling and immunostaining techniques to assess gut-specific morphology and integrity changes in cardiovascular disease and related gut microbiota mechanisms.

Introduction

Cardiometabolic disorders, including heart disease and stroke, are the leading global causes of death¹. Physical inactivity, poor nutrition, advancing age, and genetics modulate the pathophysiology of these disorders. Accumulating evidence supports the concept that gut microbiota affect cardiovascular and metabolic disorders, including type 2 diabetes², obesity³, and hypertension⁴, which may hold a key to the development of new therapeutic approaches for these diseases.

The exact mechanisms by which the microbiota cause diseases are still unknown, and current studies are highly variable, in part due to methodological differences. Fecal microbiota transplantation (FMT) is a valuable approach to delineating a direct role of the total microbiota or isolated species in disease pathophysiology. FMT is widely used in animal studies to induce or suppress a phenotype. For example, caloric intake and glucose metabolism can be modulated by transferring fecal matter from a sick donor to a healthy recipient^5,6. In humans, FMT has been shown to be a safe treatment option for patients with recurrent Clostridium difficile infection⁷. Evidence supporting its use in cardiovascular disease management is emerging; for instance, FMT from lean to metabolic syndrome patients improves insulin sensitivity⁸. Gut dysbiosis is also associated with high blood pressure in both human and rodent studies^9,10,11. FMT from mice fed a high salt diet into germ-free mice predisposes the recipients to inflammation and hypertension¹².

Despite the high rate of FMT success in rodents, translational challenges remain. Clinical trials using FMT to treat obesity and metabolic syndrome indicate minimal to no effects on these disorders^13,14,15. Thus, more studies are needed to identify additional therapeutic avenues targeting the gut microbiota for the treatment of cardiometabolic disorders. Most of the available evidence on the gut microbiota and cardiovascular disease is associative. The described protocol discusses how to utilize a combination of FMT and the Swiss-rolling technique to show both an association between disease and gut microbiota and directly assess the integrity of all parts of the gut intestine^16,17,18.

The overall goal of this method is to provide guidance for studying the effects of the gut microbiome in experimental cardiovascular disease. This protocol provides more details and key considerations in the experimental design to promote physiological translation and increase the rigor and reproducibility of the findings.

Protocol

Vanderbilt University's Institutional Animal Care and Use Committee approved all procedures described in this manuscript. C57B1/6 male mice at 3 months of age, purchased from The Jackson Laboratory, were housed and cared for in accordance with the Guide for the Care and Use of Laboratory Animals.

1. Collection, storage, and processing of human fecal samples

1. Collect a stool sample, using a sterile container if the subject is in the clinic. Refrigerate the stool samples at 4 °C within 36 h of collection until ready for processing. Alternatively, collect stool samples using a commercially available tool for easy and extended DNA stability at ambient temperature, especially for at-home use.
2. Sanitize a biosafety-level fume hood with a 10% bleach solution, or other Environmental Protection Agency-approved disinfectant.
3. Remove the stool from cool storage and bring into the fume hood; use a disposable spatula to make ~1 g aliquots and store in a -80 °C freezer until completely ready for processing.
4. Discard all disposable items in biohazard trash. Disinfect all surfaces (hood and any surfaces touched by the processor) and items being removed from the hood.

2. Aseptic collection of mouse fecal samples

NOTE: Use aseptic techniques, including sterilized instruments.

1. Euthanize the mouse by CO₂ asphyxiation. Spray the chest and sides of the mouse with 70% ethanol and carefully open the skin and peritoneal cavity to expose the gastrointestinal tract.
2. Isolate the cecum and use sterile surgical scissors to cut it in half. Briefly, expose the cecum and cut 0.5 cm proximally from the ileum and 0.5 cm distally at its junction with the colon. Transfer the isolated cecum onto a sterile Petri dish.
3. Use a sterile spatula to transfer cecal content into sterile tubes, and store aliquots in a -80 °C freezer¹⁹.
   ​NOTE: Since the majority of bacteria in the gut are anaerobes, exposure to oxygen may damage or kill the organisms during the isolation procedure in a room atmosphere. Thus, fecal samples should be isolated in anaerobic chambers to maintain the viability of the bacteria.

3. Fecal matter transplantation

1. Resuspend fresh or previously frozen fecal pellets in sterile saline in a 1:20 (w:v) proportion and vortex until homogenized.
2. Pass the homogenate through a 30 µm pore nylon filter to remove large particulate matter. Centrifuge at 79 × g for 5 min and collect the supernatant to use for transplantation.
3. Oral gavage 100 µL of the slurry per germ-free recipient mouse for 3 consecutive days, followed by gavage every 3 days for 2 weeks. Use conventional mice to study mechanisms of the gut microbiota if they have been first treated with antibiotics to eliminate the recipient's own endemic microbiota. For example, administer ceftriaxone (400 mg/kg) daily to the recipient mice for 5 consecutive days by oral gavage before gavage of the fecal slurry.
   NOTE: Studies suggest that a minimum of 2 weeks of this treatment is necessary to elicit cardiovascular changes, including blood pressure²⁰.
4. Ensure that germ-free recipient mice are single-housed in gnotobiotic film isolators and fed with sterile food and water.

4. Systolic blood pressure measurements

NOTE: Gnotobiotic mice that received FMT from conventionally housed 3-month-old C57Bl/6 mice were implanted with osmotic minipumps (Alzet, model 2002) for infusion of low-dose angiotensin II (140 ng/kg/min) for 2 weeks. Blood pressure was monitored weekly via tail cuff. The protocol for implanting osmotic minipumps has been previously reported²¹. Tail-cuff was performed as briefly summarized below. A noninvasive method of measuring blood pressure, such as tail cuff, is suitable for FMT studies in gnotobiotic mice. The detailed steps on how to perform tail cuff have been described previously²².

1. Briefly, retrieve the mice from the gnotobiotic isolators and preheat the tail-cuff machine platform and mouse holder.
2. Place the conscious mice in restraints on the heated platform and collect at least three rounds of systolic pressure measurements using tail cuff plethysmography. Perform the following steps below on 3 consecutive days prior to the proper measurement days, to train mice to being restrained in order to reduce stress.
   1. Gently place the mouse in the preheated holder and leave the tail outside. Carefully tape down the top without pinching it, so as not to stress the mouse.
   2. Allow the mouse to rest in the holder; place on the platform for 3-5 min covered by a sheet to acclimate.
3. Average the measurements from all the rounds for an average systolic pressure for each animal.

5. Assessment of FMT to cardiovascular changes

1. Following blood pressure measurement, euthanize the mice and aseptically collect cecal contents, as described in section 2 .
2. Harvest the intestines and other tissues, including the heart, aorta, liver, mesenteric arteries, and kidneys, to examine the role of the gut microbiota in cardiometabolic health. To harvest tissues, locate the tissue in the mouse and use scissors to excise them.
3. Run a metagenomic sequencing analysis on fecal samples/cecal contents collected from the donor and recipient mice to confirm engraftment of the gut microbiota after FMT²³. The first proof of successful colonization of microbiota is confirming that the donor and recipient's microbiota are similar.
4. Use the Swiss-roll technique (see section 6) on the harvested intestinal tissue, coupled with immunostaining and histology to examine morphological and cellular expression changes²⁴.

6. Making gut intestine Swiss-rolls

1. Day 1
   1. In a properly euthanized mouse sprayed with 70% ethanol, dissect the mouse gut from the anal side (fixed in retroperitoneum) to the stomach side. Place the entirety of the isolated gastrointestinal tract in a Petri dish containing phosphate-buffered saline (PBS). Gently hold the proximal end from the stomach end and remove the surrounding fat and connective tissue by hand.
   2. Isolate the small intestine (cephalad from the appendix) and make a Z-type zigzag with each length. Then, cut to obtain the duodenum, jejunum, and ileum successively, as previously described²⁵. Isolate the colon by cutting the section of the intestine below the cecum.
   3. Cut the duodenum, jejunum, ileum, and colon.
   4. Flush and wash the gut inside using PBS with a syringe and a needle with a ball tip, so as not to tear the intestine.
   5. Put the gut on filter paper. Label the paper with the name of the section (e.g., duodenum) and then 'P' at the right top corner for proximal or 'D' for distal at the right bottom corner.
   6. Cut the gut longitudinally with ball-tip scissors. Open the gut on the filter paper. Wash with more PBS as needed.
   7. Sandwich the gut between two filter papers. Staple the filter papers at four points/corners near the gut.
   8. Soak in 10% formalin neutral buffer solution (4.0 g of sodium phosphate, monobasic, 6.5 g of sodium phosphate, dibasic, 100 mL of 37% formaldehyde, 900 mL of distilled water). Shake using a platform rocker at 5 rpm at room temperature overnight.
2. Day 2
   1. Prepare 2% agarose in distilled water and heat with a stir bar in a beaker covered with aluminum foil.
   2. Retrieve the tissues; strip the upper filter paper. Roll the gut from the proximal side so that the proximal side goes inside first, and roll inward so that the lumen is inside on the slide as well. Pin with a 30 G needle or two, as needed.
   3. Aspirate 1 mL of agarose using disposable graduate transfer pipettes, and pour the agarose on a rolled gut section on a flat surface while avoiding air bubbles in the tissues.
   4. Allow the agarose to cool and solidify. Use a razor blade to trim the extra agarose around the tissue section.
   5. Put the gut sections in tissue processing/embedding cassettes (bigger than the regular ones to accommodate the increased height due to agarose). Soak in 70% ethanol at 4 °C.
   6. Prepare paraffin-embedded tissue slides and proceed to immunostaining, as outlined below.

7. Immunostaining of the gut intestinal tract

1. Deparaffinization
   1. Pass through the following baths with slides in a rack: xylene for 3 min, fresh xylene again for 3 min, xylene with 100% ethanol (1:1) for 3 min, 95% ethanol for 3 min, 70% ethanol for 3 min, and 50% ethanol for 3 min.
   2. Rinse gently with cold running tap water. Store in a bath of tap water.
2. Antigen retrieval
   1. Following deparaffinization, boil the slides in a rack in a bath of antigen retrieval buffer (0.01M trisodium citrate dihydrate at pH 6 and 0.05% Tween-20) at 100 °C for 20 min.
   2. Run under cold tap water.
3. Staining
   1. Remove the slides from the bath and place the tissue face up in a slide box with wet laboratory wipes/paper towels in the bottom. Draw an outline around the tissue with a hydrophobic marker pen.
   2. Drop Tris-buffered saline (TBS) + 0.025% Triton X-100 onto the tissue and incubate for 5 min. Repeat this step.
   3. Block with TBS + 10% fetal bovine serum (FBS) + 1% bovine serum albumin (BSA) for 2 h at room temperature. Turn the slides on their side and remove the blocking buffer on a laboratory wipe.
   4. Add primary antibody solution and incubate at 4 °C for at least 2 h or overnight. Wash gently with TBS + 0.025% Triton X-100 by gently pipetting ~200 µL over the section.
   5. Add secondary antibody solution and incubate for 1 h at room temperature. Rinse the slides 3 x 5 min with TBS by rinsing with a pipette, as in step 7.3.4.
   6. Mount with mounting medium and a coverslip.

Results

The steps described above are summarized in Figure 1. Mouse cecal contents or human feces are resuspended in sterile saline to prepare a slurry to give to germ-free mice (100 µL) by gavage, first for 3 consecutive days, then once every 3 days. At the end of the protocol, blood pressure is measured by the tail-cuff method, mice are euthanized, and tissues are harvested for assessment of changes in the gut microbiota and cardiovascular and metabolic changes.

A key s...

Discussion

A valuable approach to studying the causal role of gut microbiota in cardiovascular and metabolic disease is to transfer the total microbiota or select species of interest into germ-free mice. Here, we describe protocols to collect fecal samples from humans and conventionally housed mice into germ-free mice to study the role of gut microbiota in hypertensive disorders.

In mice, we use aseptically collected cecal contents processed in an aerobic chamber, and in humans, collect feces. FMT can be...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Alexa Fluor 488 Tyamide SuperBoost	ThermoFisher	B40932
Anaerobic chamber	COY	7150220
Apolipoprotein AI	Novus Biologicals	NBP2-52979
Artery Scissors - Ball Tip	Fine Science Tools	14086-09
Bleach solution	Fisher Scientific	14-412-53
Bovine Serum Albumin	Fisher Scientific	B14
CD3 antibody	ThermoFisher	14-0032-82
CD68 monoclonal antibody	ThermoFisher	14-0681-82
Centrifuge	Fisher Scientific	75-004-221
CODA high throughput monitor	Kent Scientic Corporation	CODA-HT8
Cryogenic vials	Fisher Scientific	10-500-26
Disposable graduate transfer pipettes	Fisher Scientific	137119AM
Disposable syringes	Fisher Scientific	14-823-2A
Ethanol	Fisher Scientific	AA33361M1
Feeding Needle	Fine Science Tools	18061-38
Filter paper sheet	Fisher Scientific	09-802
Formalin (10%)	Fisher Scientific	23-730-581
High salt diet	Teklad	TD.03142
OMNIgene.GUT	DNAgenotek	OM-200+ACP102
Osmotic mini-pumps	Alzet	MODEL 2002
PAP Pen	Millipore Sigma	Z377821-1EA
Petri dish	Fisher Scientific	AS4050
Pipette tips	Fisher Scientific	21-236-18C
Pipettes	Fisher Scientific	14-388-100
Serile Phosphate-buffered saline	Fisher Scientific	AAJ61196AP
Smart spatula	Fisher Scientific	NC0133733
Stool collection device	Fisher Scientific	50-203-7255
TBS Buffer	Fisher Scientific	R017R.0000
Triton X-100	Millipore Sigma	9036-19-5
Varimix platform rocker	Fisher Scientific	09047113Q
Vortex mixer	Fisher Scientific	02-215-41
Xylene	Fisher Scientific	1330-20-7, 100-41-4