This work was funded by Swiss National Foundation (SNF) project grant 31003A_135649, 31003A_153457 and 31003A_172994 (to JVS), and Leopoldina fellowship LPDS 2011-16 (to BS). This work benefitted from optical setups of the "Microscopy Imaging Center" (MIC) of the University of Bern.

All animal work has been approved by the Cantonal Committee for Animal Experimentation and conducted according to federal guidelines.
1. Anesthetize the Mouse
<ol>
	<li>Wear personal protective equipment, including lab coat and gloves.</li>
	<li>Mix ketamine, xylazine, and saline to a working concentration of 20 mg/mL and 1 mg/mL, respectively. Inject the working stock intraperitoneally (i.p.) at 8-10 &#181;L/g of mouse. Place the mouse back into the cage. 
	NOTE: This protocol has been tested for 6- to 40-week-old male and female mice with a C57BL/6 background.</li>
	<li>Prepare the acepromazine solution at 2.5 mg/mL and inject 30 &#181;L i.p., 15 min after the ketamine-xylazine injection (75 &#181;g/mouse).</li>
	<li>After 5-15 min, check for adequate anesthesia and analgesia by controlling for reflexes, e.g., withdrawal reflex of the paw or corneal reflex.</li>
	<li>Control for reflexes approximately every 30 min during the entire duration of the experiment. Subcutaneously inject ketamine/xylazine at 1.5 &#181;L/g of mouse, every 60 min or earlier if reflexes are positive. 
	NOTE: Inhalation anesthesia such as isoflurane cannot be used with this protocol, owing to the fixation of the mouse.</li>
</ol>
2. Remove the Fur from the Operating Area
<ol>
	<li>Use an electric razor to shave an area that expands roughly from 5 mm caudal of the mouth towards the front legs, stretching over the jaws towards both ears.</li>
	<li>Apply hair removal cream to the shaved area with a cotton swab. Carefully and thoroughly remove after 3 min, e.g., by wiping with wet tissues.</li>
</ol>
3. Fix the Mouse on the Stage
NOTE: This protocol uses a customized stage (Figure 1) equipped with two freely adjustable holders for cover slips (one attached to the stage with a screw, the other fixed permanently onto it); adjustable stereotactic ear bars; a string, which can be tightened with a screw; a freely adjustable metal heating ring; and a raised area designated to place the torso of the mouse. This stage is modified from a popliteal lymph node stage, with an added stereotactic holder and a support for the lower cover slip holding the exteriorized SMG. Detailed plans or quotes of the stage are available upon request.
<ol>
	<li>Prepare the stage by removing the holders for the heating ring and the removable cover slip holder. Loosen the screws of the ear bar holders and the fixed cover slip holder. Tape a piece of gauze to the center area of the stage, serving as bedding for the mouse.</li>
	<li>Use glue (see Table of Materials) to attach a 20 mm diameter cover glass to the top of a cylindrical metal holder (this will be the lower cover glass). Glue a 25 mm cover glass to the bottom of the flat metal holder (this will be the upper cover glass). Ensure that the cover glasses overlap approximately 2-3 mm with metal holders and that the rest of the glass remains free of grease and glue.</li>
	<li>Place the mouse on its back onto the gauze, perpendicular to the stretched string, with the head centered between the ear bars.</li>
	<li>Fix the upper extremities of the mouse onto the stage with surgical tape.</li>
	<li>Next, hook the string into the upper front incisors and tighten until the jaw reaches a horizontal position. Use an angled curved forceps to slightly pull out the tongue from the mouth, which facilitates breathing.</li>
	<li>Tightly, but not forcefully, pull the tail and tape it to the stage.</li>
	<li>Align the ear bars to achieve an angle of approximately 30&#176; between the bar and the edge of the stage. To ensure this angle, use a triangle ruler to draw a right and left 30&#176; angle on a sheet of paper, and place the transparent stage on the paper to align the holders accordingly.</li>
	<li>Fix the jaw between the ear bars by simultaneously moving the left and right bar inwards until the jaw is immotile, and tighten screws. Observe chest movement before, during, and after fixing the jaw. Reduced or absent breathing indicates that the bars are fixed at the neck, not the jaw bone; in this case, repeat the fixing procedure immediately.</li>
	<li>Provide heating to the mouse, e.g., by placing a heating lamp nearby and/or a heating pad underneath the stage.</li>
</ol>
4. Surgical Exposure of the Salivary Gland Using a Stereomicroscope
NOTE: The following steps require the use of a stereomicroscope, fine forceps, and surgical scissors. Since the procedure is terminal, it is not necessary to maintain sterile operating conditions. Ethanol-cleaned instruments suffice.
<ol>
	<li>Locate the approximate position of the right SMG lobe by looking for a slight bulge in the skin approximately 15 mm caudal of the mouth and 5 mm medial of the jaw line. Lift a small skin fold in the center of the bulge and cut it to create and opening of approximately 5 mm length.</li>
	<li>Immediately apply a 4 mm-high ring of vacuum grease on the skin surrounding the cut. (Use a vacuum grease-filled syringe with a blunted 25 G hypodermic needle).</li>
	<li>Fill the grease ring with saline to ensure the tissue remains moist during the operation.</li>
	<li>Under the stereomicroscope, disrupt the connective tissue using the following procedure: Use one fine forceps to firmly hold a piece of connective tissue directly attached to the SMG, and use a second forceps to pull the distal connective tissue until it ruptures. Ensure that some connective tissue remains attached to the SMG; this is desirable since it allows handling. Do not touch the gland itself with the forceps, since this will cause bleeding.</li>
	<li>Repeat this motion of disrupting connective tissue first on top of the gland, then on the medial side (between the right and left lobe of the SMG). Do not separate the sublingual from the submandibular gland.</li>
	<li>Continue to remove the lateral connective tissue counterclockwise around the SMG (medial to caudal, to distal, to rostral).</li>
	<li>Ensure that the blood supply, efferent lymphatics, and salivary duct, which are all located in a bundle at the cranial site of the gland, are not ruptured.</li>
	<li>Once the lateral connective tissue is disrupted, use forceps to pull the connective tissue attached to the caudal end of the SMG upwards to lift the caudal end of the SMG. Disrupt the connective tissue below the SMG lobe, starting from the caudal and moving towards the rostral end.</li>
	<li>Continue until the connective tissue to all sites of the SMG is removed (excluding the rostral site with major feeding/draining blood vessels and the WD).</li>
</ol>
5. Fixing the Gland
<ol>
	<li>Confirm that the grease ring holding the saline is at least 4 mm high and not leaking (see step 4.2).</li>
	<li>Install the lower cover glass (see step 3.2) by attaching the metal bar to its height-adjustable holder. Ensure that the cover slip holder is adjusted to the upmost position and does not touch the grease ring when attached to the stage. Position the holder from the caudal direction at an angle of approximately 60&#176; to the mouse, with the cover slip covering roughly two thirds (caudal and distal parts) of the exposed lobe. Lower the cover glass until it makes contact with the gland.</li>
	<li>Complete the grease ring on top of the cover glass to form a new saline reservoir.</li>
	<li>Carefully pull the SMG lobe on top of the cover slip with forceps, only touching the connective tissue attached to it (the organ can be flipped with the bottom site facing upwards).</li>
	<li>Ensure that the grease ring has an even height of at least 4 mm and does not leak. If necessary, refill the saline reservoir using a 30 G syringe filled with saline.</li>
	<li>Ensure that the cover glass holder for the upper cover glass is in its upmost position. Use the screw to attach the metal holder with the 25 mm upper cover glass to the holder. Use the adjustable screws of the holder to position the cover glass so that the SMG is centered in the middle.</li>
	<li>Use the screws of adjustable holder to lower the top cover glass until it directly contacts the SMG. Observe the shape and coloring of the gland during this step. If the gland surface area appears larger, or the coloring changes (for instance from pink to white), immediately put the cover glass to a higher position to relieve pressure on the SMG.</li>
	<li>Check the saline reservoir for leaks by looking for air bubbles and or drops of saline outside the reservoir. If a leak or air bubble is detected, remove both cover slips and all grease and restart from step 2.</li>
	<li>Screw shut all fixation screws.</li>
</ol>
6. Fixing the Heating Ring and Temperature Probe
<ol>
	<li>Insert a temperature probe into the reservoir and position it close to the gland, then tape the probe wire to the stage.</li>
	<li>Apply a grease ring to the top cover slip, with the SMG in the center (the diameter of the grease ring should correspond to diameter of the heating ring). Position the heating ring on top of the cover slip and seal with grease. Tape the tubing of the heating ring to the stage.</li>
</ol>
7. Imaging
NOTE: We use an upright two-photon microscope system equipped with a tunable Ti:Sa Laser and a fluorescent microscope with water-immersion 20X or 25X objectives. Real time correction of tissue drift during acquisition can be implemented by using an automated stage and VivoFollow19. This improves the stability of image acquisition, but is not required.
<ol>
	<li>Turn on the upright microscopy system using an appropriate software provided by the microscope manufacturer. Tune the Ti:Sa Laser to a 780-900 nm excitation wavelength.</li>
	<li>Connect the heating tubing to the heating ring. A peristaltic water pump circulates hot water (heated by submerging part of the tubing in an 80&#176;C water bath) through the ring to warm the SMG.</li>
	<li>Connect the temperature probe to a digital thermometer. Adjust the speed of the peristaltic pump if the temperature is not between 35-39 &#176;C. The faster the pump works, the warmer the object gets, and vice versa.</li>
	<li>Before starting image acquisition, perform a visual control of the blood flow velocities. Carefully check leukocyte passage through the thin capillaries or by fluorescent dextran injection. 
	NOTE: Intravenously injected fluorescent dextran will immediately fill out the entire SMG vasculature when blood flow is physiological. This is virtually always the case in this procedure.</li>
	<li>To follow immune cells, imaging depth is typically between 20-150 &#181;m below the surface of the SMG identified by the 2nd harmonic generation signal. Record image stacks of 150-300 &#181;m field of view with pixel resolution of approximately 512 x 512 or higher, and z-depths of 20-60 &#181;m with spacing of 2-4 &#181;m between images. Repeat stack acquisition every 20-30 s for a total duration of 20-60 min.</li>
	<li>Use the digital thermometer to control the minimum and maximum temperature levels after acquisition of one image sequence. Discard the image sequence if the minimum or maximum are not in the range of 37 &#177; 2 &#176;C. Refill the water for immersion of objective between image sequences.</li>
	<li>At the end of the imaging session, euthanize the mice by CO2 asphyxiation under anesthesia. 
	NOTE: This is in accordance to Swiss federal guidelines of animal experimentation; other guidelines for euthanasia may apply elsewhere.</li>
</ol>

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L., Shakhar, G., Dudziak, D., 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=Visualizing+dendritic+cell+networks+in+vivo.">Visualizing dendritic cell networks in vivo.</a> Nat Immunol. 5 (12), 1243-1250 (2004).</li><li>Riedl, J., Flynn, K. C., Raducanu, A., 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=Lifeact+mice+for+studying+F-actin+dynamics.">Lifeact mice for studying F-actin dynamics.</a> Nat Methods. 7 (3), 168-169 (2010).</li><li>Chtanova, T., Hampton, H. R., Waterhouse, L. A., 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=Real-time+interactive+two-photon+photoconversion+of+recirculating+lymphocytes+for+discontinuous+cell+tracking+in+live+adult+mice.">Real-time interactive two-photon photoconversion of recirculating lymphocytes for discontinuous cell tracking in live adult mice.</a> J Biophotonics. 7 (6), 425-433 (2014).</li><li>Kyratsous, N. I., Bauer, I. J., Zhang, G., 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=Visualizing+context-dependent+calcium+signaling+in+encephalitogenic+T+cells+in+vivo+by+two-photon+microscopy.">Visualizing context-dependent calcium signaling in encephalitogenic T cells in vivo by two-photon microscopy.</a> Proc Natl Acad Sci U S A. 114 (31), E6381-E6389 (2017).</li><li>Mank, M., Reiff, D. F., Heim, N., Friedrich, M. W., Borst, A., Griesbeck, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+FRET-based+calcium+biosensor+with+fast+signal+kinetics+and+high+fluorescence+change.">A FRET-based calcium biosensor with fast signal kinetics and high fluorescence change.</a> Biophys J. 90 (5), 1790-1796 (2006).</li><li>Tsyboulski, D., Orlova, N., Saggau, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amplitude+modulation+of+femtosecond+laser+pulses+in+the+megahertz+range+for+frequency-multiplexed+two-photon+imaging.">Amplitude modulation of femtosecond laser pulses in the megahertz range for frequency-multiplexed two-photon imaging.</a> Opt Express. 25 (8), 9435 (2017).</li><li>Potma, E. O., Xie, X. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+single+lipid+bilayers+with+coherent+anti-Stokes+Raman+scattering+(CARS)+microscopy.">Detection of single lipid bilayers with coherent anti-Stokes Raman scattering (CARS) microscopy.</a> J Raman Spectrosc. 34 (9), 642-650 (2003).</li></ol>

No conflicts of interest declared.

This protocol offers a straightforward approach for in vivo imaging of murine submandibular and sublingual salivary glands using upright non-linear microscopy often used in the field of immunology. The method can be adapted for the imaging of other exocrine glands in the head and neck region. For instance, our lab has performed imaging of the lacrimal gland in an analogous manner (not shown).
Removing the connective tissue around the SMG is the most critical step of this protocol, sin...

Saliva is secreted by exocrine glands to lubricate food, protect the mucosal surfaces of the oral tract, and to deliver digestive enzymes as well as antimicrobial substances1,2. In addition to minor salivary glands interspersed in the oral submucosa, there are three bilateral sets of major glands identified as parotid, sublingual, and submandibular, according to their location1,2. Pyramid-shaped epithelial cells, organized into flask-shaped sacs (acini) or demilunes that are surrounded by myoepithelial cells and a basement membrane, secrete the serous and mucous components of saliva1. The narrow luminal space of the acini drains into intercalated ducts, which unite into striated ducts until they finally join into a single excretory duct1. The main excretory duct of the SMG is called Wharton&#39;s duct (WD) and opens into the sublingual caruncle3,4. The SMG epithelial compartment therefore represents a highly arborized structure with manifold terminal endpoints, resembling a bundle of grapes1,5,6. The SMG interstitium is composed of blood and lymphatic vessels embedded in connective tissue7 containing parasympathetic nerves8 and extracellular matrix5. Normal human and rodent salivary glands also contain T cells, macrophages, and dendritic cells9, as well as plasma cells that secrete Immunoglobulin A (IgA) into the saliva9,10. Due to its multifaceted functions in health and disease, the SMG is a subject of interest for many fields of biological research, including dentistry4, immunology11, oncology12, physiology8, and cell biology3.
Imaging of dynamic cellular processes and interactions is a powerful tool in biological research13,14. The development of deep tissue imaging and innovations inmicroscopes based on nonlinear optics (NLO), which rely on scattering or absorption of multiple photons by the sample, has allowed to directly examine cellular processes in complex tissues13,15. Absorption of multiple photons involves delivery of the total excitation energy by low energy photons, which confines fluorophore excitation to the focal plane and thus allows deeper tissue penetration with reduced photodamage and noise from out of focus excitation13,15. This principle is employed by two-photon microscopy (2PM) and allows for imaging of fluorescent specimens in depths of up to 1 mm15,16. While commercially available 2PM setups have become user-friendly and reliable, the major challenge for intravital imaging is to carefully expose and stabilize the target organ of anesthetized mice, especially for imaging of time lapse series. Several methods for digital drift correction after data acquisition have been published17,18 and we recently developed &#34;VivoFollow&#34;, an automated correction system, which counteracts slow tissue drift in real time using a computerized stage19. However, it is still critical for high quality imaging to minimize tissue motion, especially fast movements caused by breathing or heartbeat19. Preparation and stabilization procedures have been published for multiple organs, including spinal cord20, liver21, skin22, lung23, and lymph node24. Furthermore, models for rat salivary gland imaging have been developed3,25 and further refined for high resolution intravital imaging of the murine SMG tailored to an inverted microscope setup26,27,28.
Here, we present a practical and adaptable protocol for intravital imaging of the murine SMG using upright nonlinear microscopy, which is commonly used for intravital imaging in the field of immunology. To this end, we modified a widely employed immobilization stage used for popliteal lymph node preparations.

<table><tbody><tr><td>Narketan 10 %&amp;nbsp; (Ketamine) 20ml (100 mg/ml)</td><td>Vetoquinol</td><td>3605877535982</td><td></td></tr><tr><td>Rompun 2% (Xylazine) 25 ml (20 mg/ml)</td><td>Bayer</td><td>680538150144</td><td></td></tr><tr><td>Saline NaCl 0.9%</td><td>B. Braun</td><td>3535789</td><td></td></tr><tr><td>Prequillan 1% (Acepromazine) 10 ml (10 mg/ml)</td><td>Fatro</td><td>6805671900029</td><td></td></tr><tr><td>Electric shaver</td><td>Wahl</td><td>9818L</td><td>or similar</td></tr><tr><td>Hair removal cream</td><td>Veet</td><td>4002448090656</td><td></td></tr><tr><td>Durapore Surgical tape (2.5 cm x 9.1 m and 1.25 cm x 9.1 m)</td><td>Durapore (3M)</td><td>1538-1</td><td></td></tr><tr><td>Durapore Surgical tape (2.5 cm x 9.1 m and 1.25 cm x 9.1 m)</td><td>Durapore (3M)</td><td>1538-0</td><td></td></tr><tr><td>Super glue Ultra gel, instantaneous glue</td><td>Pattex, Henkel</td><td>4015000415040</td><td></td></tr><tr><td>Microscope cover glass slides 20 mm and 22 mm</td><td>Menzel-Gl&amp;auml;ser</td><td>631-1343/ 631-1344</td><td></td></tr><tr><td>Grease for laboratories 60 g glisseal N</td><td>Borer (VWR supplier)</td><td>DECO514215.00-CA15</td><td></td></tr><tr><td>Surgical scissors</td><td>Fine Science Tools (F.S.T )</td><td>14090-09</td><td>or similar</td></tr><tr><td>Fine Forceps</td><td>Fine Science Tools (F.S.T )</td><td>11252-20</td><td>or similar</td></tr><tr><td>Cotton swab</td><td>Migros</td><td>617027988254</td><td>or similar</td></tr><tr><td>Gauze Gazin 5 x 5 cm</td><td>Lohmann and Rauscher</td><td>18500</td><td>or similar</td></tr><tr><td>Stereomicroscope</td><td>Leica</td><td>MZ16</td><td>or similar</td></tr><tr><td>Texas Red dextran 70kDa</td><td>&amp;nbsp;Molecular Probes</td><td>D1864</td><td></td></tr><tr><td>Cascade Blue dextran 10kDa</td><td>invitrogen</td><td>D1976</td><td></td></tr><tr><td>Two-photon system</td><td>LaVision Biotec</td><td>TrimScope I and II</td><td>or similar</td></tr><tr><td>XLUMPLANFL 20x/0.95 W objective</td><td>Olympus</td><td>n/a</td><td>or other water immersion objective&amp;nbsp;</td></tr><tr><td>Digital thermometer</td><td>Fluke</td><td>95969077651</td><td></td></tr></tbody></table>

preparation of murine submandibular salivary gland for upright intravital microscopy

The submandibular salivary gland (SMG) is one of the three major salivary glands, and is of interest for many different fields of biological research, including cell biology, oncology, dentistry, and immunology. The SMG is an exocrine gland comprised of secretory epithelial cells, myofibroblasts, endothelial cells, nerves, and extracellular matrix. Dynamic cellular processes in the rat and mouse SMG have previously been imaged, mostly using inverted multi-photon microscope systems. Here, we describe a straightforward protocol for the surgical preparation and stabilization of the murine SMG in anesthetized mice for in vivo imaging with upright multi-photon microscope systems. We present representative intravital image sets of endogenous and adoptively transferred fluorescent cells, including the labeling of blood vessels or salivary ducts and second harmonic generation to visualize fibrillar collagen. In sum, our protocol allows for surgical preparation of mouse salivary glands in upright microscopy systems, which are commonly used for intravital imaging in the field of immunology.

This protocol allows imaging of almost the entire dorsal or ventral side of the SMG. The field of view typically also includes the sublingual salivary gland, which differs slightly from the SMG in cellular composition4. Both glands are encapsulated by fibrillar collagen and subdivided into lobes. Most 2PM systems can produce a label-free image of fibrillar collagen by measuring the 2nd harmonic signal, but usually fluorescent molecules are needed for vis...

Watch this Scientific Journal Video about Preparation of Murine Submandibular Salivary Gland for Upright Intravital Microscopy at JoVE.com

Preparation of Murine Submandibular Salivary Gland for Upright Intravital Microscopy

We describe a protocol to surgically expose and stabilize the murine submandibular salivary gland for intravital imaging using upright intravital microscopy. This protocol is easily adaptable to other exocrine glands of the head and neck region of mice and other small rodents.

The submandibular salivary gland (SMG) is one of the three major salivary glands, and is of interest for many different fields of biological research, ...

preparation-murine-submandibular-salivary-gland-for-upright

Research

JoVE Journal

Immunology and Infection

8.1K Views.  University of Bern. We beschrijven een protocol om operatief bloot en stabiliseren de lymfkliertest submandibulaire speekselklier voor intravital geschikt zijn met behulp van rechtop intravital microscopie. Dit protocol is gemakkelijk aan te passen aan andere exocrine klieren van het hoofd en nek regio van muizen en andere kleine knaagdieren.

Video: Preparation of Murine Submandibular Salivary Gland for Upright Intravital Microscopy

Cannulation of the Mouse Submandibular Salivary Gland via the Wharton's Duct

Severe salivary gland hypofunction is frequently found in patients with Sj&ouml;gren's syndrome and those who receiving therapeutic 
irradiation in their head and neck regions for cancer treatment. Both groups of patients experience symptoms such as xerostomia (dry mouth), dysphagia 
(impaired chewing and swallowing), severe dental caries, altered taste, oro-pharyngeal infections (candidiasis), mucositis, pain and discomfort.
One innovative approach of regenerative medicine for the treatment of salivary gland hypo-function is speculated in RS Redman, E Mezey 
et al. 2009: stem cells can be directly deposited by cannulation into the gland as a potent method in reviving the functions of the impaired organ. Presumably, 
the migrated foreign stem cells will differentiate into glandular cells to function as part of the host salivary gland. Also, this cannulation technique is an 
expedient and effective delivery method for clinical gene transfer application.
Here we illustrate the steps involved in performing the cannulation procedure on the mouse submandibular salivary gland via the Wharton's duct 
(Fig 1). C3H mice (Charles River, Montreal, QC, Canada) are used for this experiment, which have been kept under clean conventional conditions at 
the McGill University animal resource center. All experiments have been approved by the University Animal Care Committee and were in accordance with the 
guidelines of the Canadian Council on Animal Care.
For this experiment, a trypan blue solution is infused into the gland through the opening of the Wharton's duct using a 
insulin syringe with a 29-gauge needle encased inside a polyethylene tube. Subsequently, the mouse is dissected to show that the infusions migrated 
into the gland successfully.

A protocol for the cannulation of the mouse submandibular salivary gland via the Wharton's duct is described. For this experiment, the trypan blue solution is used as a dyer to demonstrate how this technique effectively delivers infusions into the targeted gland, and to suggest the reliability of this new approach as a potential clinical drug/cell therapy for the regeneration of salivary glands.

Severe salivary gland hypofunction is frequently found in patients with Sj&ouml;gren's syndrome and those who receiving therapeutic 
irradiation in their ...

Medicine

Retroductal Nanoparticle Injection to the Murine Submandibular Gland

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

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

Two common goals of salivary gland therapeutics are prevention and cure of tissue dysfunction following either autoimmune or radiation injury. By locally ...

Bioengineering

Retroductal Submandibular Gland Instillation and Localized Fractionated Irradiation in a Rat Model of Salivary Hypofunction

Normal tissues that lie within the portals of radiation are inadvertently damaged. Salivary glands are often injured during head and neck radiotherapy. Irreparable cell damage results in a chronic loss of salivary function that impairs basic oral activities, and increases the risk of oral infections and dental caries. Salivary hypofunction and its complications gravely impact a patient's comfort. Current symptomatic management of the condition is ineffective, and newer therapies to assuage the condition are needed. 
Salivary glands are exocrine glands, which expel their secretions into the mouth via excretory ducts. Cannulation of these ducts provides direct access to the glands. Retroductal delivery of a contrast agent to major salivary glands is a routine out-patient procedure for diagnostic imaging. Using a similar procedure, localized treatment of the glands is feasible. However, performing this technique in preclinical studies with small animals poses unique challenges. In this study we describe the technique of retroductal administration in rat submandibular glands, a procedure that was refined in Dr. Bruce Baum's laboratory (NIH)1, and lay out a procedure for local gland irradiation.

Salivary gland hypofunction, a major adverse effect of head and neck radiotherapy diminishes a patient's quality of life. The demonstration of efficacy of new therapies in animal models is a prerequisite before clinical transition. This protocol describes retroductal administration and local irradiation of rat submandibular glands.

Normal tissues that lie within the portals of radiation are inadvertently damaged. Salivary glands are often injured during head and neck radiotherapy. ...

Genetic Modification and Recombination of Salivary Gland Organ Cultures

Branching morphogenesis occurs during the development of many organs, and the embryonic mouse submandibular gland (SMG) is a classical model for the study of branching morphogenesis. In the developing SMG, this process involves iterative steps of epithelial bud and duct formation, to ultimately give rise to a complex branched network of acini and ducts, which serve to produce and modify/transport the saliva, respectively, into the oral cavity1-3. The epithelial-associated basement membrane and aspects of the mesenchymal compartment, including the mesenchyme cells, growth factors and the extracellular matrix, produced by these cells, are critical to the branching mechanism, although how the cellular and molecular events are coordinated remains poorly understood 4. The study of the molecular mechanisms driving epithelial morphogenesis advances our understanding of developmental mechanisms and provides insight into possible regenerative medicine approaches. Such studies have been hampered due to the lack of effective methods for genetic manipulation of the salivary epithelium. Currently, adenoviral transduction represents the most effective method for targeting epithelial cells in adult glands in vivo5. However, in embryonic explants, dense mesenchyme and the basement membrane surrounding the epithelial cells impedes viral access to the epithelial cells. If the mesenchyme is removed, the epithelium can be transfected using adenoviruses, and epithelial rudiments can resume branching morphogenesis in the presence of Matrigel or laminin-1116,7. Mesenchyme-free epithelial rudiment growth also requires additional supplementation with soluble growth factors and does not fully recapitulate branching morphogenesis as it occurs in intact glands8. Here we describe a technique which facilitates adenoviral transduction of epithelial cells and culture of the transfected epithelium with associated mesenchyme. Following microdissection of the embryonic SMGs, removal of the mesenchyme, and viral infection of the epithelium with a GFP-containing adenovirus, we show that the epithelium spontaneously recombines with uninfected mesenchyme, recapitulating intact SMG glandular structure and branching morphogenesis. The genetically modified epithelial cell population can be easily monitored using standard fluorescence microscopy methods, if fluorescently-tagged adenoviral constructs are used. The tissue recombination method described here is currently the most effective and accessible method for transfection of epithelial cells with a wild-type or mutant vector within a complex 3D tissue construct that does not require generation of transgenic animals.

A technique to genetically manipulate epithelial cells within whole ex vivo cultured embryonic mouse submandibular glands (SMGs) using viral gene transfer is described. This method takes advantage of the innate ability of SMG epithelium and mesenchyme to spontaneously recombine after separation and infection of epithelial rudiments with adenoviral vectors.

Branching  morphogenesis occurs during the development of many organs, and the embryonic  mouse submandibular gland (SMG) is a classical model for the ...

Biology

Murine Salivary Functional Assessment via Pilocarpine Stimulation Following Fractionated Radiation

Hyposalivation is commonly observed in the autoimmune reaction of Sj&#246;gren's syndrome or following radiation injury to the major salivary glands. In these cases, questions remain regarding disease pathogenesis and effective interventions. An optimized technique that allows functional assessment of the salivary glands is invaluable for investigating exocrine gland biology, dysfunction, and therapeutics. Here, we present a step by step approach to performing pilocarpine stimulated saliva secretion, including tracheostomy and the dissection of the three major murine salivary glands. We also detail the appropriate murine head and neck anatomy accessed during these techniques. This approach is scalable, allowing for multiple mice to be processed simultaneously, thus improving the efficiency of the work flow. We aim to improve the reproducibility of these methods, each of which has further applications within the field. In addition to saliva collection, we discuss metrics for quantifying and normalizing functional capacity of these tissues. Representative data are included from submandibular glands with depressed salivary gland function 2 weeks following fractionated radiation (4 doses of 6.85 Gy).

We present a detailed approach to performing saliva collection, including murine tracheostomy and the isolation of three major salivary glands.

Hyposalivation is commonly observed in the autoimmune reaction of Sj&#246;gren's syndrome or following radiation injury to the major salivary glands. In ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

Radiation Treatment of Organotypic Cultures from Submandibular and Parotid Salivary Glands Models Key In Vivo Characteristics

Hyposalivation and xerostomia create chronic oral complications that decrease the quality of life in head and neck cancer patients who are treated with radiotherapy. Experimental approaches to understanding mechanisms of salivary gland dysfunction and restoration have focused on in vivo models, which are handicapped by an inability to systematically screen therapeutic candidates and efficiencies in transfection capability to manipulate specific genes. The purpose of this salivary gland organotypic culture protocol is to evaluate maximal time of culture viability and characterize cellular changes following ex vivo radiation treatment. We utilized immunofluorescent staining and confocal microscopy to determine when specific cell populations and markers are present during a 30-day culture period. In addition, cellular markers previously reported in in vivo radiation models are evaluated in cultures that are irradiated ex vivo. Moving forward, this method is an attractive platform for rapid ex vivo assessment of murine and human salivary gland tissue responses to therapeutic agents that improve salivary function.

Using three-dimensional organotypic cultures to visualize morphology and functional markers of salivary glands may provide novel insights into the mechanisms of tissue damage following radiation. Described here is a protocol to section, culture, irradiate, stain, and image 50&#8211;90 &#956;m thick salivary gland sections prior to and following exposure to ionizing radiation.

Hyposalivation and xerostomia create chronic oral complications that decrease the quality of life in head and neck cancer patients who are treated with ...

Real-Time Live Imaging of Mouse Submandibular Gland Regeneration

Interrogating Cell-Cell Interactions in the Salivary Gland via Ex Vivo Live Cell Imaging

Salivary gland regeneration is a complex process involving intricate interactions among various cell types. Recent studies have shed light on the pivotal role played by macrophages in driving the regenerative response. However, our understanding of this critical role has primarily relied on static views obtained from fixed tissue biopsies. To overcome this limitation and gain insights into these interactions in real time, this study outlines a comprehensive protocol for culturing salivary gland tissue ex vivo and capturing live images of cell migration.
The protocol involves several key steps: First, mouse submandibular salivary gland tissue is carefully sliced using a vibratome and then cultured at an air-liquid interface. These slices can be intentionally injured, for instance, through exposure to radiation, to induce cellular damage and trigger the regenerative response. To track specific cells of interest, they can be endogenously labeled, such as by utilizing salivary gland tissue collected from genetically modified mice where a particular protein is marked with green fluorescent protein (GFP). Alternatively, fluorescently-conjugated antibodies can be employed to stain cells expressing specific cell surface markers of interest. Once prepared, the salivary gland slices are subjected to live imaging using a high-content confocal imaging system over a duration of 12 h, with images captured at 15 min intervals. The resulting images are then compiled to create a movie, which can subsequently be analyzed to extract valuable cell behavior parameters. This innovative method provides researchers with a powerful tool to investigate and better understand macrophage interactions within the salivary gland following injury, thereby advancing our knowledge of the regenerative processes at play in this dynamic biological context.

Author Spotlight: Studying Macrophage-Epithelial Cell Interactions in Salivary Gland Regeneration After Injury 

Immunofluorescent imaging is constrained by the ability to observe complex, time-dependent biological processes in just a single snapshot in time. This study outlines a live-imaging approach conducted on precision-cut mouse submandibular gland slices. This approach allows for the real-time observation of cell-cell interactions during homeostasis and the processes of regeneration and repair.

Salivary gland regeneration is a complex process involving intricate interactions among various cell types. Recent studies have shed light on the pivotal ...

Manipulating the Murine Lacrimal Gland

The lacrimal gland (LG) secretes aqueous tears necessary for maintaining the structure and function of the cornea, a transparent tissue essential for vision. In the human a single LG resides in the orbit above the lateral end of each eye delivering tears to the ocular surface through 3 - 5 ducts. The mouse has three pairs of major ocular glands, the most studied of which is the exorbital lacrimal gland (LG) located anterior and ventral to the ear. Similar to other glandular organs, the LG develops through the process of epithelial branching morphogenesis in which a single epithelial bud within a condensed mesenchyme undergoes multiple rounds of bud and duct formation to form an intricate interconnected network of secretory acini and ducts. This elaborate process has been well documented in many other epithelial organs such as the pancreas and salivary gland. However, the LG has been much less explored and the mechanisms controlling morphogenesis are poorly understood. We suspect that this under-representation as a model system is a consequence of the difficulties associated with finding, dissecting and culturing the LG. Thus, here we describe dissection techniques for harvesting embryonic and post-natal LG and methods for ex vivo culture of the tissue.

The lacrimal gland (LG) is a branching organ that produces the aqueous components of tears necessary for maintaining vision and ocular health. Here we describe murine LG dissection and ex vivo culture techniques to decipher signaling pathways involved in LG development.

The lacrimal gland (LG) secretes aqueous tears necessary for maintaining the structure and function of the cornea, a transparent tissue essential for ...

Intravital Subcellular Microscopy of the Mammary Gland

The mammary gland constitutes a model par excellence for investigating epithelial functions, including tissue remodeling, cell polarity, and secretory mechanisms. During pregnancy, the gland expands from a primitive ductal tree embedded in a fat pad to a highly branched alveolar network primed for the formation and secretion of colostrum and milk. Post-partum, the gland supplies all the nutrients required for neonatal survival, including membrane-coated lipid droplets (LDs), proteins, carbohydrates, ions, and water. Various milk components, including lactose, casein micelles, and skim-milk proteins, are synthesized within the alveolar cells and secreted from vesicles by exocytosis at the apical surface. LDs are transported from sites of synthesis in the rough endoplasmic reticulum to the cell apex, coated with cellular membranes, and secreted by a unique apocrine mechanism. Other preformed constituents, including antibodies and hormones, are transported from the serosal side of the epithelium into milk by transcytosis. These processes are amenable to intravital microscopy because the mammary gland is a skin gland and, therefore, directly accessible to experimental manipulation. In this paper, a facile procedure is described to investigate the kinetics of LD secretion in situ, in real-time, in live anesthetized mice. Boron-dipyrromethene (BODIPY)665/676&#160;or monodansylpentane are used to label the neutral lipid fraction of transgenic mice, which either express soluble EGFP (enhanced green fluorescent protein) in the cytoplasm, or a membrane-targeted peptide fused to either EGFP or tdTomato. The membrane-tagged fusion proteins serve as markers of cell surfaces, and the lipid dyes resolve LDs &#8805; 0.7 &#181;m. Time-lapse images can be recorded by standard laser scanning confocal microscopy down to a depth of 15-25 &#181;m or by multiphoton microscopy for imaging deeper in the tissue. The mammary gland may be bathed with pharmacological agents or fluorescent dyes throughout the surgery, providing a platform for acute experimental manipulations as required.

The present protocol describes a facile technique for the intravital imaging of the lactating mouse mammary gland by laser scanning confocal and multiphoton microscopy.

The mammary gland constitutes a model par excellence for investigating epithelial functions, including tissue remodeling, cell polarity, and secretory ...

Preparation of Murine Submandibular Salivary Gland for Upright Intravital Microscopy

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