Immunostimulatory Agent Evaluation: Lymphoid Tissue Extraction and Injection Route-Dependent Dendritic Cell Activation

All experimental procedures including animal handling, sacrifice, and organ isolation were performed in strict accordance with the rules of the International Animal Care and Use Committee at Shanghai Public Health Clinical Center and Asan Medical Center. The study protocol was approved by the respective committees on the Ethics of Animal Experiments at Shanghai Public Health Clinical Center (Mouse Protocol Number: SYXK-2010-0098) and Asan Medical Center (2016-02-168).

1. Preparation of Material

NOTE: A nano-adjuvant, INP, comprised of a self-assembled lipid-DNA carrier, namely U4T, and CpG-motif containing oligonucleotide, eCpG, was employed in the current study to test the appropriate injection route for immunotherapy and vaccination in mice². Importantly, other potential therapeutic molecules such as antigens, antibodies, and adjuvant agents can be substituted for INP and tested using the same methodology described below.

1. INP formulation^2,4,7
   1. Anneal U4T (160 µM, carrier) with eCpG (80 µM, containing a TLR9 agonist sequence) in the presence of 1x phosphate-buffered saline (PBS, pH 7.4) and MgCl₂ (2 mM). The standard volume for annealing is between 50 and 110 µL in a PCR tube. Heat the mixture to 95 °C for 10 min and then slowly cool (1 °C/16 min) to 25 °C using a thermocycler. The annealing protocol requires approximately 2 h.
      CAUTION: The U4T-containing liquid is very foamy and must be carefully handled.
   2. Make 50 µL aliquots of the INP preparation; use 50 µL of the material for each injection.
      NOTE: In the case of 100 µL per injection, repeat step 1.1 with proper calculation of ingredients.

2. General Animal Procedures

Note: Alternative methods for handling the mice can be used depending on the laboratory requirements and approved animal protocols⁸. The type of mice used is 6-week-old C57BL/6 and female mice.

1. List of materials
   1. Prepare the liquid anesthetic (isoflurane), anesthetic machine, carbon dioxide (CO₂) inhalation chamber, sterilized gauze, forceps, mouse restrainer, gloves, hooked forceps, 1-inch dissecting scissors, 2-inch dissecting scissors, alcohol for sterilization, and insulin syringe.
2. Inhalant anesthesia
   NOTE: Anesthesia is mandatory for intranasal, subcutaneous, footpad, and intravenous injections of mice⁹. An induction chamber with a vaporizer machine and oxygen tank connected is used for this procedure.
   1. Place the animal in the induction chamber and make sure to tightly close the lid.
   2. Flow the oxygen at 1 L/min and set the vaporizer setting to an induction level of 4% for isoflurane.
   3. Vary the exposure time of animals in the chamber depending on animal weight; however, more than 10 s of exposure and monitoring every 5 min is necessary for anesthesia¹⁰.
      NOTE: No pedal reflex indicates successful anesthesia.
   4. Turn off the isoflurane inductor and flow 1 L/min of oxygen to prevent personnel exposure to isoflurane gas.
3. Sacrifice
   1. Prepare a chamber for CO₂ inhalation sacrifice in accordance with the NIH "Guidelines for Euthanasia of Rodents Using Carbon Dioxide"¹¹.
4. Test gag response to confirm that the animal is fully sacrificed.
   1. Verify that the mouse is properly sacrificed by pinching the tip of its foot and confirming no gag response. If there is a response, repeat step 2.2.
5. Disinfection
   1. Disinfect all surface areas used for injections or dissections with alcohol spray and sterilized gauze.
6. Organ storage
   1. Use forceps to remove all fat and blood surrounding the harvested organs. Store these harvested tissues individually in a Petri dish filled with 3 mL of PBS. The container and media may differ depending on the subsequent procedure requirements.

3. Injection Routes

1. Use the six injection methods to investigate injection-dependent immune responses: oral, intranasal, subcutaneous, footpad, intraperitoneal, and intravenous^8,12,13.

4. Isolation of Lymph Nodes and Spleen

Note: Use young and healthy lean mice (6 weeks old) for these procedures because the fats that typically build up around the lymph nodes in older mice are difficult to remove and can prevent proper visualization of the organ.

1. Pre-harvesting steps
   NOTE: This section describes the post-injection harvesting of the iLN, spleen, and mLN from mice for analyzing immune stimulation. Refer to steps 2.3-2.5 for sacrifice, gag response, and disinfection methods.
   1. Pin the limbs of the mouse to the surgical foam board and sterilize the ventral side of the mouse with 70% ethanol.
   2. Pick a location 2 cm above the genital area and use hooked forceps to pull up the outer skin cover like a tent. Make an approximately 1-cm cut at this location and slide the scissors (2-inch) underneath the skin, making further dissection cuts.
   3. For the integument, cut vertically through the outer skin with both arms of the scissors (2-inch). Expose the internal organs covered with peritoneum.
   4. Pin the outer layer of the skin.
2. Isolation of the inguinal lymph node (iLN)
   1. Locate the iLN at the left side of the leg following the conjunction of the blood vessel going down the left leg, which appears as a tilted letter 'Y'.
      NOTE: For subcutaneously injected mice, choose an iLN on the same side of the injection location.
   2. Uncover the layer with two forceps to rip the covered lipid layer and harvest the pale yellow iLN (bean shape, 2 mm). For subsequent treatment and storage of the tissue, see step 2.6.
3. Isolation of the spleen¹⁴
   1. Make a tent at the center of the peritoneum using hooked forceps in one hand and use small thin scissors (1 inch) in the other hand to cut the peritoneum.
   2. Use hooked forceps in the left hand to grasp the intestine and flip it over to locate the red bean-like spleen (length of approximately 14 mm) attached to the left upper side of the mouse abdomen.
   3. Gently pull out the spleen with another set of forceps in the right hand while detaching other organs with the hooked forceps. Follow step 2.6 to store the spleen.
      NOTE: Care is required when harvesting the spleen, as it is an easily damaged soft organ.
4. Isolation of the mediastinal lymph node (mLN)
   NOTE: The mLN is located approximately 1 cm beneath the ribs and is covered by other organs such as the heart and lungs. Because of its location and small size, it is important to carefully expose the surrounding area using forceps and scissors (1-inch).
   1. Cut the diaphragm and detach it from the ribs. Hold the body of the ribs with forceps and then cut the left and right side of the ribs with scissors (1-inch). Pin the cutoff ribs over the mouse's shoulder on the right to expose the heart and lungs.
      NOTE: Take care not to cut any of the blood vessels around the heart and lungs, as leaking blood makes it very difficult to locate the tiny translucent mLN. If any blood vessels are cut, gently absorb the blood in the area with gauze.
   2. Using hooked forceps in the right hand, flip the heart and lungs gently to the right until the backbone is exposed. Use a pair of regular forceps in the left hand to remove the iLN with the hooked forceps in the right hand to assist with this harvesting.
   3. Grab the lungs and flip them until the backbone is widely exposed. The mLN (a translucent bean-shaped structure of approximately 2 mm in size) is located between the lungs and the spine.
   4. Utilize the hooked forceps to expose the area around the mLN and the other forceps to extract the tissue.
      NOTE: The mLN is surrounded by numerous other tissues which must be removed carefully with forceps before harvest.
   5. See step 2.7 for instructions on storing the harvested mLN.

5. Preparation of samples for flow cytometry

Note: To assay the activation of DCs in the harvested iLN, spleen, and mLN, single-cell suspensions of these tissues are stained with fluorescence-conjugated antibodies as DC-specific markers, co-stimulatory molecules, and major histocompatibility complex molecules and analyzed by flow cytometry.

1. Preparation of single-cell suspensions: spleen
   1. Add 5 mL of fresh culture medium (CM) into 6-mm culture dish and place the samples on ice.
   2. Remove surrounding blood and fat tissues and place the tissue in the lid of the culture dish.
   3. Cut the tissues into small fragments (<1 mm) with curved scissors. Suspend the fragments in PBS followed by spinning down the tissue using a centrifuge at 646 × g for 5 min and removing the supernatant.
   4. Suspend the samples again in 3 mL of CM and digest the fragments with 200 µL of solution of 2% fetal bovine serum (FBS) supplemented with collagenase IV for 20 min.
   5. Gently shake and incubate the sample at 37 °C for 1 h. Filter the digested tissues through nylon mesh (100 nm) and spin them down. Wash the single cells in 5 mL of PBS and remove the supernatant by centrifugation.
   6. Re-suspend the cells in 5 mL of cell isolating solution. Re-load another layer of 5 mL of clear cell isolation solution to form an aqueous layer-by-layer stack.
   7. Centrifuge the cell preparation at 2012 × g for 10 min. Obtain the supernatant, which is the lighter density fraction (<1.077 g/cm³), for subsequent flow cytometry analysis.
   8. Count the number of cells with a hemocytometer.
2. Preparation of single-cell suspensions: lymph node
   1. Follow step 5.1.1-5.1.2.
   2. Add 5 mL of CM using a 10-mL pipet to suspend the cells by pipetting 3-4 times. After thoroughly suspending the cells, use sterile nylon mesh to filter the cells for collection in a 15-mL conical tube.
   3. Spin the cells down at 646 ×g for 7 min at 4 °C and remove the supernatant.
   4. Add an additional 5 mL of CM into 10-mL pipet to better suspend the cells by pipetting.
   5. Wash with PBS and count the number of cells.
3. Flow cytometry analysis
   1. Aliquot 0.5-1 × 10⁶ cells into each fluorescence-activated cell sorting (FACS) tube.
   2. Centrifuge at 646 × g at 4 °C for 5 min and aspirate the supernatant.
   3. Add 5 µL of each diluted fluorescence-conjugated antibody preparation to the cells to re-suspend the cell pellet and vortex.
      NOTE: It is important to shield the cell pellet from light exposure as much as possible because fluorescently labeled antibodies are light-sensitive. Generate an antibody master mix and always add Fc block.
   4. Prepare FACS buffer using 100 µL of PBS, 1% of heat-inactivated regular FBS, and 0.1% sodium azide.
   5. Place the sample in the tube at 4 °C for 30 min. Rinse the cells with 2-4 mL of FACS buffer and centrifuge the sample at 1700 rpm for 7 min at 4 °C.
   6. Re-suspend the cells in 500 µL of FACS buffer in the dark. Analyze the cells by flow cytometry.