The authors would like to thank Mr. Ray Lee, Mr. HC Leung and Mr. Wallace So for their kind assistance in making the light source and powder insufflator; and the Faculty Core Facility for the assistance in animal imaging. The work was supported by the Research Grant Council, Hong Kong (17300319).

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 &#956;m. 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 &#956;m.
1. Fabrication of dry powder insufflator and loading of dry powder
<ol>
	<li>(Optional)&#160;Neutralize the static charges of dry powder (in a vial) and the 200 &#181;L non-filter round gel-loading pipette tip. Use an anti-static gun or a balance with deionizing function according to the manufacturer&#8217;s instruction.</li>
	<li>Prepare a weighing paper with a size of around 4 cm x 4 cm. Fold the paper in half diagonally and then unfold it.</li>
	<li>Weigh 1-2 mg of dry powder on the weighing paper.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
2. Fabrication of intubation device
<ol>
	<li>Light source (Figure 2)
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Affix the optical fiber in position with clear epoxy glue.</li>
	</ol>
	</li>
	<li>Guiding cannula (Figure 3)
	<ol>
		<li>Take a 1 mL plastic Pasteur pipette (Figure 3A) and hold the pipette at both ends.</li>
		<li>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.</li>
		<li>When the plastic becomes soft and deformable, move the pipette away from the flame and stretch the pipette gently.</li>
		<li>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 &#181;L 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.</li>
		<li>(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.</li>
	</ol>
	</li>
</ol>
3. Intubation
<ol>
	<li>Anaesthetize the mouse (BALB/c, 7-9 weeks) with ketamine (100 mg/kg) and xylazine (10 mg/kg) by intraperitoneal injection.</li>
	<li>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&#176; 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Gently protrude the tongue of the mouse with a pair of forceps to expose its trachea.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Upon intubation, swiftly remove the optical fiber and leave the guiding cannula inside the trachea (Figure 5B). A normal respiration should be observed.</li>
	<li>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.</li>
</ol>
4. Powder administration
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Turn the valve of the three-way stopcock to connect the airflow between the syringe and the powder-loaded tip.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Once the administration is complete, remove the guiding cannula from the trachea.</li>
	<li>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.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dry+powder+inhalers+in+COPD,+lung+inflammation+and+pulmonary+infections.">Dry powder inhalers in COPD, lung inflammation and pulmonary infections.</a> Expert Opinion on Drug Delivery. 12 (6), 947-962 (2015).</li><li>de Boer, A. H., 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=Dry+powder+inhalation:+past,+present+and+future.">Dry powder inhalation: past, present and future.</a> Expert Opinion on Drug Delivery. 14 (4), 499-512 (2017).</li><li>Das, S., Tucker, I., Stewart, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhaled+dry+powder+formulations+for+treating+tuberculosis.">Inhaled dry powder formulations for treating tuberculosis.</a> Current Drug Delivery. 12 (1), 26-39 (2015).</li><li>Okamoto, H., 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=Stability+of+chitosan-pDNA+complex+powder+prepared+by+supercritical+carbon+dioxide+process.">Stability of chitosan-pDNA complex powder prepared by supercritical carbon dioxide process.</a> International Journal of Pharmaceutics. 290 (1-2), 73-81 (2005).</li><li>He, 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=Evaluation+of+inhaled+recombinant+human+insulin+dry+powders:+pharmacokinetics,+pharmacodynamics+and+14-day+inhalation.">Evaluation of inhaled recombinant human insulin dry powders: pharmacokinetics, pharmacodynamics and 14-day inhalation.</a> Journal of Pharmacy and Pharmacology. 71 (2), 176-184 (2019).</li><li>Durham, P. 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The authors have no conflicts of interest to disclose.

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 &#181;L 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 ...

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 vaccination2,3,4. 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 nebulizers5,6. 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&#160;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 sedation8,9. 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 loss12,13. 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 effects14,15.
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.

<table>
	<tbody>
		<tr>
			<td>BALB/c mouse</td>
			<td></td>
			<td></td>
			<td>Female; 7-9 weeks old; Body weight 20-25 g</td>
		</tr>
		<tr>
			<td>CleanCap Firefly Luciferase mRNA</td>
			<td>TriLink Biotechnology</td>
			<td>L-7602</td>
			<td></td>
		</tr>
		<tr>
			<td>Dry Powder Insufflator</td>
			<td>PennCentury</td>
			<td>Model DP-4M</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine 10%</td>
			<td>Alfasan International B.V.</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Light emitting diode (LED) torch</td>
			<td>Unilite Internation</td>
			<td>PS-K1</td>
			<td></td>
		</tr>
		<tr>
			<td>Mannitol (Pearlitol 160C)</td>
			<td>Roquette</td>
			<td>450001</td>
			<td></td>
		</tr>
		<tr>
			<td>Non-filter round gel loading pipette tip (200 &#181;L)</td>
			<td>Labcon</td>
			<td>1034-800-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylon floss</td>
			<td>Reach</td>
			<td>30017050</td>
			<td></td>
		</tr>
		<tr>
			<td>One milliliter syringe without needle</td>
			<td>Terumo</td>
			<td>SS-01T</td>
			<td></td>
		</tr>
		<tr>
			<td>Optical fibre</td>
			<td>Fibre Data</td>
			<td>OMPF1000</td>
			<td></td>
		</tr>
		<tr>
			<td>PEG12KL4 peptide</td>
			<td>EZ Biolab</td>
			<td></td>
			<td>(PEG12)-KLLLLKLLLLKLLLLKLLLLK-NH2</td>
		</tr>
		<tr>
			<td>Plastic Pasteur fine tip pipette</td>
			<td>Alpha Labotatories</td>
			<td>LW4061</td>
			<td></td>
		</tr>
		<tr>
			<td>Three-way stopcock</td>
			<td>Braun</td>
			<td>D201</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine 2%</td>
			<td>Alfasan International B.V.</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Zerostat 3 anti-static gun</td>
			<td>MILTY</td>
			<td>5036694022153</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

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...

Watch this Scientific Journal Video about Intratracheal Administration of Dry Powder Formulation in Mice at JoVE.com

Intratracheal Administration of Dry Powder Formulation in Mice

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 ...

intratracheal-administration-of-dry-powder-formulation-in-mice

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9.7K Views.  The University of Hong Kong. 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.

Video: Intratracheal Administration of Dry Powder Formulation in Mice

A Novel Surgical Approach for Intratracheal Administration of Bioactive Agents in a Fetal Mouse Model

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and acquired diseases. Apart from congenital or inherited abnormalities with the requirement for long-term expression of the delivered gene, several non-inherited perinatal conditions, where short-term gene expression or pharmacological intervention is sufficient to achieve therapeutic effects, are considered as potential future indications for this kind of approach. Candidate diseases for the application of short-term prenatal therapy could be the transient neonatal deficiency of surfactant protein B causing neonatal respiratory distress syndrome1,2 or hyperoxic injuries of the neonatal lung3. Candidate diseases for permanent therapeutic correction are Cystic Fibrosis (CF)4, genetic variants of surfactant deficiencies5 and &alpha;1-antitrypsin deficiency6.
Generally, an important advantage of prenatal gene therapy is the ability to start therapeutic intervention early in development, at or even prior to clinical manifestations in the patient, thus preventing irreparable damage to the individual. In addition, fetal organs have an increased cell proliferation rate as compared to adult organs, which could allow a more efficient gene or stem cell transfer into the fetus. Furthermore, in utero gene delivery is performed when the individual's immune system is not completely mature. Therefore, transplantation of heterologous cells or supplementation of a non-functional or absent protein with a correct version should not cause immune sensitization to the cell, vector or transgene product, which has recently been proven to be the case with both cellular and genetic therapies7.
In the present study, we investigated the potential to directly target the fetal trachea in a mouse model. This procedure is in use in larger animal models such as rabbits and sheep8, and even in a clinical setting9, but has to date not been performed before in a mouse model. When studying the potential of fetal gene therapy for genetic diseases such as CF, the mouse model is very useful as a first proof-of-concept because of the wide availability of different transgenic mouse strains, the well documented embryogenesis and fetal development, less stringent ethical regulations, short gestation and the large litter size.
Different access routes have been described to target the fetal rodent lung, including intra-amniotic injection10-12, (ultrasound-guided) intrapulmonary injection13,14 and intravenous administration into the yolk sac vessels15,16 or umbilical vein17. Our novel surgical procedure enables researchers to inject the agent of choice directly into the fetal mouse trachea which allows for a more efficient delivery to the airways than existing techniques18.

We developed a novel surgical approach for intratracheal administration of bioactive agents into the mouse fetus. The delivery route is more efficient in targeting the fetal mouse lungs than the commonly used intra-amniotic injection. This procedure has to date not been described in a mouse model.

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and ...

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 ...

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 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 ...

A Pleural Effusion Model in Rats by Intratracheal Instillation of Polyacrylate/Nanosilica

Pleural effusion is a prevalent clinical finding of many pulmonary diseases. Having a useful animal pleural effusion model is very important to study these pulmonary diseases. Previous pleural effusion models paid more attention to biological factors rather than nanoparticles in the environment. Here, we introduce a model to make pleural effusion in rats by intratracheal instillation of polyacrylate/nanosilica, and a method of nanoparticle isolation in the pleural effusion. By intratracheal instillation of polyacrylate/nanosilica with concentrations of 3.125, 6.25 and 12.5 mg/kg&#8729;mL, the pleural effusion in rats presented on day 3, peaked at days 7-10 in 6.25 and 12.5 mg/kg&#8729;mL groups, then slowly decreased and disappeared on day 14. When the concentration of polyacrylate/nanosilica increased, the pleural effusion is produced more and faster. This pleural fluid was detected by ultrasound examination or CT chest scanning and confirmed by dissection of rats. Silica nanoparticles were observed in the rats' pleural effusion by transmission electron microscope. These results showed that the exposure to polyacrylate/nanosilica leads to the induction of pleural effusion, which was consistent with our previous report in humans. Additionally, this model is beneficial for the further study of nanotoxicology and the pleural effusion diseases.

Here, we present a protocol to construct a pleural effusion model in rats by intratracheal instillation of polyacrylate/nanosilica.

Pleural effusion is a prevalent clinical finding of many pulmonary diseases. Having a useful animal pleural effusion model is very important to study ...

Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. Various approaches have been described, such as the inhalation of aerosolized LPS as well as nasal or intratracheal instillation. The presented protocol describes a detailed step-by-step procedure to induce ALI in mice by direct intratracheal LPS instillation and perform FACS analysis of blood samples, bronchoalveolar lavage (BAL) fluid, and lung tissue. After intraperitoneal sedation, the trachea is exposed and LPS is administered via a 22 G venous catheter. A robust and reproducible inflammatory reaction with leukocyte invasion, upregulation of proinflammatory cytokines, and disruption of the alveolo-capillary barrier is induced within hours to days, depending on the LPS dosage used. Collection of blood samples, BAL fluid, and lung harvesting, as well as the processing for FACS analysis, are described in detail in the protocol. Although the use of the sterile LPS is not suitable to study pharmacologic interventions in infectious diseases, the described approach offers minimal invasiveness, simple handling, and good reproducibility to answer mechanistic immunological questions. Furthermore, dose titration as well as the use of alternative LPS preparations or mouse strains allow modulation of the clinical effects, which can exhibit different degrees of ALI severity or early vs. late onset of disease symptoms.

Presented here is a step-by-step procedure to induce acute lung injury in mice by direct intratracheal lipopolysaccharide instillation and to perform FACS analysis of blood samples, bronchoalveolar lavage fluid, and lung tissue. Minimal invasiveness, simple handling, good reproducibility, and titration of disease severity are advantages of this approach.

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. ...

Establishment of a Severe Dry Eye Model Using Complete Dacryoadenectomy in Rabbits

Dry eye disease (DED) is a complex disease with multiple etiologies and variable symptoms, having ocular surface inflammation as its key pathophysiologic step. Despite advances in our understanding of DED, significant knowledge gaps remain. Advances are limited in part due to the lack of informative animal models. The authors recently reported on a method of DED induced by injecting all orbital lacrimal gland (LG) tissues with the lectin concanavalin A. Here, we report a novel model of aqueous-deficient DED based on the surgical resection of all orbital LG (dacryoadenectomy) tissues. Both methods use rabbits because of their similarity to human eyes in terms of the size and structure of the ocular surface. One week after removal of the nictitating membrane, the orbital superior LG was surgically removed under anesthesia, followed by removal of the palpebral superior LG, and finally removal of the inferior LG. Dacryoadenectomy induced severe DED, evidenced by a marked reduction in the tear break up time test and the Schirmer&#39;s tear test, and significantly increased tear osmolarity and rose bengal staining. Dacryoadenectomy-induced DED lasted at least eight weeks. There were no complications and animals tolerated the procedure well. The technique can be mastered relatively easily by those with adequate surgical experience and appreciation of the relevant rabbit anatomy. Since this model recapitulates the features of human aqueous-deficient DED, it is suitable for studies of ocular surface homeostasis, DED, and candidate therapeutics.

A novel approach is presented to induce chronic dry eye disease in rabbits by surgically removing all orbital lacrimal glands. This method, distinct from those previously reported, produces a stable, reproducible model of aqueous deficient dry eye well suited to study tear physiology and pathophysiology and ocular therapeutics.

Dry eye disease (DED) is a complex disease with multiple etiologies and variable symptoms, having ocular surface inflammation as its key pathophysiologic ...

Intratracheal Instillation of Stem Cells in Term Neonatal Rats

Prolonged exposure to high concentrations of oxygen leads to inflammation and acute lung injury, which is similar to human bronchopulmonary dysplasia (BPD). In premature infants, BPD is a major complication despite early use of surfactant therapy, optimal ventilation strategies, and noninvasive positive pressure ventilation. Because pulmonary inflammation plays a crucial role in the pathogenesis of BPD, corticosteroid use is one potential treatment to prevent it. Nevertheless, systemic corticosteroid treatment is not usually recommended for preterm infants due to long-term adverse effects. Preclinical studies and human phase I clinical trials demonstrated that use of mesenchymal stromal cells (MSCs) in hyperoxia-induced lung injuries and in preterm infants is safe and feasible. Intratracheal and intravenous MSC transplantation has been shown to protect against neonatal hyperoxic lung injury. Therefore, intratracheal administration of stem cells and combined surfactant and glucocorticoid treatment has emerged as a new strategy to treat newborns with respiratory disorders. The developmental stage of rat lungs at birth is equivalent to that in human lungs at 26&#8722;28 week of gestation. Hence, newborn rats are appropriate for studying intratracheal administration to preterm infants with respiratory distress to evaluate its efficacy. This intratracheal instillation technique is a clinically viable option for delivery of stem cells and drugs into the lungs.

Described is a protocol for performing intratracheal transplantation of mesenchymal stromal cells (MSCs) through intratracheal injection in term neonatal rats. This technique is a clinically viable option for delivery of stem cells and drugs into neonatal rat lungs to evaluate their efficacy.

Prolonged exposure to high concentrations of oxygen leads to inflammation and acute lung injury, which is similar to human bronchopulmonary dysplasia ...

A Minimally Invasive Method for Intratracheal Instillation of Drugs in Neonatal Rodents to Treat Lung Disease

Treatment of neonatal rodent with drugs instilled directly into the trachea could serve as a valuable tool to study the impact of a locally administered drug. This has direct translational impact because surfactant and drugs are administered locally into the lungs. Though the literature has many publications describing minimally invasive transoral intubation of adult mice and rats in therapeutic experiments, this approach in neonatal rat pups is lacking. The small size of orotracheal region/pharynx in the pups makes visualization of laryngeal lumen (vocal cords) difficult, contributing to the variable success rate of intratracheal drug delivery. We hereby demonstrate effective oral intubation of neonatal rat pup - a technique that is non-traumatic and minimally-invasive, so that it can be used for serial administration of drugs. We used an operating otoscope with an illumination system and a magnifying lens to visualize the tracheal opening of the rat neonates. The drug is then instilled using a 1 mL syringe connected to a pipette tip. The accuracy of the delivery method was demonstrated using Evans blue dye administration. This method is easy to get trained in and could serve as an effective way to instill drugs into trachea. This method could also be used for administration of inoculum or agents to simulate disease conditions in animals and, also, for cell-based treatment strategies for various lung diseases.

This technique of instilling drugs directly into the trachea of neonatal rodents is important in studying the impact of locally administered drugs or biologicals on neonatal lung diseases. Additionally, this method can also be used for inducing lung injury in animal models.

Treatment of neonatal rodent with drugs instilled directly into the trachea could serve as a valuable tool to study the impact of a locally administered ...

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 ...