This project was supported by Andrew McDonough B+ Foundation and the NIH (R01-HD099252, R01-HD098131.&#160;R.J.P. is a CPRIT Scholar in Cancer Research (RR150106) and V Scholar in Cancer Research (V Foundation).<span style="border-width: 1pt; border-style: none; border-color: windowtext; padding: 0in; background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-origin: initial; background-clip: initial;">&#160;The authors would also like to acknowledge the Cytometry and Cell Sorting Core at BCM for providing equipment necessary for this project.

All research and animal care procedures were approved by the Baylor College of Medicine Institutional Animal Care and Use Committee and housed in the Association for Assessment and Accreditation of Laboratory Animal Care-approved animal facility at Baylor College of Medicine. All strains were maintained on C57BL6 background.
1. Embryo dissection (E7.5-E9.5)
<ol>
	<li>Remove uterine horns from a pregnant female mouse, sterilizing the abdomen with 70% ethanol where the incision is to be made.</li>
	<li>Clean off the uterus by rinsing with phosphate buffered saline (PBS). Place the uterus into a sterile plastic 10 cm dish.</li>
	<li>Using microdissection scissors, separate each decidua containing region from one another. Peel off the uterine muscle and expose all deciduae. One by one, remove the decidua from each embryo.</li>
	<li>Remove the yolk sac from each embryo and save for genotyping.</li>
	<li>Move all embryos into a new dish of fresh PBS for imaging.</li>
	<li>Take an image of the entire litter. Image each embryo individually and count the number of somites of each embryo. Keep the magnification and exposure (for fluorescence) the same between experiments. 
	NOTE: We find that 50-200 ms exposure is adequate for fluorescence and that 20 ms is adequate for bright field setting.</li>
</ol>
2. Embryo dissociation and cell sorting
<ol>
	<li>For each embryo to be sorted from, do the following.
	<ol>
		<li>Decapitate just above the otic placode for embryos older than E8.0 (if only labeled cells of the cranial region are desired). For E7.5 embryos, remove extraembryonic structures (if only the embryo proper is desired).</li>
		<li>Move the head in a minimal volume of PBS over to a clean well of a 48-well plate. Add 250 &#181;L of papain (27 U/mL). Pipette up and down gently using a p200.</li>
		<li>Check under the microscope and look for clumps and single cells.</li>
		<li>Repeat gentle pipetting up and down until a single cell suspension is achieved (usually three rounds of pipetting up and down which should equate to between 30 s to 1 min for E7.5-E9.5).</li>
		<li>Quench with 250 &#181;L of fetal bovine serum (FBS).</li>
		<li>Repeat steps 2.1.1-2.1.5 for each embryo to be sorted.</li>
	</ol>
	</li>
	<li>Filter 500 &#181;L of cell suspension through a 35 &#181;m nylon mesh filter cap tube to remove clumps.</li>
	<li>Take the filtrate and move to new 1.5 mL tube and spin at 200 x g for 5 min. Remove supernatant carefully and transfer to new tube (save until live cells are known to be in the pellet).</li>
	<li>Resuspend cell pellet in 300 &#181;L of PBS containing 0.5-1% bovine serum albumin and keep on ice until sorting (minimize the time between now and the end of sorting).
	<ol>
		<li>If desired, take 10 &#181;L of cell suspension and combine with 10 &#181;L of Trypan blue (0.4%) and count using a hemocytometer to identify approximate cell quantification and viability as Trypan blue only stains dead cells.</li>
	</ol>
	</li>
	<li>Just before sorting, filter each sample again through a filter cap tube and add DAPI stain for live cells.</li>
	<li>Sort each sample into 500 &#181;L of RNA extraction lysis solution (see Table of Materials) on cell sorter with 70 &#181;m nozzle.</li>
	<li>Mix and store at -80 &#176;C until all samples are harvested. Proceed from this point only when all samples needed for an experiment are harvested to reduce technical variation between rounds of library preparation.</li>
</ol>
3. RNA extraction
NOTE: The protocol is adapted from the RNA isolation kit; see Table of Materials.
<ol>
	<li>Add 500 &#181;L of RNA extraction lysis solution (see Table of Materials) to each sample to make the total volume 1000 &#181;L.</li>
	<li>Thaw each sample at room temperature.</li>
	<li>Vortex each sample for 1.5 min, making sure that the cap is securely on each tube before starting homogenization. Incubate the homogenate at room temperature (15&#8211;25 &#176;C) for 5 min.</li>
	<li>Add 140 &#181;L of chloroform, cap tube securely and shake vigorously for 15 s. Incubate at room temperature for 3 min.</li>
	<li>Centrifuge for 15 min at 12,000 x g at 4 &#176;C. Transfer the upper aqueous phase to a new collection tube on ice. Do not transfer any interphase or any organic phase. When approaching the pink organic phase, tip the tube to side and remove small aliquots until no clear aqueous phase can be collected.</li>
	<li>Measure the volume of RNA containing aqueous phase collected from each sample for the correct calculation in the next step.</li>
	<li>Add 1.5 volumes (usually 525 &#181;L but may be slightly more or less) of 100% ethanol and mix thoroughly by pipetting.</li>
	<li>Pipette up to 700 &#181;L sample, including any precipitate, into column in a 2 mL collection tube. Close the lid and centrifuge at &#8805;8,000 x g for 15 s at room temperature. Discard the flow-through.</li>
	<li>Repeat step 3.8 using the remainder of the sample.</li>
	<li>Wash the column as per manufacturer instructions. Dry the column by spinning at full speed for 1 min. Transfer the column to a new 1.5 mL collection tube.</li>
	<li>Pipette 11 &#181;L of nuclease free water onto the center of the membrane. Let sit at room temperature for 1 min. Spin max speed for 1 min to elute off the column.</li>
	<li>Measure RNA concentration using a spectrophotometer and a parallel capillary electrophoresis instrument (Table of Materials).</li>
</ol>
4. Library preparation
NOTE: The protocol is adapted from small RNA library preparation kit handbook; see Table of Materials.
<ol>
	<li>Complete the denaturation and 3&#8217; adapter ligation as stated in the small RNA library preparation kit handbook using &#188; dilution of 3&#8217; adapter.</li>
	<li>Continue to excess 3&#39; adapter removal.</li>
	<li>Complete the adapter depletion as stated in the small RNA library preparation kit handbook. Be sure to use freshly prepared 80% ethanol for the bead cleanup and that all excess ethanol is completely removed just prior to the resuspension in nuclease-free water without over-drying the beads (cracking of the bead pellet is observed upon over-drying).</li>
	<li>Proceed directly to excess adapter inactivation.</li>
	<li>Complete the excess adapter inactivation as stated in the small RNA library preparation kit handbook assembling all reagents on ice.</li>
	<li>Continue to 5&#39; 4N adapter ligation.</li>
	<li>Complete 5&#39; adapter ligation as stated in the small RNA library preparation kit handbook being sure to use a &#188; dilution of the 5&#8217; adapter and assembling all reagents on ice.</li>
	<li>Proceed to reverse transcription.</li>
	<li>Complete reverse transcription-first strand synthesis as stated in the small RNA library preparation kit handbook and taking care to assemble all reagents on ice and not keeping the enzyme out of the freezer for extended periods of time. 
	NOTE: At this point, the protocol may be stopped, with samples stored overnight at 4 &#176;C. Alternatively continue to PCR amplification.</li>
	<li>Complete PCR amplification by assembling the reagents as stated in the small RNA library preparation kit handbook and minimizing the number of PCR cycles. The number of PCR cycles should be experimentally determined for each application and the minimum number of cycles should be used. Here we used 16 cycles to successfully amplify our small RNA libraries.</li>
	<li>Proceed directly to size selection.</li>
</ol>
5. Size selection
<ol>
	<li>To each sample, add 5 &#181;L of 6x gel loading dye and pipette to mix.</li>
	<li>Load 10 &#181;L of ladder to the first well of the gel, leaving the well immediately to the right empty. Load all of product from step 5.1 onto a 6% Tris/borate/ethylenediaminetetraacetic acid &#8211; polyacrylamide gel (TBE-PAGE) and leave 1 lane in between each sample. 
	NOTE: Keep the container that the gel came in for subsequent steps.</li>
	<li>Run the gel at 150 V for approximately 30-40 min in 0.5x TBE running buffer (until the lower dye band is near the bottom of the gel, 0.5-1 cm).</li>
	<li>Make 1 L of staining solution while the gel is running: SYBR Gold diluted 1:10,000 in 0.5x TBE.</li>
	<li>When the gel has finished running (i.e., lower dye band is near the bottom of the gel), carefully remove the gel from the glass plates, noting the orientation of the gel.</li>
	<li>Place gel in the tray of its original packaging. Remove the 0.5x TBE and replace with the staining solution from step 5.5. Stain gel for 15 min. Staining time may need to be increased.</li>
	<li>Place the gel on a UV transilluminator, taking care to not rip the gel and maintaining the orientation noted above (re-stain for additional time if bands of ladder cannot be clearly seen).</li>
	<li>Image the gel once staining is complete on UV transilluminator with a camera.
	<ol>
		<li>If a better image is required, first use an imaging apparatus other than the UV transilluminator to capture an image of the gel before cutting out the bands on a UV transilluminator as imaging directly on the UV transilluminator causes the variable coloring of the background as shown in Figure 3A-B. Do not move the gel too many times as it is delicate and may tear.</li>
	</ol>
	</li>
	<li>Identify and remove the ~150 bp band using a clean razor blade and place into clean 1.5 mL tube. Do not cut out the ~130 bp band (adapter dimer product).
	<ol>
		<li>Image the gel again after removing the slices containing the library product for documentation purposes.</li>
	</ol>
	</li>
	<li>Spin the microcentrifuge tubes containing the gel slice at max speed for 30 s to collect the slices at the bottom of each tube.</li>
	<li>Use a p200 tip for each sample to crush the slice of gel into the smallest bits possible. Eject each tip into each tube.</li>
	<li>To each tube, add 300 &#181;L of elution buffer, taking care to wash gel bits from side of tube and reattach the p200 tip and wash off the tip for each sample.</li>
	<li>Place the eluting samples in a shaking incubator set to 25 &#176;C. Incubate overnight with 1000 rpm shaking. Remove tubes from incubator and spin at max speed for 10 min at room temperature.</li>
	<li>Transfer the supernatant containing the library product to a 2 mL RNase free 96 well plate. Take care to not transfer any gel.</li>
	<li>Add 50 &#181;L cleanup beads to each sample and mix by pipetting. Immediately, add 350 &#181;L of isopropanol and mix by pipetting. Incubate at room temperature for 10 minutes with rocking.</li>
	<li>Pulse spin plate to pellet beads and then magnetize for 2 min. Carefully remove and discard the supernatant.</li>
	<li>Add 950 &#181;L of 80% ethanol. Incubate for 30 s. Remove all of the supernatant. Repeat for a total of two ethanol washes.</li>
	<li>Dry the sample for 3 min. At this step take care to remove any residual ethanol that collects at the bottom of each well (not more than once) and to not over-dry the beads (over-drying will result in cracking of the bead pellet).</li>
	<li>Remove all residual liquid at the bottom of the well. Remove the plate from the magnetic stand.</li>
	<li>Resuspend the pellet in 13 &#181;L of resuspension buffer. Mix by pipette until homogeneous. Incubate for 2 min and then magnetize for 3 min.</li>
	<li>Transfer 12 &#181;L of supernatant to a clean tube or well of clean plate for subsequent storage. Store all samples at 4 &#176;C overnight or -20 &#176;C long term.</li>
	<li>Check the size and concentration of fragments present in each library using the capillary electrophoresis high sensitivity DNA assay. Confirm the concentration with more than one method. 
	NOTE: We have occasionally used the standard sensitivity kit to avoid overloading the capillary electrophoresis.</li>
</ol>

<ol>
	<li>Sauka-Spengler, T., Bronner-Fraser, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+gene+regulatory+network+orchestrates+neural+crest+formation.">A gene regulatory network orchestrates neural crest formation.</a> Nature Reviews Molecular Cell Biology. 9, 557-568 (2008).</li><li>Abrahante, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Caenorhabditis+elegans+hunchback-like+gene+lin-57/hbl-1+controls+developmental+time+and+is+regulated+by+microRNAs.">The Caenorhabditis elegans hunchback-like gene lin-57/hbl-1 controls developmental time and is regulated by microRNAs.</a> Developmental Cell. 4 (5), 625-637 (2003).</li><li>Lin, S. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+C+elegans+hunchback+homolog,+hbl-1,+controls+temporal+patterning+and+is+a+probable+microRNA+target.">The C elegans hunchback homolog, hbl-1, controls temporal patterning and is a probable microRNA target.</a> Developmental Cell. 4 (5), 639-650 (2003).</li><li>Reinhart, B. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+21-nucleotide+let-7+RNA+regulates+developmental+timing+in+Caenorhabditis+elegans.">The 21-nucleotide let-7 RNA regulates developmental timing in Caenorhabditis elegans.</a> Nature. 403 (6772), 901 (2000).</li><li>Slack, F. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+lin-41+RBCC+gene+acts+in+the+C.+elegans+heterochronic+pathway+between+the+let-7+regulatory+RNA+and+the+LIN-29+transcription+factor.">The lin-41 RBCC gene acts in the C. elegans heterochronic pathway between the let-7 regulatory RNA and the LIN-29 transcription factor.</a> Molecular Cell. 5 (4), 659-669 (2000).</li><li>Vella, M. C., Choi, E. Y., Lin, S. Y., Reinert, K., Slack, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+C.+elegans+microRNA+let-7+binds+to+imperfect+let-7+complementary+sites+from+the+lin-41+3'UTR.">The C. elegans microRNA let-7 binds to imperfect let-7 complementary sites from the lin-41 3'UTR.</a> Genes &amp; Development. 18 (2), 132-137 (2004).</li><li>Ventura, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+deletion+reveals+essential+and+overlapping+functions+of+the+miR-17+92+family+of+miRNA+clusters.">Targeted deletion reveals essential and overlapping functions of the miR-17 92 family of miRNA clusters.</a> Cell. 132 (5), 875-886 (2008).</li><li>Eberhart, J. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroRNA+Mirn140+modulates+Pdgf+signaling+during+palatogenesis.">MicroRNA Mirn140 modulates Pdgf signaling during palatogenesis.</a> Nature Genetics. 40 (3), 290 (2008).</li><li>Li, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+regulation+of+small+RNAome+during+the+early+stage+of+cardiac+differentiation+from+pluripotent+embryonic+stem+cells.">Dynamic regulation of small RNAome during the early stage of cardiac differentiation from pluripotent embryonic stem cells.</a> Genomics Data. 12, 136-145 (2017).</li><li>Chugh, P., Dittmer, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Potential+pitfalls+in+microRNA+profiling.">Potential pitfalls in microRNA profiling.</a> Wiley Interdisciplinary Reviews: RNA. 3 (5), 601-616 (2012).</li><li>Werber, M., Wittler, L., Timmermann, B., Grote, P., Herrmann, B. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+tissue-specific+transcriptomic+landscape+of+the+mid-gestational+mouse+embryo.">The tissue-specific transcriptomic landscape of the mid-gestational mouse embryo.</a> Development. 141 (11), 2325-2330 (2014).</li><li>Yang, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+sensitive+sequencing+reveals+dynamic+modifications+and+activities+of+small+RNAs+in+mouse+oocytes+and+early+embryos.">Highly sensitive sequencing reveals dynamic modifications and activities of small RNAs in mouse oocytes and early embryos.</a> Science Advances. 2, (2016).</li><li>Ohnishi, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Small+RNA+class+transition+from+siRNA/piRNA+to+miRNA+during+pre-implantation+mouse+development.">Small RNA class transition from siRNA/piRNA to miRNA during pre-implantation mouse development.</a> Nucleic Acids Research. 38 (15), 5141-5151 (2010).</li><li>Loontiens, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Purification+of+high-quality+RNA+from+a+small+number+of+fluorescence+activated+cell+sorted+zebrafish+cells+for+RNA+sequencing+purposes.">Purification of high-quality RNA from a small number of fluorescence activated cell sorted zebrafish cells for RNA sequencing purposes.</a> BMC Genomics. 20 (1), 228 (2019).</li></ol>

The authors have nothing to disclose.

Developmental processes can proceed rapidly, and cells are undergoing many sequential specifications such that to capture a comprehensive view of the miRNAs contributing to early fate decisions, more specific staging is needed than the widely used half-day increment. A recent study has performed RNA sequencing from Theiler stage 12 embryos which range from having 3 to 6 somites11. We find that during this period of time, the neural crest cells are specified (3 somites of age), delaminate (4 somites of age), and migrate (5-6 somites of age). We also find that besides the Cre-driver used to lineage trace cell populations, age is the largest source of variation between biological samples, and only somite matched embryos should be considered as replicates. This should also be taken into consideration when comparing transgenic embryos to wildtype controls.
Previous methods to profile miRNAs during early development have used between 10-100 ng of RNA input for small RNA sequencing library preparation and have pooled multiple embryos into one sample13,14. We demonstrate RNA isolation and library preparation from a single E7.5 embryo or from sorted neural crest cells at E8.5 and E9.5 using approximately 100 ng of total RNA as input. When dissociating embryos for sorting, one should take care to watch the dissociation under a microscope and quench the reaction to observe when single cells are obtained. We find that dissociation of the cranial region of E8.5-E9.5 embryos is almost instant with gentle manual pipetting as described in the protocol. For larger tissues and increasingly older embryos, dissociation time may be longer depending on the portion of the embryo being dissociated. For E7.5-E9.5 embryos, clumps of cells are easily visible under the microscope and the dissociation should continue until no more clumps are visible. Single cells are visible in solution if you adjust the focus through the solution in your well anywhere from 5-10x. Previous methods sort cells directly into lysis buffer for RNA sequencing to prep bulk RNA from a low number of cells14. Here we sort directly into RNA extraction lysis solution so that RNA can be isolated before the start of library preparation. Use of mini columns with 11 &#181;L elution volume allowed for a high enough RNA concentration such that a single RNA prep could be split between small RNA and bulk RNA sequencing.
One current limitation of most small RNA sequencing methods is the PCR amplification of converted cDNA. Our method does not overcome this limitation, but we were able to minimize the number of PCR cycles from the 25-maximum recommended down to 16 cycles. This reduction in amplification decreases artificial amplification bias introduced by PCR. Another source of bias is the ligation of adapters, where it is known that specific sequences located at the ends of adapters and miRNAs can ligate together with greater efficiency than other sequences. To avoid this, the adapters used in this protocol have 4 random bases incorporated at the end of each adapter to prevent bias in ligation reactions. Additionally, another common issue is the amount of adapter dimers that form when the RNA input is low. The library preparation kit does include steps to reduce adapter dimer formation such as adapter inactivation and bead cleanups to remove excess adapters after each ligation. We also diluted the 3&#8217; and 5&#8217; 4N adapters by &#188; to reduce the amount of adapter dimer that can form. We found that when not diluted, the 130 bp band intensity increases making it difficult to distinguish from the 150 bp band containing the desired small RNA libraries on a gel.
Another current challenge of preparing sequencing libraries is the accurate quantification of product prior to sequencing. We have found that different methods give varying results on the same library. We suggest that the researchers use multiple methods of quantification to get an accurate estimation of concentration.
This protocol can be widely applied to genetic, developmental studies, or other applications where RNA is being harvested from a low number of cells. This approach simplifies temporal studies by avoiding the pooling of embryos and can easily be applied to both non-sorted and sorted cells.

A central question of developmental biology is how a single undifferentiated cell can give rise to an entire organism with numerous complex cell types. During embryogenesis, the developmental potential of cells becomes progressively restricted as the organism develops. One example is the neural crest lineage, which progressively differentiates from a multipotent cell population into various terminal derivatives, such as peripheral neurons, glia, cranial bone, and cartilage. Neural crest cells are specified from the ectoderm during gastrulation and then undergo an epithelial to mesenchymal transition and migrate through the embryo to discrete locations throughout the body where they will terminally differentiate1. Decades of work have uncovered a transcriptional gene regulatory network, but far less is known about the mechanisms of post-transcriptional regulation that control the timing of neural crest development.
Previous work suggests that microRNAs (miRNAs) repress gene expression for proper developmental timing and cell fate decisions2,3,4,5,6. Studies of miRNAs in neural crest development have largely focused on later stages of craniofacial development. For example, miR-17~92 and miR-140 are critical for palatogenesis during craniofacial development in mouse and zebrafish, respectively7,8. The contribution of miRNAs to the earliest neural crest fate decisions of the embryo has not been thoroughly investigated. Studies of miRNAs in early fate decisions have been limited by technical challenges such as the low cell number present in early embryos.
MiRNAs have been profiled in vitro from cell lines using embryoid bodies at different stages of differentiation to model early mouse development9. The investigation of small RNAs in vivo during early mammalian development has been relatively limited. Previous methods to profile miRNAs have led to bias as a known sequence is used to analyze expression of a specific miRNA in methods such as qPCR, microarrays, and northern blots10. Next generation sequencing and ever improving molecular tools now allow for relatively unbiased analysis of miRNA expression to study their contribution to early mammalian development and cell fate decisions.
Here, we report a technique to harvest and sequence small RNAs expressed in neural crest cells from early mouse embryos spanning gastrulation (E7.5) to the beginning of organogenesis (E9.5). This technique is straightforward and combines lineage tracing, cell sorting, and gel-based size selection to prepare small RNA sequencing libraries from a minimal number of cells for next generation sequencing. We highlight the importance for strict somite stage matching of embryos to resolve 6-hour time intervals to obtain a comprehensive view of miRNAs during the rapid changes of early development. This method can be widely applied to genetic and developmental studies and avoids the pooling of embryos. We describe a way to overcome challenges of current methods such as miRNA enrichment using gel-based purification, library quantification, and minimizing bias introduced from PCR. This method has been used to identify miRNA expression patterns over time to study how miRNAs control developmental timing in the neural crest lineage of mouse embryos.

<table>
	<tbody>
		<tr>
			<td>#5 Forceps Fine Science Tools</td>
			<td>Fisher Scientific</td>
			<td>NC9277114</td>
			<td></td>
		</tr>
		<tr>
			<td>0.4% Trypan blue</td>
			<td>ThermoFisher</td>
			<td>15250061</td>
			<td></td>
		</tr>
		<tr>
			<td>10 cm Petri dishes</td>
			<td>Fisher Scientific</td>
			<td>07-202-011</td>
			<td></td>
		</tr>
		<tr>
			<td>2-Propanol</td>
			<td>Miilapore Sigma</td>
			<td>I9516-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>5 % Criterion TBE Polyacrylamide Gel, 12+2 well, 45 &#181;L</td>
			<td>Bio-Rad</td>
			<td>3450047</td>
			<td></td>
		</tr>
		<tr>
			<td>5200 Fragment Analyzer</td>
			<td>Agilent</td>
			<td>M5310AA</td>
			<td></td>
		</tr>
		<tr>
			<td>96 well PCR plates high profile semi skirted clear/clear</td>
			<td>Bio-Rad</td>
			<td>HSS9601</td>
			<td></td>
		</tr>
		<tr>
			<td>BD FACSAria II</td>
			<td>bdbiosciences</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin, fraction V</td>
			<td>Fisher Scientific</td>
			<td>BP1600100</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Phosphate-Buffered Saline (DPBS)</td>
			<td>Gendepot</td>
			<td>CA008-300</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf tubes</td>
			<td>Fisher Scientific</td>
			<td>05-408-129</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol Pure, 200 proof anhydrous</td>
			<td>Sigma Aldrich</td>
			<td>E7023-500ml</td>
			<td></td>
		</tr>
		<tr>
			<td>FC-404-2001</td>
			<td>Illumina</td>
			<td>FC-404-2001</td>
			<td></td>
		</tr>
		<tr>
			<td>HS NGS Fragment kit</td>
			<td>Agilent</td>
			<td>DNF-474-0500</td>
			<td></td>
		</tr>
		<tr>
			<td>HS RNA Kit</td>
			<td>Agilent</td>
			<td>DNF-472-0500</td>
			<td></td>
		</tr>
		<tr>
			<td>miRNeasy Mini Kit (50)</td>
			<td>Qiagen</td>
			<td>217004</td>
			<td></td>
		</tr>
		<tr>
			<td>NEXTflex Small RNA-Seq Kit v3 (48 barcodes)</td>
			<td>Fisher Scientific</td>
			<td>NC1289113</td>
			<td></td>
		</tr>
		<tr>
			<td>NextSeq 500/550 Mid Output Kit v2 (150 cycles)</td>
			<td>Illumina</td>
			<td>FC-404-2001</td>
			<td></td>
		</tr>
		<tr>
			<td>NGS Fragment kit</td>
			<td>Agilent</td>
			<td>DNF-473-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Papain</td>
			<td>Worthington</td>
			<td>LK003176</td>
			<td>resuspend in 5 mL of Ca/Mg free PBS for a final concentration of 27 U/mL</td>
		</tr>
		<tr>
			<td>SYBR Gold nucleic acid gel stain</td>
			<td>ThermoFisher</td>
			<td>S11494</td>
			<td></td>
		</tr>
		<tr>
			<td>Trizol LS</td>
			<td>ThermoFisher</td>
			<td>10296028</td>
			<td></td>
		</tr>
		<tr>
			<td>UVP Benchtop UV Transilluminators: Single UV</td>
			<td>Fisher Scientific</td>
			<td>UVP95044701</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas Scissors Straight Fine Science Tools #91500-09</td>
			<td>Fisher Scientific</td>
			<td>NC9609583</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Using the procedure demonstrated here, we have harvested embryos at E7.5, E8.5, and E9.5. Extraembryonic structures were removed from all embryos and then embryos were somite staged to resolve 6-hour time intervals (Figure 1A-1B). Using principle component analysis to group samples based on similarity, we find that samples cluster by age, highlighting the variation as a result of embryonic age and the need for careful somite matching for biological replicates (Figure 1C). We profiled the pluripotent epiblast at E7.5, lineage traced premigratory and migratory neural crest cells using Wnt1-Cre at E8.5 and migratory neural crest cells using Sox10-Cre at E9.5 (Figure 1D). Here we specifically harvest the cranial neural crest by decapitating the embryo just above the otic placode. A comparison of the two Cre-drivers (Wnt1 and Sox10) that are frequently used to label neural crest cells confirms they mark different populations in early mouse embryos (Figure 1E). Gating strategies were used to obtain live single RFP positive cells which were sorted directly into RNA extraction lysis solution (see Table of Materials) and stored at -80 &#176;C (Figure 1F). It is important to keep track of how many cells were harvested from each sample.
RNA isolation was performed using a modified version of the RNA kit protocol. Specifically, we use the mini RNA columns that are to be stored at 4 &#176;C. These columns are useful for eluting in a small volume (11 &#181;L) to obtain the highest possible concentration of RNA from samples where cell input is limiting. For this same reason, it is important to obtain all of the aqueous phase after the phenol chloroform extraction. Slow pipetting and tilting the tube to one side while collecting are critical to maximize yield. In this procedure, we quantify the RNA using both a spectrophotometer, to obtain information on salt/protein contamination, and a capillary electrophoresis to measure concentration (Figure 2). The spectrophotometer trace reveals that RNA was isolated with no contaminating proteins but has high salt content (Figure 2A-2B). Ideally the 260/280 ratio should be ~1.8 and the 260/230 ratio should be &#62;2.0. The total RNA yield as measured by the spectrophotometer did not increase with age between E8.5-E9.5. This is due to the resolution of the spectrophotometer not being sensitive enough to detect changes in RNA concentration from the number of cells that we are harvesting, and we recommend using the concentration information obtained from the capillary electrophoresis (Figure 2C). The capillary electrophoresis trace can be used to estimate concentration and size of the RNA fragments. Peaks &#60;1000 nucleotides are indicative of degradation. The representative trace is consistent with RNA that is not degraded. Peaks at 2000 nucleotides and 5100 nucleotides are 18s and 28s rRNA, respectively. The small RNA region is located at ~150 nucleotides (Figure 2D).
Small RNA sequencing libraries were prepped using the small RNA library prep kit described in the Table of Materials. Here we use just less than half of the RNA obtained from each sample (4 &#181;L of the 11 &#181;L elution, ~80-120 ng) that enables RNA-sequencing libraries to be synthesized from the remaining RNA from each sample. Here we dilute the 3&#8217; and 5&#8217; small RNA adapters &#188; to lower the amount of adapter dimers present and suggest that the adapter ligation, cleanup, and reverse transcription steps are completed as much as possible in a continuous manner. We have used 16 PCR cycles to amplify the libraries and suggest that the minimum number of PCR cycles for any experiment be empirically determined. By completing excess PCR cycles, one could artificially inflate the read count of a lowly expressed miRNA.
Size selection is imperative to enrich for libraries of miRNAs and not adapter dimers, which are typically present. The library product size (150 bp) and adapter dimers (130 bp) are very similar in size. Gel extraction is used to isolate the small RNA sequencing libraries away from the adapter dimers (Figure 3). An image of a PAGE gel before excision shows that a multitude of product sizes are present in each sample (Figure 3A). It is important to leave one lane open between any two samples or a ladder that is loaded onto the gel. The migration front at the bottom of the gel is slightly curved indicating that running at a slower speed may be necessary if the 150 bp band is difficult to distinguish for all samples. This representative image also shows an abundance of 150 bp product from the higher concentration of positive control RNA (total RNA from the brain of a rat) that went into the library prep as compared to that which went into each embryonic sample. The negative control reveals that the reagents were free of contaminating nucleic acid species (Figure 3A). Excision of the 150 bp band should be done with a clean razor blade and the area collected is shown in Figure 3B. Capillary electrophoresis traces before and after size selection show the dramatic improvement of the purity of the 150 bp library product with gel purification (Figure 3C-3D). It is important to note that the traces in Figure 3C-3D have sample peaks higher than the markers, indicative of overloading. In these cases, the size of the fragments can be accurate, however the concentration may not. Quantification can be obtained by diluting the libraries into the range of the markers or using alternative methods. We find some variation across alternative methods of library quantification and recommend that multiple methods of quantification be empirically tested. Accurate quantification of concentration is essential when maximizing the number of samples to be sequenced at one time. Libraries will be diluted down to 1.3 pM for sequencing and generally anywhere from 15-25 samples can be sequenced at one time on a 150 cycle sequencing kit that provides ~140 million reads. This results in about 5 million reads per sample. Generally mapping rates are between 60-80%.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61086/61086fig01.jpg" /> 
Figure 1: Harvesting cells from mouse embryos for small RNA sequencing.&#160;(A) E8.5 mouse embryo to highlight the removal of the yolk sac and somites used to determine the stage of the embryo (B) Somite staging of mouse embryos to capture the stages of neural crest development (C) Principle component analysis of libraries prepped from sorted wildtype neural crest cells showing that samples group by age (D) Schematic showing how samples were harvested using Wnt1-Cre at E8.5 to label premigratory and migratory neural crest cells and Sox10-Cre at E9.5 to label only migratory neural crest cells (E) Principle component analysis of libraries prepped from sorted wildtype neural crest cells showing samples group together by Cre-driver regardless of age (F) Gating strategy used to isolate RFP+ neural crest cells from E8.5 and E9.5 embryos using FACS sorting. <a href="https://www.jove.com/files/ftp_upload/61086/61086fig01large.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/61086/61086fig02.jpg" /> 
Figure 2: RNA isolation from E7.5-E9.5 mouse embryos.&#160;(A) Representative spectrophotometer trace of an RNA isolation from the youngest stage of embryo that we have applied this protocol. (B) Table showing the representative values of RNA quality obtained from the spectrophotometer. (C) Example of the RNA harvested from sorted neural crest cells from embryos of each age (D) Capillary electrophoresis trace showing good quality RNA. <a href="https://www.jove.com/files/ftp_upload/61086/61086fig02large.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/61086/61086fig03.jpg" /> 
Figure 3: miRNA libraries before and after size selection.&#160;(A) TBE-PAGE gel showing representative small RNA libraries for four samples and controls before and (B) after 150 bp band excision. The arrows represent the 150 bp band to be excised (C) Capillary electrophoresis trace of small RNA library before and (D) after size selection. <a href="https://www.jove.com/files/ftp_upload/61086/61086fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Preparation of Small RNA Libraries for Sequencing from Early Mouse Embryos at JoVE.com

Preparation of Small RNA Libraries for Sequencing from Early Mouse Embryos

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 ...

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JoVE Journal

Developmental Biology

5.3K Views.  Baylor College of Medicine. 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.

Video: Preparation of Small RNA Libraries for Sequencing from Early Mouse Embryos

Methods for Precisely Localized Transfer of Cells or DNA into Early Postimplantation Mouse Embryos

Manipulation and culture of early mouse embryos is a powerful yet largely under-utilized technology enhancing the value of this model system. Conversely, cell culture has been widely used in developmental biology studies. However, it is important to determine whether in vitro cultured cells truly represent in vivo cell types. Grafting cells into embryos, followed by an assessment of their contribution during development is a useful method to determine the potential of in vitro cultured cells. In this study, we describe a method for grafting cells into a defined site of early postimplantation mouse embryos, followed by ex vivo culture. We also introduce an optimized electroporation method that uses glass capillaries of known diameter, allowing precise localization and adjustment of the number of cells receiving exogenous DNA with both high transfection efficiency and low cell death. These techniques, which do not require any specialized equipment, render experimental manipulations of the gastrulation and early organogenesis-stage mouse embryo possible, allowing analysis of commitment in cultured cell subpopulations and the effect of genetic manipulations in situ on cell differentiation.

We demonstrate a method for grafting cultured cells into defined sites of early mouse embryos to determine their in vivo potential. We also introduce an optimized electroporation method that uses glass capillaries of known diameter, allowing the precise delivery of exogenous DNA into a few cells in the embryos.

Manipulation and culture of early mouse embryos is a powerful yet largely under-utilized technology enhancing the value of this model system. Conversely, ...

Loss- and Gain-of-function Approach to Investigate Early Cell Fate Determinants in Preimplantation Mouse Embryos

Gene silencing and overexpression techniques are instrumental for the identification of genes involved in embryonic development. Direct target gene modification in preimplantation embryos provides a means to study the underlying mechanisms of genes implicated in, for instance, cellular differentiation into the trophectoderm (TE) and the inner cell mass (ICM). Here, we describe a protocol that examines the role of neogenin as an authentic receptor for initial cell fate determination in preimplantation mouse embryos. First, we discuss the experimental manipulations that were used to produce gain and loss of neogenin function by microinjecting neogenin cDNA and shRNA; the effectiveness of this approach was confirmed by a strong correlation between the pair-wise expression levels of either red fluorescent protein (RFP) or green fluorescent protein (GFP) and the immunocytochemical quantification of neogenin expression. Secondly, overexpression of neogenin in preimplantation mouse embryos leads to normal ICM development while neogenin knockdown causes the ICM to develop abnormally, implying that neogenin could be a receptor that relays extracellular cues to drive blastomeres to early cell fates. Given the success of this detailed protocol in investigating the function of a novel embryonic developmental stage-specific receptor, we propose that it has the potential to aid in exploration and identification of other stage-specific genes during embryogenesis.

The goal of this protocol is to describe a loss- and gain-of function method that is applicable to identify neogenin as a stage-specific receptor that leads to trophectoderm and inner cell mass differentiation in preimplantation mouse embryos.

 Gene silencing and overexpression techniques are instrumental for the identification of genes involved in embryonic development. Direct target gene ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and behave by observing the surface of a tissue. This method provides a means to visualize the complex 3-dimensional vascular architecture of the lung.
To accomplish this, a catheter is inserted into the pulmonary artery and the vasculature is simultaneously flushed of blood and chemically dilated to limit resistance. Lungs are then inflated through the trachea at a standard pressure and the polymer compound is infused into the vascular bed at a standard flow rate. Once the entire arterial network is filled and allowed to cure, the lung vasculature may be visualized directly or imaged on a micro-CT (&#181;CT) scanner.
When performed successfully, one can appreciate the pulmonary arterial network in mice ranging from early postnatal ages to adults. Additionally, while demonstrated in the pulmonary arterial bed, this method can be applied to any vascular bed with optimized catheter placement and endpoints.

The aim of this technique is ex vivo visualization of pulmonary arterial networks of early postnatal and adult mice through lung inflation and injection of a radio-opaque polymer-based compound via the pulmonary artery. Potential applications for casted tissues are also discussed.

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and ...

En Face Endocardial Cushion Preparation for Planar Morphogenesis Analysis in Mouse Embryos

The study of the cellular and molecular mechanisms underlying the development of the mammalian heart is essential to address human congenital heart disease. The development of the primitive cardiac valves involves the epithelial-to-mesenchymal transition (EMT) of endocardial cells from the atrioventricular canal (AVC) and outflow tract (OFT) regions of the heart in response to local inductive myocardial and endocardial signals. Once the cells delaminate and invade the extracellular matrix (cardiac jelly) located between the endocardium and the myocardium, the primitive endocardial cushions (EC) are formed. This process implies that the endocardium has to fill the gaps left by the delaminated cells and has to reorganize itself to converge (narrow) or extend (lengthen) along an axis. Current research has implicated the planar cell polarity (PCP) pathway in regulating the subcellular localization of the factors involved in this process. Classically, the initial phases of cardiac valve development have been studied in cross-sections of embryonic hearts or in ex vivo AVC or OFT explants cultured on collagen gels. These approaches allow the analysis of apico-basal polarity but do not allow the analysis of cell behavior within the plane of the epithelium or of the morphological changes of migrating cells. Here, we show an experimental approach that allows the visualization of the endocardium at valvulogenic regions as a planar field of cells. This experimental approach provides the opportunity to study PCP, planar topology, and intercellular communication within the endocardium of the OFT and AVC during valve development. Deciphering new cellular mechanisms involved in cardiac valve morphogenesis may contribute to understanding congenital heart disease associated with endocardial cushion defects.

En Face Endocardial Cushion Preparation for Planar Morphogenesis Analysis in Mouse Embryos

Classically, the endocardium of the mouse embryonic valve primordium has been analyzed using transversal, coronal, or sagittal sections. Our novel approach for en face, two-dimensional imaging of the endocardium at valvulogenic regions allows planar polarity and cell rearrangement analysis of the endocardium during valve development.

The study of the cellular and molecular mechanisms underlying the development of the mammalian heart is essential to address human congenital heart ...

Processing Gut Cells from Zebrafish Larvae for Single-Cell RNA Sequencing

Gut Isolation from Zebrafish Larvae for Single-cell RNA Sequencing

The gastrointestinal (GI) tract performs a range of functions essential for life. Congenital defects affecting its development can lead to enteric neuromuscular disorders, highlighting the importance to understand the molecular mechanisms underlying GI development and dysfunction. In this study, we present a method for gut isolation from zebrafish larvae at 5 days post fertilization to obtain live,&#160;viable cells which can be used for single-cell RNA sequencing (scRNA-seq) analysis. This protocol is based on the manual dissection of the zebrafish intestine, followed by enzymatic dissociation with papain. Subsequently, cells are submitted to fluorescence-activated cell sorting, and viable cells are collected for scRNA-seq. With this method, we were able to successfully identify different intestinal cell types, including epithelial, stromal, blood, muscle, and immune cells, as well as enteric neurons and glia. Therefore, we consider it to be a valuable resource for studying the composition of the GI tract in health and disease, using the zebrafish.

Author Spotlight: Isolating and Analyzing Intestinal Cells of Zebrafish Larvae for Investigating Transcriptomic Aspects of Gastrointestinal Development 

Here, we describe a method for gut isolation from zebrafish larvae at 5 days post fertilization, for single-cell RNA sequencing analysis.

The gastrointestinal (GI) tract performs a range of functions essential for life. Congenital defects affecting its development can lead to enteric ...

Single Cell and Nuclei Extraction from Human Brain Organoids

Generation and Downstream Analysis of Single-Cell and Single-Nuclei Transcriptomes in Brain Organoids

Over the past decade, single-cell transcriptomics has significantly evolved and become a standard laboratory method for simultaneous analysis of gene expression profiles of individual cells, allowing the capture of cellular diversity. In order to overcome limitations posed by difficult-to-isolate cell types, an alternative approach aiming at recovering single nuclei instead of intact cells can be utilized for sequencing, making transcriptome profiling of individual cells universally applicable. These techniques have become a cornerstone in the study of brain organoids, establishing them as models of the developing human brain. Leveraging the potential of single-cell and single-nucleus transcriptomics in brain organoid research, this protocol presents a step-by-step guide encompassing key procedures such as organoid dissociation, single-cell or nuclei isolation, library preparation&#160;and sequencing. By implementing these alternative approaches, researchers can obtain high-quality datasets, enabling the identification of neuronal and non-neuronal cell types, gene expression profiles, and cell lineage trajectories. This facilitates comprehensive investigations into cellular processes and molecular mechanisms shaping brain development.

Author Spotlight: Integrating Organoid Models with Single-Cell and Spatial Transcriptomics Technologies

Here, we introduce a comprehensive protocol for the generation and downstream analysis of human brain organoids using single-cell and single-nucleus RNA sequencing.

Over the past decade, single-cell transcriptomics has significantly evolved and become a standard laboratory method for simultaneous analysis of gene ...

Isolation and Genotyping of Gastrulating Mouse Embryo for Single-Cell Sequencing

Single-Cell RNA Sequencing of Mutant Whole Mouse Embryos: From the Epiblast to the End of Gastrulation

Over the last decade, single-cell approaches have become the gold standard for studying gene expression dynamics, cell heterogeneity, and cell states within samples. Before single-cell advances, the feasibility of capturing the dynamic cellular landscape and rapid cell transitions during early development was limited. In this paper, a robust pipeline was designed to perform single-cell and nuclei analysis on mouse embryos from embryonic day E6.5 to E8, corresponding to the onset and completion of gastrulation. Gastrulation is a fundamental process during development that establishes the three germinal layers: mesoderm, ectoderm, and endoderm, which are essential for organogenesis. Extensive literature is available on single-cell omics applied to wild-type perigastrulating embryos. However, single-cell analysis of mutant embryos is still scarce and often limited to FACS-sorted populations. This is partially due to the technical constraints associated with the need for genotyping, timed pregnancies, the count of embryos with desired genotypes per pregnancy, and the number of cells per embryo at these stages. Here, a methodology is presented designed to overcome these limitations. This method establishes breeding and timed pregnancy guidelines to achieve a higher chance of synchronized pregnancies with desired genotypes. Optimization steps in the embryo isolation process coupled with a same-day genotyping protocol (3 h) allow for microdroplet-based single-cell to be performed on the same day, ensuring the high viability of cells and robust results. This method further includes guidelines for optimal nuclei isolations from embryos. Thus, these approaches increase the feasibility of single-cell approaches of mutant embryos at the gastrulation stage. We anticipate that this method will facilitate the analysis of how mutations shape the cellular landscape of the gastrula.

Author Spotlight: A Pipeline to Analyze Lineage-Specific Mutant Embryos at Single-Cell Resolution

This paper establishes a pipeline for high-quality single-cell and nuclei suspensions of gastrulating mouse embryos for sequencing of single cells and nuclei.

Over the last decade, single-cell approaches have become the gold standard for studying gene expression dynamics, cell heterogeneity, and cell states ...