Name: dry powder and nebulized aerosol inhalation of pharmaceuticals delivered to mice using a nose-only exposure system
Uploaded: 2017-04-06T12:30:00.000+00:00
Duration: 7 min 28 s
Description: Watch this Scientific Journal Video about Dry Powder and Nebulized Aerosol Inhalation of Pharmaceuticals Delivered to Mice Using a Nose-only Exposure System at JoVE.com

We acknowledge: Dr. Thomas Budiman at TSE Systems GmbH for his technical expertise and equipment customization. John Fry (Battelle Inc.) and Dr. Rudy Jaeger (CH Technologies Inc.) for their helpful discussions. Tian Wu, Sam Mboggo, April Miller, and Sean Davis (Amgen) for help with the experiments.

The mice used in these studies were cared for in accordance with the Guide for the Care and Use of Laboratory Animals, eighth edition24. Mice were group housed at an Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC) international-accredited facility in sterile ventilated microisolator housing on corn cob bedding. When measuring bronchoconstriction, mice were anesthetized with 100 mg/kg i.p. pentobarbital and depth of anesthesia was monitored by lack of toe pinch reflex and maintained by i.p. anesthesia as necessary. At the end of the experiments, mice were euthanized by cervical dislocation after barbiturate overdose. Verification of euthanasia was confirmed by lack of respiration. No survival surgeries were performed on the mice. All research protocols were approved by the Institutional Animal Care and Use Committee (IACUC).
1. Formulation and Device Selection for Generation of Pharmaceutical Aerosols
NOTE: Formulation and device selection are dependent on the physicochemical properties of the individual drug to be aerosolized, therefore general protocols are presented below and the reader is referred to the reviews by Zeng25 and O&#39;Riordan26.
<ol>
	<li>Dry powder aerosols
	<ol>
		<li>Micronize drug dry powders in a ball mill, jet mill or similar device27and ensure the micronized particle size distribution (PSD) contains particles of respirable (0.5-5 &#956;m mass median aerodynamic diameter, MMAD) particle size. Blend potent test compounds that require dilution with micronized lactose. 
		NOTE: If there is not enough micronized powder to determine the PSD with a cascade impactor, the PSD of a small (sub-milligram) sample of the micronized powder can be measured by light scattering to confirm it contains small/respirable particles.</li>
		<li>Generate the dry powder aerosol using a Wright dust feed dry powder aerosol generator. Pack the micronized drug/lactose powder into the cylindrical reservoir using a manual hydraulic press at approximately 1000 pounds per square inch (psi) to produce compacted cakes of powder used as input by the Wright dust feed aerosol generator17.</li>
		<li>Screw the cylindrical reservoir onto the Wright dust feed advancing the reservoir until the scraper blade is in contact with the drug cake.</li>
		<li>Connect the outlet of the Wright dust feed to a cyclone and the inlet to a compressed air source set to 15 L/min airflow rate (maximum pressure 90 psi).</li>
		<li>Set the feed rate control to 0.7 revolution per minute (rpm) and turn on the Wright dust feed aerosol generator. 
		NOTE: 0.7 rpm corresponds to a test article cake feed rate of 1 g/h when using the small Wright dust feed cylindrical reservoir. The Wright dust feed scrapes a thin layer of compacted powder off the test article cake by rotating the reservoir. Air carries the dust out of the Wright dust feed, through a sonic nozzle for de-agglomeration, and into a cyclone to remove non-respirable particles and agglomerates.</li>
		<li>Connect the outlet of the cyclone to the central aerosol plenum of the inhalation unit (Figure 1). 
		NOTE: Compounds can be compressed from 300 to 1500 psi in the reservoir of the Wright dust feed. The object is to compress the particles enough so they will be retained in the reservoir when inverted, but not so much that the Wright dust feed cannot scrape off a thin layer for re-aerosolization. It must be remembered that the gauge on the manual hydraulic press reads in pounds and the plunger on the small dust feed reservoir has a surface area of about 0.25 square-inch. Therefore, 250 pounds of compression force on 0.25 square inch is equivalent to 1000 psi.</li>
	</ol>
	</li>
	<li>Nebulized liquid aerosols
	<ol>
		<li>Dissolve the drug in 100 mL of water or physiologic saline.</li>
		<li>Load a 100 mL syringe with the drug solution and place the syringe into the syringe pump with a flow rate set at 1 mL/min.</li>
		<li>Connect the syringe pump to the jet nebulizer and purge the air from the feed line leading to the nebulizer.</li>
		<li>Connect the pressurized air source to the jet nebulizer and set the air flow meter to 10 L/min.</li>
		<li>Insert the jet nebulizer into the pre-separator. The pre-separator connects the nebulizer to the central aerosol plenum of the inhalation unit (Figure 1). 
		NOTE: Many drug compounds have limited aqueous solubility and are better formulated as dry powder aerosols. If a stable suspension can be made from micronized (MMAD &#60; 5 &#956;M) compound, it may be used with the jet nebulizer. Caution should be used as the suspension may clog the nebulizer. Nebulizer feed concentrations from 1 mg/mL for potent compounds (like the bronchodilator ipratropium) to 40 mg/mL suspensions (for less potent compounds like salbutamol sulfate) have been used. The syringe pump feed rate is set at 1 mL/min for a practical reason; to allow the aerosol concentration equilibration period and 45-min-long exposure to be completed without the need to reload the syringe.</li>
	</ol>
	</li>
</ol>
2. Aerosol Exposure Experiment Setup
<ol>
	<li>Measure the drug concentration (by collecting aerosol on an absolute filter) and particle size distribution (by collecting aerosol using a cascade impactor) of the aerosol that enters the inhalation unit post pre-separator/cyclone. Use these parameters along with the animal&#39;s minute ventilation, body weight, and exposure time to estimate the dose of drug deposited in the lungs.
	<ol>
		<li>Weigh the absolute filter and record the filter weight. Place the filter into the filter holder and assemble the filter holder. Connect the inlet of the absolute filter holder to a central aerosol plenum sample port and the outlet to a vacuum source set to sample the aerosol at a flow rate of 1 L/min for the duration of the experiment. 
		NOTE: The mass of drug on the filter after sampling for 45 min can be in the sub-microgram range and/or be mixed with lactose, NaCl salt, or other vehicle. A microbalance that reads to 0.1 microgram is necessary. To get an accurate weight of drug deposited on the filter, the filter must be equilibrated and weighed in a humidity controlled environment. The weight of drug on the filters can only be used in dose calculation if there is no vehicle in the formulation or the vehicle is water. When there is a vehicle in the formulation other than water, the weight of substance on the filter only gives an estimated starting point for further analysis of drug content by high performance liquid chromatography (HPLC).</li>
		<li>Weigh and record the weight of the 7 cascade impactor stage filters and one final &#34;tail piece&#34; filter. Place a filter on each of the seven stages of the cascade impactor and assemble the Mercer cascade impactor28. Connect the inlet of the cascade impactor to a central aerosol plenum sample port and the outlet to a vacuum source set to sample the aerosol at the flow rate the cascade impactor is calibrated at (usually 0.5 or 1 L/min) for the duration of the experiment.</li>
		<li>Monitor the aerosol content of the inhalation unit with the real time aerosol monitor (see Table of Materials and Reagents) to confirm the aerosol generators are functional and producing a stable aerosol throughout the experiment. Connect the inlet of the real time aerosol monitor to a central aerosol plenum sample port and the outlet to a vacuum source set to sample the aerosol at a flow rate of 1 L/min for the duration of the experiment. 
		NOTE: The signal from the real time aerosol monitor is reported in &#956;g/L, but is calibrated for road dust and must be recalibrated for each individual drug aerosol to yield correct aerosol concentration values. Calibration is not needed to use the monitor to confirm the presence or absence and temporal stability of aerosol concentration.</li>
		<li>Set the process control parameters (air flow, vacuum, pressure, aerosol generator power) to the values required that depends on the number of animals to be connected to the inhalation unit. The inhalation unit and aerosol generators are continuously controlled/monitored by a computerized process control/data acquisition system (DACO) supplied by the manufacturer (see Table of Materials and Reagents). Air flow rate into the inhalation unit should be minimally approximately 2 times the total minute ventilation rate of all animals in the inhalation unit in order to avoid a buildup of CO2.</li>
	</ol>
	</li>
	<li>Load mice into nose-only restrainers before exposing to aerosol in the nose-only inhalation unit. Also load restraint stress control animals into restrainers to breathe room air. 
	NOTE: A dose/response experiment consists of multiple groups of mice exposed to aerosol for different amounts of time. Exposure time is used to control the dose each group receives during a dose/response experiment.
	<ol>
		<li>Angle the restraining tube up towards the ceiling while trying to load the animals, as they tend to run upward trying to escape. Pointing the tubes down while loading will encourage turning around and escaping out the back of the tube. Ensure the mouse&#39;s nose is oriented into the pointed end of the tube and fix the variable-position plunger into the back end of the restrainer. 
		NOTE: The plunger is fashioned to allow the tail of the mouse to protrude from the holder, which allows the mouse to regulate its body temperature while in the restrainer.</li>
		<li>Adjust the plunger to allow the mice to rotate, but not turn head to tail; to ensure the aerosol is inhaled.</li>
		<li>Continuously monitor the mice while in the restraining tubes. After positioning the plunger, small mice (&#60;20 grams) often attempt to turn head to tail in the tubes and adopt a U-position where they have trouble breathing. This turning behavior is most prevalent in the first 5 minutes of restraint, after that the mice rarely attempt to turn head to tail.</li>
	</ol>
	</li>
</ol>
3. Aerosol Delivery
<ol>
	<li>Insert stoppers to plug the delivery ports of the inhalation unit and activate the aerosol generator, compressed air flow controller, and inhalation unit vacuum pump from within the process control software.</li>
	<li>Once the readings from the real time aerosol monitor demonstrate the aerosol concentration has to come to equilibrium (~30 min, Figure 2), begin removing stoppers and inserting the nose-only restraining tubes containing mice into the inhalation unit. Repeat until all mice to be exposed to drug are connected to the inhalation unit. 
	NOTE: In the example experiment, the total air flow supplied to the inhalation unit via the aerosol generator and dilution air is set to supply a 0.5 L/min flow rate to each of animal exposure ports in use. For example, a 15 L/min total air flow is enough to supply each of the 30 ports in the inhalation unit. This is much more air flow than required by the minute ventilation of the mice, but a larger air flow is necessary to supply the energy (pressure drop across the aerosol generator) to produce (atomize/de-agglomerate) the aerosol.</li>
	<li>Once all animals are loaded into the exposure unit, turn on the vacuum sampling pumps that are connected to the absolute filter and cascade impactor using the process control software.</li>
	<li>When all exposures are completed, turn off the aerosol generator and remove remaining mice from inhalation unit. 
	NOTE: Animals will be handled in a change station hood or by personnel wearing a facemask. After conclusion of aerosol delivery, the mice are removed from the tubes and the tubes are sanitized after each use.</li>
</ol>
4. Calculation of the Deposited Dose
<ol>
	<li>To calculate the deposited dose29 in &#956;g/kg (Equation 1) multiply the concentration of the drug in the aerosol (&#956;g/L) x minute ventilation (L/min) x duration of exposure (min) x inhalable fraction x pulmonary deposition fraction and divide by the body weight (kg).</li>
	<li>Calculate the aerosol concentration (&#956;g/L) by dividing the mass of drug on the absolute filter (&#956;g) by the air flow rate through the filter (L/min) multiplied by the sampling time (min). Estimate the mouse&#39;s minute ventilation by an allometric equation based on body weight30. The inhalable fraction is 1 as the aerosol was passed through a pre-separator to remove non-respirable particles, and the pulmonary deposition fraction is determined from the drug aerosol MMAD (Figure 3) using experimental calibration curves from monodisperse aerosols (Figure 4)31.</li>
</ol>
<img alt="Equation" src="/files/ftp_upload/55454/55454eq1.jpg" />

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Species+comparison+of+drug+absorption+from+the+lung+after+aerosol+inhalation+or+intratracheal+injection.">Species comparison of drug absorption from the lung after aerosol inhalation or intratracheal injection.</a> Drug Metab Dispos. 14 (1), 79-88 (1986).</li><li>Biju, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Steroidal+C-21+mercapto+derivatives+as+dissociated+steroids:+discovery+of+an+inhaled+dissociated+steroid.">Steroidal C-21 mercapto derivatives as dissociated steroids: discovery of an inhaled dissociated steroid.</a> Bioorg Med Chem Lett. 21 (21), 6343-6347 (2011).</li><li>West, L. J., Pierce, C. M., Thomas, W. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lysergic+acid+diethylamide:+Its+effects+on+a+male+asiatic+elephant.">Lysergic acid diethylamide: Its effects on a male asiatic elephant.</a> Science. 138 (3545), 1100-1103 (1962).</li><li>Boxenbaum, H., DiLea, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=First-time-in-human+dose+selection:+allometric+thoughts+and+perspectives.">First-time-in-human dose selection: allometric thoughts and perspectives.</a> J Clin Pharmacol. 35 (10), 957-966 (1995).</li><li>Spina, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Current+and+novel+bronchodilators+in+respiratory+disease.">Current and novel bronchodilators in respiratory disease.</a> Curr Opin Pulm Med. 20 (1), 73-86 (2014).</li><li>Degeorge, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Considerations+for+toxicology+studies+of+respiratory+drug+products.">Considerations for toxicology studies of respiratory drug products.</a> Regul.Toxicol.Pharmacol. 25 (2), 189-193 (1997).</li><li>McClellan, R. O., Henderson, R. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Concepts in Inhalation Toxicology. , (1989).</li><li>Patton, J. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+particle+has+landed--characterizing+the+fate+of+inhaled+pharmaceuticals.">The particle has landed--characterizing the fate of inhaled pharmaceuticals.</a> J Aerosol Med Pulm Drug Deliv. 23 Suppl 2, S71-S87 (2010).</li><li>Depreter, F., Pilcer, G., Amighi, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhaled+proteins:+Challenges+and+perspectives.">Inhaled proteins: Challenges and perspectives.</a> Int J Pharm. 447 (1-2), 251-280 (2013).</li></ol>

All authors employed by Amgen. Publication fees for this this video-article are paid by Amgen.

A nose-only inhalation system and its operation to deliver pharmaceutical aerosols to the lungs of rodents has been described. Restraining animals in nose-only holders is a commonly used method for exposing animals to airborne material. An experiment was conducted demonstrating the anticholinergic bronchodilator ipratropium bromide34 can potently reverse methacholine induced bronchoconstriction when delivered by nose-only inhalation to mice with an ED50 of 0.1 &#956;g/kg deposited dose. A greater than 10-fold increase in effective dose of ipratropium was required for bronchodilation following intratracheal delivery (ED50intratracheal = 1.3 &#956;g/kg, data not shown) delivery. This is due to the non-homogeneous deposition pattern of drug in the lungs produced by intratracheal dosing3,5,10. 10-fold and greater effective dose differentials between inhaled and intratracheal dosing techniques for other drugs have been observed previously by others4.
The non-homogeneous deposition pattern of drug in the lungs produced by intratracheal dosing also slows drug absorption by the lung35, decreasing the rate at which the drug enters the systemic circulation and decreasing the chances of seeing systemic side effects. Therefore, to optimize the safety/efficacy (therapeutic index)36 of novel inhaled drugs, nose-only inhalation delivery should be used. Intratracheal deposition will give an inaccurate estimate of the TI by generating erroneously high effective doses in the lung and low doses to the systemic circulation.
As new drugs delivered by the inhaled route are developed, it is essential to appropriately translate the efficacious drug dose from preclinical efficacy studies to predict an efficacious human dose for clinical trials. The infamous death of Tusko the elephant37 is commonly cited in the literature to remind us to use allometric scaling to predict interspecies drug doses and that interspecies drug doses should not be linearly extrapolated on the basis of a simple comparison of body masses. It is common to use an allometric approach with an allometric exponent b of 0.67 to predict human drug doses from preclinical efficacy studies38. Using the mouse inhaled ED90 of 0.9 &#956;g/kg for ipratropium, an allometric exponent of 0.67, a mouse body mass of 0.03 kg, and a human body mass of 60 kg; an estimated human ED90 of 0.07 &#956;g/kg ((0.9*(0.03/60)(1-0.67) = 0.07) can be calculated as the predicted human deposited dose from our mouse data. This predicted value is comparable to the actual human efficacious deposited dose of 0.26 &#956;g/kg, which can be calculated from the delivered dose of 40 &#956;g39 divided by the human body mass of 60 kg multiplied by a human oral inhalation pulmonary deposition fraction16 of 0.4 ((40/60)*0.4 = 0.26). An estimate of the effective human inhaled deposited dose also helps to plan the delivered doses used in inhalation toxicology studies required for clinical trials40.
The specialized equipment and large amount of test compound (gram quantities) required for nose-only inhaled drug delivery can be significant limitations when developing the nose-only inhalation technique. A calibration run without animals present is necessary if a specific dose is required for a (usually toxicology and not an efficacy dose/response) study, and this calibration run will require more drug to be available. This calibration run is necessary because the physicochemical properties of each drug/formulation can vary enough to greatly affect the deposited dose of drug in the lungs. Optimization of the air and drug feed rates to the aerosol generator during the calibration run is required to achieve an aerosol particle size and concentration for the specific dose required. While the aerosol generator air and drug feed rates suggested in the methods are a reasonable starting point, there is potential that for a specific drug formulation a non-respirable (particle size &#62;5 mm) aerosol will be generated. At the preclinical stage, there is often not enough drug available to do a calibration run and it is impossible to know a priori what doses (if any) are getting into the lungs. Also, static electricity is produced during the aerosol generation process and can influence the time required for the aerosol concentration to equilibrate. It is important to properly ground the equipment to minimize static charge. Another option to minimize static electricity is to add humidity to the air supplied to the inhalation unit, to increase conductivity and dissipate electrostatic charges on particles25. Humidifying the air supplied to the aerosol generator is not required for animal comfort during short (&#60; 1 h) dosing sessions but should be considered if longer dosing times are used.
Drugs can be delivered to the lungs of animals by passive nose-only inhalation or direct intratracheal administration methods that bypass nasopharyngeal deposition. Nose-only inhalation delivery is commonly used in the field of inhalation toxicology41 but is sparingly used early in the drug discovery process. A multidisciplinary research team is needed for conducting nose-only inhalation studies due to the: need for large quantities of drug, specialized knowledge required to formulate, generate, and characterize aerosols; operate complex equipment, and measure drug efficacy in animal models of respiratory disease. The delivery techniques described herein are used to develop small molecule inhaled drugs but in the future can be applied to develop inhaled biologics42,43. Hopefully the procedures and tips documented in this manuscript will facilitate new pre-clinical drug discovery and toxicology researchers to acquire the skills necessary to deliver drugs to rodents by aerosol inhalation.

There are many advantages to inhaled administration of drugs for the treatment of respiratory disease. Inhaled delivery applies the therapeutic agent directly to the site of action, the lungs. A high local concentration of the drug in the lungs offers a significant advantage that minimizes dose and systemic exposure, and maximizes efficacy. This is an important advantage that can greatly increase the therapeutic index (TI, ratio of dose of drug that causes a side-effect over the dose of drug that provides efficacy) of a drug. Inhaled &#946;2-adrenergic agonists, corticosteroids, and anti-cholinergic drugs have proven effective at improving lung function in patients with asthma and COPD, while minimizing the systemic side effects (tachycardia, immunosuppression, and constipation) observed when these drugs are taken orally. New drug classes (for example, PDE4 inhibitors1 and BTK inhibitors2) have recently proven effective at improving lung function in pre-clinical animal disease models but, similar to &#946;2-agonists, corticosteroids, and anti-cholinergic drugs, suffer from systemic side effects that may be minimized by inhaled delivery. Due to the added cost of development of inhaled vs. oral drugs, an inhaled formulation should only be considered for respiratory indications after successful oral/systemic administration reveals dose limiting mechanism-based systemic side effects.
Pre-clinically, inhaled compounds are optimized to increase the TI, which requires in vivo efficacy and side effect measurements. Initially these measurements can be made in separate assays, usually a topically delivered efficacy measurement and a systemically delivered side-effect measurement, but to truly compare compounds, efficacy and side effects should be measured in the same animals after inhaled administration. This requires dose/response studies that achieve enough compound administered to the lungs to induce a measureable side effect. The only way currently to uniformly distribute large doses of drug into the lungs of multiple small animals simultaneously is nose-only inhalation3,4,5. The strengths and weaknesses of different inhalation exposure techniques have been recently reviewed6,7,8. Specialized equipment and a large amount of test compound (gram quantities) are required for nose-only inhaled drug delivery, but proof-of-concept studies may be possible by other means.
When the amount of drug is limited (mg quantities), direct administration methods are an option but all suffer from non-homogeneous deposition, with more drug concentrated along the central airways and less well represented in the parenchymal/alveolar regions3,4,5. The effective dose delivered by direct instillation is always higher and can never be directly compared to inhaled doses4. Direct instillation methods including intranasal9, intratracheal liquid10,11 and spray instillation12, or dry-powder insufflation13,14 can be used as a screening tool to determine the approximate dose range for later nose-only inhalation studies, or to determine the ranking of efficacy/toxicity for a series of structurally similar drugs15. Due to the central airway deposition pattern, direct administration methods may be more useful to determine the effects of compounds that act on the central airways (bronchodilators or mast cell inhibitors) than in the peripheral lung (anti-inflammatories).
Unlike humans, who can inhale a substantial dose of concentrated aerosol from an inhaler in a single deep breath, continuous generation of a respirable (0.5-5 &#956;m mass median aerodynamic diameter, MMAD) aerosol, for up to an hour, is required to deposit an efficacious drug dose into a spontaneously breathing rodent&#39;s lungs in a nose-only inhalation system16. Aerosol generators (jet nebulizer or the Wright dust feed17) that can continuously produce the required aerosol particle size and concentration for nose-only inhalation studies are not very efficient at generating high quality (respirable) aerosols. The drug feed rates to these aerosol generators for potent (IC50 pM to nM in functional cell based assays) compounds are commonly in the 1 to 10 mg/min range and usually less than 1% of that drug aerosol makes it to the breathing zone of the animals (Figure 1). Many of the particles generated are too big (&#62;5 &#956;m) to enter the lungs and are removed by an aerosol classifier (a pre-separator or cyclone with a 5 &#956;m cut point) to avoid a large dose of drug in the nose. Adding to the inefficiency of nose-only inhalation systems is the small particle size range (0.5 to 5 &#956;m MMAD) for respirable particles. Many of the aerosol particles less than 0.5 &#181;m are exhaled (like cigarette smoke) and not deposited in the lungs18. Also, many of the &#34;larger&#34; aerosol particles (~ 5 &#181;m) deposit in the nose are absorbed or transported by mucociliary clearance towards the back of the throat where they are swallowed into the stomach19. When using nose-only inhalation, the dose deposited in the nose is always larger than the dose deposited in the lungs and the nasal dose can contribute to systemic exposure and side-effects20. Inherently, inhaled drug doses are small (in the microgram range) minimizing any systemic side effect potential of drug absorbed by the nasal, lung, or gastrointestinal tissues. Even when the particle size of the aerosol supplied to the animals is in the respirable range, an average of only 4% of the aerosol particles that make it to the breathing zone of the animals deposit in the lungs. More efficient aerosol generators are available, but the jet nebulizer and Wright dust feed are unmatched for their ability to produce continuous aerosols from diverse liquid and dry powder formulations, respectively.
Respirable aerosol from the pre-separator passes into the nose-only exposure inhalation unit21 that is based on a flow-past the nose design22. The inhalation unit has 3 tiers (only two tiers are shown in Figure 1) and each tier contains 10 exposure ports for animals and aerosol sampling. The ports are located peripherally around the central aerosol plenum. Conscious mice are placed in glass restraining holders (6 inches long by 1.2 inch diameter) and breathe the aerosol contents of the inhalation unit. The mice are not acclimated to the restraint devices23. Previous experience has shown that mice tolerate tube restraint of less than an hour duration similarly, with or without adaptation2.
The inhalation unit is designed to deliver drug aerosols directly to the lungs of the animals while avoiding exposure to the operators. The potency/toxicity of these drugs is usually unknown and multiple engineering safety controls are used to avoid exposure to the operators. The operators must always wear personal protective equipment (gloves, laboratory coats, respirators, and safety glasses). The outer plenum of the inhalation unit is under negative pressure at all times during operation, allowing for the safe removal of single or groups of animals without shutting off the aerosol generator. The inhalation unit is also contained in a secondary enclosure maintained at a negative pressure by an exhaust port in the ceiling to prevent any escape of aerosol into the room in case of malfunction. All effluent air from the inhalation system is filtered by a HEPA filter prior to release into the environment. The nose-only exposure system used in this manuscript was purchased from a single vendor (see supplemental Table of Materials).

<table><tbody><tr><td>Nose-only exposure inhalation unit</td><td>TSE systems</td><td>700100-KNES-040-ss</td><td>Custom configurations available</td></tr><tr><td>DACO data acquisition system</td><td>TSE systems</td><td>700400-PRO-C-D/1</td><td></td></tr><tr><td>MC One Jet Mill</td><td>Jetpharma</td><td>DEC MicroJet 10</td><td></td></tr><tr><td>Turbula Mixer</td><td>GlenMills Inc</td><td>T2F</td><td></td></tr><tr><td>Micronized Lactose</td><td>DFE Pharma</td><td>Lactohale 200</td><td></td></tr><tr><td>Hydraulic press</td><td>Specac</td><td>GS15011</td><td></td></tr><tr><td>Cascade impactor filters</td><td>Pall Life Sciences</td><td>7219</td><td>Emfab filter</td></tr><tr><td>Absolute filters</td><td>Whatman</td><td>10370302</td><td>5 cm diameter</td></tr><tr><td>Real time aerosol monitor&lt;br/&gt; Microdust Pro Monitor</td><td>Casella</td><td>CEL-712</td><td></td></tr><tr><td>Ipratropium bromide</td><td>Spectrum Chemical</td><td>I1178</td><td>pre-micronized</td></tr><tr><td>flexiVent FX1 system</td><td>scireq</td><td>FV-FXCS</td><td></td></tr></tbody></table>

dry powder and nebulized aerosol inhalation of pharmaceuticals delivered to mice using a nose-only exposure system

Obstructive respiratory diseases like asthma and chronic obstructive pulmonary disease (COPD) are currently treated by inhaled anti-inflammatory and bronchodilator drugs. Despite the availability of multiple treatments, both diseases are growing public health concerns. The majority of asthma patients are well controlled on current inhaled therapies but a substantial number of patients with severe asthma are not. Asthma affects an estimated 300 million people worldwide and approximately 20 percent have a severe form of the disease. In contrast to asthma, there are few effective therapies for COPD. An estimated 10% of the population has COPD and the trend in death rates is increasing for COPD while decreasing for other major diseases. Although developing drugs for inhaled delivery is challenging, the nose-only inhalation unit enables direct delivery of novel drugs to the lung of rodents for pre-clinical efficacy and safety/toxicology studies. Inhaled drug delivery has multiple advantages for respiratory diseases, where high concentration in the lung improves efficacy and minimizes systemic side effects. Inhaled corticosteroids and bronchodilators benefit from these advantages and inhaled delivery may also hold potential for future biologic therapies. The inhalation unit described herein can generate, sample for characterization, and uniformly deposit a drug aerosol in the lungs of rodents. This enables the pre-clinical determination of the efficacy and safety of drug doses deposited in the lungs of rodents, key data required before initiating clinical development.

The bronchodilator ipratropium bromide was dissolved in 0.9% normal saline at a concentration of 1 mg/mL. A 100 mL syringe was filled with the ipratropium formulation solution and the syringe was inserted into a syringe pump set to feed the jet nebulizer (Figure 1) at a flow rate of 1 mL/min. Activating the control system for the inhalation unit initiates a 10 L/min air flow to the jet nebulizer, 5 L/min dilution air flow to the dilution chamber, and 15.5 L/min of vacuum air flow to draw the drug aerosol out of the inhalation unit once it passes the nose of the mice. The syringe pump and real time aerosol monitor sample pump were activated. Twenty-four mice were inserted into the inhalation unit after the real time aerosol monitor confirms the aerosol concentration in the inhalation unit had come to equilibrium (15-30 min, Figure 2). The sample pumps for the absolute filter (1 L/min) and cascade impactor (0.5 L/min) were turned on once all mice were connected to the inhalation unit. The first group of 8 mice was removed from the inhalation unit after 5 min, the second group after 15 min, and the third group after 45 min. Bronchoconstriction32 was measured as the increase in respiratory system resistance (Rrs) to a single nebulized dose of methacholine (30 mg/mL) delivered 2 h after dosing ipratropium using the rodent respirator 33. The percent increase in Rrs for each animal, calculated as the maximum Rrs over the 3-min interval after nebulized methacholine (Rmax) minus the Rrs value of the baseline measurement before methacholine (Rbase) divided by Rbase (% increase in Rrs = (Rmax-Rbase)/Rbase), was used to quantitate the bronchoconstriction.
The aerosol deposited on the cascade impactor filters and absolute filter during the 45 min of aerosol exposure was dissolved in 50% acetonitrile and the mass of ipratropium quantitated by HPLC (Table 1). The MMAD x/ GSD of the ipratropium aerosol was calculated to be 1.7 x/ 1.5 &#956;m (Figure 3) and a deposition fraction (DF) of 0.037 for mice was used for an aerosol with an MMAD of 1.7 &#956;m (Figure 4). The inhalable fraction (IF) was 1 as the aerosol was passed through a pre-separator to remove non-respirable particles. The average concentration of ipratropium in the aerosol (0.5 &#956;g/L) was calculated from mass of ipratropium on the absolute filter (22 &#956;g) divided by the flow rate of air pulled through the filter (1 L/min) multiplied by the sampling time (45 min). The average body weight of the mice was 0.019 kg and their predicted minute ventilation was calculated as 0.021 L/min. Using Equation 1, the deposited doses for the 5, 15, and 45 min exposure groups were 0.1, 0.3, and 0.9 &#956;g/kg; respectively.
Ipratropium inhibited nebulized methacholine induced bronchoconstriction in 8 week old C57Bl/6 mice. Methacholine increased Rrs within seconds of nebulized administration (Figure 5, top panel). The Rrs of the control group (exposed to room air instead of ipratropium aerosol) increased from a baseline value of 0.62 &#177; 0.03 cmH2O*s/mL to a maximum value of 1.66 &#177; 0.12 cmH2O*s/mL, representing a 168 &#177; 9% increase in Rrs, 70 s after methacholine aerosol administration. The percent increase in respiratory system resistance was inhibited in a dose dependent manner by inhaled doses of ipratropium. The percent increase in Rrs was inhibited by 51 &#177; 9%, 79 &#177; 14%, and 89 &#177; 2% (Figure 5, bottom panel) at inhaled deposited doses of ipratropium of 0.1, 0.3, and 0.9 &#181;g/kg, respectively.
<img alt="Figure 1" src="/files/ftp_upload/55454/55454fig1.jpg" /> 
Figure 1: Inhalation unit with dry powder aerosol generator attached. The nebulizer used to generate aerosol from liquid formulations is shown on the right. 1) Wright dust feed, 2) cyclone, 3) syringe pump, 4) jet nebulizer inserted into pre-separator, 5) dilution air mixer. When delivering nebulized aerosols, the dust feed and cyclone are replaced by the syringe pump, jet nebulizer/pre-separator, and dilution air mixer. The enlarged view (modified from Oldham21) shows the aerosol flow path at the breathing zone around animal&#39;s nose that is created by inserting the restraining tube into the inhalation unit. The aerosol fills the central aerosol (gray) plenum of the inhalation unit, flows out to the animal&#39;s nose where it is pulled back by a slight vacuum in the outer (white) plenum and into the waste collection filters (not shown). <a href="http://cloudflare.jove.com/files/ftp_upload/55454/55454fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55454/55454fig2.jpg" /> 
Figure 2: Real time aerosol monitor measurements confirm the aerosol concentration in the inhalation unit reaches equilibrium ~30 min after turning on the aerosol generator. <a href="http://cloudflare.jove.com/files/ftp_upload/55454/55454fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55454/55454fig3.jpg" /> 
Figure 3: Ipratropium mass collected on each stage of the cascade impactor (blue bar graph) overlaid with curve fit (black curve) used to calculate the MMAD and GSD of the aerosol particle size distribution. <a href="http://cloudflare.jove.com/files/ftp_upload/55454/55454fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/55454/55454fig4.jpg" /> 
Figure 4: Deposition fraction for mouse lungs as a function of MMAD for aerosols delivered by nose-only inhalation (as modified from Hsieh31). For particle sizes less than 0.5 &#181;m most of the aerosol is exhaled and does not deposit in the lungs (like cigarette smoke). For particle sizes greater than 5 &#181;m, most of the aerosol is filtered out by the nose. <a href="http://cloudflare.jove.com/files/ftp_upload/55454/55454fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/55454/55454fig5.jpg" /> 
Figure 5: The rapid increase in respiratory system resistance induced by nebulized methacholine administered to mice is blocked by ipratropium delivered by nose-only inhalation to mice (n = 8 per group, top panel). Ipratropium inhibited the methacholine induced increase in respiratory system resistance with an ED50 of 0.1 &#181;g/kg deposited dose (*p &#60;0.05 vs. control, bottom panel). <a href="http://cloudflare.jove.com/files/ftp_upload/55454/55454fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="3">Cascade impactor data used to calculate MMAD and GSD</td>
		</tr>
		<tr>
			<td>Stage Filter</td>
			<td>Ipratropium on filter 
			by HPLC (mg)</td>
			<td>Stage Cut 
			Diameter (mm)</td>
		</tr>
		<tr>
			<td>1</td>
			<td>0.002</td>
			<td>6</td>
		</tr>
		<tr>
			<td>2</td>
			<td>0.19</td>
			<td>4.5</td>
		</tr>
		<tr>
			<td>3</td>
			<td>1.43</td>
			<td>2.5</td>
		</tr>
		<tr>
			<td>4</td>
			<td>2.55</td>
			<td>1.5</td>
		</tr>
		<tr>
			<td>5</td>
			<td>2.95</td>
			<td>1.2</td>
		</tr>
		<tr>
			<td>6</td>
			<td>1.03</td>
			<td>0.7</td>
		</tr>
		<tr>
			<td>7</td>
			<td>0.37</td>
			<td>0.5</td>
		</tr>
	</tbody>
</table>
Table 1: Cascade impactor data used to calculate MMAD and GSD

Watch this Scientific Journal Video about Dry Powder and Nebulized Aerosol Inhalation of Pharmaceuticals Delivered to Mice Using a Nose-only Exposure System at JoVE.com

Dry Powder and Nebulized Aerosol Inhalation of Pharmaceuticals Delivered to Mice Using a Nose-only Exposure System

The inhalation unit described herein can generate, sample for characterization, and uniformly deposit a drug aerosol in the lungs of rodents. This enables the pre-clinical determination of the efficacy and safety of drug doses deposited in the lungs; key data enabling clinical inhaled drug development.

Obstructive respiratory diseases like asthma and chronic obstructive pulmonary disease (COPD) are currently treated by inhaled anti-inflammatory and ...

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40.5K Views.  Amgen. The inhalation unit described herein can generate, sample for characterization, and uniformly deposit a drug aerosol in the lungs of rodents. This enables the pre-clinical determination of the efficacy and safety of drug doses deposited in the lungs; key data enabling clinical inhaled drug development.

Video: Dry Powder and Nebulized Aerosol Inhalation of Pharmaceuticals Delivered to Mice Using a Nose-only Exposure System

Intraoperative Detection of Subtle Endometriosis: A Novel Paradigm for Detection and Treatment of Pelvic Pain Associated with the Loss of Peritoneal Integrity

Endometriosis is a common disease affecting 40 to 70% of reproductive-aged women with chronic pelvic pain (CPP) and/or infertility. The purpose of this study was to demonstrate the use of a blue dye (methylene blue) to stain peritoneal surfaces during laparoscopy (L/S) to detect the loss of peritoneal integrity in patients with pelvic pain and suspected endometriosis. Forty women with CPP and 5 women without pain were evaluated in this pilot study. During L/S, concentrated dye was sprayed onto peritoneal surfaces, then aspirated and rinsed with Lactated Ringers solution. Areas of localized dye uptake were evaluated for the presence of visible endometriotic lesions. Areas of intense peritoneal staining were resected and some fixed in 2.5% buffered gluteraldehyde and examined by scanning (SEM) electron microscopy. Blue dye uptake was more common in women with endometriosis and chronic pelvic pain than controls (85% vs. 40%). Resection of the blue stained areas revealed endometriosis by SEM and loss of peritoneal cell-cell contact compared to normal, non-staining peritoneum. Affected peritoneum was associated with visible endometriotic implants in most but not all patients. Subjective pain relief was reported in 80% of subjects. Based on scanning electron microscopy, we conclude that endometrial cells extend well beyond visible implants of endometriosis and appear to disrupt the underlying mesothelium. Subtle lesions of endometriosis could therefore cause pelvic pain by disruption of peritoneal integrity, allowing menstrual or ovulatory blood and associated pain factors access to underlying sensory nerves. Complete resection of affected peritoneum may provide a better long-term treatment for endometriosis and CPP. This simple technique appears to improve detection of subtle or near invisible endometriosis in women with CPP and minimal visual findings at L/S and may serve to elevate diagnostic accuracy for endometriosis at laparoscopy.

Loss of peritoneal integrity provides a new paradigm to understand and treat chronic pelvic pain in women with mild forms of endometriosis and can be easily detected using intraoperative instillation of dye at the time of laparoscopy.

Endometriosis is a  common disease affecting 40 to 70% of reproductive-aged women with chronic  pelvic pain (CPP) and/or infertility. The purpose of this ...

Intranasal Administration of CNS Therapeutics to Awake Mice

Intranasal administration is a method of delivering therapeutic agents to the central nervous system (CNS). It is non-invasive and allows large molecules that do not cross the blood-brain barrier access to the CNS. Drugs are directly targeted to the CNS with intranasal delivery, reducing systemic exposure and thus unwanted systemic side effects1. Delivery from the nose to the CNS occurs within minutes along both the olfactory and trigeminal neural pathways via an extracellular route and does not require drug to bind to any receptor or axonal transport2. Intranasal delivery is a widely publicized method and is currently being used in human clinical trials3.
Intranasal delivery of drugs in animal models allows for initial evaluation of pharmacokinetic distribution and efficacy. With mice, it is possible to administer drugs to awake (non-anesthetized) animals on a regular basis using a specialized intranasal grip. Awake delivery is beneficial because it allows for long-term chronic dosing without anesthesia, it takes less time than with anesthesia, and can be learned and done by many people so that teams of technicians can dose large numbers of mice in short periods. Efficacy of therapeutics administered intranasally in this way to mice has been demonstrated in a number of studies including insulin in diabetic mouse models 4-6 and deferoxamine in Alzheimer's mouse models. 7,8
The intranasal grip for mice can be learned, but is not easy and requires practice, skill, and a precise grip to effectively deliver drug to the brain and avoid drainage to the lung and stomach. Mice are restrained by hand using a modified scruff in the non-dominant hand with the neck held parallel to the floor, while drug is delivered with a pipettor using the dominant hand. It usually takes 3-4 weeks of acclimating to handling before mice can be held with this grip without a stress response. We have prepared this JoVE video to make this intranasal delivery technique more accessible.

A method to intranasally administer drugs to awake mice for the purpose of targeting the brain is described. This method allows for repeat dosing over long periods using intranasal administration of drug without anesthesia, and nose-to-brain delivery with minimal systemic exposure.

Intranasal administration is a method of delivering  therapeutic agents to the central nervous system (CNS). It is  non-invasive and allows large ...

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are necessary for whole animal experiments that require surgery under deep anesthesia or measurements of lung function. Tracheostomy has been the standard for intubating the airway in these mice to allow mechanical ventilation. Orotracheal intubation has been reported but has not been successfully used in many studies because of the substantial technical difficulty or a requirement for highly specialized and expensive equipment. Here we report a technique of direct laryngoscopy using an otoscope fitted with a 2.0 mm speculum and using a 20 G&#160;intravenous catheter as an endotracheal tube. We have used this technique extensively and reliably to intubate and conduct accurate assessments of lung function in mice. This technique has proven safe, with essentially no animal loss in experienced hands. Moreover, this technique can be used for repeated studies of mice in chronic models.

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

We have developed a simple, reliable, and relatively inexpensive method for endotracheal intubation in mice via direct laryngoscopy using an otoscope with a 2.0 mm speculum. This technique is atraumatic and can be used for repeated measurements in chronic experiments. We find it superior to tracheostomy or previously reported nonsurgical techniques.

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are ...

Flexible Colonoscopy in Mice to Evaluate the Severity of Colitis and Colorectal Tumors Using a Validated Endoscopic Scoring System

The use of modern endoscopy for research purposes has greatly facilitated our understanding of gastrointestinal pathologies. In particular, experimental endoscopy has been highly useful for studies that require repeated assessments in a single laboratory animal, such as those evaluating mechanisms of chronic inflammatory bowel disease and the progression of colorectal cancer. However, the methods used across studies are highly variable. At least three endoscopic scoring systems have been published for murine colitis and published protocols for the assessment of colorectal tumors fail to address the presence of concomitant colonic inflammation. This study develops and validates a reproducible endoscopic scoring system that integrates evaluation of both inflammation and tumors simultaneously. This novel scoring system has three major components: 1) assessment of the extent and severity of colorectal inflammation (based on perianal findings, transparency of the wall, mucosal bleeding, and focal lesions), 2) quantitative recording of tumor lesions (grid map and bar graph), and 3) numerical sorting of clinical cases by their pathological and research relevance based on decimal units with assigned categories of observed lesions and endoscopic complications (decimal identifiers). The video and manuscript presented herein were prepared, following IACUC-approved protocols, to allow investigators to score their own experimental mice using a well-validated and highly reproducible endoscopic methodology, with the system option to differentiate distal from proximal endoscopic colitis (D-PECS).

Over the past few years, new generation endoscopes have emerged as important diagnostic research aids for evaluating murine colitis and colorectal tumors. We present herein a detailed protocol for endoscopic assessment of inflammation and colorectal tumors in mice, as well as a novel scoring system that uses decimal identifiers to document the endoscopic severity of colitis and colorectal tumors.

The use of modern endoscopy for research purposes has greatly facilitated our understanding of gastrointestinal pathologies. In particular, experimental ...

Use of a Low-flow Digital Anesthesia System for Mice and Rats

A traditional vaporizer depends on flowing gas and atmospheric pressure for passive anesthetic vaporization. Newly developed direct injection vaporizers utilize a syringe pump to directly administer volatile anesthetics into a gas stream. Unlike a traditional vaporizer, it can be used at very low flow rates, making it ideal for use on mice and rats. 
The equipment's capability to use low flow rates could result in a substantial cost savings due to the reduced need for anesthetic agents, compressed gas, and charcoal scavenging filters1. A lower flow rate means less waste of anesthetic gas and likely reduces the risk of anesthetic exposure to laboratory personnel. Thus, the high levels of precision and safety associated with direct injection vaporizers, along with a reduced need for anesthetic agents, compressed gas, and charcoal filters are beneficial for research requiring small animal anesthesia. 
The goal of this protocol is to demonstrate the use of a syringe-driven direct injection vaporizer as part of a digital, low-flow anesthesia system. The direct injection vaporizer is capable of accurately delivering anesthesia at very low flow rates compared to a traditional vaporizer, making it a promising alternative for controlled gas anesthetic delivery to rodents.

Here, we present a protocol to more safely and efficiently administer anesthetic gas to mice using a digital, low flow anesthesia system utilizing a syringe-driven direct injection vaporizer.

A traditional vaporizer depends on flowing gas and atmospheric pressure for passive anesthetic vaporization. Newly developed direct injection vaporizers ...

Disposable Dosators for Pulmonary Insufflation of Therapeutic Agents to Small Animals

Development of new therapeutic products requires efficacy testing in an animal model. The pulmonary route of administration can be utilized to deliver drugs locally and systemically. Evaluation of dry powder aerosols necessitates an efficient dispersion mechanism to maintain high concentrations in an exposure chamber or for direct endotracheal administration. While solutions exist to expose animals by passive inhalation to dry powder aerosols, most require masses of powder in large excess of the dose delivered. This precludes conducting early feasibility studies as insufficient drug is available at the research or early development stage to support the dose delivery requirements for conventional aerosol delivery systems. When designing an aerosol drug product, aerodynamic performance can relate directly to delivery efficiency and efficacy. Dispersion of powder into an aerosol requires energy input sufficient to overcome interparticulate forces, and particle engineering approaches can substantially improve aerosol performance. We have developed a dispersion system (dosator) which can aerosolize engineered dry powder aerosols efficiently for the purpose of direct pulmonary insufflation, dispersion into an exposure system or generation for analytical purposes.

During development of drugs for pulmonary delivery, it is necessary to evaluate pharmacokinetics and efficacy in an animal model. We present a method to build a disposable aerosol dispersion system from of-the-shelf components that can be used to administer intrapulmonary dry powder aerosol to rodents.

Development of new therapeutic products requires efficacy testing in an animal model. The pulmonary route of administration can be utilized to deliver ...

Novel Methods for Intranasal Administration Under Inhalation Anesthesia to Evaluate Nose-to-Brain Drug Delivery

Intranasal administration has been reported to be a potential pathway for nose-to-brain delivery of therapeutic agents that circumvents the blood-brain barrier. However, there have been few reports regarding not only the quantitative analysis but also optimal administration conditions and dosing regimens for investigations of nose-to-brain delivery. The limited progress in research on nose-to-brain pathway mechanisms using rodents represents a significant impediment in terms of designing nose-to-brain delivery systems for candidate drugs.
To gain some headway in this regard, we developed and evaluated two novel methods of stable intranasal administration under inhalation anesthesia for experimental animals. We also describe a method for the evaluation of drug distribution levels in the brain via the nose-to-brain pathway using radio-labeled [14C]-inulin (molecular weight: 5,000) as a model substrate of water-soluble macromolecules.
Initially, we developed a pipette-based intranasal administration protocol using temporarily openable masks, which enabled us to perform reliable administration to animals under stable anesthesia. Using this system, [14C]-inulin could be delivered to the brain with little experimental error.
We subsequently developed an intranasal administration protocol entailing reverse cannulation from the airway side through the esophagus, which was developed to minimize the effects of mucociliary clearance (MC). This technique led to significantly higher levels of [14C]-inulin, which was quantitatively detected in the olfactory bulb, cerebrum, and medulla oblongata, than the pipette method. This appears to be because retention of the drug solution in the nasal cavity was substantially increased by active administration using a syringe pump in a direction opposite to the MC into the nasal cavity.
In conclusion, the two methods of intranasal administration developed in this study can be expected to be extremely useful techniques for evaluating pharmacokinetics in rodents. The reverse cannulation method, in particular, could be useful for evaluating the full potential of nose-to-brain delivery of drug candidates.

Here, we describe two novel methods of stable intranasal administration under inhalation anesthesia with minimal physical stress for experimental animals. We also describe a method for quantitative evaluation of drug distribution levels in the brain via the nose-to-brain pathway using radiolabeled [14C]-inulin as a model substrate of water-soluble macromolecules.

Intranasal administration has been reported to be a potential pathway for nose-to-brain delivery of therapeutic agents that circumvents the blood-brain ...

Intratracheal Administration of Dry Powder Formulation in Mice

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 &#181;L 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.

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.

In the development of inhalable dry powder formulations, it is essential to evaluate their biological activities in preclinical animal models. This paper ...

Customizable Disposable Dosators to Deliver Dry Powder Aerosol Drugs to Mice

Disposable Dosators Intended for Dry Powder Delivery to Mice

Dry powder inhalers offer numerous advantages for delivering drugs to the lungs, including stable solid-state drug formulations, device portability, bolus metering and dosing, and a propellant-free dispersal mechanism. To develop pharmaceutical dry powder aerosol products, robust in vivo testing is essential. Typically, initial studies involve using a murine model for preliminary evaluation before conducting formal studies in larger animal species. However, a significant limitation in this approach is the lack of suitable device technology to accurately and reproducibly deliver dry powders to small animals, hindering such models&#39; utility. To address these challenges, disposable syringe dosators were developed specifically for intrapulmonary delivery of dry powders in doses appropriate for mice. These dosators load and deliver a predetermined amount of powder obtained from a uniform bulk density powder bed. This discrete control is achieved by inserting a blunt needle to a fixed depth (tamping) into the powder bed, removing a fixed quantity each time. Notably, this dosing pattern has proven effective for a range of spray-dried powders. In experiments involving four different model spray-dried powders, the dosators demonstrated the ability to achieve doses within the range of 30 to 1100 &#181;g. The achieved dose was influenced by factors such as the number of tamps, the size of the dosator needle, and the specific formulation used. One of the key benefits of these dosators is their ease of manufacturing, making them accessible and cost-effective for delivering dry powders to mice during initial proof-of-concept studies. The disposable nature of the dosators facilitates use in animal procedure rooms, where cleaning and refilling reusable systems and weighing materials is inconvenient. Thus, developing disposable syringe dosators has addressed a significant hurdle in murine dry powder delivery for proof-of-concept studies, enabling researchers to conduct more accurate and reproducible preliminary studies in small animal models for pulmonary drug delivery.

Author Spotlight: Developing a Disposable Dosator for Preclinical Testing of Dry Powder Inhalers in Small Animal Models 

Pharmaceutical dry powder development necessitates reliable in vivo testing, often using a murine model. Device technology for accurately and reproducibly delivering dry powder aerosols to mice is restricted. This study presents disposable dosators for pulmonary drug delivery at mouse-relevant doses, aiding initial proof-of-concept research.

Dry powder inhalers offer numerous advantages for delivering drugs to the lungs, including stable solid-state drug formulations, device portability, bolus ...

A Technique to Generate and Deliver Drug Aerosols Directly to Mouse Lungs

Source: Phillips, J. E. et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/55454">Dry Powder and Nebulized Aerosol Inhalation of Pharmaceuticals Delivered to Mice Using a Nose-only Exposure System.</a>&#160;J. Vis. Exp. (2017)
This video demonstrates the generation of drug aerosols and the targeted nose-only delivery of these aerosols to mice using an inhalation unit equipped with an aerosol generator. The mouse is restrained in a nose-only restrainer and connected to the inhalation unit, which delivers the drug directly into the mouse&#39;s nasal cavity. Subsequently, the drug passes through the trachea and enters the lungs.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Dry Powder and Nebulized Aerosol Inhalation of Pharmaceuticals Delivered to Mice Using a Nose-only Exposure System

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Chapters in this video

Intraoperative Detection of Subtle Endometriosis: A Novel Paradigm for Detection and Treatment of Pelvic Pain Associated with the Loss of Peritoneal Integrity

Intranasal Administration of CNS Therapeutics to Awake Mice

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Flexible Colonoscopy in Mice to Evaluate the Severity of Colitis and Colorectal Tumors Using a Validated Endoscopic Scoring System

Use of a Low-flow Digital Anesthesia System for Mice and Rats

Disposable Dosators for Pulmonary Insufflation of Therapeutic Agents to Small Animals

Novel Methods for Intranasal Administration Under Inhalation Anesthesia to Evaluate Nose-to-Brain Drug Delivery

Intratracheal Administration of Dry Powder Formulation in Mice

Disposable Dosators Intended for Dry Powder Delivery to Mice

A Technique to Generate and Deliver Drug Aerosols Directly to Mouse Lungs