This work was supported by research grants (NIH/NIDCD 009269, 012831 and ARRA administrative supplement NIH grants) to Weihong Lin. We especially thank Mr. Tim Ford at UMBC for his technical assistance in videotaping and processing. We also wish to thank Dr. Daphne Blumberg, Ms. Chere Petty at UMBC and Mr. Nicholas McCollum from Olympus America Inc. for their equipment assistance in videotaping.

1. Mouse Nose Preparation
We used adult C57BL/6 background mice in this study. All animal care and procedures are approved by the Animal Care and Use Committees (IACUC) of University of Maryland, Baltimore County.

 1.1 Acquiring the nose from paraformaldahyde-fixed mice
<img src="/files/ftp_upload/50538/50538fig1.jpg" alt="Figure 1" fo:content-width="6in" fo:src="/files/ftp_upload/50538/50538fig1highres.jpg"/> 
Figure 1. Bones of a mouse skull. A: Dorsal view of the skull. B: Ventral view of the skull with the mandible removed. The skull was prepared from a 40-day old mouse. Individual bones are colored for better visualization. <a href="https://www.jove.com/files/ftp_upload/50538/50538fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<ol>
 <li>Perfuse transcardially to fix individual mice following the protocol of Lin, et al., (2008)16. Briefly, mice were deeply anesthetized with tribromoethanol (Avertin 250 &mu;g/g body weight), perfused transcardially with 0.1M phosphate buffer (PB, 30-50 ml), followed by a phosphate buffered fixative containing 3% paraformaldehyde, 19 mM L-lysine monohydrochloride, and 0.23% sodium m-periodate (approximately 35-50 ml). One can also follow the steps in the JoVE article for animal perfusion 35. </li>
 <li>Use a pair of scissors to cut off mandible (or the lower jaw) and remove the skin on the head. </li>
 <li>Separate the entire head from the rest of the body. </li>
 <li>Remove the palate. Also, clean and remove the remaining connective tissue and muscle on the surface of the skull to obtain the specimen shown in Figures 1A and 1B.</li>
 <li>Under a dissection microscope, remove the skull bone covering the brain and olfactory bulbs. Trim off excess tissue and bones. Note, for the extended tissue preparation in which the brain and nose remain connected, only the skull bones are removed. For immunohistochemical experiments, the tissue was post-fixed for 1.5 hr and transferred to 0.1M phosphate buffered saline (PBS) with 25% sucrose overnight. Nasal tissue should be kept humidified throughout the dissection by dipping it in the buffered sucrose solution several times. </li>
</ol>
1.2 Acquiring the nose from freshly euthanized mice
<ol>
 <li>Individual mice were transferred to a clean cage and exposed to CO2 gas, which was followed by cervical dislocation 5 min after the final breath. To reduce the blood in the nose tissue, use a pair of scissors to open the chest and cut the heart to allow blood to drain. </li>
 <li>Repeat steps 1.1.2 to 1.1.5, except the post-fixation and cryoprotection with 25% sucrose. The specimen should be kept humidified and maintained with Tyrode&#x2019;s saline containing (in mM): 140 NaCl, 5 KCl, 1 MgCl2, 1 CaCl2, 10 Na pyruvate, 10 D-glucose, and 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid buffer (HEPES, adjusted at pH 7.4). Alternatively, fold a piece of Kimwipes, soak it with Tyrode&#x2019;s solution, and place beneath the specimen while dissecting and occasionally dip the specimen in the Tyrode's solution or drop some solution onto the tissue to keep it humidified to maintain the viability of the cells and tissues. </li>
</ol>
2. Incisor, Anterior Vomer and Maxilla Bone Removal
<ol>
 <li>Start from a ventral viewpoint. Locate the vomer bone and break the vomer bone along its length using a rongeur or forceps with teeth to break the ventral most part of the bone. </li>
 <li>Using serrated forceps, remove the broken segments of the vomer bone by gently plying the bone fragments away from the vomeronasal organ (VNO). Note, for mice less than a month old, or if only interested in the MOE, these two steps can be skipped.</li>
 <li>Hold the head firmly with forceps. Use the rongeur to break the front portion of both incisors and the jointed region of anterior maxilla between the two incisors and the vomer bone. </li>
 <li>Break the ventral portion of the right maxilla at the region just anterior to the zygomatic arch up to the level of the dorsal zygomatic plate. </li>
 <li>Flip over the nose for a dorsal viewpoint. Use the rongeur to break the right maxilla anterior of the dorsal zygomatic plate. The entire right maxilla anterior of the zygomatic plate should be loose. Use the fine forceps to gently separate any nasal tissue underlying the maxilla, and then gently lift away the bone fragment. </li>
 <li>Repeat steps 2.4 and 2.5 to loosen the left incisor and left anterior maxilla and remove them after the nasal bone has been removed. </li>
</ol>
3. Nasal Bone and Dorsal Zygomatic Plate Removal 
<ol>
 <li>Use the serrated forceps or the rongeur to remove the remainder of the frontal bones just caudal to the nasal bones. After removing this piece of bone, the caudal part of the nasal bone can be gripped using forceps.</li>
 <li>Use the rongeur to break the anterior portion of the zygomatic arch, which is connected to the zygomatic plate. </li>
 <li>Take the fine forceps at the lateral edge of the dorsal end of the zygomatic plate and gently flip the bone and remove it. If the bone is not loose, use the rongeur to clamp it gently to loosen it for removal. </li>
 <li>Use the fine forceps or a razor blade to loosen the medial suture between the right and left nasal bones. </li>
 <li>Use the serrated forceps to grip the caudal end of the right nasal bone. Gently move the bone from side to side to separate it from underlying tissue. It is useful to move the forceps along the caudal third of the nasal bone for moving the bone side to side. As the nasal bone separates, slowly lift the bone from the caudal end. </li>
 <li>While the nasal bone is slightly lifted, tilt the bone laterally to reveal a lateral outcropping of the bone that is lined with thin respiratory epithelial tissue. Use fine forceps to gently release this tissue from the nasal bone. Continue to lift the nasal bone. When the bone is completely separated from underlying tissue, use scissors to cut off the nasal bone at the rostral end. </li>
 <li>Repeat steps 3.1, 3.5 and 3.6 for the removal of left nasal bone.</li>
 <li> Repeat steps 3.2 and 3.3 for the left side of the nose. </li>
</ol>
4. Lateral Zygomatic Plate Removal 
<ol>
 <li>Remove the zygomatic plate one side at a time. Either zygomatic plate can be removed first.</li>
 <li>From a ventral viewpoint, break the maxilla inferior to the zygomatic arch until break reaches the zygomatic arch. </li>
 <li>From a dorsal viewpoint, gently grab the zygomatic arch and lift forward and lateral. If the zygomatic plate is still attached to any tissue, take the fine forceps and gently sever the connections between the tissue and the bone. </li>
 <li>Repeat for the zygomatic plate on the other side of the nose. </li>
</ol>
5. Orbit Bone Removal
<ol>
 <li>From a ventral viewpoint, break the palantine bone between the molars of the nose. </li>
 <li>Use the rongeur to break the 3 molars and the maxilla on each side of the nose. </li>
 <li>Break and remove any remaining thick pieces of bone ventral and posterior to the turbinates on each side of the nose. </li>
</ol>
6. Ethmoid Bone Removal
<ol>
 <li>Break any portion of the ethmoid bone protruding caudal to the turbinates. This is necessary to avoid the loss of turbinate tissue when removing thin pieces of the ethmoid bone covering the turbinates.</li>
 <li>For the right side of the nose, place the fine forceps at the anterior edge of the ethmoid bone and gently remove it. If a portion of the bone remains, repeat this procedure until all the thin bone covering the turbinates has been removed. The turbinate bones do not need to be removed in most of the preparations. In aged mice, the cribriform plate becomes brittle. If cryosectioning of the nasal tissue is needed, remove small pieces of the plate with fine forceps to reduce the potential damage caused by the bone. </li>
 <li>Repeat step 6.2) for the left side of the nose. </li>
 <li>Remove any remaining bone fragments prior to sectioning. Note: in animals more than a year-old, the posterodorsal region of the septum bone is somewhat thick and hard. One can remove this portion using fine forceps. Insert the tip of the forceps into both side of the bone to separate the dorsal portion of the septum bone and the lining epithelial tissue. Use one pair of forceps to grab the bone and hold the specimen. Use another pair of forceps to break the upper bony portion from the lower cartilaginous portion of the septum and gently remove it. </li>
</ol>
7. Nose Preparation for Cryosectioning
<ol>
 <li>Set up the aspirator vacuum pump. </li>
 <li>Place the nose in an embedding mold. Submerge the nose in OCT media.</li>
 <li>Use a vacuum to remove air bubbles trapped within the nose tissue. This process takes up to 5 min.</li>
 <li>After removing air bubbles, set the tissue in the desired orientation.</li>
 <li>Freeze the OCT and tissue in the mold using dry ice. The embedded tissue can then be cryosectioned immediately or stored at -80 &deg;C for future use. </li>
</ol>

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F., Lin, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+profile+of+G-protein+&#946;&#947;+subunit+gene+transcripts+in+the+mouse+olfactory+sensory+epithelia.">Expression profile of G-protein &#946;&#947; subunit gene transcripts in the mouse olfactory sensory epithelia.</a> Frontiers in Cellular Neuroscience. 7, 84 (2013).</li></ol>

The authors declare that they have no competing financial interests.

Here, we demonstrated a step-by-step procedure for isolating intact olfactory and respiratory tissue from the mouse nose by sequentially removing the surrounding bones while sparing the tissue below. We show that careful bone removal can preserve even the most delicate tissues in their entirety. We also share insight into possible modifications of this technique, in which we isolate both the brain and nose tissue together to preserve the nerve connection. This new method provides a means for isolating whole olfactory ...

The mammalian nasal cavity contains various types of tissues and organs that serve distinct functions. The nasal cavity makes up the entry portion of the upper respiratory tract, which allows air travel into and out of the lungs. Inhaled air passes through the nasal cavity where it undergoes temperature and humidity conditioning 1 as well as cleaning or filtering to remove irritating and toxic substances and infectious microorganisms 2. Both treatments are carried out by nasal epithelia and subepithelial tissues, including glands and vessels and are critical for protecting the lower airways and the lungs. In addition to its role in respiration and epithelial defense, the nasal tissue also contains peripheral sensory apparatuses of the olfactory and trigeminal systems, which detect a wide range of chemical substances in the passing air. Depending on which system is activated, sensory detection of chemicals in the nose can elicit either a sense of smell, irritation, or pain 3,4. 
The peripheral olfactory system is complex and made up of several anatomically separated olfactory sensory organs within the nasal cavity. Among them, the main olfactory epithelium (MOE) is the largest, which makes up approximately 45-52% of the nasal epithelia in rodents 5 and is located in the posterior region. In the anteroventral region, there is a pair of tubular structures known as the vomeronasal organ 6, which sit along each side of the nasal septum. Two additional small groupings of olfactory sensory neurons, known as the septal organ of Masera 7,8 and the Gruneberg ganglion 9, reside along the ventral septum and the dorsal entry region of the nasal cavity, respectively. These peripheral organs contain neuro-epithelia with distinctive features in morphology, cell marker expression, and physiological function. Together they detect thousands of odor molecules with exquisite sensitivity 10-12. 
In addition to the olfactory sensory organs, the nasal cavity also houses other sensory systems. It is known that peptidergic trigeminal nerve fibers are present in the nasal epithelium, especially the respiratory epithelium 13,14. Some of these fibers detect irritating and toxic chemicals and are responsible for initiating protective reflexes such as coughing and sneezing 4,15. Irritating odorous and bitter compounds can also be detected by a recently discovered population of solitary chemosensory cells (SCCs), many of which are innervated by trigeminal nerve fibers 16-19. These SCCs are located in higher density in the entry region of the nasal cavity and vomeronasal entry ducts, hinting that they may also serve a protective function 16-18. Thus, nasal epithelia can differ substantially in function, morphology, and cell composition depending on their anatomical locations.
Even within a single and specialized epithelium, there are regional differences. The MOE is one such example. The MOE lines various turbinates, which are complicated and curled structures. Because of them, different regions of the MOE experience different air flow rates, and thus, different diffusion and clearance rates of airborne odor molecules 20. Also, it is known that olfactory sensory neurons (OSNs) expressing a given odor receptor are located in one of four circumvented zones of the MOE 21,22. How this location difference affects an OSN&#x2019;s response to odorants is largely not known. In addition, some OSN populations exhibit regional preference. Guanylyl cyclase-D (GC-D)-expressing OSNs have zonal distributions favoring the cul-de-sac regions of the ectoturbinates 23,24. More recently, we found a subpopulation of canonical OSNs that expresses transient receptor potential channel M5 (trpM5) and is preferentially located in the lateral and ventral regions 25. These results indicate that MOE is not uniform. However, how these regional differences affect olfactory coding is not understood. This is in part because thorough physiological investigation of the MOE and the nose has been limited by the difficulty of obtaining intact nasal epithelia with preserved anatomical organization using current methods.
The nasal epithelia are predominantly surrounded by the anterior bones of the skull, including the nasal, maxilla, palatine, zygomatic, and ethmoid bones. In adult mice and other rodent models, these bones are hard and difficult to remove without damaging the closely associated nasal tissue, particularly the delicate turbinates. Often, chemical-based decalcification is used to soften bones to allow cryosectioning of nasal tissues for morphological, immunohistochemical, and in situ hybridization studies; however, depending on the age of the animal, the decalcification process can last overnight up to 7 days 24,26-28. This treatment is also limited because it requires tissue be fixative-preserved. Additionally, chemical decalcification can be harsh and affect the immunolabeling of some sensitive antibodies 29,30. For physiological studies, live tissue is required, and thus, these experiments are often conducted on isolated OSNs or MOE slices obtained from neonates whose skull bones are thin and soft 17,31,32. Physiological studies can also utilize whole mount preparations by splitting the head 25,33,34, but usually only the medial surface of the nose is easily accessible, limiting physiological recordings on other areas. 
Here, we describe an effective, manual deboning method to prepare intact nasal tissues with preserved original anatomical organization and morphology. We sequentially remove the major bones of the anterior skull under a dissection microscope to expose an almost entirely intact nasal epithelium while keeping the thin turbinate bones intact unless the mice are very old and cryosectioning is needed. We also extend the method to preserve the connection between the nasal tissues and olfactory bulbs, as well as the rest of the brain, thus facilitating simultaneous examination of both peripheral and central circuits. Our method can be used to prepare paraformaldehyde-fixed, as well as fresh, live nasal tissue. Thus, our method is expected to facilitate morphological, immunohistochemical and physiological studies of respiration, olfaction, and nasal damage and illness.

<table border="1">
 <tr>
 <td>Name</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments </td>
 </tr>
 <tr>
 	<td></td>
 <td></td>
 <td></td>
 <td>Dissection</td>
 </tr>
 <tr>
 <td>Rongeur, 1.0 mm Jaw width</td>
 <td>World Precision Instruments (WPI)</td>
 <td>501333</td>
 <td></td>
 </tr>
 <tr>
 <td>Fine forceps, Dumont 3 </td>
 <td>WPI</td>
 <td>503235 </td>
 <td></td>
 </tr>
 <tr>
 <td>Fine forceps, Dumont 55</td>
 <td>WPI</td>
 <td>14099</td>
 <td></td>
 </tr>
 <tr>
 <td>Fine forceps, Dumont AA</td>
 <td>Fine Science Tools (FST)</td>
 <td>11210-20</td>
 <td></td>
 </tr>
 <tr>
 <td>Specimen forceps, Serrated</td>
 <td>VWR</td>
 <td>82027-440</td>
 <td></td>
 </tr>
 <tr>
 <td>Operating scissors</td>
 <td>WPI</td>
 <td>501753</td>
 <td></td>
 </tr>
 <tr>
 <td>Iris scissors, Straight</td>
 <td>Miltex</td>
 <td>V95-304</td>
 <td></td>
 </tr>
 <tr>
 <td>Dissection microscope</td>
 <td>Olympus</td>
 <td>SZ40</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>[header]</td>
 </tr>
 <tr>
 	<td></td>
 <td></td>
 <td></td>
 <td>Tissue embedding</td>
 </tr>
 <tr>
 <td>Optimum cutting temperature (OCT) compound</td>
 <td>Sakura Finetek</td>
 <td>4583</td>
 <td></td>
 </tr>
 <tr>
 <td>Plastic embedding mold</td>
 <td>VWR</td>
 <td>15160-215 </td>
 <td></td>
 </tr>
 <tr>
 <td>Aspirator vacuum pump</td>
 <td>Fisher Scientific</td>
 <td>09-960-2</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>[header]</td>
 </tr>
 <tr>
 	 <td></td>
 <td></td>
 <td></td>
 <td>Section staining </td>
 </tr>
 <tr>
 <td>Neutral red</td>
 <td>ACROS Organic</td>
 <td>CAS 553-24-2</td>
 <td>Nuclei staining</td>
 </tr>
</table>

an effective manual deboning method to prepare intact mouse nasal tissue with preserved anatomical organization

The mammalian nose is a multi-functional organ with intricate internal structures. The nasal cavity is lined with various epithelia such as olfactory, respiratory, and squamous epithelia which differ markedly in anatomical locations, morphology, and functions. In adult mice, the nose is covered with various skull bones, limiting experimental access to internal structures, especially those in the posterior such as the main olfactory epithelium (MOE). Here we describe an effective method for obtaining almost the entire and intact nasal tissues with preserved anatomical organization. Using surgical tools under a dissecting microscope, we sequentially remove the skull bones surrounding the nasal tissue. This procedure can be performed on both paraformaldehyde-fixed and freshly dissected, skinned mouse heads. The entire deboning procedure takes about 20-30 min, which is significantly shorter than the experimental time required for conventional chemical-based decalcification. In addition, we present an easy method to remove air bubbles trapped between turbinates, which is critical for obtaining intact thin horizontal or coronal or sagittal sections from the nasal tissue preparation. Nasal tissue prepared using our method can be used for whole mount observation of the entire epithelia, as well as morphological, immunocytochemical, RNA in situ hybridization, and physiological studies, especially in studies where region-specific examination and comparison are of interest.

Using this method, we can reliably obtain almost entirely intact nasal tissue. Figure 2A shows an image of adult nasal specimen from a paraformaldehyde-fixed head. In this specimen, all four sub-olfactory sensory organs, including the MOE, septal organ, the Gruneberg ganglion, and VNO, are intact. Also, the respiratory epithelia and subepithelial tissues, such as glands and vessels, are preserved. We have successfully used this method in a number of studies in which we investigated morphology, distri...

Watch this Scientific Journal Video about An Effective Manual Deboning Method To Prepare Intact Mouse Nasal Tissue With Preserved Anatomical Organization at JoVE.com

An Effective Manual Deboning Method To Prepare Intact Mouse Nasal Tissue With Preserved Anatomical Organization

The mammalian nose is a multi-functional organ with intricate internal structures.  The nasal cavity is lined with various epithelia such as olfactory, ...

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JoVE Journal

Biology

34.4K Görüntüleme.  University of Maryland Baltimore County. 

Video: An Effective Manual Deboning Method To Prepare Intact Mouse Nasal Tissue With Preserved Anatomical Organization

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

In vivo Görüntüleme ve Yumurtalık Kanseri Ortotopik Fare Modeli Tedavi Tedaviler

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Biyoloji

Combined Immunofluorescence and DNA FISH on 3D-preserved Interphase Nuclei to Study Changes in 3D Nuclear Organization

Fluorescent in situ hybridization using DNA probes on 3-dimensionally preserved nuclei followed by 3D confocal microscopy (3D DNA FISH) represents the most direct way to visualize the location of gene loci, chromosomal sub-regions or entire territories in individual cells. This type of analysis provides insight into the global architecture of the nucleus as well as the behavior of specific genomic loci and regions within the nuclear space. Immunofluorescence, on the other hand, permits the detection of nuclear proteins (modified histones, histone variants and modifiers, transcription machinery and factors, nuclear sub-compartments, etc). The major challenge in combining immunofluorescence and 3D DNA FISH is, on the one hand to preserve the epitope detected by the antibody as well as the 3D architecture of the nucleus, and on the other hand, to allow the penetration of the DNA probe to detect gene loci or chromosome territories 1-5. Here we provide a protocol that combines visualization of chromatin modifications with genomic loci in 3D preserved nuclei.

Here we describe a protocol for simultaneous detection of histone modifications by immunofluorescence and DNA sequences by DNA FISH followed by 3D microscopy and analyses (3D immuno-DNA FISH).

3D Nükleer teşkilatında değişiklikler Eğitim için 3D-korunmuş Tane Çekirdekle ilgili Kombine İmmünofloresan ve DNA FISH

Fluorescent in situ hybridization using DNA probes  on 3-dimensionally preserved nuclei followed by 3D confocal microscopy (3D DNA  FISH) represents the ...

Hybrid-Cut: An Improved Sectioning Method for Recalcitrant Plant Tissue Samples

Maintaining plant section integrity is essential for studying detailed anatomical structures at the cellular, tissue, or even organ level. However, some plant cells have rigid cell walls, tough fibers and crystals (calcium oxalate, silica, etc.), and high water content that often disrupt tissue integrity during plant tissue sectioning.
This study establishes a simple Hybrid-Cut tissue sectioning method. This protocol modifies a paraffin-based sectioning technique and improves the integrity of tissue sections from different plants. Plant tissues were embedded in paraffin before sectioning in a cryostat at -16 &#176;C. Sectioning under low temperature hardened the paraffin blocks, reduced tearing and scratching, and improved tissue integrity significantly. This protocol was successfully applied to calcium oxalate-rich Phalaenopsis orchid tissues as well as recalcitrant tissues such as reproductive organs and leaves of rice, maize, and wheat. In addition, the high quality of tissue sections from Hybrid-Cut could be used in combination with in situ hybridization (ISH) to provide spatial expression patterns of genes of interest. In conclusion, this protocol is particularly useful for recalcitrant plant tissue containing high crystal or silica content. Good quality tissue sections enable morphological and other biological studies.

This protocol describes a simple Hybrid-Cut tissue sectioning method that is useful for recalcitrant plant tissues. Good quality tissue sections enable anatomical studies and other biological studies including in situ hybridization (ISH).

Hibrid-Cut: Dirençli Bir Bitki Doku Örnekleri Bir Geliştirilmiş Kesit Yöntemi

Maintaining plant section integrity is essential for studying detailed anatomical structures at the cellular, tissue, or even organ level. However, some ...

Trans-inner Cell Mass Injection of Embryonic Stem Cells Leads to Higher Chimerism Rates

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current blastocyst injection protocol. Here we report that a simple rotation of the embryo, and injection through Trans-Inner cell mass (TICM) increased the percentage of chimeric mice from 31% to 50%, with no additional equipment or further specialized training. 26 different inbred clones, and 35 total clones were injected over a period of 9 months. There was no significant difference in either pregnancy rate or recovery rate of embryos between traditional injection techniques and TICM. Therefore, without any major alteration in the injection process and a simple positioning of the blastocyst and injecting through the ICM, releasing the ES cells into the blastocoel cavity can potentially improve the quantity of chimeric production and subsequent germline transmission.

Here we present a protocol to increase chimera production without the use of new equipment. A simple orientation change of the embryo for injection can increase the number of embryos produced, and potentially reduce the timeline to germline transmission.

Trans-iç hücre embriyonik kök hücrelerin kitle enjeksiyon daha yüksek Chimerism oranları için yol açar.

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current ...

Isolation of Intact Eyeball to Obtain Integral Ocular Surface Tissue for Histological Examination and Immunohistochemistry

Ocular surface (OS) consists of an epithelial sheet with three connected parts: palpebral conjunctiva, bulbar conjunctiva and corneal epithelium. Disruption of OS would lead to keratitis, conjunctivitis or both (keratoconjunctivitis). In experimental animal models with certain genetic modifications or artificial operations, it is useful to examine all parts of the OS epithelial sheet to evaluate relative pathogenetic changes of each part in parallel. However, dissection of OS tissue as a whole without distortion or damage has been challenging, primarily due to the softness and thinness of the OS affixed to physically separate yet movable eyelids and eyeball. Additionally, the deep eye socket formed by the hard skull/orbital bones is fully occupied by the eyeball leaving limited space for operating dissections. As a result, direct dissection of the eyeball with associated OS tissues from the facial side would often lead to tissue damages, especially the palpebral and bulbar conjunctiva. In this protocol, we described a method to remove the skull and orbital bones sequentially from a bisected mouse head, leaving eyelids, ocular surface, lens and retina altogether in one piece. The integrity of the OS sheet was well preserved and could be examined by histology or immunostaining in a single section.

This protocol describes a method for the isolation of the mouse eyeball with eyelid, ocular surface, anterior and posterior segments in relatively intact position.

Histolojik Muayene ve İmmünohistokimya için İntegral Göz Yüzey Dokusu Elde Etmek Için Bozulmamış Göz Küresinin İzolasyon

Ocular surface (OS) consists of an epithelial sheet with three connected parts: palpebral conjunctiva, bulbar conjunctiva and corneal epithelium. ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Ultrayapısal Olarak Korunmuş Hücrelerde Membran Protein Lokalizasyonunun Elektron Mikroskobu Analizi için CryoAPEX Yöntemi

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...

Application of Passive Head Motion to Generate Defined Accelerations at the Heads of Rodents

Exercise is widely recognized as effective for various diseases and physical disorders, including those related to brain dysfunction. However, molecular mechanisms behind the beneficial effects of exercise are poorly understood. Many physical workouts, particularly those classified as aerobic exercises such as jogging and walking, produce impulsive forces at the time of foot contact with the ground. Therefore, it was speculated that mechanical impact might be implicated in how exercise contributes to organismal homeostasis. For testing this hypothesis on the brain, a custom-designed ''passive head motion'' (hereafter referred to as PHM) system was developed that can generate vertical accelerations with controlled and defined magnitudes and modes and reproduce mechanical stimulation that might be applied to the heads of rodents during treadmill running at moderate velocities, a typical intervention to test the effects of exercise in animals. By using this system, it was demonstrated that PHM recapitulates the serotonin (5-hydroxytryptamine, hereafter referred to as 5-HT) receptor subtype 2A (5-HT2A) signaling in the prefrontal cortex (PFC) neurons of mice. This work provides detailed protocols for applying PHM and measuring its resultant mechanical accelerations at rodents' heads.

The present protocol describes a custom-designed ''passive head motion'' system, which reproduces mechanical accelerations at rodents' heads generated during their treadmill running at moderate velocities. It allows dissecting mechanical factors/elements from the beneficial effects of physical exercise.

Kemirgenlerin Kafalarında Tanımlanmış İvmelenmeler Oluşturmak için Pasif Kafa Hareketinin Uygulanması

Exercise is widely recognized as effective for various diseases and physical disorders, including those related to brain dysfunction. However, molecular ...

A Mouse Model of Pulmonary Fibrosis Induced by Nasal Bleomycin Nebulization

Pulmonary fibrosis is characteristic of several human lung diseases that arise from various causes. Given that treatment options are fairly limited, mouse models continue to be an important tool for developing new anti-fibrotic strategies. In this study, intrapulmonary administration of bleomycin (BLM) is carried out by nasal nebulization to create a mouse model of pulmonary fibrosis that closely mimics clinical disease characteristics. C57BL/6 mice received BLM (7 U/mL, 30 min/day) by nasal nebulization for 3 consecutive days and were sacrificed on day 9, 16, or 23 to observe inflammatory and fibrotic changes in lung tissue. Nasal aerosolized BLM directly targeted the lungs, resulting in widespread and uniform lung inflammation and fibrosis. Thus, we successfully generated an experimental mouse model of typical human pulmonary fibrosis. This method could easily be used to study the effects of the administration of various nasal aerosols on lung pathophysiology and validate new anti-inflammatory and anti-fibrotic treatments.

<meta charset="utf8"></head><body>Nazal Bleomisin Nebulizasyonunun Neden Olduğu Pulmoner Fibrozisin Bir Fare Modeli

Various animal models of pulmonary fibrosis have been established using bleomycin to clarify the pathogenesis of pulmonary fibrosis and find new drug targets. However, most pulmonary fibrosis models targeting lung tissue have uneven drug administration. Here, we propose a model of uniform pulmonary fibrosis induced by nasal bleomycin nebulization.

Nazal Bleomisin Nebulizasyonunun Neden Olduğu Pulmoner Fibrozisin Bir Fare Modeli

Pulmonary fibrosis is characteristic of several human lung diseases that arise from various causes. Given that treatment options are fairly limited, mouse ...

Investigating Cardiac Metabolism in the Isolated Perfused Mouse Heart with Hyperpolarized [1-13C]Pyruvate and 13C/31P NMR Spectroscopy

Metabolism is the basis of important processes in cellular life. Characterizing how metabolic networks function in living tissues provides crucial information for understanding the mechanism of diseases and designing treatments. In this work, we describe procedures and methodologies for studying in-cell metabolic activity in a retrogradely perfused mouse heart in real-time. The heart was isolated in situ,&#160;in conjunction with cardiac arrest to minimize the myocardial ischemia and was perfused inside a nuclear magnetic resonance (NMR) spectrometer. While in the spectrometer and under continuous perfusion, hyperpolarized [1-13C]pyruvate was administered to the heart, and the subsequent hyperpolarized [1-13C]lactate and [13C]bicarbonate production rates served to determine, in real-time, the rates of lactate dehydrogenase and pyruvate dehydrogenase production. This metabolic activity of hyperpolarized [1-13C]pyruvate was quantified with NMR spectroscopy in a model free-manner using the product selective saturating-excitations acquisition approach. 31P spectroscopy was applied in between the hyperpolarized acquisitions to monitor the cardiac energetics and pH. This system is uniquely useful for studying metabolic activity in the healthy and diseased mouse heart.

Investigating Cardiac Metabolism in the Isolated Perfused Mouse Heart with Hyperpolarized [1-13C]Pyruvate and 13C/31P NMR Spectroscopy

We describe an experimental setup for administrating hyperpolarized 13C-labeled metabolites in continuous perfusion mode to an isolated perfused mouse heart. A dedicated 13C-NMR acquisition approach enabled the quantification of metabolic enzyme activity in real-time, and a multiparametric 31P-NMR analysis enabled the determination of the tissue ATP content and pH.

Hiperpolarize [1-13 C] Piruvat ve13 C/31P NMR Spektroskopisi ile İzole Perfüze Fare Kalbinde Kardiyak Metabolizmanın İncelenmesi