Mouse Models Of Helicobacter Infection And Gastric Pathologies

1. Growth and Preparation of Bacterial Inocula

1. Thaw glycerol stocks of H. pylori or H. felis¹⁵ from -80 °C and subculture on horse blood agar (HBA) plates comprising: Blood Agar Base No.2 (see Table of Materials); a modified “Skirrow’s antibiotic selective supplement” (consisting of vancomycin, 10 μg/mL; polymyxin B, 25 ng/mL; trimethoprim, 5 µg/mL; amphotericin B, 2.5 μg/mL); and 5–10% (v/v) horse blood^15,16. The bacteria grow well under microaerobic conditions in 2.5 or 3.5 L anaerobic jars containing the appropriate gas packs (see Table of Materials), at 37 °C.
   NOTE: H. pylori strains grown under these conditions must be subcultured after 1–1.5 days of incubation, whereas for H. felis, at least 2 days of incubation is generally required. Suitable culture and storage media have been described in detail previously¹⁵.
2. Prepare bacterial inocula for mouse infection from early-to-mid logarithmic phase cultures. Harvest bacteria gently from agar plates by flooding each plate with 1–2 mL of brain heart infusion (BHI) broth. Aspirate suspensions from plates with Pasteur pipettes.
   1. Alternatively, prepare inocula from Helicobacter bacteria that have been propagated in BHI broth for 16–18 h¹⁵. In this case, collect bacteria by low speed centrifugation at 2,200 x g for 10 min at 4 °C.
3. Assess the viability and motility of the bacteria by examining wet mount preparations under phase contrast microscopy (100X objective). Prepare wet mounts by resuspending a loopful of bacteria in a 10–20 μL droplet of BHI broth on a glass microscope slide. In the case of H. felis, which is a much larger bacterium than H. pylori, use a haemocytometer to accurately count the numbers of viable bacteria. Confirm culture purity by performing a Gram-stain.
   NOTE: Only use H. pylori inocula if the majority of bacteria have a bacillary or spiral shape (the morphology can vary depending on the strain). H. felis inocula should primarily contain helical-shaped bacteria. Do not use inocula if most bacteria have a coccoid morphology, as these forms are not viable and will not establish an infection in mice.
4. Estimate the number of bacteria in the inoculum by counting under phase contrast microscopy the approximate number of bacteria per field (100X objective) and by using the following guide: 1 bacterium per field = approximately 10⁶ colony-forming units (CFU) of Helicobacter/mL; 10 bacteria per field = approximately 10⁷ CFU/mL; 100 bacteria per field = approximately 10⁸ CFU/mL, etc.
   1. When using a haemocytometer, calculate CFU/mL using the following formula:
      CFU/mL = (average number of bacteria in a 4 x 4 field) x (dilution factor) x (10⁴).
5. Adjust the bacterial cell density of the inoculum to approximately 10⁷–10⁸ CFU/mL by dilution in BHI broth, if necessary.
   1. To ensure maximal bacterial viability, use inocula for intragastric gavage as soon as possible after preparation.
   2. Always confirm H. pylori cell density and viability by performing viable counting of inocula immediately after the gavage procedure (see below). This is not always possible for H. felis, as it does not usually form isolated colonies on culture media. The numbers of viable Helicobacters in inocula cannot be determined by optical density measurement (A₆₀₀) alone as this method does not discriminate between viable (i.e., bacillary/spiral/helical) and non-viable (i.e., coccoid) bacteria.
   3. Use optical density values as a means of estimating the numbers of viable H. pylori bacteria in inocula, but in this case, it is first necessary to generate a growth curve. For this, the A₆₀₀ values of H. pylori cultures are monitored over time and correlated directly against the numbers of viable bacteria, determined by plate counting.
      NOTE: A convenient method for performing such growth curve determinations is to culture bacteria in liquid medium (Section 1.2), using standard flat bottom tissue culture flasks placed in a 10% CO₂ incubator. The numbers of CFUs, determined from aliquots of the cultures obtained every 4–6 h over 2–3 days, are then compared to the corresponding A₆₀₀ values¹⁷. Importantly, growth curves must be generated for each H. pylori strain, as these may grow at different rates and, furthermore, not all strains grow well in 10% CO₂.

2. Intragastric Gavage of Mice with Helicobacter

NOTE: This method of intragastric gavage can be applied to other bacterial species that colonize the gut e.g. S. Typhimurium, C. rodentium, Listeria monocytogenes.

1. Use 6–8-week-old, specific pathogen-free (SPF) and Helicobacter-free male or female mice. Use animals with a C57BL/6 genetic background for infection experiments with H. pylori or H. felis. In the present study, we used wild-type (WT) and genetically modified C57BL/6 mice lacking a key innate immune receptor (termed knock-out or KO animals).
   Note: Mice on other genetic backgrounds can also be used for Helicobacter infections, however, colonization levels and disease severity may be impacted by the type of host background^18,19.
2. Aspirate the bacterial inoculum (Step 1.5) into disposable 1 mL syringes and replace the supplied needles with 23 gauge needles onto which are affixed disposable polyethylene catheters (length, 6–8 cm; internal diameter, 0.58 mm). Fasten catheters to the needles by the application of small strips of plastic film (see Table of Materials). Alternatively, replace the needle/catheter assembly by using sterile plastic feeding tubes (20 gauge x 38 mm).
3. Physically restrain mice with a firm grip at the scruff of the neck and tail.
   NOTE: This procedure can be performed without anaesthesia, or alternatively, with the use of an inhaled anaesthetic, such as methoxyflurane or isoflurane¹⁵.
4. Insert catheter into the center of the open jaw and guide in a caudal direction towards the esophagus. Extend the neck of the mouse to allow ease of access to the stomach through the esophagus (and away from the trachea) until most or all of the catheter is no longer visible and a resistance is felt, corresponding to the base of the stomach. Deliver a specific aliquot, usually 100 μL per inoculation (Figure 1).
   NOTE: Mice should be gavaged with ≥ 10⁵ CFU to ensure optimal colonization and disease pathology.
5. House mice in an SPF animal facility for the duration of the experiment.
   NOTE: Severe pathology and adenocarcinoma in WT C57BL/6 mice is only observed at approximately 24 months’ post-infection²⁰. However, this effect may be accelerated in some genetically modified animals, or in mice with other genetic backgrounds.
6. Upon completion of the gavage procedure, perform a modification of the Miles and Misra technique to determine the numbers of viable H. pylori bacteria administered to mice. For this, the inoculum is serially diluted (from 10⁻¹ to 10⁻⁵) in BHI using the method described in detail previously¹⁵.
   NOTE: In order to ensure isolation of single colonies, HBA plates should be warmed and dried in a biological safety cabinet or 37 °C incubator for 10–15 min prior to use.

3. Harvesting Tissues from Mice Post-experiment

1. Euthanize mice by either carbon dioxide inhalation or cervical dislocation, according to the relevant ethics committee for animal experimentation.
2. Open the abdominal cavity and excise the stomach using fine, curved scissors.
   NOTE: Sera can also be collected by cardiac puncture to aid in the investigation of systemic responses to Helicobacter infection. Additionally, the collection of spleens and paragastric lymph nodes are useful in studying adaptive immune responses.
3. Cut the stomach along the greater curvature and remove residual food by gentle washing in sterile phosphate buffered saline (PBS) in a 50 mL tubes.
4. Wash the stomach again in sterile PBS and then record the wet weight using tared 6 cm plastic Petri dishes.
5. Flatten the stomach and dissect sagittally into two equal tissue fragments, each comprising: the antrum, body and non-glandular forestomach regions (Figure 2). Remove the non-glandular region and weigh one half of each stomach before adding to either 1 mL of sterile BHI (for viable counting) or snap freezing in liquid nitrogen (for DNA/RNA extraction).
   Note: Tubes containing tissue in BHI must be stored on ice until they are ready to be processed. Snap frozen stomach tissues can also be used to extract RNA or proteins for qPCR (quantitative PCR) or western blotting analyses, respectively.
6. Add the other stomach half to a 15 mL tube containing 10% formalin. Immerse tissues in 10% formalin for 10 s and then flatten to the top side of tubes. Allow tissues to fix before re-immersing in 10% formalin solution for a minimum of 24 h.
   NOTE: Tissues can remain in 10% formalin for many weeks prior to processing for histology. Prolonged storage of tissue may, however, affect its architecture and/or antigenicity resulting in sub-optimal results in downstream analyses.

4. Confirmation of Bacterial Colonization in the Stomach Post-infection

1. Viable Counting of H. pylori in the Stomach
   1. Supplement sterile HBA plates with additional antibiotics (200 μg/mL bacitracin and 10 μg/mL naladixic acid) prior to performing colony counts from infected mouse stomachs¹⁶.
   2. Homogenize stomach sections either manually, using autoclavable polypropylene micropestles, or using a mechanical dissociation instrument (see Table of Materials).
   3. Prepare duplicate serial dilutions (10⁻¹ to 10⁻²) of the resulting gastric homogenates in sterile BHI.
      NOTE: Dilutions should be decided based on the typical bacterial loads obtained for a given H. pylori strain used for infection, as well as the duration of infection. Undiluted samples can also be used.
   4. Divide pre-dried HBA plates (see Note above) into three or four segments. Using an adaptation of the Miles and Misra technique, add 10–100 mL of each dilution onto a segment of the agar plate and spread using sterile plastic loops¹⁵.
   5. Allow the plates to dry and then place them in an inverted position (lid side down) in anaerobe gas jars. To maintain humidity in the jars, include a Petri dish containing water.
   6. Incubate jars at 37 °C until colonies form (typically 4–7 days).
   7. Enumerate segment(s) containing between 10 and 100 isolated colonies.
      NOTE: H. pylori colonies and H. felis growth on plates can be distinguished from those of other members of the murine gastric microbiota using standard urease, catalase and oxidase tests. H. pylori and H. felis are positive for all three tests.
   8. Calculate the bacterial loads as (CFU/g of tissue), using the following formula:
      [(Average number of colonies counted) × (dilution factor) x (volume plated)]/(stomach weight).
2. Detection of H. felis Infection in Gastric Tissues by the Polymerase Chain Reaction (PCR)
   1. Extract DNA from mouse stomachs using standard DNA isolation protocols, or a commercially available kit.
   2. Determine the DNA concentration of samples using a fluorometric quantitation technique (see Table of Materials).
   3. Set up PCR reactions targeting a 325-base pairs (bp) fragment of the H. felis urease B gene (ureB)²¹. Each reaction should contain: 100 ng of genomic DNA; 1 mM each of forward (5’-AAA ATC CAC GAA GAC TGG GG-3’) and reverse (5’-CTT TTA TCC AAG TGG TGG CAC ACC-3’) primers; 200 mM dNTPs; 0.5 units of Taq Polymerase and the appropriate amounts of buffer and nuclease-free water.
      NOTE: This oligonucleotide pair has been designed to recognize and bind to homologous sequences in both H. pylori and H. felis ureB genes but when subjected to the PCR conditions below, not those present in the ureB genes of enterohepatic Helicobacter spp.
   4. Perform PCR amplification using the following thermal profile: heating at 94 °C for 5 min, followed by 35 cycles of 94 °C for 30 s, 61 °C for 30 s and 72 °C for 1 min, before holding at 20 °C.
   5. Run PCR products on a 2% agarose gel for 30 min at 100 V.

5. Histological Analyses of Helicobacter-infected Mouse Stomach Sections

1. Processing of Stomach Tissues
   1. Remove stomach tissues from formalin and place in clean Petri dishes.
   2. Cut stomach tissues with a scalpel into several equal-sized longitudinal strips (each 2–3 mm thick) and place in labelled embedding cassettes containing foam padding.
   3. Fix stomach tissues by placing embedding cassettes in a jar filled with 80% ethanol.
      NOTE: Stomachs can be processed immediately or stored for up to 1–2 days prior to proceeding to the next step.
   4. Process stomachs on an automated tissue processor, programmed with the following settings:
      Dehydration: 70% ethanol – 1 cycle, 20 min; 90% ethanol – 1 cycle, 20 min; 100% ethanol – 2 cycles, 20 min each + 1 cycle, 40 min + 1 cycle, 1 h.
      Clearing: xylene – 2 cycles, 30 min each + 1 cycle, 45 min.
      Impregnation: paraffin wax at 60 °C – 1 cycle, 45 min + 1 cycle, 1 h + 1 cycle, 1.25 h.
   5. Remove the processed samples from the machine and store at room temperature for paraffin embedding.
2. Paraffin Embedding of Processed Gastric Tissues
   1. Place stainless steel base molds on the stage of the embedding unit to warm the bases of the molds.
      NOTE: Embedding machine should be set at 60 °C for efficient paraffin embedding.
   2. Place waxed cassettes containing the samples into a warm wax bath/hot plate area of the embedding unit until wax fully dissolves.
      NOTE: It is recommended that the sample be embedded shortly after wax dissolves. This ensures that hardening of tissues does not occur.
   3. Fill half of the stainless-steel molds with paraffin wax. Using warm forceps, remove stomach tissue strips from the cassettes and gently push the stomach strips through the paraffin to the base of the molds. Carefully orientate tissue strips at a right angle to the base of the molds so that their sliced ends are facing upwards.
      NOTE: The following steps should be performed as quickly as possible to avoid hardening and separation of the paraffin layers within embedded blocks. The proper orientation of stomach tissues is critical for downstream analyses.
   4. Place molds on the cold plate of the embedding unit to fix the specimens in place and re-orientate tissues if necessary.
      NOTE: If the tissues become dislodged and the paraffin begins to harden, place molds back onto the hot plate to melt the wax and re-embed tissues in molds.
   5. Place half of the labelled embedding cassettes (which were used for the tissue processing) on to the top of the molds and gently fill with warm wax. Do not allow paraffin to overflow.
   6. Gently place molds on to a cold plate and allow to cool.
   7. Once the paraffin has fully set, separate the embedded blocks from molds. Clean excess paraffin wax on cassette edges using a hot plate (set above 80 °C) or a scraper. Blocks can be stored at room temperature until sectioning is performed.
3. Sectioning of Tissues
   1. Chill paraffin-embedded tissue blocks on ice and heat a water bath filled with ultrapure water to 40–45 °C.
   2. Secure the blade in the holder of the microtome and set the clearance angle between 1–5° to prevent contact between the block face and knife facet, before inserting paraffin blocks.
      NOTE: Ensure blocks are clear of excess paraffin acquired from the embedding, as this may hinder the fit of the block.
   3. Orientate the blade for a straight cut across the block. Gently cut 2–3 thin sections to ensure correct positioning of the block.
   4. Trim blocks by a thickness of approximately 10–30 mm. This step ensures that a maximal surface area of each tissue strip will be cut.
   5. Cut 10 µm sections and discard any that contain holes caused by trimming.
   6. Carefully pick up sections using tweezers and float them in the water bath for flattening. Use tweezers to separate each section.
   7. Collect sections from the water bath and place onto charged glass slides (see Table of Materials).
   8. Store slides upright in a slide rack and place in an incubator at 37 °C. Dry sections overnight.
   9. Store sections at room temperature indefinitely for subsequent analyses.
4. Haematoxylin and Eosin (H&E) Staining of Stomach Tissues
   1. Dewax slides using 3 washes of xylene for 5 min each, followed by 3 washes in 100% ethanol, for 3 min each. Ensure fresh solutions are used at each stage.
   2. Rinse slides in tap water for 30–60 s.
   3. Remove excess water by gently tapping the bottom of the slides on a paper towel. Stain with filtered Haematoxylin for 3 min. Ensure that sections are sufficiently covered with the solution.
   4. Rinse slides under running tap water until water runs clear.
   5. Dip slides in Scott’s Tap water for 8–10 s. Do not expose slides to the solution more than 10 s as this will result in darker and intense stains. Staining of sections can be viewed under a microscope. Efficient staining at this stage will result in a ‘baby blue’ color.
      NOTE: Prepare Scott’s Tap water by dissolving 2 g of sodium hydrogen carbonate (NaHCO₃) and 20 g of magnesium sulphate (MgSO₄) in 1 L of distilled water.
   6. Rinse slides in tap water for 30–60 s.
   7. Remove excess water as described in step 3, and then stain with filtered 1% aqueous Eosin for 3 min.
   8. Rinse slides under running tap water until water runs clear.
      NOTE: Staining can be assessed using a light microscope. If staining is too dark, slides can be dehydrated in 100% ethanol for longer than the specified time. If darker staining is required, slides can be stained with Eosin for 1–2 min longer, prior to proceeding to subsequent steps.
   9. Dehydrate slides using 3 washes of 100% ethanol for 30 s each, followed by 3 washes of xylene for 2 min each. Ensure fresh solutions are used at each stage.
   10. Mount slides with mounting medium. Add a drop of mounting medium in the center of a clean coverslip prior to gently placing slide on top with sections facing downwards.
       NOTE: Do not dry slides prior to cover slipping.
   11. Place slides on a flat surface and allow to air dry. Slides can also be dried in a fume hood to accelerate drying time.
5. Giemsa Staining of Stomach Tissues
   1. Dewax slides using 2 washes of histolene for 5 min each, followed by 2 washes of 100% ethanol for 3 min each and then a final wash in 70% ethanol for 3 min. Ensure fresh solutions are used for each wash.
   2. Rinse slides in tap water for 30–60 s.
   3. Prepare Giemsa solution by mixing 20% Giemsa stain with 80% distilled water. Stain slides with Giemsa solution for 1 h.
   4. Place slides in 100 mL of distilled water containing 3–4 drops of acetic acid for 2–3 s.
      NOTE: Solution must be mixed well prior to use. At this stage, slides should appear pale blue in color.
   5. Wash slides in 96% ethanol for 30 s.
   6. Wash slides in 3 baths of isopropanol for 2 min each, followed by 3 baths of histolene for 2 min each. Use fresh isopropanol and histolene for each wash.
   7. Coverslip slides with mounting medium, as described earlier.