We acknowledge the UCLA Advanced Light Microscopy/Spectroscopy Laboratory and Leica Microsystems Center of Excellence at the California NanoSystems Institute (RRID:SCR_022789) for providing two-photon microscopy. AS was supported by ISF 604-21 from the Israel Science Foundation. JH was supported by R03DE030205 and R01DE030471 from the NIH/NIDCR. AS and JH were also supported by grant 2021007 from the United States-Israel Binational Science Foundation (BSF).

All mice were maintained in pathogen-free animal facilities at the University of California Los Angeles (UCLA) or the Hebrew University of Jerusalem (HUJI). All experiments involving mice were performed according to regulations and protocols approved by the respective Institutional Animal Care and Use Committee (IACUC) (ARC-2019-013; UCLA) or (MD-23-17184-3; HUJI). A general workflow of the experimental steps is shown in Figure 2A. See the Table of Materials for details related to all instruments, reagents, and materials used in this protocol.
1. Preparation of solutions and gels
<ol>
	<li>Dissection medium: Prepare fresh DMEM/F12 with 0.5% glucose and keep it warm at 37 &#176;C until needed in step 2.4. 
	NOTE: We use DMEM/F12 without phenol red to reduce autofluorescence during live imaging.</li>
	<li>1x Culture medium: Prepare fresh medium using 50% DMEM/F12, 50% rat serum, 1x glutamine substitute, 1x MEM Non-Essential Amino Acids, 1% glucose, 0.1 mg/mL L-ascorbic acid, and 0.5% penicillin-streptomycin. Keep it warm at 37 &#176;C until needed in step 5.5. This medium is used for culturing incisor explants during live imaging. 
	NOTE: The rat serum should be of high quality and specially prepared for whole tissue culturing by researchers or from a commercial source. In particular, the blood must be centrifuged (for 5 min at 1,200 &#215; g at room temperature) before clotting begins to form. After centrifugation, the resulting fibrin clot should be squeezed and discarded33.</li>
	<li>Culture gel: The purpose of the gel is to immobilize samples during imaging and is prepared fresh.
	<ol>
		<li>Make 2% gel in DMEM/F12 by dissolving 200 mg of low melting point agarose in 10 mL of DMEM/F12 using a microwave. Keep the 2% gel at 37 &#176;C.</li>
		<li>Make 2x culture medium (without DMEM/F12) by mixing 50% rat serum, 1x glutamine substitute, 1x MEM Non-Essential Amino Acids, 1% glucose, 0.1 mg/mL L-ascorbic acid, and 0.5% penicillin-streptomycin. Warm the 2x culture medium (without DMEM/F12) to 37 &#176;C.</li>
		<li>Make 1% culture gel by mixing equal volumes of 2% gel and 2x culture medium (without DMEM/F12). Keep the 1% culture gel at 37 &#176;C until needed in step 5.1. 
		NOTE: Make sure all solutions are warm before mixing to avoid gelling. We found 1% gel to be suitable for culturing the adult mouse incisor. Gel percentage should be empirically determined if other tissues are to be cultured.</li>
	</ol>
	</li>
</ol>
2. Extraction of the adult mouse mandibles
<ol>
	<li>Euthanize mice at the desired age using standard procedures approved by the IACUC. 
	NOTE: Here we use CO2&#160;asphyxiation followed by cervical dislocation. Regulations for animal euthanasia may vary in different regions. Researchers should obtain necessary institutional approval prior to performing experiments and ensure compliance with local animal care regulations.</li>
	<li>Disinfect the mouse using 70% ethanol.</li>
	<li>Decapitate the mouse and extract the left and right mandibles.
	<ol>
		<li>Lay the mouse on its belly, then use a #9 single-edge industrial razor blade to cut at the neck region and separate the mouse head from the rest of the body. 
		NOTE: If tissues from the pharynx or the neck region are to be collected as well, decapitation should not be performed, and one can directly proceed to step 2.3.2.</li>
		<li>Turn the mouse over so its ventral side is facing up and the mandibles are easily accessible.</li>
		<li>Secure the animal&#39;s head by holding it gently between the thumb and the index finger.</li>
		<li>Use the razor blade to make a mid-sagittal incision that cuts over the skin of the lower jaw from the lower lip toward the neckline. 
		NOTE: A #15 surgical blade can also be used to make the incision and subsequent dissections (steps 2.3.6-2.3.9).</li>
		<li>As the incision is made, spread open the cut skin using the thumb and the index finger to expose the muscles and jawbone underneath.</li>
		<li>Use the razor blade to sever the masseter muscles on the buccal side of the lower jaw, such that the outside of the left and right hemimandibles are now free from muscle attachments.</li>
		<li>Sever the mylohyoid muscles along the inner side of the mandible to remove muscle attachments there.</li>
		<li>Make another incision at the mandibular symphysis that connects the two hemimandibles. Once cut, the mandible will be separated into the left and right halves.</li>
		<li>Wedge the razor blade between the mandibular condyle and the temporal mandibular joint and carefully dissect out the hemimandible from the rest of the head. 
		NOTE: We found it helpful to simultaneously pull gently on the incisor while cutting. Care should be taken to avoid breaking the mandible and damaging the soft tissues within.</li>
	</ol>
	</li>
	<li>Immediately transfer dissected mandibles to a Petri dish with the prewarmed dissection media (step 1.1) and remove the remaining muscle tissues using a #15 surgical blade. 
	NOTE: Keeping samples in cold media slows down cell activities, resulting in delayed or even failed recoveries of certain cell activities, such as proliferation or cell movement, during live imaging.</li>
</ol>
3. Isolation of the whole mouse incisors
NOTE: Further isolation of the incisor is done under a bright field dissection microscope.
<ol>
	<li>Visually identify the oval region of the mandible that covers the incisor socket and houses the apical portion of the incisor.</li>
	<li>Position the mandible so the inner (lingual) surface is facing upwards.</li>
	<li>While holding the mandible in place with a pair of serrated forceps, generate a window at the oval region by shaving off the overlying membrane bone using a #15 surgical blade, in a direction that is from the condyle towards the molars. This exposes the soft tissue of the apical incisor on the inner surface.</li>
	<li>Turn over the mandible so the outer (buccal) surface is facing upwards.</li>
	<li>Generate a window at the oval region on the outer mandible as described in step 3.3&#160;and use the tip of the scalpel to pick away any remaining bone fragments at the edge. Ensure that the apical end of the tooth is visible from both sides. 
	NOTE: Avoid excessive pressure while shaving the bones to prevent damage to the underlying soft tissue.</li>
	<li>Systematically cut away the bones surrounding the incisor to isolate the entire tooth.
	<ol>
		<li>Make a clean cut at a plane that is immediately adjacent to the apical incisor to first remove the condylar process. 
		NOTE: Be careful not to cut into the incisor.</li>
		<li>Make a second cut just posterior to the 3rd molar, but dorsal to the incisor without damaging the tooth. This removes the bone that includes the coronoid process.</li>
		<li>Serially cut from the tip of the angular process towards the incisor to gradually remove the ventral mandible in a stepwise manner. 
		NOTE: The dental epithelium and associated periodontal tissues are often stuck to the bone and easily ripped off if a large portion of the ventral bone is cut at once. To separate the bone from the incisor soft tissue, one can insert the scalpel (or a pair of sharp non-serrated forceps) between the two tissues at the apical end of the incisor and delicately slide the instrument forward.</li>
		<li>Cut away the alveolar bone with molars and any remaining bones that are still attached to the incisor.</li>
		<li>The entire incisor is now isolated. Transfer the fully dissected incisor to a dish with clean warm dissection media (step 1.1).</li>
	</ol>
	</li>
	<li>Repeat steps 3.2-3.6&#160;to isolate additional incisors as required. 
	NOTE: The maxillary/upper incisors of the mouse also maintain adult stem cells and can be used to study tooth regeneration34. Dental researchers typically focus on the lower incisors because they are more accessible and can be more easily dissected than the upper incisors. Differences between mandibular and maxillary incisor progenitor cells remain to be determined.</li>
</ol>
4.&#160;Removal of periodontal tissues to expose the incisor epithelial cervical loop
<ol>
	<li>With the whole incisor lying down on its lingual side and while holding the tooth in place with a pair of serrated forceps, use a pair of #5 fine forceps to start tucking on the periodontal tissues covering the apical incisor and the cervical loop region.</li>
	<li>Carefully peel off the periodontal tissue from the apical bud, such that the lateral side of the cervical loop (or other regions of interest) becomes visible under the scope. 
	NOTE: Care should be taken to avoid damaging the dental epithelium or detaching it from the mesenchyme. If the incisor has fluorescence signals in the region of interest, and a fluorescent dissection microscope is available, fluorescence signals can be used to help distinguish between tissue types and aid dissections. It is important to efficiently carry out sections 2-4 so samples can be quickly transferred into the culture media and adequately maintained through explant culturing (see below).</li>
</ol>
5. Tissue embedding for explant culture
<ol>
	<li>Add 500 &#181;L of warm, unsolidified culture gel to a well in a 24-well plate and quickly transfer the dissected whole incisors to the well. Swirl the plate a few times to rinse the incisors.</li>
	<li>Add 400 &#181;L of warm, unsolidified culture gel to a culture dish and transfer the rinsed incisors to the dish. 
	NOTE: We use a commercially available culture dish that is compatible with the perfusion setup. See step 6.4 for alternative options.</li>
	<li>Orient the incisor to position the cervical loop region (or other regions of interest) at the center of the dish (Figure 2A,B) and adjust the tilt of the apical incisor for the desired imaging plane. 
	NOTE: Tissue orientation should be done quickly before complete gelation.</li>
	<li>After the gel is set, use a pair of fine forceps to remove any gel on top of the region of interest so it is not covered by gel.</li>
	<li>Add a sufficient volume of warm culture media by slowly pipetting it against the edge of the dish to just cover the sample (~150 &#181;L).</li>
	<li>Move the explant culture to a 37 &#176;C cell culture incubator to allow tissue settling and adjustment to the culture condition for 1 h.</li>
</ol>
6. Timelapse microscopy of incisor explants
NOTE: In this experiment, we used an upright microscope equipped with a 25x water-dipping objective that has a numerical aperture of 1. In general, a water dipping lens with a high numerical aperture is best suited for deep tissue imaging.
<ol>
	<li>Turn on the microscope and the two-photon laser.</li>
	<li>Secure the culture dish to the stage adapter and mount the perfusion adapter ring on top (Figure 2C).</li>
	<li>Connect the stage adapter to a temperature controller set to maintain the culture at 37 &#176;C.</li>
	<li>Connect the inlet and the outlet of the adapter ring to a micro-perfusion pump and begin perfusion of the culture medium, with the perfusion speed set at 20 on the pump. This generates a slow flow (~5 mL/h) of the culture medium over the top of the samples (Figure 2C). 
	NOTE: See the Table of Materials for details about the system to maintain the tissue in a stably controlled environment. Other systems can be used as well. A combination of a 37 &#176;C heating plate and a homemade perfusion culture dish (Supplemental Figure S1) with an inlet and an outlet connected to syringe pumps can also be used.</li>
	<li>Place an Atmospheric Control Barrier Ring (ACBR) on top of the adapter ring and lower the objective through the ACBR to just make contact with the culture medium (Figure 2D).</li>
	<li>Tune the laser wavelength to 920 nm to visualize both GFP and red fluorescence (e.g., tdTomato) signals.</li>
	<li>Locate samples through eyepieces and then on the microscope&#39;s software.</li>
	<li>Set up Z-stacks, multi-position imaging, and time intervals using the software. To follow this protocol, use a z-step size of 4 &#956;m and a time interval of 5 min for 14 h. 
	NOTE: We found that a time interval of longer than 5 min is often insufficient to capture smooth cell movements and divisions.</li>
	<li>Initiate the timelapse imaging. 
	NOTE: Sample position may shift during the first hour and further adjustments may be required.</li>
	<li>Save files for downstream data processing and analyses.</li>
</ol>

Live Tracking of Dental Cells via Two-Photon Microscopy

The authors have no conflicts of interest to disclose.

Live tissue imaging is an important technique that allows us to study the dynamic processes and behaviors of cells when they are maintained in their niche environment41. Ideally, live imaging is performed in vivo with high spatiotemporal resolution. However, in vivo imaging for mammalian organs can be challenging due to tissue inaccessibility, optical opaqueness, and difficulty in immobilizing the animal or the organ for a prolonged period42. Tissue explan...

Over the past two decades, the mouse incisor has emerged as an invaluable platform for investigating the principles of adult stem cell regulation and tooth regeneration1,2. The mouse incisor grows continuously and renews itself throughout the animal&#39;s life. It does so by maintaining both epithelial and mesenchymal stem cells, which can self-renew and differentiate into different cell types of the tooth1,2. While dental epithelial stem cells give rise to ameloblasts, which secrete the enamel matrix, dental mesenchymal stem cells give rise to odontoblasts, cementoblasts, and fibroblasts, which form dentin, cementum, and periodontal ligament, respectively3,4,5,6. This constant supply of new cells maintains tissue homeostasis and allows the replacement of old cells that are lost due to masticatory wear or injuries7,8. Elucidating the cellular and molecular mechanisms that regulate the maintenance and differentiation of dental stem cells is therefore central to understanding dental regeneration, an area of growing interest.
Anatomically, a large portion of the adult mouse incisor is encased in the jawbone. While the incisal edge of the tooth is exposed, the apical end of the incisor fits within a socket and is firmly attached to the surrounding bone through periodontal ligaments and connective tissues (Figure 1A,B). The incisor&#39;s apical end is also the growth region of the tooth and maintains dental stem and progenitor cells in both the epithelial layer and the mesenchymal pulp9,10,11,12,13. Specifically, dental epithelial stem cells are maintained at the bulbous end of the epithelium, known as the apical bud, also referred to as the labial cervical loop (Figure 1C). Similar to the intestinal epithelium and the epidermis, epithelial renewal in the incisor is primarily supported by actively cycling stem cells and their highly proliferative intermediate descendants, called transit-amplifying cells14,15,16,17, both residing in the inner part of the cervical loop. However, whether the incisor epithelium contains and utilizes quiescent stem cells during regeneration remains to be determined. In contrast, both active and quiescent dental mesenchymal stem cells have been identified in the apical pulp, and the quiescent stem cells function as a reserve population that becomes activated during injury repair13,18.
Many of the discoveries on the biology of the mouse incisor renewal and regeneration have resulted from histological investigations, in which samples are obtained at distinct temporal junctures, fixed, processed, and then sectioned into micron-thin slices along a particular plane. Through detailed analysis of histological sections from different mouse models that enable lineage tracing or genetic perturbations, scientists have identified the cell lineages of different progenitor populations, as well as the genetic and signaling pathways that control incisor homeostasis and injury repair19,20,21. However, the static two-dimensional (2D) images of non-vital cells in sections cannot capture the full spectrum of cellular behaviors and spatial organizations in living tissue, such as cell shape changes, movements, and cellular kinetics. Detecting and measuring these rapid cellular changes, which occur at a timescale that is unresolvable through tissue sectioning, require a different strategy. Moreover, acquiring such information is also critical for understanding how dental cells interact with each other, react to different signaling stimuli, and self-organize to maintain tissue structures and functions.
The advent of four-dimensional (4D) deep tissue imaging using two-photon microscopy22, a technology that integrates three spatial dimensions with temporal resolution, overcomes the inherent limitations of histological analysis by enabling spatiotemporal examination of cultured tissue explants, organoids, or even tissues in situ23,24,25,26. For instance, 4D live imaging of the developing tooth epithelium has unveiled the spatiotemporal patterns of cell divisions and migrations that coordinate tissue growth, signaling center formation, and dental epithelial morphogenesis27,28,29,30,31,32. In the adult mouse incisor, 4D imaging has been recently adapted to study cellular behaviors during dental epithelial injury repair. Live imaging revealed that stratum intermedium cells in the suprabasal layer can be directly converted into ameloblasts in the basal layer to regenerate the damaged epithelium, challenging the traditional paradigm of epithelial injury repair15.
Here, we describe the dissection, culturing, and imaging of the adult mouse incisor, focusing on epithelial cells in the labial cervical loop (Figure 1). This technique preserves dental cell vitality for more than 12 h and permits live tracking of fluorescently labeled cells at single-cell resolution. This approach allows investigation of cell motion and migration as well as dynamic changes in cell shape and division orientation under normal culture conditions, or in responses to genetic, physical, and chemical perturbations.

<table><tbody><tr><td>24 well, flat bottom tissue culture plate</td><td>Olympus plastics</td><td>25-107</td><td></td></tr><tr><td>25x HC IRAPO motCORR water dipping objective</td><td>Leica</td><td>11507704</td><td></td></tr><tr><td>Ascorbic acid (Vitamin C)</td><td>Acros Organics</td><td>352685000</td><td></td></tr><tr><td>D-(+)-Glucose bioxtra&amp;nbsp;</td><td>Sigma Aldrich</td><td>G7528</td><td></td></tr><tr><td>Delta T system&amp;nbsp;</td><td>Bioptechs</td><td>0420-4</td><td>Including temperature control, culture dishes, and perfusion setup</td></tr><tr><td>Dissection microscope- LEICA S9E</td><td>Leica</td><td>LED300 SLI</td><td></td></tr><tr><td>DMEM/F12</td><td>Thermo Scientific</td><td>11039047</td><td>Basal media without phenol red</td></tr><tr><td>Feather surgical blade (#15)</td><td>Feather</td><td>72044-15</td><td></td></tr><tr><td>Fine forceps</td><td>F.S.T</td><td>11252-23</td><td></td></tr><tr><td>Glutamax&amp;nbsp;</td><td>Thermo Scientific</td><td>35050-061</td><td>Glutamine substitute</td></tr><tr><td>Leica SP8-DIVE equipped with a 25X HC IRAPO motCORR water dipping objective&amp;nbsp;</td><td>Leica</td><td>n/a</td><td></td></tr><tr><td>low-melting agarose</td><td>NuSieve</td><td>50080</td><td></td></tr><tr><td>non-essential amino acids (100x)</td><td>Thermo Scientific</td><td>11140-050</td><td></td></tr><tr><td>penicillin&amp;ndash;streptomycin</td><td>Thermo Scientific</td><td>15140122</td><td>10,000 U/mL&amp;nbsp;</td></tr><tr><td>Petri dish</td><td>Gen Clone</td><td>32-107G</td><td>90 mm&amp;nbsp;</td></tr><tr><td>Rat serum</td><td>Valley Biomedical</td><td>AS3061SC</td><td>Processed for live imaging</td></tr><tr><td>Razor blade #9</td><td>VWR</td><td>55411-050</td><td></td></tr><tr><td>Scalpel handle</td><td>F.S.T</td><td>10003-12</td><td></td></tr><tr><td>Scissors</td><td>F.S.T</td><td>37133</td><td></td></tr><tr><td>serrated forceps</td><td>F.S.T</td><td>11000-13</td><td></td></tr><tr><td>spring scissors</td><td>F.S.T</td><td>91500-09</td><td></td></tr></tbody></table>

using ex vivo live imaging to investigate cell divisions and movements during mouse dental renewal

The continuously growing mouse incisor is emerging as a highly tractable model system to investigate the regulation of adult epithelial and mesenchymal stem cells and tooth regeneration. These progenitor populations actively divide, move, and differentiate to maintain tissue homeostasis and regenerate lost cells in a responsive manner. However, traditional analyses using fixed tissue sections could not capture the dynamic processes of cellular movements and interactions, limiting our ability to study their regulations. This paper describes a protocol to maintain whole mouse incisors in an explant culture system and live-track dental epithelial cells using multiphoton timelapse microscopy. This technique adds to our existing toolbox for dental research and allows investigators to acquire spatiotemporal information on cell behaviors and organizations in a living tissue. We anticipate that this methodology will help researchers further explore mechanisms that control the dynamic cellular processes taking place during both dental renewal and regeneration.

The apical region of the adult mouse incisor is encased within the mandible (Figure 1) and hence, not directly accessible for visualizing and live-tracking the progenitor cells residing within the growth region. Therefore, we have developed a method to extract the whole incisor from the jawbone and maintain it in an explant culture system for two-photon timelapse microscopy (Figure 2). Here we describe representative results that capture the dynamic process of c...

Watch this Scientific Journal Video about Understanding Dynamic Cellular Behaviors in Adult Mouse Dental Tissue Renewal and Repairment at JoVE.com

Author Spotlight: Understanding Dynamic Cellular Behaviors in Adult Mouse Dental Tissue Renewal and Repairment 

Ex vivo live imaging is a powerful technique for studying the dynamic processes of cellular movements and interactions in living tissues. Here, we present a protocol that implements two-photon microscopy to live track dental epithelial cells in cultured whole adult mouse incisors.

Using Ex Vivo Live Imaging to Investigate Cell Divisions and Movements During Mouse Dental Renewal

The continuously growing mouse incisor is emerging as a highly tractable model system to investigate the regulation of adult epithelial and mesenchymal ...

using-ex-vivo-live-imaging-to-investigate-cell-divisions-movements

Research

JoVE Journal

Developmental Biology

1.3K Views.  University of California Los Angeles. Om de juiste cellulaire organisatie te behouden tijdens orgaanvernieuwing, beweegt weefsel dat de cellen nadert en deelt het specifieke oriëntaties en patronen. Het is daarom belangrijk om de regels en voorschriften van deze dynamische cellulaire gebeurtenissen te begrijpen. De muizensnijtand is een opkomend model voor onderzoek naar volwassen stamcellen en er zijn veel genetische hulpmiddelen beschikbaar in het systeem om genfuncties te bestuderen en cellen te labelen voor het traceren van ...

Video: Auteur in de schijnwerpers: dynamisch cellulair gedrag begrijpen bij vernieuwing en herstel van tandweefsel bij volwassen muizen 

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a rapid and high-throughput manner. S2 cells have proven amenable to both fixed- and live-cell imaging applications. Notably, live-cell imaging can yield valuable information about how loss or knockdown of a gene can affect the kinetics and dynamics of key events during cell division, including mitotic spindle assembly, chromosome congression, and segregation, as well as overall cell cycle timing. Here we utilize S2 cells stably transfected with fluorescently tagged mCherry:&#945;-tubulin to mark the mitotic spindle and GFP:CENP-A (referred to as &#8216;CID&#8217; gene in Drosophila) to mark the centromere to analyze the effects of key mitotic genes on the timing of cell divisions, from prophase (specifically at Nuclear Envelope Breakdown; NEBD) to the onset of anaphase. This imaging protocol also allows for the visualization of the spindle microtubule and chromosome dynamics throughout mitosis. Herein, we aim to provide a simple yet comprehensive protocol that will allow readers to easily adapt S2 cells for live imaging experiments. Results obtained from such experiments should expand our understanding of genes involved in the cell division by defining their role in several simultaneous and dynamic events. Observations made in this cell culture system can be validated and further investigated in vivo using the impressive toolkit of genetic approaches in flies.

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and quantified.

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a ...

Genetics

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of existing mouse lines for genetic cell lineage tracing: a tamoxifen-inducible Cre line and a Cre reporter line expressing dsRed upon Cre-mediated recombination. By using a relatively low level of tamoxifen, we were able to induce recombination in a small number of cells, permitting us to follow individual cell divisions. Additionally, we observed the transcriptional response to Sonic Hedgehog (Shh) signaling using an Olig2-eGFP transgenic line 1-3 and we monitored formation of cilia by infecting the cultured slice with virus expressing the cilia marker, Sstr3-GFP 4. In order to image the neuroepithelium, we harvested embryos at E8.5, isolated the neural tube, mounted the neural slice in proper culturing conditions into the imaging chamber and performed time-lapse confocal imaging. Our ex vivo live imaging method enables us to trace single cell divisions to assess the relative timing of primary cilia formation and Shh response in a physiologically relevant manner. This method can be easily adapted using distinct fluorescent markers and provides the field the tools with which to monitor cell behavior in situ and in real time.

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

Here we develop the tools necessary for ex vivo live imaging to trace single cell divisions in the mouse E8.5 neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of ...

Neuroscience

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Intravital Imaging of Neutrophil Priming Using IL-1&#946; Promoter-driven DsRed Reporter Mice

Neutrophils are the most abundant leukocytes in human blood circulation and are quickly recruited to inflammatory sites. Priming is a critical event that enhances the phagocytic functionality of neutrophils. Although extensive studies have unveiled the existence and importance of neutrophil priming during infection and injury, means of visualizing this process in vivo have been unavailable. The protocol provided enables monitoring of the dynamic process of neutrophil priming in living animals by combining three methodologies: 1) DsRed reporter signal &#8212; used as a measure of priming 2) in vivo neutrophil labeling &#8212; achieved by injection of fluorescence-conjugated anti-lymphocyte antigen 6G (Ly6G) monoclonal antibody (mAb) and 3) intravital confocal imaging. Several critical steps are involved in this protocol: oxazolone-induced mouse ear skin inflammation, appropriate sedation of animals, repeated injections of anti-Ly6G mAb, and prevention of focus drift during imaging. Although a few limitations have been observed, such as the limit of continuous imaging time (~ 8 hr) in one mouse and the leakage of fluorescein isothiocyanate-dextran from blood vessels in the inflammatory state, this protocol provides a fundamental framework for intravital imaging of primed neutrophil behavior and function, which can easily be expanded to examination of other immune cells in mouse inflammation models.

This current protocol employs fluorescent reporters, in vivo labeling, and intravital imaging techniques to enable monitoring of the dynamic process of neutrophil priming in living animals.

Neutrophils are the most abundant leukocytes in human blood circulation and are quickly recruited to inflammatory sites. Priming is a critical event that ...

Immunology and Infection

Isolation and Time-Lapse Imaging of Primary Mouse Embryonic Palatal Mesenchyme Cells to Analyze Collective Movement Attributes

Development of the palate is a dynamic process, which involves vertical growth of bilateral palatal shelves next to the tongue followed by elevation and fusion above the tongue. Defects in this process lead to cleft palate, a common birth defect. Recent studies have shown that palatal shelf elevation involves a remodeling process that transforms the orientation of the shelf from a vertical to a horizontal one. The role of the palatal shelf mesenchymal cells in this dynamic remodeling has been difficult to study. Time-lapse-imaging-based quantitative analysis has been recently used to show that primary mouse embryonic palatal mesenchymal (MEPM) cells can self-organize into a collective movement. Quantitative analyses could identify differences in mutant MEPM cells from a mouse model with palate elevation defects. This paper describes methods to isolate and culture MEPM cells from E13.5 embryos-specifically for time-lapse imaging-and to determine various cellular attributes of collective movement, including measures for stream formation, shape alignment, and persistence of direction. It posits that MEPM cells can serve as a proxy model for studying the role of palatal shelf mesenchyme during the dynamic process of elevation. These quantitative methods will allow investigators in the craniofacial field to assess and compare collective movement attributes in control and mutant cells, which will augment the understanding of mesenchymal remodeling during palatal shelf elevation. Furthermore, MEPM cells provide a rare mesenchymal cell model for investigation of collective cell movement in general.

We present a protocol for isolation and culture of primary mouse embryonic palatal mesenchymal cells for time-lapse imaging of two-dimensional (2D) growth and wound-repair assays. We also provide the methodology for analysis of the time-lapse imaging data to determine cell-stream formation and directional motility.

Development of the palate is a dynamic process, which involves vertical growth of bilateral palatal shelves next to the tongue followed by elevation and ...

Live Imaging Followed by Single Cell Tracking to Monitor Cell Biology and the Lineage Progression of Multiple Neural Populations

Understanding the mechanisms that control critical biological events of neural cell populations, such as proliferation, differentiation, or cell fate decisions, will be crucial to design therapeutic strategies for many diseases affecting the nervous system. Current methods to track cell populations rely on their final outcomes in still images and they generally fail to provide sufficient temporal resolution to identify behavioral features in single cells. Moreover, variations in cell death, behavioral heterogeneity within a cell population, dilution, spreading, or the low efficiency of the markers used to analyze cells are all important handicaps that will lead to incomplete or incorrect read-outs of the results. Conversely, performing live imaging and single cell tracking under appropriate conditions represents a powerful tool to monitor each of these events. Here, a time-lapse video-microscopy protocol, followed by post-processing, is described to track neural populations with single cell resolution, employing specific software. The methods described enable researchers to address essential questions regarding the cell biology and lineage progression of distinct neural populations.

A robust protocol to monitor neural populations by time-lapse video-microscopy followed by software-based post-processing is described. This method represents a powerful tool to identify biological events in a selected population during live imaging experiments.

Understanding the mechanisms that control critical biological events of neural cell populations, such as proliferation, differentiation, or cell fate ...

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, ...

Biology

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 ...

3D Imaging of PDL Collagen Fibers during Orthodontic Tooth Movement in Mandibular Murine Model

Orthodontic tooth movement is a complex biological process of altered soft and hard tissue remodeling as a result of external forces. In order to understand these complex remodeling processes, it is critical to study the tooth and periodontal tissues within their 3D context and therefore minimize any sectioning and tissue artefacts. Mouse models are often utilized in developmental and structural biology, as well as in biomechanics due to their small size, high metabolic rate, genetics and ease of handling. In principle this also makes them excellent models for dental related studies. However, a major impediment is their small tooth size, the molars in particular. This paper is aimed at providing a step by step protocol for generating orthodontic tooth movement and two methods for 3D imaging of the periodontal ligament fibrous component of a mouse mandibular molar. The first method presented is based on a micro-CT setup enabling phase enhancement&#160;imaging of fresh collagen tissues. The second method is a bone clearing method using ethyl cinnamate that enables imaging through the bone without sectioning and preserves endogenous fluorescence. Combining this clearing method with reporter mice like Flk1-Cre;TdTomato provided a first of its kind opportunity to image the 3D vasculature in the PDL and alveolar bone.

We present a protocol for generating orthodontic tooth movement in mice and methods for 3D visualization of the collagen fibers and blood vessels of periodontal ligament without sectioning.

Using Ex Vivo Live Imaging to Investigate Cell Divisions and Movements During Mouse Dental Renewal

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