Intratracheal Administration of Dry Powder Formulation in Mice

Summary

Dry powder formulations for inhalation have great potential in treating respiratory diseases. Before entering human studies, it is necessary to evaluate the efficacy of the dry powder formulation in preclinical studies. A simple and noninvasive method of the administration of dry powder in mice through the intratracheal route is presented.

Abstract

In the development of inhalable dry powder formulations, it is essential to evaluate their biological activities in preclinical animal models. This paper introduces a noninvasive method of intratracheal delivery of dry powder formulation in mice. A dry powder loading device that consists of a 200 µL gel loading pipette tip connected to an 1 mL syringe via a three-way stopcock is presented. A small amount of dry powder (1-2 mg) is loaded into the pipette tip and dispersed by 0.6 mL of air in the syringe. Because pipette tips are disposable and inexpensive, different dry powder formulations can be loaded into different tips in advance. Various formulations can be evaluated in the same animal experiment without device cleaning and dose refilling, thereby saving time and eliminating the risk of cross-contamination from residual powder. The extent of powder dispersion can be inspected by the amount of powder remaining in the pipette tip. A protocol of intubation in mouse with a custom-made light source and a guiding cannula is included. Proper intubation is one of the key factors that influences the intratracheal delivery of dry powder formulation to the deep lung region of the mouse.

Introduction

The pulmonary route of administration offers various benefits in delivering therapeutics for both local and systemic actions. For the treatment of lung diseases, high local drug concentration can be achieved by pulmonary delivery, thereby reducing the required dose and lowering the incidence of systemic side effects. Moreover, the relatively low enzymatic activities in the lung can reduce premature drug metabolism. The lungs are also efficient for drug absorption for systemic action due to the large and well-perfused surface area, the extremely thin epithelial cell layer and the high blood volume in pulmonary capillaries¹.

Inhaled dry powder formulations have been widely investigated for the prevention and treatment of various diseases such as asthma, chronic obstructive pulmonary disease, diabetes mellitus and pulmonary vaccination^2,3,4. Drugs in the solid state are generally more stable than in the liquid form, and dry powder inhalers are more portable and user-friendly than nebulizers^5,6. In the development of inhaled dry powder formulations, the safety, the pharmacokinetic profile and the therapeutic efficacy need to be evaluated in preclinical animal models following pulmonary administration⁷. Unlike humans who can inhale dry powder actively, pulmonary delivery of dry powder to small animals is challenging. It is necessary to establish an efficient protocol of delivering dry powder to the lungs of animals.

Mice are widely used as research animal models because they are economical and they breed well. They are also easy to handle and many disease models are well-established. There are two major approaches to administer dry powder to the lung of mouse: inhalation and intratracheal administration. For inhalation, the mouse is placed in a whole-body or nose-only chamber where dry powder is aerosolized and the animals breathe in the aerosol without sedation^8,9. Expensive equipment is required and the drug delivery efficiency is low. While the whole-body chamber may be technically less challenging, the nose-only exposure chamber could minimize exposure of drugs to the body surface. Regardless, it is still difficult to precisely control and determine the dose delivered to the lungs. The dry powder is mainly deposited in the nasopharynx region where mucociliary clearance is prominent¹⁰. Moreover, mice inside the chamber are under significant stress during the administration process because they are constrained and deprived of food and water supply¹¹. For intratracheal administration, it generally refers to the introduction of the substance directly into the trachea. There are two different techniques to achieve this: tracheotomy and orotracheal intubation. The former requires a surgical procedure that makes an incision in the trachea, which is invasive and seldom used for powder administration. Only the second technique is described here. Compared to the inhalation method, intratracheal administration is the more commonly used method for pulmonary delivery in the mouse because of its high delivery efficiency with minimal drug loss^12,13. It is a simple and fast method to precisely deliver a small amount of powder within a few milligrams to the mouse. Although the mouse is anatomically and physiologically distinct to humans and anesthetization is required during the intubation process, intratracheal administration bypasses the upper respiratory tract and offers a more effective way to assess the biological activities of the dry powder formulation such as the pulmonary absorption, bioavailability and therapeutic effects^14,15.

To administer dry powder intratracheally, the mouse has to be intubated, which could be challenging. In this paper, the fabrication of a custom-made dry powder insufflator and an intubation device is described. The procedures of intubation and insufflation of dry powder in the lung of the mouse are demonstrated.

Protocol

The experiments conducted in this study have been approved by the Committee on the Use of Live Animals for Teaching and Research (CULATR), The University of Hong Kong. Dry powder formulations prepared by spray freeze drying (SFD) containing 0.5% of luciferase messenger RNA (mRNA), 5% synthetic peptide PEG₁₂KL4 and 94.5% of mannitol (w/w) are used in this study to demonstrate mRNA expression in the lung¹⁶. The mass median aerodynamic diameter (MMAD) of SFD powder is 2.4 μm. Spray dried (SD) mannitol powder are used to investigate the effect of air volume used in powder dispersion¹⁶. The MMAD of SD powder is 1.5 μm.

1. Fabrication of dry powder insufflator and loading of dry powder

1. (Optional) Neutralize the static charges of dry powder (in a vial) and the 200 µL non-filter round gel-loading pipette tip. Use an anti-static gun or a balance with deionizing function according to the manufacturer’s instruction.
2. Prepare a weighing paper with a size of around 4 cm x 4 cm. Fold the paper in half diagonally and then unfold it.
3. Weigh 1-2 mg of dry powder on the weighing paper.
4. Fill a gel-loading pipette tip with powder through the wider opening of the tip. Tap gently to pack the powder until the powder forms loose agglomerates near the narrow end of the tip (Figure 1A). Avoid packing the powder too tightly as it may hamper powder dispersion.
5. Connect the powder-loaded tip to a 1 mL syringe through a three-way stopcock (Figure 1B). The size of syringe can be changed according to the volume of air used to disperse the powder. Hold the tip and syringe vertically during connection to prevent spillage of powder. If administration is not performed immediately, use parafilm to seal the openings of the tip and store it temporarily under suitable condition until administration.

2. Fabrication of intubation device

1. Light source (Figure 2)
   1. Prepare a custom-made light source with a light emitting diode (LED) torch and a flexible optical fiber with a diameter of 0.8-1 mm.
   2. Make a centered orifice on the clear lens of the LED torch with a hand drill or a drill bit so that the optical fiber can barely pass through.
   3. Insert the optical fiber through the orifice. Switch on the LED torch to adjust the position and the depth of insertion for maximum brightness at the other end of the optical fiber.
   4. Affix the optical fiber in position with clear epoxy glue.
2. Guiding cannula (Figure 3)
   1. Take a 1 mL plastic Pasteur pipette (Figure 3A) and hold the pipette at both ends.
   2. Use an alcohol lamp (or other heat sources in the laboratory such as a Bunsen burner) to heat the middle of the pipette by placing it at 5-10 cm above the flame (Figure 3B). Rotate the pipette to make sure it is heated evenly.
   3. When the plastic becomes soft and deformable, move the pipette away from the flame and stretch the pipette gently.
   4. Cut the stretched pipette in the middle with a pair of scissor into Part A and Part B (Figure 3C-E). Use Part A as a fine tip pipette and Part B as a guiding cannula. To increase the chance of successful intubation with the guiding cannula, make a bevel (not too sharp which may increase the risk of injuring the animal) at the end of Part B (Figure 3F). When a 200 µL gel-loading pipette tip (for powder loading) is inserted into the guiding cannula, it should protrude the cannula by 1-2 mm.
      NOTE: A guiding cannula (Part B) with the appropriate dimension (internal and external diameter) for intubation could have a 21 gauge needle fit inside it while it can also fit inside a 17 gauge needle. Multiple attempts may be needed in stretching the pipettes to achieve the appropriate dimension.
   5. (Optional): Cut a small opening at the wider end of the guiding cannula to make it more flexible so that it is easier to hold the optical fiber (Figure 3F). This opening also allows the fitting of a microsprayer for the administration of liquid aerosol.

3. Intubation

1. Anaesthetize the mouse (BALB/c, 7-9 weeks) with ketamine (100 mg/kg) and xylazine (10 mg/kg) by intraperitoneal injection.
2. Prepare a platform made of Plexiglass and mount it to a stand with a clamp (Figure 4A). Place the anaesthetized mouse on the platform (at around 60° of inclination) in a supine position. The height and angle of inclination of the platform could be adjusted by the position of the clamp on the stand.
3. Suspend the mouse by hooking its incisors on a nylon floss (Figure 4B). Secure the position of the mouse by a piece of tape or a rubber band.
4. Insert the optical fiber into the guiding cannula before intubation with the tip of the fiber level with the opening of the guiding cannula. Turn on the LED torch to illuminate.
5. Gently protrude the tongue of the mouse with a pair of forceps to expose its trachea.
6. Use the other hand to hold the guiding cannula with optical fiber inside. Insert them through the oral cavity. With the illumination from the optical fiber, the opening of the trachea can be visualized as an orifice between the vocal cords.
7. Align the bevel of the guiding cannula towards the midline of the opening (Figure 5A). Gently intubate the guiding cannula with optical fiber into the trachea by aiming the finest tip of the cannula at the tracheal opening.
8. Upon intubation, swiftly remove the optical fiber and leave the guiding cannula inside the trachea (Figure 5B). A normal respiration should be observed.
9. Hold the fine tip pipette (Part A) at the opening of the guiding cannula and insufflate a small puff of air (about 0.2 mL) into the lung of the mouse. A slight inflation in the chest of the mouse indicates proper intubation. Remove the fine tip pipette prior to powder administration.

4. Powder administration

1. Hold the powder loaded-tip that is connected to the syringe as described in step 1.5. Ensure the airflow between the syringe and the tip is disconnected.
2. Pull the syringe plunger backward to withdraw 0.6 mL of air.
   NOTE: The volume of air used to disperse the powder is dependent on the properties of the powder and the amount of powder loaded. This is further described in the result section.
3. Turn the valve of the three-way stopcock to connect the airflow between the syringe and the powder-loaded tip.
4. Insert the powder-loaded tip into the guiding cannula which has already been placed in the trachea of the mouse (Figure 5C). Hold the guiding cannula and push the syringe plunger forcefully in one continuous action to disperse the powder as aerosols into the lung.
   NOTE: Any forward motion of the device should be minimized to avoid injuring the animal.
5. Remove the tip and check if the powder inside the tip has been emptied. If not, repeat step 4.1 to 4.4.
   NOTE: If the powder is packed too tightly due to excessive tapping, it might not be dispersed properly.
6. Once the administration is complete, remove the guiding cannula from the trachea.
7. Allow the mouse to recover by positioning it horizontally in a supine position with its tongue half protruded to avoid the blockade of the airways.

Results

When a dry powder insufflator is used to deliver powder aerosol to the lung of an animal, the volume of air used is critical as it affects the safety as well as the powder dispersion efficiency. To optimize the method, different volumes of air (0.3 mL, 0.6 mL and 1.0 mL) were used to disperse the dry powder (1 mg of spray dried mannitol) and the weight of mice was monitored for 48 hours after administration (Figure 6). The use of 0.3 mL and 0.6 mL of air did not cause weight loss of the mice...

Discussion

In this paper, custom-made devices for dry powder insufflation and intratracheal intubation are presented. In the powder loading step, dry powder are loaded into a 200 µL gel-loading pipette tip. It is important to gently tap the tip to allow the loose packing of powder at the narrow end of the tip. However, if the powder are packed too tightly, they will get stuck in the tip and cannot be properly dispersed. It is recommended to neutralize the static charges of the powder and the pipette tip in order to facilitate ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
BALB/c mouse			Female; 7-9 weeks old; Body weight 20-25 g
CleanCap Firefly Luciferase mRNA	TriLink Biotechnology	L-7602
Dry Powder Insufflator	PennCentury	Model DP-4M
Ketamine 10%	Alfasan International B.V.	NA
Light emitting diode (LED) torch	Unilite Internation	PS-K1
Mannitol (Pearlitol 160C)	Roquette	450001
Non-filter round gel loading pipette tip (200 µL)	Labcon	1034-800-000
Nylon floss	Reach	30017050
One milliliter syringe without needle	Terumo	SS-01T
Optical fibre	Fibre Data	OMPF1000
PEG12KL4 peptide	EZ Biolab		(PEG12)-KLLLLKLLLLKLLLLKLLLLK-NH2
Plastic Pasteur fine tip pipette	Alpha Labotatories	LW4061
Three-way stopcock	Braun	D201
Xylazine 2%	Alfasan International B.V.	NA
Zerostat 3 anti-static gun	MILTY	5036694022153