J.K. was supported by the Grant Agency of Masaryk University (MUNI/H/1615/2018) and by funds from the Faculty of Medicine MU to junior researcher. J.L. was supported by the Grant Agency of Masaryk University, (MUNI/IGA/1532/2020) and is a Brno Ph.D. Talent Scholarship Holder - Funded by the Brno City Municipality. T.B. was supported by the Austrian Science Fund (Lise Meitner grant: M2688-B28). We thank to Lydie Izakovicova Holla and Veronika Kovar Matejova for their help with the obtaining of human teeth. Finally, we thank Radek Fedr and Karel Soucek for their kind assistance with FACS sorting.

All animal experiments were performed according to the International and local regulations and approved by the Ministry of Education, youth and sports, Czech Republic (MSMT-8360/2019-2; MSMT-9231/2020-2; MSMT-272/2020-3). This protocol was tested with both male and female wildtype C57BL/6 and CD-1 mice and with genetically modified Sox10::iCreERT2 mice17 (combined with various reporter systems) on a C57BL/6 background. Experiments with human samples were performed with the approval of the Committees for Ethics of the Medical Faculty, Masaryk University Brno &#38; St. Anne&#180;s Faculty Hospital in Brno, Czech republic.
1. Experimental set-up and preparation of solutions
<ol>
	<li>Instrument set-up
	<ol>
		<li>Cool down the centrifuge to 4 &#176;C.</li>
		<li>Heat the incubation chamber to 37 &#176;C.</li>
		<li>Start the Fluorescence-activated cell sorting (FACS) machine and set all the temperatures of the sorter (including collection tube holder) to 4 &#176;C. Perform the instrument quality control, set the drop delay.</li>
		<li>Set the preliminary gating strategy and perform the test sorting. 
		NOTE: When using FACS, use the 100 &#181;m nozzle.</li>
	</ol>
	</li>
	<li>Prepare the solutions (steps 1.2.1-1.2.3).
	<ol>
		<li>Wash solution: Prepare fresh 2% FBS (Fetal Bovine Serum) in HBSS (Hanks&#39; Balanced Salt Solution).</li>
		<li>Digestion mixture: Prepare fresh collagenase P (3 U/mL) fully dissolved in HBSS.</li>
		<li>Optional: Prepare fresh HBSS + BSA (0.04%) and chill analytical grade methanol in a -20 &#176;C freezer for storing single-cell suspension at -80 &#176;C. 
		&#8203;NOTE: Step 1 needs to be performed before the start of the experiment. The composition of the utilized solutions is summarized in Supplementary Table 1.</li>
	</ol>
	</li>
</ol>
2. Preparation of experimental animal/s and human tooth
<ol>
	<li>Prepare the experimental animals (mouse).
	<ol>
		<li>Euthanize the mouse according to the local regulations; e.g. by anaesthetics overdose as described previously1. 
		CAUTION: Regulations for humane euthanizing of experimental animals varies locally. Always follow valid local regulations.</li>
		<li>Immediately proceed to the tissue dissection step (step 3). 
		NOTE: If the tissue from the experimental animals cannot be dissected immediately (e.g., because of transfer from animal housing facility), place the experimental animals on ice and perform tissue dissection as soon as possible. To obtain more cells from mouse molar pulps, use younger (6 weeks and less) animals. With increasing age, the size of dental pulp decreases. Mouse incisors mostly keep their structure with increasing age so animals of various ages can be used for tissue dissection.</li>
	</ol>
	</li>
	<li>Prepare the human tooth 
	NOTE: Human teeth were extracted for a clinically relevant reason. Every diagnosis was treated individually, and an experienced dental surgeon always performed the tooth extraction.
	<ol>
		<li>Put the freshly extracted tooth immediately into a 50 mL tube with ice-cold HBSS and keep the tube on ice until further processing. 
		NOTE: Using retained wisdom teeth from patients until age 25-30 is recommended for the highest cell yield.</li>
	</ol>
	</li>
</ol>
3. Tissue dissection
<ol>
	<li>Hold the experimental animal behind its head, looking on the ventral aspect of its head so that the tail points away.</li>
	<li>Using small, sharp scissors, quickly remove the skin from the mandible to expose the mandibular arch, the soft tissue between each half of the mandibles, and the adjacent facial muscles.
	<ol>
		<li>Make a deep cut from each side of the mandible; firstly, through m. masseter along the buccal side of the mandible up to the temporomandibular joint, and then along the inner part of each half of the mandible through the base of the oral cavity (see Supplementary Figure 1).</li>
	</ol>
	</li>
	<li>Cut all the muscles and ligaments along the mandible up to the temporomandibular joint from both outside and inside of the oral cavity. 
	NOTE: Avoid cutting bones. This might damage the most apical part of the incisor.</li>
	<li>Grasp the mandible using bent tip tweezers and remove it. Then, split the dissected mandible into two halves with scissors by cutting through mandibular symphysis (see Supplementary Figure 1).</li>
	<li>Use an industrial low lint wipe to chafe the remaining soft tissue from each half of the mandible. After both parts of the mandible are cleaned, place them into a pre-prepared Petri dish with ice-cold HBSS. 
	NOTE: From this point forward, work on ice. Further dissection of mouse incisors and molars is performed under a stereomicroscope with a black background.</li>
	<li>Mouse mandibular incisors
	<ol>
		<li>For the dissection of mandibular incisors, remove the alveolar ridge with all three molars and transversally crack the mandibular arch in the place corresponding to the position between the first and second molar. 
		NOTE: A sharp scalpel blade no. 11 and tweezers are used to perform this step (see Table of Materials).</li>
		<li>Carefully pull the incisor out of the rest of the dental socket. 
		NOTE: If successful, the extracted incisor will contain intact apical parts, including epithelial tissue with complete cervical loops.</li>
		<li>If needed, remove the remaining fragments of bone still attached to the incisor with tweezers and a scalpel.</li>
		<li>Place the dissected incisors into fresh, ice-cold HBSS and dissect the tissue of interest: cervical loop, dental pulp, or other parts of the tooth.</li>
		<li>Place the dissected soft tissue into a droplet of fresh, ice-cold HBSS in the middle of a 10 cm Petri dish. Keep on ice.</li>
	</ol>
	</li>
	<li>Mouse mandibular molars
	<ol>
		<li>For dissecting mandibular molars, completely remove the alveolar ridge from the rest of the mandible using a scalpel blade.</li>
		<li>Move the dissected alveolar crest into fresh, ice-cold HBSS in a Petri dish and carefully remove all the remaining fragments of alveolar bones attached to the roots.</li>
		<li>Place the dissected molars without alveolar bone into fresh, ice-cold HBSS. To expose the pulp, start to remove parts of the dentin from the apical side using fine, sharp tip tweezers until you reach the pulp cavity.</li>
		<li>Once the pulp cavity is reached, carefully dissect dental pulp using a pair of sharp tip tweezers and place the soft tissue of the dental pulp into a droplet of fresh, ice-cold HBSS in the middle of a 10 cm Petri dish kept on ice. 
		NOTE: Since mouse molar pulps are extremely small, adapt magnification on the stereomicroscope accordingly.</li>
	</ol>
	</li>
	<li>Human tooth
	<ol>
		<li>Wash human tooth once again in ice-cold HBSS to remove the remaining blood.</li>
		<li>Place the tooth into three thick-walled sterile plastic bags and use a cast iron benchtop engineer&#39;s vise to crack the tooth. Use vise jaws with a flat surface to avoid penetrating the bags.</li>
		<li>Slowly tighten the vise until you hear the tooth cracking; remove it from the bags and place it into fresh, ice-cold HBSS in a Petri dish.</li>
		<li>Using two tweezers, take out the dental pulp, clean it from all the remnants of hard tissue and place it into one droplet of ice-cold HBSS in the middle of a 10 cm Petri dish kept on ice. 
		&#8203;CAUTION: Human tissue might potentially be infectious. When working with human tissue, use protective equipment to avoid direct contact with the tissue.</li>
	</ol>
	</li>
</ol>
4. Preparation of single-cell suspension
<ol>
	<li>Prepare a 15 mL tube with 2.5 mL of digestion mixture composed of Collagenase P (3 U/mL) fully dissolved in HBSS. Keep the solution on ice until use. 
	NOTE: Always use freshly prepared Collagenase P; do not freeze the aliquots. Collagenase P activity varies from batch-to-batch. Check the activity of your batch before diluting. Collagenase P is activated by calcium, whose amount is already sufficient in the lyophilized powder. Therefore, it is unnecessary to enrich the enzyme mix with calcium. Collagenase P is not inactivated by FBS but can be inactivated by chelating agents (e.g., Ethylenediaminetetraacetic - EDTA). Stopping the dissociation process is ensured by diluting the Collagenase P in Wash solution (2% FBS in HBSS). It has been shown that adding FBS increases cell viability and the final number of cells after sorting18.</li>
	<li>Using a round-shaped scalpel blade no. 10, cut the tissue in all directions into the smallest possible pieces. Reaggregate the tissue pieces in the middle of the droplet and repeatedly cut the tissue aggregates using a round-shaped (No. 10) scalpel blade. Repeat this process several times until the material is sufficiently minced.</li>
	<li>Transfer the shredded tissue pieces using a 1 mL pipette tip into the prepared digestion mixture. 
	NOTE: In the case of a larger number of animals being processed at once, split the tissue into several tubes with Collagenase P. The maximum amount per tube are pulps obtained from approximately 10 mouse incisors. In the case of molars, where pulps are much smaller, the number of processed pulps can be increased adequately. When using human teeth, use a maximum of 2-3 pulps per tube, depending on the pulp size.</li>
	<li>Place the tube into a 37 &#176;C preheated incubator with a shaker. Tilt the tube inside the shaker to an angle of 60&#176; and set the speed to 150-200 rpm to ensure constant suspension movements inside the tube.</li>
	<li>Vigorously triturate the suspension every 3-4 min with a 1 mL pipette tip to disintegrate all the clumps. 
	NOTE: With time, the clumps will become smaller and softer until they almost disappear.</li>
	<li>In total, incubate for 15-20 min. At the end of the incubation, triturate for the last time, and then slowly add ice cold Wash solution to a final volume of 12 mL.</li>
	<li>Remove the remaining clumps or pieces of calcified tissue by filtering the suspension using a 50 &#181;m cell strainer.</li>
	<li>Take 10-20 &#181;L of the filtered cell suspension and count the cells during centrifugation using a cell counting chamber (Hemocytometer). Centrifuge for 5 min at 300 x g at 4 &#176;C. Remove the supernatant using a 10 mL serological pipette. 
	NOTE: If the number of cells is limited that cannot be counted in diluted suspension, skip the cell counting and use all the cells for further processing.</li>
	<li>Optional: Proceed for fixation and storage21.
	<ol>
		<li>Re-suspend the pellets (up to 107 cells) in 100 uL of HBSS + BSA (0.04%).</li>
		<li>Add 400 &#181;L of chilled&#160;methanol and mix slowly.</li>
		<li>Incubate the cell suspension for 15 min on ice.</li>
		<li>Store at -80 &#176;C until processed (no longer than 1 month).</li>
	</ol>
	</li>
	<li>Resuspend the pellet in the Wash solution. Aim for 700-1200 cells/&#181;L of the Wash solution.</li>
	<li>Keep the tube on ice until further processing. 
	NOTE: If necessary, perform the fixation and storage at -80 &#176;C. However, immediate processing of the cell suspension for scRNA-seq is recommended. Further steps will depend based on the single-cell RNA seq protocol. When the scRNA-seq protocol uses a microfluidic system (some sc-RNA-seq companies do), load the cells based on the manufacturer&#39;s guidelines either directly or after cell sorting.</li>
	<li>For FACS, proceed to step 5.</li>
</ol>
5. Fluorescence-activated cell sorting (FACS)
<ol>
	<li>Before sorting, prepare the cell sorting instrument.
	<ol>
		<li>Cool down the whole system to 4 &#176;C, perform the instrument quality control, set the drop delay and the voltage of the deflection plates. Put collection tubes in the collection tube holder. 
		NOTE: To avoid additional centrifugation steps resulting in an inevitable cell loss, sort cells into a microtube (1.5 mL) with a small amount (25-50 &#181;L) of the Wash solution.</li>
	</ol>
	</li>
	<li>Optional: perform viability staining before FACS.
	<ol>
		<li>Add Propidium iodide into cell suspension before FACS to a final concentration 0.5 &#181;g/mL and incubate for 15 min at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Load the sample into the cell sorter. Set a strict gating strategy to remove cell debris, doublets, and dead cells. Sort the cells into a prepared tube or multi-well plates. An example of a gating strategy is shown in Figure 2. 
	NOTE: To minimize manipulation steps and accelerate the protocol, applying a strict gating strategy is recommended (suggested in Figure 2) rather than using a viability staining. To separate the immune cells from the isolated population, CD45 antibody staining can be performed. To perform this, resuspend cell suspension in 100 &#181;L of staining solution (PBS + 2% FBS + anti-CD45-APC conjugated antibody 1/100 dilution). Incubate for 15 min on ice protected from light and perform FACS directly.</li>
</ol>

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Authors declare no conflicts of interest.

Studying teeth and bones on the cellular or molecular level is generally challenging since cells forming these tissues are surrounded by different kinds of hard matrices19. One of the main goals for performing single-cell RNA-seq on dental tissue is the need to obtain cells of interest fast and without any artificial changes in their transcriptomes. To accomplish this, a highly efficient protocol suitable for isolating cells from mouse and human tooth pulps was developed, which allows for quick generation of single-cell suspensions for all transcriptomic applications. This was ensured by fast tissue isolation, minimizing the steps of tissue and cell manipulations, and streamlining the mechanical and enzymatic digestion.
The most critical steps of this protocol are fast tissue processing and adequate single-cell suspension preparation8,9. A manual approach is used to obtain dental pulps without utilizing a dental drill or other heat-generating devices. Overheating may cause an artificial expression of heat shock proteins and other genes, ultimately leading to the analyzed gene expression patterns being unrepresentative of the original tissue20. Manual tissue harvesting may be a challenging step that will likely need some training beforehand. The pulp is then cut into small pieces and enzymatically digested at 37 &#176;C. Except for the 15-20 min of enzymatic digestion, the whole protocol is performed at 4 &#176;C. The tissue processing and especially the enzymatic digestion were minimized to the shortest possible time since more prolonged incubation at 37 &#176;C can cause changes in gene expression patterns10. Mechanical removing of the dentin is recommended before enzymatic digestion. Dentin and the pulp-attached predentin contain a large amount of collagen, and its excessive presence might decrease the effectiveness of the digesting solution. After being removed from the body (or death of organisms), it was shown that cells start to modify their gene expression patterns&#160;quickly12. Therefore, cell isolation and processing should be carried out as fast as possible. The current protocol reduces the processing time to 35-45 min from isolating the tissue (euthanizing animal) to preparing single-cell suspension.
One alternative modification of this technique is cell preservation for later use. This is achieved by methanol fixation. Methanol-fixed cell suspension can be stored for up to 1 month at -80 &#176;C, as described in the protocol21. However, whenever possible, perform scRNA-seq directly, since it was shown that the single-cell data from methanol-fixed single-cell suspensions might suffer from increased expression of stress-related genes and contamination with ambient RNA22. This step might need additional modification according to the manufacturer&#39;s protocols.
Before the first application of this protocol, performing several validation steps are recommended to test the technique. From our experience, we suggest testing the aforementioned critical steps of the protocol. Additionally, we suggest testing the effectiveness of the collagenase P solution and testing the handling of the tissue dissociation step. Specifically, around the first 5 min after the initiation of collagenase P incubation, the pieces of tissue should aggregate together. This is a common situation. Aggregates are disintegrated every 3-4 min using a 1 mL pipette, and with increasing time, they should become smaller until barely visible.
Furthermore, it is recommended to perform cell counting in a cell counting chamber before centrifugation and before and after filtering to detect possible cell losses due to suboptimal supernatant removal. If the final single-cell suspension needs to be purified, FACS can be used. Cell sorting enables not only to remove debris or dead cells but importantly enables to enrich final suspension with fluorescently labeled cells13,19. To avoid shear stress or clogging of the cell sorter, a wide nozzle (85 &#181;m or 100 &#181;m) is used. This will further improve the viability of the sorted cells.
This technique was designed and tested on both mouse and human teeth. The major limiting factor is the small number of cells in the reduced dental pulps of the teeth of older mice (molars) and humans. Suppose a larger number of cells need to be obtained or cells from the teeth of older patients are to be acquired. One possible solution is to process a higher number of teeth and merge them into a single batch, subsequently processed as one sample.
Living cells of human dental pulp were firstly isolated more than twenty years ago using an enzyme mixture of collagenase I and dispase23. Since then, isolations of dental pulp cells became widely utilized, and several techniques have been used5,6,7,8. The critical significance of the method presented here is the adaption of all isolation steps to make the isolation fast and gentle to ensure the high quality of the final cell suspension for scRNA-seq. Higher cell yield can be obtained by more prolonged incubation with enzymes. This protocol provides an efficient solution for quickly obtaining single cells from mouse and human teeth of suitable quality for single-cell RNA-sequencing. This technique is expected to be widely used for other tissues or organisms with just slight technical modifications.

Single-cell RNA sequencing is a powerful tool for deciphering in vivo cell population structure, hierarchy, interactions, and homeostasis1,2. However, its results strongly depend on the first step of this advanced analysis - the preparation of a single-cell suspension of perfect quality out of the complex, well-organized tissue. This encompasses keeping cells alive and preventing unwanted, artificial changes in gene expression profiles of the cells3,4. Such changes might lead to the inaccurate characterization of population structure and misinterpretation of the collected data.
Specific protocols for the isolation out of a wide range of tissues have been developed5,6,7,8. They usually employ mechanical dissociation in combination with further incubation with various proteolytic enzymes. These typically include trypsin, collagenases, dispases, papain6,7,8,9, or commercially available enzyme mixtures such as Accutase, Tryple, etc.5. The most critical part affecting the transcriptome quality is enzymatic digestion. It was shown that prolonged incubation with enzymes at 37 &#176;C influences the gene expression and causes the upregulation of many stress-related genes10,11,12,13. The other critical parameter of the isolation process is its overall length, as it has been shown that cell transcriptomes change after the tissue ischemia14. This protocol presents an efficient protocol for gentle isolation of single cells from mouse and human teeth, faster than other, previously utilized protocols for isolation of cells from complex tissues5,6,9,11,13,15,16.
This protocol presents how to quickly dissect soft tissue from the hard tooth and prepare a single-cell suspension suitable for scRNA-seq. This method employs only one centrifugation step and minimizes the effect of unwanted transcriptional changes by reducing the tissue handling and digestion time and keeping the tissue and cells at 4 &#176;C most of the time. The procedure showcases the isolation of cells from mouse incisors, molar, and human wisdom teeth as an example, but principally should work&#160;for other teeth in various organisms. The complete protocol is schematically visualized in Figure 1. This protocol has been recently used to generate a dental cell type atlas obtained from mouse and human teeth1.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>APC anti-mouse CD45 Antibody</td>
			<td>BioLegend</td>
			<td>103112</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin Fraction V</td>
			<td>Roche</td>
			<td>10735078001</td>
			<td></td>
		</tr>
		<tr>
			<td>CellTrics 50 &#181;m, sterile</td>
			<td>Sysmex</td>
			<td>04-004-2327</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase P (COLLP-PRO)</td>
			<td>Roche</td>
			<td>11213857001</td>
			<td></td>
		</tr>
		<tr>
			<td>CUTFIX Scalpel Blades, Fig. 10</td>
			<td>AESCULAP</td>
			<td>16600495</td>
			<td></td>
		</tr>
		<tr>
			<td>CUTFIX Scalpel Blades, Fig. 11</td>
			<td>AESCULAP</td>
			<td>16600509</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (South America), Ultra low Endotoxin</td>
			<td>Biosera</td>
			<td>FB-1101/500</td>
			<td></td>
		</tr>
		<tr>
			<td>Hank&#39;s balanced salt solution (HBSS) without Ca2+ and Mg2+</td>
			<td>SIGMA</td>
			<td>H6648</td>
			<td></td>
		</tr>
		<tr>
			<td>Industrial low lint wipes, MAX60</td>
			<td>Dirteeze</td>
			<td>MAX60B176</td>
			<td></td>
		</tr>
		<tr>
			<td>Industrial strong tweezers, style 660, bent serrated pointed tips, 150mm</td>
			<td>Value-Tec</td>
			<td>50-014366</td>
			<td></td>
		</tr>
		<tr>
			<td>Methyl alcohol A.G.</td>
			<td>Penta</td>
			<td>21210-11000</td>
			<td></td>
		</tr>
		<tr>
			<td>Propidium Iodide Solution</td>
			<td>BioLegend</td>
			<td>421301</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel handle</td>
			<td>CM Instrumente</td>
			<td>AG-013-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Serological pipettes 10mL, individually wrapped, 200 pcs.</td>
			<td>CAPP</td>
			<td>SP-10-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereo microscope</td>
			<td>Leica</td>
			<td>EZ4</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical scissors (9cm)</td>
			<td>CM Instrumente</td>
			<td>AI-430-09Y</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue culture dish &#216; 100 mm</td>
			<td>TPP</td>
			<td>93100</td>
			<td></td>
		</tr>
	</tbody>
</table>

rapid isolation of single cells from mouse and human teeth

Mouse and human teeth represent challenging organs for quick and efficient cell isolation for single-cell transcriptomic or other applications. The dental pulp tissue, rich in the extracellular matrix, requires a long and tedious dissociation process that is typically beyond the reasonable time for single-cell transcriptomics. For avoiding artificial changes in gene expression, the time elapsed from euthanizing an animal until the analysis of single cells needs to be minimized. This work presents a fast protocol enabling to obtain single-cell suspension from mouse and human teeth in an excellent quality suitable for scRNA-seq (single-cell RNA-sequencing). This protocol is based on accelerated tissue isolation steps, enzymatic digestion, and subsequent preparation of final single-cell suspension. This enables fast and gentle processing of tissues and allows using more animal or human samples for obtaining cell suspensions with high viability and minimal transcriptional changes. It is anticipated that this protocol might guide researchers interested in performing the scRNA-seq not only on the mouse or human teeth but also on other extracellular matrix-rich tissues, including cartilage, dense connective tissue, and dermis.

Exemplary isolation of single cells was performed from two mandibular incisors from one 6-week-old C57BL/6 mouse male. Following this protocol, a single-cell suspension was prepared, and subsequently, single-cell sequencing was performed. The prepared single-cell suspension was analyzed and sorted using FACS (Figure 2). Firstly, the FSC-A (forward scatter, area) and SSC-A (side scatter, area) plotting was applied, and an appropriate gating strategy was used to select a population with expected size and granularity to filter our cell debris and cell doublets or aggregates (Figure 2A). This selected population (P1), counting 38% of all events, was further used, and FSC-A and FSC-H (forward scatter, height) parameters were applied to remove the remaining cell doublets (Figure 2B). The population without cell doublets (P2) counting 95% of P1 can be subsequently used for scRNA-seq. Alternatively, additional gating can be used to select the population of interest (e.g., expression of fluorescent proteins or live/dead staining). To check the number of live/dead cells in the final suspension, the PI (propidium iodide) staining was performed (Figure 2C). The P3 population containing PI- (living) cells was 98.4% out of the parent P2 population and 35.5% out of the total events. The total number of filtered out, dead (PI+) cells was 1887.
The final number of cells obtained without viability staining suitable for RNA-seq (P2) counted 118,199 cells from two mouse incisor pulps. This means the number of almost 60,000 living cells from one mandibular incisor.
To clarify the number of immune cells in the final single-cell suspension, two approaches were used. Firstly, the CD45 antibody staining and subsequent FACS analysis were used. As a complementary method, the total number of immune cells (CD45+) in scRNA-seq data was analysed. FACS analysis showed 14.44% of CD45+ cells (13.20% alive and 1.24% dead) (Figure 3A). Analysis of scRNA-seq data showed 10.90% of CD45-expressing cells (Figure 3B). The decrease of CD45+ cells in scRNA-seq data can be caused by additional thresholding during scRNA-seq analysis.
These representative data on the example of mouse incisor show that the given protocol in combination with strict gating strategy is efficient in obtaining a high number of cells out of a single mouse tooth without the necessity of additional use of viability staining. The ratio of immune (CD45+) cells was minor (13.2%). Moreover, it was previously shown that the immune cells are essential in maintaining tooth homeostasis, so removing them from scRNA-seq analysis during the FACS would be counterproductive in some applications.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63043/63043fig01.jpg" /> 
Figure 1: Schematic representation of the protocol. Different steps, including temperature conditions and expected time, are represented to prepare single-cell suspension from mouse and human teeth. <a href="https://www.jove.com/files/ftp_upload/63043/63043fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63043/63043fig02.jpg" /> 
Figure 2: Example of the gating strategy. FSC-A and SSC-A gating was used to produce the P1 gate, reflecting the cell population with expected size and granularity and filtering out the cell and extracellular matrix debris and most large events (A). Subsequently, the P1 population was plotted in FSC-H and FCS-A plot, which filtered out cell doublets (B). This P2 population was then analyzed for the presence of dead cells by propidium iodide (C). The number of events/cells per gate are represented in (D). (FSC-A - forward scatter, area; SSC-A - side scatter, area; FSC-H - forward scatter, height; PI - propidium iodide). <a href="https://www.jove.com/files/ftp_upload/63043/63043fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63043/63043fig03.jpg" /> 
Figure 3: Quantification of the immune cells. Quantification of the immune cells was performed by FACS analysis of cells stained with anti-CD45 antibody and Live/Dead analysis using propidium iodide staining (A). Further quantification of immune cells was performed during scRNA-seq analysis (B). (CD45-APC - anti-CD45 allophycocyanin conjugated antibody; PI - propidium iodide; t-SNE - t-distributed stochastic neighbor embedding). <a href="https://www.jove.com/files/ftp_upload/63043/63043fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Overview of the mandible dissection process. Dashed lines illustrate suggested cuts. TMJ - temporomandibular joint, m. masseter - musculus masseter. <a href="https://www.jove.com/files/ftp_upload/63043/Supplementary figure 1.png" target="_blank">Please click here to download this File.</a>
Supplementary Table 1: The compositions of the solutions used in the study. <a href="https://www.jove.com/files/ftp_upload/63043/Supplemental table 1.docx" target="_blank">Please click here to download this Table.</a>

Watch this Scientific Journal Video about Rapid Isolation of Single Cells from Mouse and Human Teeth at JoVE.com

Rapid Isolation of Single Cells from Mouse and Human Teeth

The current protocol presents a fast, efficient, and gentle method for isolating single cells suitable for single-cell RNA-seq analysis from a continuously growing mouse incisor, mouse molar, and human teeth.

Mouse and human teeth represent challenging organs for quick and efficient cell isolation for single-cell transcriptomic or other applications. The dental ...

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3.4K Views.  Masaryk University. The current protocol presents a fast, efficient, and gentle method for isolating single cells suitable for single-cell RNA-seq analysis from a continuously growing mouse incisor, mouse molar, and human teeth.

Video: Rapid Isolation of Single Cells from Mouse and Human Teeth

Isolation and Quantitative Immunocytochemical Characterization of Primary Myogenic Cells and Fibroblasts from Human Skeletal Muscle

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated from human muscle biopsy samples using enzymatic digestion and their myogenic properties studied in culture. Quantitatively, the two main adherent cell types obtained from enzymatic digestion are: (i) the satellite cells (termed myogenic cells or muscle precursor cells), identified initially as CD56+ and later as CD56+/desmin+ cells and (ii) muscle-derived fibroblasts, identified as CD56&#8211; and TE-7+. Fibroblasts proliferate very efficiently in culture and in mixed cell populations these cells may overrun myogenic cells to dominate the culture. The isolation and purification of different cell types from human muscle is thus an important methodological consideration when trying to investigate the innate behavior of either cell type in culture. Here we describe a system of sorting based on the gentle enzymatic digestion of cells using collagenase and dispase followed by magnetic activated cell sorting (MACS) which gives both a high purity (&#62;95% myogenic cells) and good yield (~2.8 x 106 &#177; 8.87 x 105 cells/g tissue after 7 days in vitro) for experiments in culture. This approach is based on incubating the mixed muscle-derived cell population with magnetic microbeads beads conjugated to an antibody against CD56 and then passing cells though a magnetic field. CD56+ cells bound to microbeads are retained by the field whereas CD56&#8211; cells pass unimpeded through the column. Cell suspensions from any stage of the sorting process can be plated and cultured. Following a given intervention, cell morphology, and the expression and localization of proteins including nuclear transcription factors can be quantified using immunofluorescent labeling with specific antibodies and an image processing and analysis package.

The main adherent cell types derived from human muscle are myogenic cells and fibroblasts. Here, cell populations are enriched using magnetic-activated cell sorting based on the CD56 antigen. Subsequent immunolabelling with specific antibodies and use of image analysis techniques allows quantification of cytoplasmic and nuclear characteristics in individual cells.

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated ...

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Isolation and Expansion of Mesenchymal Stem/Stromal Cells Derived from Human Placenta Tissue

Mesenchymal stem/stromal cells (MSC) are promising candidates for use in cell-based therapies. In most cases, therapeutic response appears to be cell-dose dependent. Human term placenta is rich in MSC and is a physically large tissue that is generally discarded following birth. Placenta is an ideal starting material for the large-scale manufacture of multiple cell doses of allogeneic MSC. The placenta is a fetomaternal organ from which either fetal or maternal tissue can be isolated. This article describes the placental anatomy and procedure to dissect apart the decidua (maternal), chorionic villi (fetal), and chorionic plate (fetal) tissue. The protocol then outlines how to isolate MSC from each dissected tissue region, and provides representative analysis of expanded MSC derived from the respective tissue types. These methods are intended for pre-clinical MSC isolation, but have also been adapted for clinical manufacture of placental MSC for human therapeutic use.

Herein we describe methods for the dissection of fetal and maternal tissues from human term placenta, followed by isolation and expansion of mesenchymal stem/stromal cells (MSC) from these tissues.

Mesenchymal stem/stromal cells (MSC) are promising candidates for use in cell-based therapies. In most cases, therapeutic response appears to be cell-dose ...

Isolation and Characterization of Mesenchymal Stromal Cells from Human Umbilical Cord and Fetal Placenta

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly gained attention because they do not pose any ethical or moral concerns. Current methods to isolate MSCs from UC yield low amounts of cells with variable proliferation potentials. Since UC is an anatomically-complex organ, differences in MSC properties may be due to the differences in the anatomical regions of their isolation. In this study, we first dissected the cord/placenta samples into three discrete anatomical regions: UC, cord-placenta junction (CPJ), and fetal placenta (FP). Second, two distinct zones, cord lining (CL) and Wharton&#39;s jelly (WJ), were separated. The explant culture technique was then used to isolate cells from the four sources. The time required for the primary culture of cells from the explants varied depending on the source of the tissue. Outgrowth of the cells occurred within 3 - 4 days of the CPJ explants, whereas growth was observed after 7 - 10 days and 11 - 14 days from CL/WJ and FP explants, respectively. The isolated cells were adherent to plastic and displayed fibroblastoid morphology and surface markers, such as CD29, CD44, CD73, CD90, and CD105, similarly to bone marrow (BM)-derived MSCs. However, the colony-forming efficiency of the cells varied, with CPJ-MSCs and WJ-MSCs showing higher efficiency than BM-MSCs. MSCs from all four sources differentiated into adipogenic, chondrogenic, and osteogenic lineages, indicating that they were multipotent. CPJ-MSCs differentiated more efficiently in comparison to other MSC sources. These results suggest that the CPJ is the most potent anatomical region and yields a higher number of cells, with greater proliferation and self-renewal capacities in vitro. In conclusion, the comparative analysis of the MSCs from the four sources indicated that CPJ is a more promising source of MSCs for cell therapy, regenerative medicine, and tissue engineering.

Here, we present a protocol for the dissection of human umbilical cord (UC) and fetal placenta sample into cord lining (CL), Wharton&#39;s jelly (WJ), cord-placenta junction (CPJ), and fetal placenta (FP) for the isolation and characterization of mesenchymal stromal cells (MSCs) using the explant culture technique.

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly ...

Isolation of Endothelial Progenitor Cells from Human Umbilical Cord Blood

The existence of endothelial progenitor cells (EPCs) in peripheral blood and its involvement in vasculogenesis was first reported by Ashara and colleagues1. Later, others documented the existence of similar types of EPCs originating from bone marrow2,3. More recently, Yoder and Ingram showed that EPCs derived from umbilical cord blood had a higher proliferative potential compared to ones isolated from adult peripheral blood4,5,6. Apart from being involved in postnatal vasculogenesis, EPCs have also shown promise as a cell source for creating tissue-engineered vascular and heart valve constructs7,8. Various isolation protocols exist, some of which involve the cell sorting of mononuclear cells (MNCs) derived from the sources mentioned earlier with the help of endothelial and hematopoietic markers, or culturing these MNCs with specialized endothelial growth medium, or a combination of these techniques9. Here, we present a protocol for the isolation and culture of EPCs using specialized endothelial medium supplemented with growth factors, without the use of immunosorting, followed by the characterization of the isolated cells using Western blotting and immunostaining.

The goal of this protocol is to isolate endothelial progenitor cells from umbilical cord blood. Some of the applications include using these cells as a biomarker for identifying patients with cardiovascular risk, treating ischemic diseases, and creating tissue-engineered vascular and heart valve constructs.

The existence of endothelial progenitor cells (EPCs) in peripheral blood and its involvement in vasculogenesis was first reported by Ashara and ...

Rapid, Directed Differentiation of Retinal Pigment Epithelial Cells from Human Embryonic or Induced Pluripotent Stem Cells

We describe a robust method to direct the differentiation of pluripotent stem cells into retinal pigment epithelial cells (RPE). The purpose of providing a detailed and thorough protocol is to clearly demonstrate each step and to make this readily available to researchers in the field. This protocol results in a homogenous layer of RPE with minimal or no manual dissection needed. The method presented here has been shown to be effective for induced pluripotent stem cells (iPSC) and human embryonic stem cells. Additionally, we describe methods for cryopreservation of intermediate cell banks that allow long-term storage. RPE generated using this protocol might be useful for iPSC disease-in-a-dish modeling or clinical application.

This protocol describes how to produce retinal pigment epithelial cells (RPE) from pluripotent stem cells. The method uses a combination of growth factors and small molecules to direct the differentiation of stem cells into immature RPE in fourteen days and mature, functional RPE after three months.

We describe a robust method to direct the differentiation of pluripotent stem cells into retinal pigment epithelial cells (RPE). The purpose of providing ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Isolation and Characterization of Human Umbilical Cord-derived Mesenchymal Stem Cells from Preterm and Term Infants

Mesenchymal stem cells (MSCs) have considerable therapeutic potential and attract increasing interest in the biomedical field. MSCs are originally isolated and characterized from bone marrow (BM), then acquired from tissues including adipose tissue, synovium, skin, dental pulp, and fetal appendages such as placenta, umbilical cord blood (UCB), and umbilical cord (UC). MSCs are a heterogeneous cell population with the capacity for (1) adherence to plastic in standard culture conditions, (2) surface marker expression of CD73+/CD90+/CD105+/CD45-/CD34-/CD14-/CD19-/HLA-DR- phenotypes, and (3) trilineage differentiation into adipocytes, osteocytes, and chondrocytes, as currently defined by the International Society for Cellular Therapy (ISCT). Although BM is the most widely used source of MSCs, the invasive nature of BM aspiration ethically limits its accessibility. Proliferation and differentiation capacity of MSCs obtained from BM generally decline with the age of the donor. In contrast, fetal MSCs obtained from UC have advantages such as vigorous proliferation and differentiation capacity. There is no ethical concern for UC sampling, as it is typically regarded as medical waste. Human UC starts to develop with continuing growth of the amniotic cavity at 4-8 weeks of gestation and keeps growing until reaching 50-60 cm in length, and it can be isolated during the whole newborn delivery period. To gain insight into the pathophysiology of intractable diseases, we have used UC-derived MSCs (UC-MSCs) from infants delivered at various gestational ages. In this protocol, we describe the isolation and characterization of UC-MSCs from fetuses/infants at 19-40 weeks of gestation.

Human umbilical cord (UC) can be obtained during the perinatal period as a result of preterm, term, and postterm delivery. In this protocol, we describe the isolation and characterization of UC-derived mesenchymal stem cells (UC-MSCs) from fetuses/infants at 19-40 weeks of gestation.

Mesenchymal stem cells (MSCs) have considerable therapeutic potential and attract increasing interest in the biomedical field. MSCs are originally ...

In Vitro Generation of Somite Derivatives from Human Induced Pluripotent Stem Cells

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give rise to multiple cell types, including the myotome (MYO), sclerotome (SCL), dermatome (D), and syndetome (SYN), which in turn develop into skeletal muscle, axial skeleton, dorsal dermis, and axial tendon/ligament, respectively. Therefore, the generation of SMs and their derivatives from human induced pluripotent stem cells (iPSCs) is critical to obtain pluripotent stem cells (PSCs) for application in regenerative medicine and for disease research in the field of orthopedic surgery. Although the induction protocols for MYO and SCL from PSCs have been previously reported by several researchers, no study has yet demonstrated the induction of SYN and D from iPSCs. Therefore, efficient induction of fully competent SMs remains a major challenge. Here, we recapitulate human SM patterning with human iPSCs in vitro by mimicking the signaling environment during chick/mouse SM development, and report on methods of systematic induction of SM derivatives (MYO, SCL, D, and SYN) from human iPSCs under chemically defined conditions through the presomitic mesoderm (PSM) and SM states. Knowledge regarding chick/mouse&#160;SM development was successfully applied to the induction of SMs with human iPSCs. This method could be a novel tool for studying human somitogenesis and patterning without the use of embryos and for cell-based therapy and disease modeling.

We present here a protocol for the differentiation of human induced pluripotent stem cells into each somite derivative (myotome, sclerotome, dermatome, and syndetome) in chemically defined conditions, which has applications in future disease modeling and cell-based therapies in orthopedic surgery.

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give ...

Preparation of Small RNA Libraries for Sequencing from Early Mouse Embryos

MicroRNAs (miRNAs) are important for the complex regulation of cell fate decisions and developmental timing. In vivo studies of the contribution of miRNAs during early development are technically challenging due to the limiting cell number. Moreover, many approaches require a miRNA of interest to be defined in assays such as northern blotting, microarray, and qPCR. Therefore, the expression of many miRNAs and their isoforms have not been studied during early development. Here, we demonstrate a protocol for small RNA sequencing of sorted cells from early mouse embryos to enable relatively unbiased profiling of miRNAs in early populations of neural crest cells. We overcome the challenges of low cell input and size selection during library preparation using amplification and gel-based purification. We identify embryonic age as a variable accounting for variation between replicates and stage-matched mouse embryos must be used to accurately profile miRNAs in biological replicates. Our results suggest that this method can be broadly applied to profile the expression of miRNAs from other lineages of cells. In summary, this protocol can be used to study how miRNAs regulate developmental programs in different cell lineages of the early mouse embryo.

We describe a technique for profiling microRNAs in early mouse embryos. This protocol overcomes the challenge of low cell input and small RNA enrichment. This assay can be used to analyze changes in miRNA expression over time in different cell lineages of the early mouse embryo.

MicroRNAs (miRNAs) are important for the complex regulation of cell fate decisions and developmental timing. In vivo studies of the contribution of miRNAs ...