Retroductal Nanoparticle Injection to the Murine Submandibular Gland

Summary

Local drug delivery to the submandibular glands is of interest in understanding salivary gland biology and for the development of novel therapeutics. We present an updated and detailed retroductal injection protocol, designed to improve delivery accuracy and experimental reproducibility. The application presented herein is the delivery of polymeric nanoparticles.

Abstract

Two common goals of salivary gland therapeutics are prevention and cure of tissue dysfunction following either autoimmune or radiation injury. By locally delivering bioactive compounds to the salivary glands, greater tissue concentrations can be safely achieved versus systemic administration. Furthermore, off target tissue effects from extra-glandular accumulation of material can be dramatically reduced. In this regard, retroductal injection is a widely used method for investigating both salivary gland biology and pathophysiology. Retroductal administration of growth factors, primary cells, adenoviral vectors, and small molecule drugs has been shown to support gland function in the setting of injury. We have previously shown the efficacy of a retroductally injected nanoparticle-siRNA strategy to maintain gland function following irradiation. Here, a highly effective and reproducible method to administer nanomaterials to the murine submandibular gland through Wharton's duct is detailed (Figure 1). We describe accessing the oral cavity and outline the steps necessary to cannulate Wharton's duct, with further observations serving as quality checks throughout the procedure.

Introduction

Salivary gland dysfunction has many etiologies, including Sjögren's syndrome, an autoimmune mediated loss of functional secretory tissue, and radiation induced hyposalivation (RIH), a common sequella of head and neck cancer radiotherapy¹. Loss of salivary function due to either condition predisposes individuals to oral and systemic infection, tooth decay, digestive and swallowing dysfunction, speech impairment, and major depression^1,2,3. As a result, quality of life significantly suffers, with interventions limited to palliation of symptoms rather than cure⁴. To investigate novel therapies in vivo, it is of interest to administer bioactive compounds directly to the salivary gland.

Retroductal injection is a valuable method to deliver bioactive compounds directly to the salivary glands and test the efficacy in disease, injury, or under normal tissue homeostasis. The three major salivary glands are the parotid (PG), the submandibular (SMG), and the sublingual (SLG), all of which empty into the oral cavity through excretory ducts. The anatomy of the murine SMG permits direct access through cannulation of Wharton's duct, located in the floor of the mouth beneath the tongue⁵. Following the cannulation, solvated drugs can be administered directly to the SMG. Following retroductal delivery, extra-glandular diffusion is restricted by the surrounding tissue capsule which regulates the exchange of material with surrounding structures⁶. The SMG and its duct are similarly structured in humans, and are routinely accessed during SMG surgery and sialoendoscopy⁷. In humans and mice, the PG is likewise accessible via Stensen's duct in the buccal mucosa⁸.

In murine models of RIH, SMG retroductal injection has been used to deliver therapeutics including growth factors, primary cells, adenoviral vectors, cytokines, and antioxidant compounds to modulate the cellular response to injury, and reduce the resulting tissue damage^{5,9,10,11,12,13,14,15,16}. The most notable clinical success of retroductal injection is the administration of adenoviral vector to direct expression of a water channel (Aquaporin 1; AQP1) in patients following the radiation for head and neck cancer¹⁷.

Previously, we have developed and shown the efficacy of a retroductally injected polymeric nanoparticle-siRNA system to protect salivary gland function from RIH^11,18,19,20. As an extension of our past work, here, we demonstrate our protocol for retroductal SMG injection using a fluorescently labeled nanoparticle (NP) capable of loading and delivering otherwise poorly soluble drugs^21,22,23.

We have synthesized the NP from a diblock copolymer comprised of poly(styrene-alt-maleic anhydride)-b-poly(styrene) (PSMA) through reversible addition chain fragmentation (RAFT) polymerization, as described previously²¹. Through solvent exchange, these polymers spontaneously self-assemble into micelle NP structures with a hydrophobic interior and hydrophilic exterior²¹. The NPs are labeled with Texas-Red fluorophore to permit the verification of NP delivery into the glands without sacrificing the animal. Live animal imaging and SMG immunohistochemistry is shown at 1 h and 1 day following the injection.

This updated and reproducible cannulation protocol should enable others to achieve retroductal injection. We expect that this refined technique will become critical for in vivo studies and therapeutic development^24,25.

Protocol

All in vivo procedures outlined below were approved by the University Committee on Animal Resources at the University of Rochester, Rochester, NY.

1. Preparation

1. Using 32G intracranial catheter tubing with wire inset, cut 3 cm of the tubing to form a beveled end, approximately 45° to the long axis. Confirm that the wire is at least 1 cm longer than the tubing.
2. Load 50 µL of PSMA nanoparticle solution (Figure 1), or other injection material, into a Hamilton syringe. To reduce the probability of barotrauma during injection, attach the catheter tubing, with the stylet removed, to the syringe and expel dead volume.
3. Inspect the injection solution to ensure the nanoparticle is fully solvated to prevent ductal obstruction following the administration.
4. Prepare atropine solution at 0.1 mg/mL.
   NOTE: Because atropine is light sensitive and degrades over time, this solution should be made the day of injection, and protected from light until administered.

2. Accessing and Visualizing Ductal Entry Point

1. Weigh C57/BL6 mice using an analytical balance.
2. Using a 0.5 mL syringe with 29G x ½" needle, anesthetize mice with an intraperitoneally injected sterile saline solution of 100 mg/kg ketamine and 10 mg/kg xylazine. Proceed to the following step when the mouse no longer responds to stimuli, which generally occurs within 5 to 10 min following the injection.
   NOTE: This procedure may also be performed under isoflurane, but will require a custom nose cone that permits access to the oral cavity.
3. To prevent dryness during the procedure, apply lubricant to eyes and place the mouse in a prone position on a custom stage.
   NOTE: To maintain appropriate conditions for intra-oral procedure, tools should be disinfected or sterilized prior to each use.
4. Open the oral cavity by securing the maxillary incisors over a metal beam, and use an elastic band to apply downward tension behind the mandibular incisors (Figure 2A).
5. Align the mouse beneath the dissecting microscope such that the base of the jaw is visualized.
6. To widen the mouth, use a custom, curved steel retractor to apply tension bilaterally to the buccal mucosa.
7. To visualize the submandibular papillae, grasp and gently lift the tongue from the floor of the mouth using blunt forceps.
   NOTE: The papillae will appear as two pale protrusions beneath the tongue (Figure 2B).
8. To ease the visualization and further manipulation within the oral cavity, place cotton between the tongue and the buccal mucosa.

3. Ductal Cannulation and Line Placement

1. Using fine, curved forceps, grasp catheter tubing with the wire inset. For optimum manual control during cannulation, align the tubing with the curvature of the forceps (Figure 3A).
2. Using the dissecting microscope, move the forceps and wire into the field of view.
   NOTE: The wire should be protruding from the tubing.
3. Gently apply pressure into the base of one submandibular papilla using the wire inset to produce a small, superficial, mucosal puncture (0.076 mm diameter) that will facilitate later entry of the catheter tubing (0.25 mm diameter). If resistance is encountered, cut fresh beveled tips on both the tubing and wire inset with sharp dissecting scissors.
4. Following the entry, withdraw the stylet and, using the dissecting microscope, confirm the presence of saliva at the puncture site. Avoid forceful or sudden movement (either withdrawal or insertion) of the stylet that may cause bleeding or compromise ductal integrity.
5. Retract the stylet within the tubing (Figure 3B).
6. To ensure that injection tubing will fit into Wharton's duct opening, insert tubing containing the stylet as a rigid guide into the previously made puncture (Figure 2C).
   NOTE: If not performed quickly, local swelling may prevent re-insertion.
7. To prevent back pressure from prolonged ductal obstruction, withdraw the tubing. Inspect to verify that an opening, visible under microscopy, can be seen in the submandibular papilla. If visible bleeding occurs, remove the stylet and reattempt from step 3.2 on the opposing submandibular papillae.
8. Without moving the mouse, administer intraperitoneal injection of 1 mg/kg atropine solution, to reduce salivation during the procedure. Wait 5 - 10 min.
9. Grasp the end of the syringe tubing, and insert into the orifice using the dissecting microscope (Figure 3C). If resistance is encountered, cut a fresh beveled end to the tubing and reattempt.
10. Once the tubing is in place within the submandibular papilla, slowly advance 3 - 5 mm into the duct. Release the tubing from the forceps.
11. To improve the seal between the tubing and the submandibular papilla, dry the interface by gently blotting with gauze for 1 min.
12. Inspect to confirm that the position of the tubing has not shifted during drying.

4. Injection

1. Inject material at a rate of 10 µL/min. Inspect to confirm that the mouse remains sedated and is not showing signs of distress (Figure 2D).
   NOTE: Injections of 15 - 50 µL are well tolerated. Injection of larger volumes can result in barotrauma.
2. Following the injection, maintain syringe pressure for 5 min to improve the retention of material within Wharton's duct and SMG (Figure 4). Inspect the submandibular papilla periodically to ensure that tubing does not exit the ductal orifice.
3. Using fine forceps, grasp and gently withdraw the tubing from the submandibular papillae.
   NOTE: It is normal to observe some fluid egress from the papillae.
4. Remove the retractor and cotton from the oral cavity before moving the mouse from the stage.
   NOTE: The animal should not be left unattended until it has regained sufficient consciousness to maintain sternal recumbency. Furthermore, ensure that the mouse is not housed with other mice until fully recovered.

5. Verification and Analysis

Note: An in vivo Imaging System (IVIS) can be used to assess retention of fluorescently labeled nanoparticles following injection (as shown 1 h and 24 h after injection in Figure 5).

1. To better visualize fluorescent signal within the SMG through the skin, remove the ventral fur overlying the SMGs either by shaving or using a chemical depilatory.
   Note: Following euthanasia, SMG tissue can also be harvested, fixed (overnight in 4% paraformaldehyde), and stained using immunohistochemistry to confirm persistence of fluorescently labeled NP one day following injection (Figure 6).

Results

Retroductal injection can be used to administer NPs to the murine SMG (Figure 1). Here, we deliver 50 µg PSMA NPs labeled with Texas Red fluorophore.

Proper placement of the mouse allows facile access and visualization of the floor of the mouth (Figure 2A-B). The submandibular papillae are identified as two fleshy protrusions beneath the tongue. Followi...

Discussion

Retroductal injection is critical for localized drug delivery to the salivary gland. This technique has applications in screening therapeutic agents for conditions including Sjogren's syndrome and RIH^9,10,28. Direct drug delivery into the SMG via retroductal injection provides a key advantage over systemic administration in its potential to reduce off-target effects, including immune activation¹¹. The abi...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Pilocarpine hydrochloride	Sigma Aldrich	P6503	Pilocarpine
Student Vannas Spring Scissors	Fine Science Tools	91500-9	Spring Scissors for Tracheostomy
Sterile Saline Solution	Medline	RDI30296H	Saline
Dumont #7 Forceps	Fine Science Tools	11274-20	Curved Forceps
Dumont #5 Forceps	Fine Science Tools	11251-10	Straight Forceps
Standard Pattern Forceps	Fine Science Tools	11000-12	Blunt Forceps
Fine Scissors- Tungsten Carbide	Fine Science Tools	14568-09	Dissection Scissors
Microhematocrit Heparinized Capillary Tubes	Fisher Scientific	22362566	Capillary tubes
Lubricant Eye Ointment	Refresh	N/A	Refresh Lacri-Lube
Goat polyclonal anti-Nkcc1	Santa Cruz Biotech	SC-21545	Nkcc1 Antibody
DAPI (4',6-Diamidino-2-Phenylindole, Dihydrochloride)	Thermo Fisher Scientific	D1306	DAPI
GraphPad Prism	GraphPad	ver6.0	Statistical Software
Cotton tipped applicator	Medline	MDS202000	Applicator for eye ointment
0.5cc Insulin Syringe, 29G x 1/2"	BD	7629	Syringe for intraperitoneal injection