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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F., 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=A+novel+aerosol+generator+for+homogenous+distribution+of+powder+over+the+lungs+after+pulmonary+administration+to+small+laboratory+animals.">A novel aerosol generator for homogenous distribution of powder over the lungs after pulmonary administration to small laboratory animals.</a> European Journal of Pharmaceutics and Biopharmaceutics. 88 (3), 1056-1063 (2014).</li><li>Hoppentocht, M., Hoste, C., Hagedoorn, P., Frijlink, H. W., de Boer, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+evaluation+of+the+DP-4M+PennCentury+insufflator.">In vitro evaluation of the DP-4M PennCentury insufflator.</a> European Journal of Pharmaceutics and Biopharmaceutics. 88 (1), 153-159 (2014).</li><li>Liao, Q., 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=Porous+and+highly+dispersible+voriconazole+dry+powders+produced+by+spray+freeze+drying+for+pulmonary+delivery+with+efficient+lung+deposition.">Porous and highly dispersible voriconazole dry powders produced by spray freeze drying for pulmonary delivery with efficient lung deposition.</a> International Journal of Pharmaceutics. 560, 144-154 (2019).</li><li>Ito, T., Okuda, T., Takashima, Y., Okamoto, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Naked+pDNA+Inhalation+Powder+Composed+of+Hyaluronic+Acid+Exhibits+High+Gene+Expression+in+the+Lungs.">Naked pDNA Inhalation Powder Composed of Hyaluronic Acid Exhibits High Gene Expression in the Lungs.</a> Molecular Pharmaceutics. 16 (2), 489-497 (2019).</li><li>Chaurasiya, B., Zhou, M., Tu, J., Sun, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Design+and+validation+of+a+simple+device+for+insufflation+of+dry+powders+in+a+mice+model.">Design and validation of a simple device for insufflation of dry powders in a mice model.</a> European Journal of Pharmaceutical Sciences. 123, 495-501 (2018).</li></ol>

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 powder loading, particularly for powder with low density and in low relative humidity. The guiding cannula is a critical component of the device. It is used to facilitate the intubation of powder-loaded pipette tip into the trachea of the mouse. The diameter of the guiding cannula should not be too wide; otherwise it will be difficult to insert it into the trachea and may injure the mouse. The diameter of the guiding cannula should be just wide enough to fit the optical fiber and the powder-loaded pipette tip, and the pipette tip should protrude the guiding cannula by approximately 1-2 mm.
The ability to visualize the opening of the trachea is crucial in the intubation process, allowing the guiding cannula to be correctly inserted. The tracheal opening consists of white arytenoid cartilage with regular opening and closing motion at the back of the throat. With the fiber optic illumination, the opening of trachea could be easily visualized. By puffing a tiny volume of air through the fine tip plastic pipette, an inflation at the chest indicates a proper intubation. If inflation at the chest is not observed or resistance is felt during insertion, retract the guiding cannula swiftly and repeat the steps again.
There was a widely used commercially available dry powder insufflator12,17,18 (Table of Materials; this device is now discontinued). The dry powder is loaded into the sample chamber of the device and dispersed by air from a 3 mL plastic air syringe or an air pump. To measure the emitted dose, the device has to be weighed before and after powder administration, which leads to inaccuracy considering the dose of powder is usually very small (relative to the mass of the device). Compared to the commercial insufflator, the biggest advantage of the custom-made device is that the success of powder dispersion could be observed by the absence of powder in the transparent gel-loading pipette tips. Since the pipette tip is light, it can also be weighed accurately before and after administration to measure the emitted dose. The pipette tip is inserted into the guiding cannula rather than being exposed to the trachea of the animal. There is minimal risk of contaminating the tip with the moisture or secretion in the trachea (which may affect the accuracy of emitted dose measurement). As the 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 the need of device cleaning and dose refilling, thereby saving time and eliminating the risk of cross-contamination from residual powder. Moreover, the powder dispersion pattern generated by the commercial insufflator varied among different formulations. A number of studies reported that dry powder dispersed by the commercial insufflator were easily agglomerated and failed to reach the deep lung upon administration19,20. In contrast, other formulations dispersed by devices similar to ours are reported to have a high lung deposition15,21,22.
There are other similar custom-made devices reported in the literature for the administration of powder aerosol to the lung of animal. For instance, Chaurasiya et al. described the use of a cannula tube for intubation as well as powder loading, with a syringe connected to the cannula tube after intubation for powder dispersion23. While their approach uses standardized equipment and material (e.g., otoscope, cannula and syringe) with less customization, the method here offers some distinct advantages. Firstly, it allows confirmation of correct intubation before drug administration. This step is particularly helpful for less experienced user. Secondly, the guiding cannula can act as a protecting shield to prevent any secretion or moisture in the trachea from contaminating the gel-loading pipette tip, allowing a more accurate emitted dose measurement by weighing. Lastly, the more flexible guiding cannula together with the optical fiber may enable easier intubation.
In summary, a custom-made dry powder insufflator which is inexpensive, disposable, reproducible and efficient in dispersing small amount of powder precisely is introduced in this paper. The intubation process mentioned is noninvasive, quick and could deliver powder formulations to the mice safely and accurately. It can also be adopted to administer liquid formulation for pulmonary delivery.

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 up to 48 h post-administration. Dispersing the powder with 1 mL of air resulted in over 5% of weight loss within 24 h, which was not fully recovered after 48 h. In this protocol, BALB/c mice of 7-9 weeks old were used. Depending on the species, the strain and age of animal, the powder properties (e.g., particle size distribution, cohesiveness and density) and the mass of powder to be administered, the volume of air to be used for efficient powder dispersion and animal tolerance may require optimization by investigators for different animal models.
Dry powder formulation prepared by spray freeze drying (SFD) was delivered to the mice using the method described above. The SFD formulation contained 0.5% of mRNA expressing luciferase protein, 5% of synthetic peptide as delivery vector and 94.5% of mannitol16. BALB/c mice were intratracheally administered with 1 mg of SFD powder containing 5 &#181;g of mRNA and the luciferase expression in the lungs was evaluated at 24 h post-administration using in vivo imaging system (IVIS) (Figure 7). The SFD powder were dispersed in the deep lung and luciferase expression was observed. As a comparison, the SFD powder were reconstituted in PBS (to a final volume of 75 &#181;L) and administered to mice as liquid with the same intubation procedure but a microsprayer was used instead to generate liquid aerosol16. The luciferase expression of the reconstituted formulation was significantly higher than the dry powder formulation, which could be due to the powder dissolution issue or different pharmacokinetic profile between powder and liquid form. The histological characteristics of the lungs treated with mRNA dry powder aerosol were compared with untreated control and lipopolysaccharide (LPS) treated groups (Figure 8). The lungs without any treatment illustrated a healthy presentation while the lung treated with 10 &#181;g of LPS intratracheally showed irregular distribution of air space and inflammatory cell infiltration into the interstitial and alveolar spaces. The lungs treated with SFD powder did not show any signs of inflammation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61469/61469fig01.jpg" /> 
Figure 1: Custom-made dry powder insufflator. 
(A) Powder is packed near the narrow end of the tip. (B) A gel-loading pipette tip is connected to a 1 mL syringe via a three-way stopcock. The figure is adapted from Liao et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61469/61469fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61469/61469fig02.jpg" /> 
Figure 2: Custom-made light source for intubation. 
A flexible optical fiber is connected to a LED torch by creating a small hole on the lens. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61469/61469fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61469/61469fig03.jpg" /> 
Figure 3: Guiding cannula. 
(A) A 1 mL plastic Pasteur pipette is used to make a guiding cannula. (B) The pipette is softened by heating. (C) The heated pipette is stretched and cut. (D) Part A of the pipette is used as fine-tip pipette. (E&#38;F) Part B of the pipette is used as a guiding cannula. A bevel is created to facilitate intubation procedure. A small opening (optional) can be made to increase the flexibility of the cannula. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61469/61469fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61469/61469fig04.jpg" /> 
Figure 4: Intubation platform. 
(A) The platform for intubation consists of a Plexiglass plate which is mounted to a stand. (B) An anaesthetized mouse is placed on the platform in a supine position, suspended by hooking its incisors with a nylon floss. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61469/61469fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61469/61469fig05.jpg" /> 
Figure 5: Schematic diagram illustrating the intubation procedure. 
(A) The bevel of the guiding cannula is aligned with the midline of the tracheal opening. (B) The guiding cannula is inserted into the trachea and ready for powder administration. (C) The powder-loaded tip (connected to the syringe through a three-way stopcock) is inserted into the guiding cannula which has already been placed in the trachea of the mouse. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61469/61469fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61469/61469fig06.jpg" /> 
Figure 6: Intratracheal administration of dry powder with different volume of air. 
BALB/c mice were administered intratracheally with spray dried (SD) mannitol powder dispersed by 0.3 mL, 0.6 mL and 1.0 mL of air. Body weight of the mice was monitored before administration and at 18 h, 24 h and 48 h post-administration. The data was presented as mean value of percentage of weight change (n=2). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61469/61469fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61469/61469fig07.jpg" /> 
Figure 7: Intratracheal administration of mRNA formulation as dry powder and reconstituted liquid aerosol. 
BALB/c mice were administered intratracheally with spray freeze dried (SFD) 0.5% mRNA (luciferase) formulation as powder aerosol (1 mg) using custom-made dry powder insufflator or reconstituted liquid aerosol (1 mg in 75 &#956;L PBS) using microsprayer. Each mouse received a dose of 5 &#956;g of mRNA. PBS (75 &#181;L) was used as control. At 24 h post-administration (A) the lungs were isolated for bioluminescence imaging; (B) luciferase protein expression of the lung tissues were measured. The data was expressed as the mean value of relative light unit (RLU) per mg of protein, analyzed by one-way ANOVA followed by Tukey&#8217;s post-hoc test, ***p &#60; 0.001 (n=4). The figure is adapted from Qiu et al.16. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61469/61469fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/61469/61469fig08.jpg" /> 
Figure 8: Histology of the lungs of mice following intratracheal administration of mRNA dry powder formulation. 
(A) untreated control; mice were intratracheally administered with (B) LPS (10 mg in 25 &#956;L PBS), and (C) spray freeze dried mRNA powder (1 mg). Slides were viewed using an upright microscope at 20x magnification (scale bar = 100 mm). The figure is adapted from Qiu et al.16. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61469/61469fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Medicine

9.8K 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

Improving IV Insulin Administration in a Community Hospital

Diabetes mellitus is a major independent risk factor for increased morbidity and mortality in the hospitalized patient, and elevated blood glucose concentrations, even in non-diabetic patients, predicts poor outcomes.1-4 The 2008 consensus statement by the American Association of Clinical Endocrinologists (AACE) and the American Diabetes Association (ADA) states that "hyperglycemia in hospitalized patients, irrespective of its cause, is unequivocally associated with adverse outcomes."5 It is important to recognize that hyperglycemia occurs in patients with known or undiagnosed diabetes as well as during acute illness in those with previously normal glucose tolerance.
The Normoglycemia in Intensive Care Evaluation-Survival Using Glucose Algorithm Regulation (NICE-SUGAR) study involved over six thousand adult intensive care unit (ICU) patients who were randomized to intensive glucose control or conventional glucose control.6 Surprisingly, this trial found that intensive glucose control increased the risk of mortality by 14% (odds ratio, 1.14; p=0.02). In addition, there was an increased prevalence of severe hypoglycemia in the intensive control group compared with the conventional control group (6.8% vs. 0.5%, respectively; p&lt;0.001). From this pivotal trial and two others,7,8 Wyoming Medical Center (WMC) realized the importance of controlling hyperglycemia in the hospitalized patient while avoiding the negative impact of resultant hypoglycemia. 
Despite multiple revisions of an IV insulin paper protocol, analysis of data from usage of the paper protocol at WMC shows that in terms of achieving normoglycemia while minimizing hypoglycemia, results were suboptimal. Therefore, through a systematical implementation plan, monitoring of patient blood glucose levels was switched from using a paper IV insulin protocol to a computerized glucose management system. By comparing blood glucose levels using the paper protocol to that of the computerized system, it was determined, that overall, the computerized glucose management system resulted in more rapid and tighter glucose control than the traditional paper protocol. Specifically, a substantial increase in the time spent within the target blood glucose concentration range, as well as a decrease in the prevalence of severe hypoglycemia (BG &lt; 40 mg/dL), clinical hypoglycemia (BG &lt; 70 mg/dL), and hyperglycemia (BG &gt; 180 mg/dL), was witnessed in the first five months after implementation of the computerized glucose management system. The computerized system achieved target concentrations in greater than 75% of all readings while minimizing the risk of hypoglycemia. The prevalence of hypoglycemia (BG &lt; 70 mg/dL) with the use of the computer glucose management system was well under 1%.

When compared to the previous paper protocol, implementation of a computerized glucose management system results in a substantial increase in blood glucose concentration measurements within the target range. Using a computerized glucose management system to monitor blood glucose levels, decreases in severe hypoglycemia (&lt;40 mg/dL), clinical hypoglycemia (&lt;70 mg/dL), and hyperglycemia (&gt;180 mg/dL) also can be observed.

Diabetes mellitus is a major independent risk factor for increased morbidity and mortality in the hospitalized patient, and elevated blood glucose ...

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 ...

Carotid Artery Infusions for Pharmacokinetic and Pharmacodynamic Analysis of Taxanes in Mice

When proposing the use of a drug, drug combination, or drug delivery into a novel system, one must assess the pharmacokinetics of the drug in the study model. As the use of mouse models are often a vital step in preclinical drug discovery and drug development1-8, it is necessary to design a system to introduce drugs into mice in a uniform, reproducible manner. Ideally, the system should permit the collection of blood samples at regular intervals over a set time course. The ability to measure drug concentrations by mass-spectrometry, has allowed investigators to follow the changes in plasma drug levels over time in individual mice1, 9, 10. In this study, paclitaxel was introduced into transgenic mice as a continuous arterial infusion over three hours, while blood samples were simultaneously taken by retro-orbital bleeds at set time points. Carotid artery infusions are a potential alternative to jugular vein infusions, when factors such as mammary tumors or other obstructions make jugular infusions impractical. Using this technique, paclitaxel concentrations in plasma and tissue achieved similar levels as compared to jugular infusion. In this tutorial, we will demonstrate how to successfully catheterize the carotid artery by preparing an optimized catheter for the individual mouse model, then show how to insert and secure the catheter into the mouse carotid artery, thread the end of the catheter out through the back of the mouse&#8217;s neck, and hook the mouse to a pump to deliver a controlled rate of drug influx. Multiple low volume retro-orbital bleeds allow for analysis of plasma drug concentrations over time.

This method was developed with the goal of delivering a steady drug solution via the carotid artery, to assess the pharmacokinetics of novel drugs in mouse models.

When proposing the use of a drug, drug combination, or drug delivery into a novel system, one must assess the pharmacokinetics of the drug in the study ...

Intubation-mediated Intratracheal (IMIT) Instillation: A Noninvasive, Lung-specific Delivery System

Respiratory disease studies typically involve the use of murine models as surrogate systems. However, there are significant physiologic differences between the murine and human respiratory systems, especially in their upper respiratory tracts (URT). In some models, these differences in the murine nasal cavity can have a significant impact on disease progression and presentation in the lower respiratory tract (LRT) when using intranasal instillation techniques, potentially limiting the usefulness of the mouse model to study these diseases. For these reasons, it would be advantageous to develop a technique to instill bacteria directly into the mouse lungs in order to study LRT disease in the absence of involvement of the URT. We have termed this lung specific delivery technique intubation-mediated intratracheal (IMIT) instillation. This noninvasive technique minimizes the potential for instillation into the bloodstream, which can occur during more invasive traditional surgical intratracheal infection approaches, and limits the possibility of incidental digestive tract delivery. IMIT is a two-step process in which mice are first intubated, with an intermediate step to ensure correct catheter placement into the trachea, followed by insertion of a blunt needle into the catheter to mediate direct delivery of bacteria into the lung. This approach facilitates a &#62;98% efficacy of delivery into the lungs with excellent distribution of reagent throughout the lung. Thus, IMIT represents a novel approach to study LRT disease and therapeutic delivery directly into the lung, improving upon the ability to use mice as surrogates to study human respiratory disease. Furthermore, the accuracy and reproducibility of this delivery system also makes it amenable to Good Laboratory Practice Standards (GLPS), as well as delivery of a wide range of reagents which require high efficiency delivery to the lung.

Intubation-mediated Intratracheal IMIT Instillation: A Noninvasive, Lung-specific Delivery System

Intubation-mediated intratracheal (IMIT) instillation of reagents is an excellent, noninvasive method for studying respiratory disease, as well as a method for instilling therapeutic reagents directly into the lung. It is a rapid and highly reproducible method which is suitable for preclinical testing.

Respiratory disease studies typically involve the use of murine models as surrogate systems. However, there are significant physiologic differences ...

Transarterial Administration of Oncolytic Viruses for Locoregional Therapy of Orthotopic HCC in Rats

Hepatocellular carcinoma (HCC) is a disease with limited treatment options and poor prognosis. In recent years, oncolytic virotherapies have proven themselves to be potentially powerful tools to fight malignancy. Due to the unique dual blood supply in the liver, it is possible to apply therapies locally to orthotopic liver tumors, which are predominantly fed by arterial blood flow. We have previously demonstrated that hepatic arterial delivery of oncolytic viruses results in safe and efficient transduction efficiency of multifocal HCC lesions, resulting in significant prolongation of survival in immune competent rats. This procedure closely mimics the application of transarterial embolization in patients, which is the standard palliative care provided to many HCC patients. The ability to administer tumor therapies through the hepatic artery in rats allows for a highly sophisticated preclinical model for evaluating novel viral vectors under development. Here we describe the detailed protocol for microdissection of the hepatic artery for infusion of oncolytic virus vectors to treat orthotopic HCC.

Oncolytic virotherapies are under development as novel therapeutics for the treatment of hepatocellular carcinoma (HCC). Here we describe a method for locoregional therapy of HCC via hepatic arterial administration of oncolytic virus.

Hepatocellular carcinoma (HCC) is a disease with limited treatment options and poor prognosis.  In recent years, oncolytic virotherapies have proven ...

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 ...