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

Summary

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.

Abstract

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.

Introduction

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 regeneration^1,2. The mouse incisor grows continuously and renews itself throughout the animal'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 tooth^1,2. 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, respectively^3,4,5,6. This constant supply of new cells maintains tissue homeostasis and allows the replacement of old cells that are lost due to masticatory wear or injuries^7,8. 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'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 pulp^{9,10,11,12,13}. 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 cells^14,15,16,17, 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 repair^13,18.

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 repair^19,20,21. 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 situ^23,24,25,26. 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 morphogenesis^{27,28,29,30,31,32}. 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.

Protocol

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

1. Dissection medium: Prepare fresh DMEM/F12 with 0.5% glucose and keep it warm at 37 °C until needed in step 2.4.
   NOTE: We use DMEM/F12 without phenol red to reduce autofluorescence during live imaging.
2. 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 °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 × g at room temperature) before clotting begins to form. After centrifugation, the resulting fibrin clot should be squeezed and discarded³³.
3. Culture gel: The purpose of the gel is to immobilize samples during imaging and is prepared fresh.
   1. 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 °C.
   2. 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 °C.
   3. 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 °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.

2. Extraction of the adult mouse mandibles

1. Euthanize mice at the desired age using standard procedures approved by the IACUC.
   NOTE: Here we use CO₂ 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.
2. Disinfect the mouse using 70% ethanol.
3. Decapitate the mouse and extract the left and right mandibles.
   1. 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.
   2. Turn the mouse over so its ventral side is facing up and the mandibles are easily accessible.
   3. Secure the animal's head by holding it gently between the thumb and the index finger.
   4. 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).
   5. As the incision is made, spread open the cut skin using the thumb and the index finger to expose the muscles and jawbone underneath.
   6. 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.
   7. Sever the mylohyoid muscles along the inner side of the mandible to remove muscle attachments there.
   8. 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.
   9. 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.
4. 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.

3. Isolation of the whole mouse incisors

NOTE: Further isolation of the incisor is done under a bright field dissection microscope.

1. Visually identify the oval region of the mandible that covers the incisor socket and houses the apical portion of the incisor.
2. Position the mandible so the inner (lingual) surface is facing upwards.
3. 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.
4. Turn over the mandible so the outer (buccal) surface is facing upwards.
5. Generate a window at the oval region on the outer mandible as described in step 3.3 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.
6. Systematically cut away the bones surrounding the incisor to isolate the entire tooth.
   1. 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.
   2. 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.
   3. 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.
   4. Cut away the alveolar bone with molars and any remaining bones that are still attached to the incisor.
   5. The entire incisor is now isolated. Transfer the fully dissected incisor to a dish with clean warm dissection media (step 1.1).
7. Repeat steps 3.2-3.6 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.

4. Removal of periodontal tissues to expose the incisor epithelial cervical loop

1. 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.
2. 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).

5. Tissue embedding for explant culture

1. Add 500 µL 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.
2. Add 400 µL 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.
3. 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.
4. 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.
5. Add a sufficient volume of warm culture media by slowly pipetting it against the edge of the dish to just cover the sample (~150 µL).
6. Move the explant culture to a 37 °C cell culture incubator to allow tissue settling and adjustment to the culture condition for 1 h.

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.

1. Turn on the microscope and the two-photon laser.
2. Secure the culture dish to the stage adapter and mount the perfusion adapter ring on top (Figure 2C).
3. Connect the stage adapter to a temperature controller set to maintain the culture at 37 °C.
4. 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 °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.
5. 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).
6. Tune the laser wavelength to 920 nm to visualize both GFP and red fluorescence (e.g., tdTomato) signals.
7. Locate samples through eyepieces and then on the microscope's software.
8. Set up Z-stacks, multi-position imaging, and time intervals using the software. To follow this protocol, use a z-step size of 4 μm 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.
9. Initiate the timelapse imaging.
   NOTE: Sample position may shift during the first hour and further adjustments may be required.
10. Save files for downstream data processing and analyses.

Results

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...

Discussion

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...

Materials

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