We thank Taylor AuBuchon-Elder, Jordan Teisher, and Kristina Zudock for assistance in sampling herbarium specimens, and the Missouri Botanical Garden for access to herbarium specimens for destructive sampling. This work was support by a grant from the National Science Foundation (DEB-1457748).

1. Prior to Start
<ol>
	<li>Make fresh cetyl trimethylammonium bromide (CTAB) buffer17 by adding 20 g of CTAB, 10 g of polyvinylpyrrolidone (PVP) 40, 100.0 mL 1 M Tris pH 8.0, 40 mL of 0.5 M ethylenediaminetetraacetic acid (EDTA) pH 8.0, 280.0 mL of 5 M NaCl, and 400.0 mL of reagent water together, and bring the total volume to 1 L using reagent-grade water. Adjust the pH to 8.0. 
	NOTE: Additional reagents can be added to CTAB depending on secondary compounds in individual taxa. See Allen et al.18 for a thorough list of additive reagents.</li>
	<li>Add 10 &#181;L of &#946;-mercaptoethanol per 5 mL of CTAB buffer. 
	NOTE: this can be prepared in batches of 50 mL and stored at room temperature for 3&#8211;4 weeks.
	<ol>
		<li>Heat the CTAB solution in a 65 &#176;C water bath.</li>
	</ol>
	</li>
	<li>Chill mortars and pestles in -20 &#176;C for at least 20 min.</li>
	<li>Label 4 sets of 2 x n 2-mL tubes (where n = the number of samples). Put 1 set of the labeled tubes on ice.</li>
	<li>Chill isopropanol on ice or in a -20 &#176;C freezer.</li>
	<li>Remove solid phase reversible immobilization (SPRI) beads (see Table of Materials) from the fridge, and allow them to equilibrate to room temperature (at least 30 min).</li>
	<li>Prepare 80% ethanol.</li>
	<li>Select herbarium specimens for extraction and retrieve ~1 cm2, or 10 mg, of tissue per specimen, preferably leaf material.</li>
</ol>
2. DNA Extraction
<ol>
	<li>Grind ~1 cm2 of preselected herbarium tissue using prechilled mortars and pestles. Add liquid nitrogen and 30&#8211;50 mg sterilized sand. Grind until tissue turns into fine powder. 
	NOTE: 10 mg or more tissue is desirable, but less has also worked in some cases. Once the liquid nitrogen evaporates, add more as needed until tissue is fully ground. Another common method for disrupting cells and tissues is the use of a bead beater. However, this method was found to not work well for the specimens used in these experiments.</li>
	<li>Transfer the frozen powder into two 2 mL tubes (add no more than half of the tube&#39;s volume). Add 600 &#181;L of warm CTAB solution to each tube and mix the tubes thoroughly by inversion and vortexing. 
	NOTE: Since the quantity and quality of herbarium material is often low, performing DNA isolation in two repetitions helps to obtain higher yields.</li>
	<li>Incubate the samples in a 65 &#176;C water bath for 1&#8211;1.5 h, vortexing every 15 min.</li>
	<li>Centrifuge samples at 10,000 x g for 5 min. Transfer the supernatant to a fresh set of labeled tubes (~500 &#181;L). Discard the pellet using a standard non-chlorinated disposal procedure.</li>
	<li>Add 4 &#181;L of RNase-A (10 mg/mL) to each tube and mix by inverting or pipetting. Incubate the samples at 37 &#176;C in a heat block or water bath for 15 min.</li>
	<li>Add an equal volume (~500 &#181;L) of a 25:24:1 phenol:chloroform:isoamyl alcohol mixture once the tubes have reached room temperature. Mix thoroughly by pipetting up and down and/or with inversion. Centrifuge the tubes at 12,000 x g for 15 min. Transfer the aqueous layer (upper layer) to a fresh set of labeled tubes (~400 &#181;L). Discard the organic layer into a chlorinated waste container. 
	NOTE: Step&#160;2.6 may be repeated if large quantities of secondary compounds are expected in plant material.</li>
	<li>Add an equal volume (~400 &#181;L) of a 24:1 chloroform:isoamyl alcohol mixture. Mix thoroughly by pipetting up and down and/or with inversion. Centrifuge the tubes at 12,000 x g for 15 min. Transfer the aqueous layer (upper layer) to a fresh set of prechilled, labeled tubes (~300 &#181;L). Discard the organic layer into a chlorinated waste container.</li>
	<li>Add an equal volume (~300 &#181;L) of prechilled isopropanol and 12 &#181;L of 2.5 M sodium acetate to each tube. Incubate samples at -20 &#176;C for 30&#8211;60 min. 
	NOTE: The incubation times can be extended (up to overnight incubation), but DNA quality will decrease the longer the samples incubate.</li>
	<li>Take the samples out of the freezer and centrifuge the tubes at 12,000 x g for 15 min. Remove and discard the supernatant gently without disturbing the pellet. Wash the pellet by suspension with fresh 70% ethanol (approximately 300&#8211;500 &#181;L). For each doubled sample, consolidate the two individual pellets into one with accompanying ethanol. 
	NOTE: Samples should be consolidated into one tube using ethanol first and then proceed. It is not necessary to wash each sample with ethanol separately.</li>
	<li>Centrifuge the tubes at 12,000 x g for 10 min. Remove and discard the supernatant gently without disturbing the pellet. Air dry the pellets. 
	NOTE: Samples can be dried faster using a dry heat block (do not exceed 65 &#176;C). Make sure that the samples do not over-dry, as this can decrease the final yield of DNA.</li>
	<li>Suspend the isolated DNA in 50 &#181;L of 1x TE. Store in the -20 &#176;C freezer for long term storage or 4 &#176;C for use in the following week.</li>
</ol>
3. Quality Control (QC)
<ol>
	<li>Run an agarose gel for quality check.
	<ol>
		<li>Prepare a 1x Tris/Borate/EDTA (TBE) buffer by adding 54 g Tris base, 27.5 g boric acid, and 3.75 g EDTA disodium salt, bringing the total volume to 5 L using reagent grade water.</li>
		<li>Prepare a 1% agarose gel by adding 1 g agarose to 100 mL of 1x TBE. Microwave the solution until no agarose is visible. Add 0.01% nucleic acid gel stain (see Table of Materials). Let the flask cool until it is warm to the touch. Mix well by stirring. Pour agarose in a gel tray and let it sit until it solidifies.</li>
		<li>Mix 3 &#181;L of sample, 2 &#181;L of reagent grade water, and 1 &#181;L of 6x loading dye. Load the samples in the gel matrix, noting their order.</li>
		<li>Run the samples for 60&#8211;70 min at 60&#8211;70 V. Image the gel under UV light with correct exposure and focus. 
		NOTE: Presence of a clear high molecular weight band is a sign of high quality DNA, while smears usually indicate DNA degradation. Most herbarium specimens are degraded.</li>
	</ol>
	</li>
	<li>Run a dsDNA quantification analysis (see Table of Materials) to determine the quantity of double stranded DNA.
	<ol>
		<li>Use 2 &#181;L of sample for analysis. 
		NOTE: Dilutions are not needed for the quantification analysis of herbarium material as they tend to be in minimal amounts. Successful libraries have been made from as little as 1.26 ng total dsDNA from this step.</li>
	</ol>
	</li>
</ol>
4. DNA Shearing
NOTE: This is an optimized version of a commercial double-stranded fragmentase protocol (see Table of Materials).
<ol>
	<li>Place dsDNA fragmentation enzyme on ice after vortexing for 3 s.</li>
	<li>In a sterile 0.2 mL polymerase chain reaction (PCR) tube, mix 1&#8211;16 &#181;L isolated DNA with 2 &#181;L of accompanying fragmentation reaction buffer. Bring the total volume to 18 &#181;L by adding nuclease free water. Add 2 &#181;L of dsDNA fragmentation enzyme and vortex the mixture for 3 s. 
	NOTE: The amount of DNA needed varies depending on DNA concentration (aim for 200 ng total in the tube).</li>
	<li>Incubate the samples at 37 &#176;C for 8.5 min. Then add 5 &#181;L of 0.5 M EDTA to the tubes. 
	NOTE: This step needs to be performed as soon as the incubation period is over to terminate the reaction and prevent DNA samples from over-shearing.</li>
</ol>
5. Bead Clean-up
<ol>
	<li>Homogenize the SPRI beads by vortexing.</li>
	<li>Bring the total volume of the sheared DNA to 50 &#181;L by adding 25 &#181;L of nuclease free water. Add 45 &#181;L of room temperature SPRI beads (90% volume) to 50 &#181;L of sheared DNA and mix thoroughly by pipetting up and down. 
	NOTE: Adding beads at 90% of total sample volume is done to remove the smallest of DNA fragments, often below 200 base pairs.</li>
	<li>Let the samples incubate for 5 min. Put the tubes on a magnetic plate and let them sit for 5 min. Carefully remove and discard the supernatant. 
	NOTE: Be careful not to disturb the beads, as they contain the desired DNA targets.</li>
	<li>Add 200 &#181;L of fresh 80% ethanol to the tubes while on the magnetic stand. Incubate at room temperature for 30 s and then carefully remove and discard the supernatant. Repeat this step once. Air dry the beads for 5 min while the tube is on the magnetic stand with its lid open. 
	NOTE: Avoid over-drying the beads. This can result in lower recovery of DNA.</li>
	<li>Remove the tubes from the magnet. Elute the DNA from the beads into 55 &#181;L of 0.1x TE and mix thoroughly by pipetting up and down. Incubate at room temperature for 5 min. Place tubes on the magnetic stand and wait for the solution to turn clear (~2 min).</li>
	<li>Pull off 52 &#181;L of the supernatant. Run a DNA quantification analysis on the samples to check the recovery and the initial concentration that goes into library prep. 
	NOTE: Libraries have been made with total dsDNA estimated to be less than 1.25 ng, though in each case reamplification was required.</li>
</ol>
6. Library Preparation
NOTE: This is a modified version of a commercially available library kit (see Table of Materials protocol).
<ol>
	<li>End Prep
	<ol>
		<li>Add 3 &#181;L of endonuclease and phosphate tailing enzymes, and 7 &#181;L of accompanying reaction buffer to 50 &#181;L of cleaned, sheared DNA. Mix thoroughly by pipetting up and down. Spin the tubes to remove bubbles. 
		NOTE: The total volume should be 60 &#181;L.</li>
		<li>Place the samples in a thermocycler with the following program: 30 min at 20 &#176;C, 30 min at 65 &#176;C, then hold at 4 &#176;C. 
		NOTE: Heated lid was set to &#8805;75 &#176;C</li>
	</ol>
	</li>
	<li>Adaptor Ligation
	<ol>
		<li>Dilute the adaptor 25&#8211;50 fold (working adaptor concentration of 0.6&#8211;0.3 &#181;M). Add 30 &#181;L of ligation master mix, 1 &#181;L of ligation enhancer, and 2.5 &#181;L of adaptor for high-throughput short read sequencing to the tubes. 
		NOTE: The total volume should be 93.5 &#181;L.</li>
		<li>Mix thoroughly by pipetting up and down. Spin the tubes to remove bubbles. Incubate the tubes at 20 &#176;C for 15 min.</li>
		<li>Add 3 &#181;L of commercial mixture of uracil DNA glycosylase (UDG) and the DNA glycosylase-lyase Endonuclease VIII (see Table of Materials) to the tubes. Ensure that the total volume is 96.5 &#181;L. Mix thoroughly and incubate at 37 &#176;C for 15 min using a thermocycler. 
		NOTE: The lid should be set to &#8805;47 &#176;C. The original version of the commercial protocol has size selection after the adaptor ligation step, followed by a bead cleanup as the final step. This protocol, which achieves higher yields, switches the order of these steps and implements size selection as a final step.</li>
	</ol>
	</li>
	<li>Cleanup to Remove Enzymes and Small Fragments
	<ol>
		<li>Homogenize magnetic beads by vortexing.</li>
		<li>Add 78 &#181;L of SPRI magnetic beads (80% volume) and mix thoroughly by pipetting up and down. 
		NOTE: Adding beads at 80% of total sample volume is done to remove the smallest DNA fragments, which are often shorter than 250 base pairs. The more stringent removal of small DNA fragments is to (i) remove surplus adaptors and (ii) emphasize amplification of larger fragments in the following steps.</li>
		<li>Let the samples incubate for 5 min. Put the tubes on a magnetic plate for 5 min. Carefully remove and discard the supernatant that contains the DNA outside the desired size range. 
		NOTE: Be careful not to disturb the beads that contain the desired DNA targets.</li>
		<li>Add 200 &#181;L of fresh 80% ethanol to the tubes while on the magnetic stand. Incubate at room temperature for 30 s and then carefully remove and discard the supernatant. Repeat this step once. Air dry the beads for 5 min while the tube is on the magnetic stand with lid open. 
		NOTE: Avoid over drying the beads. This can result in lower recovery of DNA.</li>
		<li>Remove the tubes from the magnet. Elute the DNA target from the beads by adding 17 &#181;L of 0.1x TE and mix thoroughly by pipetting up and down.</li>
		<li>Incubate at room temperature for 5 min. Place tubes on the magnetic stand and wait for the solution to turn clear (~2 min).</li>
		<li>Pull off 15 &#181;L of the supernatant.</li>
	</ol>
	</li>
	<li>PCR amplification
	<ol>
		<li>Add 25 &#181;L of high fidelity PCR master mix, 5 &#181;L of high-throughput short read sequencing library prep 5&#39; primer, and 5 &#181;L of high-throughput short read sequencing library prep 3&#39; primer, to 15 &#181;L of the cleaned adaptor-ligated DNA. 
		NOTE: Total volume should be 50 &#181;L.</li>
		<li>Mix well by vortexing. Place the samples into a thermocycler using the settings found in Table 1: Thermocycler amplification setting. 
		NOTE: A large number of cycles is needed due to the low quantity of input DNA.</li>
	</ol>
	</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cycle Step</td>
			<td>Temp.&#160;</td>
			<td>Time</td>
			<td>Cycles</td>
		</tr>
		<tr>
			<td>Initial Denaturation</td>
			<td>98 &#176;C</td>
			<td>30 s</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Denaturation</td>
			<td>98 &#176;C</td>
			<td>10 s</td>
			<td>12</td>
		</tr>
		<tr>
			<td>Annealing/Extension</td>
			<td>65 &#176;C</td>
			<td>75 s</td>
			<td>12</td>
		</tr>
		<tr>
			<td>Final Extension</td>
			<td>65 &#176;C</td>
			<td>5 min</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Hold</td>
			<td>4 &#176;C</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1: PCR protocol denaturation, annealing, and extension times and temperatures. Temperature and times were optimized for the reagents presented in this protocol. If reagents are altered, temperatures and times should be optimized again.
<ol start="5">
	<li>Size Selection for Desired Library Size 
	NOTE: This bead step will remove fragments both above and below the target range.
	<ol>
		<li>Homogenize SPRI beads by vortexing.</li>
		<li>Add 25 &#181;L (50% volume) of room temperature magnetic beads and mix thoroughly by pipetting up and down. Let the samples incubate for 5 min. Put the tubes on a magnetic plate and let them sit for 5 min. Carefully remove and transfer the supernatant to a new set of labeled tubes. 
		NOTE: This volume can be adjusted based on desired library size. The supernatant contains DNA fragments of the desired size. In the first bead incubation, the beads are binding larger library fragments. These are removed to focus on those in the 400&#8211;600 base pair range. The supernatant contains smaller fragments.</li>
		<li>Add 6 &#181;L of room temperature SPRI beads to the supernatant and mix thoroughly by pipetting up and down. Let the samples incubate for 5 min. Put the tubes on a magnetic plate and let them sit for 5 min. 
		NOTE: This volume can be adjusted based on desired library size in accordance with step 6.5.2.</li>
		<li>Carefully remove and discard the supernatant. 
		NOTE: Be careful not to disturb the beads that contain the desired DNA. In the second bead incubation, the beads are binding to the fragments left after the initial removal of the largest DNA fragments. This set of fragments is usually in the desired size range.</li>
		<li>Add 200 &#181;L of fresh 80% ethanol to the tubes while on the magnetic stand. Incubate at room temperature for 30 s, then carefully remove and discard the supernatant. Repeat. Air dry the beads for 5 min while the tube is on the magnetic stand with lid open. 
		NOTE: Avoid over-drying the beads. This can result in lower recovery of DNA.</li>
		<li>Remove the tubes from the magnet. Elute the DNA target from the beads into 33 &#181;L of 0.1x TE and mix thoroughly by pipetting up and down.</li>
		<li>Incubate at room temperature for 5 min. Place tubes on the magnetic stand and wait for the solution to turn clear (~2 min). Pull off 30 &#181;L of the supernatant and transfer to 2 mL tubes (see Table of Materials) for storage. 
		NOTE: The libraries can be kept at -20 &#730;C for long-term storage.</li>
	</ol>
	</li>
	<li>Quality control
	<ol>
		<li>Run a quality control test on the DNA Libraries. Refer to steps 3.1 and 3.2. 
		NOTE: For DNA libraries, run the gel for ~45 min at 96 V.</li>
	</ol>
	</li>
	<li>Library reamplification: Optional if library quantity is not sufficient. 
	NOTE: Samples with library concentrations below 10 nM can be reamplified using the following steps. Reamplification of low concentration libraries can achieve workable results for sequencing, but reamplification may cause a modest shift in base composition diversity, though gathered data (Table 3) suggest that this is negligible for certain metrics.
	<ol>
		<li>Dilute the universal reamplification primers 10-fold using 0.1x TE.</li>
		<li>Add 25 &#181;L of high fidelity PCR master mix, 5 &#181;L of diluted universal reamplification primer 1 (AATGATACGGCGACCACCGA), and 5 &#181;L of diluted universal reamplification primer 2 (CAAGCAGAAGACGGCATACGA) to 15 &#181;L of low concentration libraries. NOTE: Total volume should be at 50 &#181;L.</li>
		<li>Mix well by vortexing. Place the samples into a thermocycler using the settings found in Table 1: Thermocycler amplification setting 
		NOTE: The large number of cycles is needed due to low quantity of input DNA.</li>
		<li>Bead clean-up</li>
		<li>Homogenize SPRI beads by vortexing. Add 45 &#181;L of room temperature SPRI beads (90% volume) and mix thoroughly by pipetting up and down.</li>
		<li>Let the samples incubate for 5 min. Put the tubes on a magnetic plate for 5 min.</li>
		<li>Carefully remove and discard the supernatant that contains the unwanted DNA. 
		NOTE: Be careful not to disturb the beads that contain the desired DNA targets.</li>
		<li>Add 200 &#181;L of fresh 80% ethanol to the tubes while on the magnetic stand. Incubate at room temperature for 30 s, and then carefully remove and discard the supernatant. Repeat this step once.</li>
		<li>Air dry the beads for 5 min while the tube is on the magnetic stand with its lid open. 
		NOTE: Avoid over-drying the beads. This can result in lower recovery of the DNA target.</li>
		<li>Remove the tubes from the magnet. Elute the DNA target from the beads into 33 &#181;L of 0.1x TE and mix thoroughly by pipetting up and down.</li>
		<li>Incubate at room temperature for 5 min. Place tubes on the magnetic stand and wait for the solution to turn clear (~2 min).</li>
		<li>Pull off 30 &#181;L of the supernatant. The libraries can be stored at -20 &#176;C for long-term storage.</li>
	</ol>
	</li>
	<li>Quality control
	<ol>
		<li>Run a quality control test on the DNA Libraries. Refer to steps 3.1 and 3.2. For DNA libraries, run the gel for ~45 min at 96 V. 
		NOTE: If a double band is seen in the gel, this is likely a consequence of primer exhaustion from the reamplification step. The bands can be removed by repeating 6.9, but using only one cycle in the PCR program depicted in Table 1.</li>
	</ol>
	</li>
</ol>

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T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Herbarium+genomics:+skimming+and+plastomics+from+archival+specimens.">Herbarium genomics: skimming and plastomics from archival specimens.</a> Webbia. 72 (1), 35-45 (2017).</li><li>Doyle, J. J., Doyle, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rapid+DNA+isolation+procedure+for+small+quantities+of+fresh+leaf+tissue.">A rapid DNA isolation procedure for small quantities of fresh leaf tissue.</a> Phytochem Bul. 19, 11-15 (1987).</li><li>Allen, G. C., Flores-Vergara, M. A., Krasynanski, S., Kumar, S., Thompson, W. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+modified+protocol+for+rapid+DNA+isolation+from+plant+tissue+using+cetryltrimethylammonium+bromide.">A modified protocol for rapid DNA isolation from plant tissue using cetryltrimethylammonium bromide.</a> Nat Prot. 1, 2320-2325 (2006).</li><li>Twyford, A. D., Ness, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strategies+for+complete+plastid+genome+seqeuncing.">Strategies for complete plastid genome seqeuncing.</a> Mol Eco Resour. , (2016).</li><li>Aird, D., 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=Analyzing+and+minimizing+PCR+amplification+bias+in+Illumina+sequencing+libraries.">Analyzing and minimizing PCR amplification bias in Illumina sequencing libraries.</a> Genome Bio. 12 (2), R18 (2011).</li><li>Bolger, A. M., Lohse, M., Usadel, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Trimmomatic:+A+flexible+trimmer+for+Illumina+sequence+data.">Trimmomatic: A flexible trimmer for Illumina sequence data.</a> Bioinf. 30, 2114-2120 (2014).</li><li>Grigoriev, I. V., 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=MycoCosm+portal:+gearing+up+for+1000+fungal+genomes.">MycoCosm portal: gearing up for 1000 fungal genomes.</a> Nucl Acids Res. 42 (1), D699-D704 (2014).</li><li>Langmead, B., Salzberg, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+gapped-read+alignment+with+Bowtie+2.">Fast gapped-read alignment with Bowtie 2.</a> Nat Meth. 9 (4), 357-359 (2012).</li><li>Herbarium Genomics. Available from: <a target="_blank" href="https://github.com/mrmckain/">https://github.com/mrmckain/</a> (2017)</li><li>. Fast-Plast: Rapid de novo assembly and finishing for whole chloroplast genomes Available from: <a target="_blank" href="https://github.com/mrmckain/">https://github.com/mrmckain/</a> (2017)</li><li>McKain, M. R., McNeal, J. R., Kellar, P. R., Eguiarte, L. E., Pires, J. C., Leebens-Mack, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Timing+of+rapid+diversification+and+convergent+origins+of+active+pollination+within+Agavoideae+(Asparagaceae).">Timing of rapid diversification and convergent origins of active pollination within Agavoideae (Asparagaceae).</a> Am J Bot. 103 (10), 1717-1729 (2016).</li><li>McKain, M. R., Hartsock, R. H., Wohl, M. M., Kellogg, E. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Verdant:+automated+annotation,+alignment,+and+phylogenetic+analysis+of+whole+chloroplast+genomes.">Verdant: automated annotation, alignment, and phylogenetic analysis of whole chloroplast genomes.</a> Bioinf. , (2016).</li><li>Staton, S. E., Burke, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transposome:+A+toolkit+for+annotation+of+transposable+element+families+from+unassembled+sequence+reads.">Transposome: A toolkit for annotation of transposable element families from unassembled sequence reads.</a> Bioinf. 31 (11), 1827-1829 (2015).</li><li>Bao, W., Kojima, K. K., Kohany, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Repbase+Update,+a+database+of+repetitive+elements+in+eukaryotic+genomes.">Repbase Update, a database of repetitive elements in eukaryotic genomes.</a> Mobile DNA. 6 (1), 11 (2015).</li><li>. Transposons Available from: <a target="_blank" href="https://github.com/mrmckain/">https://github.com/mrmckain/</a> (2017)</li><li>Wei&#223;, C. L., 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=Temporal+patterns+of+damage+and+decay+kinetics+of+DNA+retrieved+from+plant+herbarium+specimens.">Temporal patterns of damage and decay kinetics of DNA retrieved from plant herbarium specimens.</a> Royal Soc Open Sci. 3 (6), 160239 (2016).</li><li>Sawyer, S., Krause, J., Guschanski, K., Savolainen, V., P&#228;&#228;bo, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+patterns+of+nucleotide+misincorporations+and+DNA+fragmentation+in+ancient+DNA.">Temporal patterns of nucleotide misincorporations and DNA fragmentation in ancient DNA.</a> PLoS ONE. 7 (3), e34131 (2012).</li><li>Head, S. R., 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=Library+construction+for+next-generation+sequencing:+overviews+and+challenges.">Library construction for next-generation sequencing: overviews and challenges.</a> BioTechniques. 56 (2), 61-64 (2014).</li><li>Grover, C. E., Salmon, A., Wendel, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+sequence+capture+as+a+powerful+tool+for+evolutionary+analysis.">Targeted sequence capture as a powerful tool for evolutionary analysis.</a> Am J Bot. 99, 312-319 (2012).</li></ol>

The authors declare they have no competing interests.

The protocol presented here is a comprehensive and robust method for DNA isolation and sequencing library preparation from dried plant specimens. The consistency of the method and minimal need to alter it based on specimen quality make it scalable for large herbarium-based sequencing projects. The inclusion of an optional reamplification step for low yield libraries allows the inclusion of low quality, low quantity, rare, or historically important samples that would otherwise not be suitable for sequencing.
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Herbarium collections are a potentially valuable source of both species and genomic diversity for studies including phylogenetics1,2,3, population genetics4,5, conservation biology6, invasive species biology7, and trait evolution8. The ability to obtain a rich diversity of species, populations, geographical locations, and time points highlights the &#34;treasure chest&#34;9 that is the herbarium. Historically, the degraded nature of herbarium-derived DNA has hindered PCR-based projects, often relegating researchers to using only markers found in high copy, such as regions of the chloroplast genome or the internal transcribed spacer (ITS) of the ribosomal RNA. Quality of specimens and DNA vary extensively based on methods of preservation9,10, with double-stranded breaks and fragmentation from heat used in the drying process being the most common forms of damage, creating the so-called 90% DNA lock-up that has encumbered PCR-based studies11. Aside from fragmentation, the second most prevalent issue in herbarium genomics is contamination, such as that derived from endophytic fungi13 or fungi acquired postmortem after collection but before mounting in the herbarium12, though this problem can be solved bioinformatically given the right fungal database (see below). A third, and less common, problem is sequence modification through cytosine deamination (C/G&#8594;T/A)14, although it is estimated to be low (~0.03%) in herbarium specimens11. With the advent of high-throughput sequencing (HTS), the issue of fragmentation can be overcome with short reads and sequencing depth12,15, allowing genomic-level data acquisition from numerous specimens with low quality DNA, and even sometimes permitting whole genome sequencing15.
Herbarium samples are becoming more frequently used and are a larger component of phylogenetic projects16. A current challenge of using herbarium specimens for HTS is consistently obtaining sufficient double stranded DNA, a necessary prerequisite for sequencing protocols, from numerous species in a timely manner, without needing to optimize methods for individual specimens. In this paper, a protocol for DNA extraction and library preparation of herbarium specimens is demonstrated that takes advantage of existing methods and modifies them to allow for fast and replicable results. This method allows for complete processing from specimen to a library of 24 samples in 13 h, with 8 h hands-on time, or 16 h, with 9 h hands-on time, when the optional reamplification step is required. Simultaneous processing of more samples is achievable, though the limiting factor is centrifuge capacity and technical skill. The protocol is designed to require only typical laboratory equipment (thermocycler, centrifuge, and magnetic stands) instead of specialized equipment, such as a nebulizer or sonicator, for shearing DNA.
DNA quality, fragment size, and quantity are limiting factors for the use of herbarium specimens in high-throughput sequencing experiments. Other methods for isolating herbarium DNA and creating high-throughput sequencing libraries have demonstrated the utility of using as little as 10 ng of DNA16; however they require experimentally determining the optimum number of PCR cycles required for library preparation. This becomes impractical when dealing with exceedingly small amounts of viable double stranded DNA (dsDNA), as some herbarium specimens produce only enough DNA for a single library preparation. The method presented here uses a single number of cycles regardless of sample quality, so no DNA is lost in library optimization steps. Instead, a reamplification step is invoked when libraries do not meet the minimum amounts needed for sequencing. Many herbarium samples are rare and possess little material making it difficult to justify destructive sampling in many cases. To counter this, the presented protocol allows dsDNA input sizes less than 1.25 ng into the library preparation process, expanding the scope of viable samples for high-throughput sequencing and minimizing the need for destructive sampling of specimens.
The following protocol has been optimized for grasses and tested on hundreds of different species from herbarium samples, although we expect that the protocol can be applied to many other plant groups. It includes an optional recovery step that can be used to save low quality and/or rare specimens. Based on over two hundred herbarium specimens tested, this protocol works on specimens with low tissue input and quality, allowing for the preservation of rare specimens through minimal destructive sampling. Here it is shown that this protocol can provide high quality libraries that can be sequenced for phylogenomics-based projects.

<table><tbody><tr><td>Veriti Thermal Cycler</td><td>Applied Biosystems</td><td>4452300</td><td>96 well&amp;nbsp;</td></tr><tr><td>Gel Imaging System</td><td>Azure Biosystems</td><td>c300</td><td></td></tr><tr><td>Microfuge 20 Series</td><td>Beckman Coulter</td><td>B30137</td><td></td></tr><tr><td>Digital Dry Bath</td><td>Benchmark Scientific</td><td>BSH1001</td><td></td></tr><tr><td>Electrophoresis System</td><td>EasyCast</td><td>B2</td><td></td></tr><tr><td>PURELAB flex 2 (Ultra pure water)</td><td>ELGA&amp;nbsp;</td><td>89204-092</td><td></td></tr><tr><td>DNA LoBind Tube&amp;nbsp;</td><td>Eppendorf</td><td>30108078</td><td>2 ml</td></tr><tr><td>Mini centrifuge</td><td>Fisher Scientific</td><td>12-006-901</td><td></td></tr><tr><td>Vortex-Genie 2</td><td>Fisher Scientific</td><td>12-812</td><td></td></tr><tr><td>Mortar</td><td>Fisher Scientific</td><td>S02591</td><td>porcelain</td></tr><tr><td>Pestle</td><td>fisher Scientific</td><td>S02595</td><td>porcelain</td></tr><tr><td>Centrifuge tubes</td><td>fisher Scientific</td><td>21-403-161</td><td></td></tr><tr><td>Microwave</td><td>Kenmore</td><td>405.7309231</td><td></td></tr><tr><td>Qubit Assay Tubes</td><td>Invitrogen</td><td>Q32856</td><td></td></tr><tr><td>0.2 ml Strip tube and Cap for PCR</td><td>VWR</td><td>20170-004</td><td></td></tr><tr><td>Qubit 2.0 Fluorometer</td><td>Invitrogen</td><td>Q32866</td><td></td></tr><tr><td>Balance</td><td>Mettler Toledo</td><td>PM2000</td><td></td></tr><tr><td>Liquid Nitrogen Short-term Storage</td><td>Nalgene</td><td>F9401</td><td></td></tr><tr><td>Magnetic-Ring Stand&amp;nbsp;</td><td>ThermoFisher Scientific&amp;nbsp;</td><td>AM10050</td><td>96 well&amp;nbsp;</td></tr><tr><td>Water Bath</td><td>VWR</td><td>89032-210</td><td></td></tr><tr><td>Hot Plate Stirrers</td><td>VWR</td><td>97042-754</td><td></td></tr><tr><td>Liquid Nitrogen</td><td>Airgas</td><td>UN1977</td><td></td></tr><tr><td>1 X TE Buffer</td><td>Ambion</td><td>AM9849</td><td>pH 8.0</td></tr><tr><td>CTAB</td><td>AMRESCO</td><td>0833-500G</td><td></td></tr><tr><td>2-MERCAPTOETHANOL</td><td>AMRESCO</td><td>0482-200ML</td><td></td></tr><tr><td>Ribonuclease A</td><td>AMRESCO</td><td>E866-5ML</td><td>10 mg/ml solution</td></tr><tr><td>Agencourt AMPure XP</td><td>Beckman Coulter</td><td>A63882</td><td></td></tr><tr><td>Sodium Chloride</td><td>bio WORLD</td><td>705744</td><td></td></tr><tr><td>Isopropyl Alcohol</td><td>bio WORLD</td><td>40970004-1</td><td></td></tr><tr><td>Nuclease Free water</td><td>bio WORLD</td><td>42300012-2</td><td></td></tr><tr><td>Isoamyl Alcohol</td><td>Fisher Scientific</td><td>A393-500</td><td></td></tr><tr><td>Sodium Acetate Trihydrate</td><td>Fisher Scientific</td><td>s608-500</td><td></td></tr><tr><td>LE Agarose</td><td>GeneMate</td><td>E-3120-500</td><td></td></tr><tr><td>100bp PLUS DNA Ladder</td><td>Gold Biotechnology</td><td>D003-500</td><td></td></tr><tr><td>EDTA, Disodium Salt</td><td>IBI Scientific</td><td>IB70182</td><td></td></tr><tr><td>Qubit dsDNA HS Assay Kit</td><td>Life Technologies</td><td>Q32854</td><td></td></tr><tr><td>TRIS</td><td>MP Biomedicals</td><td>103133</td><td>ultra pure</td></tr><tr><td>Gel Loading Dye Purple (6 X)</td><td>New England BioLabs</td><td>B7024S</td><td></td></tr><tr><td>NEBNext dsDNA Fragmentase</td><td>New England BioLabs</td><td>M0348L</td><td></td></tr><tr><td>NEBNext Ultra II DNA Library Prep Kit for Illumina&amp;nbsp;</td><td>New England BioLabs</td><td>E7645L</td><td></td></tr><tr><td>NEBNext Multiplex Oligos for Illumina</td><td>New England BioLabs</td><td>E7600S</td><td>Dual Index Primers Set 1</td></tr><tr><td>NEBNext Q5 Hot Start HiFi PCR Master Mix</td><td>New England BioLabs</td><td>M0543L</td><td></td></tr><tr><td>Mag-Bind RXNPure Plus</td><td>Omega bio-tek</td><td>M1386-02</td><td></td></tr><tr><td>GelRed 10000 X</td><td>Pheonix Research</td><td>41003-1</td><td></td></tr><tr><td>Phenol solution</td><td>SIGMA Life Science</td><td>P4557-400ml</td><td></td></tr><tr><td>PVP40</td><td>SIGMA-Aldrich</td><td>PVP40-50G</td><td></td></tr><tr><td>Chloroform</td><td>VWR</td><td>EM8.22265.2500</td><td></td></tr><tr><td>Ethanol</td><td>Koptec</td><td>V1016</td><td>200 Proof</td></tr><tr><td>Silica sand</td><td>VWR</td><td>14808-60-7</td><td></td></tr><tr><td>Reamplification primers</td><td>Integrated DNA Technologies</td><td>see text</td><td></td></tr><tr><td>Sequencher v.5.0.1</td><td>GeneCodes</td><td></td><td></td></tr></tbody></table>

robust dna isolation and high-throughput sequencing library construction for herbarium specimens

Herbaria are an invaluable source of plant material that can be used in a variety of biological studies. The use of herbarium specimens is associated with a number of challenges including sample preservation quality, degraded DNA, and destructive sampling of rare specimens. In order to more effectively use herbarium material in large sequencing projects, a dependable and scalable method of DNA isolation and library preparation is needed. This paper demonstrates a robust, beginning-to-end protocol for DNA isolation and high-throughput library construction from herbarium specimens that does not require modification for individual samples. This protocol is tailored for low quality dried plant material and takes advantage of existing methods by optimizing tissue grinding, modifying library size selection, and introducing an optional reamplification step for low yield libraries. Reamplification of low yield DNA libraries can rescue samples derived from irreplaceable and potentially valuable herbarium specimens, negating the need for additional destructive sampling and without introducing discernible sequencing bias for common phylogenetic applications. The protocol has been tested on hundreds of grass species, but is expected to be adaptable for use in other plant lineages after verification. This protocol can be limited by extremely degraded DNA, where fragments do not exist in the desired size range, and by secondary metabolites present in some plant material that inhibit clean DNA isolation. Overall, this protocol introduces a fast and comprehensive method that allows for DNA isolation and library preparation of 24 samples in less than 13 h, with only 8 h of active hands-on time with minimal modifications.

DNA Isolation and Final Library Yield 
In this study, the efficacy of the protocol for the isolation of herbarium DNA and the recovery of high quality sequencing libraries was demonstrated using fifty different samples with the oldest from 1920 and the youngest from 2012 (Table 2). For each sample, approximately 10 mg of leaf tissue was used for DNA isolation. Greener leaf tissue was favored if available, and no tissue with obvious fungal contaminati...

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Robust DNA Isolation and High-throughput Sequencing Library Construction for Herbarium Specimens

This article demonstrates a detailed protocol for DNA isolation and high-throughput sequencing library construction from herbarium material including rescue of exceptionally poor-quality DNA.

Herbaria are an invaluable source of plant material that can be used in a variety of biological studies. The use of herbarium specimens is associated with ...

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10.5K Views.  Donald Danforth Plant Science Center. This article demonstrates a detailed protocol for DNA isolation and high-throughput sequencing library construction from herbarium material including rescue of exceptionally poor-quality DNA.

Video: Robust DNA Isolation and High-throughput Sequencing Library Construction for Herbarium Specimens

Automated Gel Size Selection to Improve the Quality of Next-generation Sequencing Libraries Prepared from Environmental Water Samples

Next-generation sequencing of environmental samples can be challenging because of the variable DNA quantity and quality in these samples. High quality DNA libraries are needed for optimal results from next-generation sequencing. Environmental samples such as water may have low quality and quantities of DNA as well as contaminants that co-precipitate with DNA. The mechanical and enzymatic processes involved in extraction and library preparation may further damage the DNA. Gel size selection enables purification and recovery of DNA fragments of a defined size for sequencing applications. Nevertheless, this task is one of the most time-consuming steps in the DNA library preparation workflow. The protocol described here enables complete automation of agarose gel loading, electrophoretic analysis, and recovery of targeted DNA fragments. 
In this study, we describe a high-throughput approach to prepare high quality DNA libraries from freshwater samples that can be applied also to other environmental samples. We used an indirect approach to concentrate bacterial cells from environmental freshwater samples; DNA was extracted using a commercially available DNA extraction kit, and DNA libraries were prepared using a commercial transposon-based protocol. DNA fragments of 500 to 800 bp were gel size selected using Ranger Technology, an automated electrophoresis workstation. Sequencing of the size-selected DNA libraries demonstrated significant improvements to read length and quality of the sequencing reads.

This manuscript describes an automated gel size selection approach for purifying DNA fragments for next-generation sequencing. The Ranger Technology provides complete automation of the entire process of agarose gel loading, electrophoretic analysis, and recovery of targeted DNA fragments allowing for high-throughput and high quality next-generation sequencing libraries.

Next-generation sequencing of environmental samples can be challenging because of the variable DNA quantity and quality in these samples. High quality DNA ...

Environment

Purification of High Molecular Weight Genomic DNA from Powdery Mildew for Long-Read Sequencing

The powdery mildew fungi are a group of economically important fungal plant pathogens. Relatively little is known about the molecular biology and genetics of these pathogens, in part due to a lack of well-developed genetic and genomic resources. These organisms have large, repetitive genomes, which have made genome sequencing and assembly prohibitively difficult. Here, we describe methods for the collection, extraction, purification and quality control assessment of high molecular weight genomic DNA from one powdery mildew species, Golovinomyces cichoracearum. The protocol described includes mechanical disruption of spores followed by an optimized phenol/chloroform genomic DNA extraction. A typical yield was 7 &#181;g DNA per 150 mg conidia. The genomic DNA that is isolated using this procedure is suitable for long-read sequencing (i.e., &#62; 48.5 kbp). Quality control measures to ensure the size, yield, and purity of the genomic DNA are also described in this method. Sequencing of the genomic DNA of the quality described here will allow for the assembly and comparison of multiple powdery mildew genomes, which in turn will lead to a better understanding and improved control of this agricultural pathogen.

Described here is a method for the extraction, purification, and quality control of genomic DNA from the obligate biotrophic fungal pathogen, powdery mildew, for use in long-read genome sequencing.

The powdery mildew fungi are a group of economically important fungal plant pathogens. Relatively little is known about the molecular biology and genetics ...

Optimization and Comparative Analysis of Plant Organellar DNA Enrichment Methods Suitable for Next-generation Sequencing

Plant organellar genomes contain large, repetitive elements that may undergo pairing or recombination to form complex structures and/or sub-genomic fragments. Organellar genomes also exist in admixtures within a given cell or tissue type (heteroplasmy), and an abundance of subtypes may change throughout development or when under stress (sub-stoichiometric shifting). Next-generation sequencing (NGS) technologies are required to obtain deeper understanding of organellar genome structure and function. Traditional sequencing studies use several methods to obtain organellar DNA: (1) If a large amount of starting tissue is used, it is homogenized and subjected to differential centrifugation and/or gradient purification. (2) If a smaller amount of tissue is used (i.e., if seeds, material, or space is limited), the same process is performed as in (1), followed by whole-genome amplification to obtain sufficient DNA. (3) Bioinformatics analysis can be used to sequence the total genomic DNA and to parse out organellar reads. All these methods have inherent challenges and tradeoffs. In (1), it may be difficult to obtain such a large amount of starting tissue; in (2), whole-genome amplification could introduce a sequencing bias; and in (3), homology between nuclear and organellar genomes could interfere with assembly and analysis. In plants with large nuclear genomes, it is advantageous to enrich for organellar DNA to reduce sequencing costs and sequence complexity for bioinformatics analyses. Here, we compare a traditional differential centrifugation method with a fourth method, an adapted CpG-methyl pulldown approach, to separate the total genomic DNA into nuclear and organellar fractions. Both methods yield sufficient DNA for NGS, DNA that is highly enriched for organellar sequences, albeit at different ratios in mitochondria and chloroplasts. We present the optimization of these methods for wheat leaf tissue and discuss major advantages and disadvantages of each approach in the context of sample input, protocol ease, and downstream application.

The comparison and optimization of two plant organellar DNA enrichment methods are presented: traditional differential centrifugation and fractionation of the total gDNA based on methylation status. We assess the resulting DNA quantity and quality, demonstrate performance in short-read next-generation sequencing, and discuss the potential for use in long-read single-molecule sequencing.

Plant organellar genomes contain large, repetitive elements that may undergo pairing or recombination to form complex structures and/or sub-genomic ...

Empirical, Metagenomic, and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered &#34;bee-safe,&#34; there is mounting evidence that fungicide residues in pollen are associated with bee declines (for both honey and bumble bee species). While the mechanisms remain relatively unknown, researchers have speculated that bee-microbe symbioses are involved. Microbes play a pivotal role in the preservation and/or processing of pollen, which serves as nutrition for larval bees. By altering the microbial community, it is likely that fungicides disrupt these microbe-mediated services, and thereby compromise bee health. This manuscript describes the protocols used to investigate the indirect mechanism(s) by which fungicides may be causing colony decline. Cage experiments exposing bees to fungicide-treated flowers have already provided the first evidence that fungicides cause profound colony losses in a native bumble bee (Bombus impatiens). Using field-relevant doses of fungicides, a series of experiments have been developed to provide a finer description of microbial community dynamics of fungicide-exposed pollen. Shifts in the structural composition of fungal and bacterial assemblages within the pollen microbiome are investigated by next-generation sequencing and metagenomic analysis. Experiments developed herein have been designed to provide a mechanistic understanding of how fungicides affect the microbiome of pollen-provisions. Ultimately, these findings should shed light on the indirect pathway through which fungicides may be causing colony declines.

Microbial consortia within bumble bee hives enrich and preserve pollen for bee larvae. Using next generation sequencing, along with laboratory and field-based experiments, this manuscript describes protocols used to test the hypothesis that fungicide residues alter the pollen microbiome, and colony demographics, ultimately leading to colony loss.

Growers often use fungicide sprays during bloom to protect crops against disease, which exposes bees to fungicide residues. Although considered ...

Ultralow Input Genome Sequencing Library Preparation from a Single Tardigrade Specimen

Tardigrades are microscopic animals that enter an ametabolic state called anhydrobiosis when facing desiccation and can return to their original state when water is supplied. The genomic sequencing of microscopic animals such as tardigrades risks bacterial contamination that sometimes leads to erroneous interpretations, for example, regarding the extent of horizontal gene transfer in these animals. Here, we provide an ultralow input method to sequence the genome of the tardigrade, Hypsibius dujardini, from a single specimen. By employing rigorous washing and contaminant exclusion along with an efficient extraction of the 50 ~ 200 pg genomic DNA from a single individual, we constructed a library sequenced with a DNA sequencing instrument. These libraries were highly reproducible and unbiased, and an informatics analysis of the sequenced reads with other H. dujardini genomes showed a minimal amount of contamination. This method can be applied to unculturable tardigrades that could not be sequenced using previous methods.

Contamination during the genomic sequencing of microscopic organisms remains a large problem. Here, we show a method to sequence the genome of a tardigrade from a single specimen with as little as 50 pg of genomic DNA without whole genome amplification to minimize the risk of contamination.

Tardigrades are microscopic animals that enter an ametabolic state called anhydrobiosis when facing desiccation and can return to their original state ...

Hybrid De Novo Genome Assembly for the Generation of Complete Genomes of Urinary Bacteria using Short- and Long-read Sequencing Technologies

Complete genome sequences provide valuable data for the understanding of genetic diversity and unique colonization factors of urinary microbes. These data may include mobile genetic elements, such as plasmids and extrachromosomal phage, that contribute to the dissemination of antimicrobial resistance and further complicate treatment of urinary tract infection (UTI). In addition to providing fine resolution of genome structure, complete, closed genomes allow for the detailed comparative genomics and evolutionary analyses. The generation of complete genomes de novo has long been a challenging task due to limitations of available sequencing technology. Paired-end Next Generation Sequencing (NGS) produces high quality short reads often resulting in accurate but fragmented genome assemblies. On the contrary, Nanopore sequencing provides long reads of lower quality normally leading to error-prone complete assemblies. Such errors may hamper genome-wide association studies or provide misleading variant analysis results. Therefore, hybrid approaches combining both short and long reads have emerged as reliable methods to achieve highly accurate closed bacterial genomes. Reported herein is a comprehensive method for the culture of diverse urinary bacteria, species identification by 16S rRNA gene sequencing, extraction of genomic DNA (gDNA), and generation of short and long reads by NGS and Nanopore platforms, respectively. Additionally, this method describes a bioinformatic pipeline of quality control, assembly, and gene prediction algorithms for the generation of annotated complete genome sequences. Combination of bioinformatic tools enables the selection of high quality read data for hybrid genome assembly and downstream analysis. The streamlined approach for the hybrid de novo genome assembly described in this protocol may be adapted for the use in any culturable bacteria.

Hybrid De Novo Genome Assembly for the Generation of Complete Genomes of Urinary Bacteria using Short- and Long-read Sequencing Technologies

This protocol details a comprehensive approach for the culturing, sequencing, and de novo hybrid genome assembly of urinary bacteria. It provides a reproducible procedure for the generation of complete, circular genome sequences useful in studying both chromosomal and extrachromosomal genetic elements contributing to urinary colonization, pathogenesis, and antimicrobial resistance dissemination.

Complete genome sequences provide valuable data for the understanding of genetic diversity and unique colonization factors of urinary microbes. These data ...

DNA Barcoding for Medicinal Plants: Identification and Phylogenetic Analysis

Application of DNA Barcoding to Identify Medicinal Plants

Medicinal plants are valuable resources globally and are used worldwide to maintain health and treat disease; however, the presence of adulteration obstructs their development. DNA barcoding, a technique for species identification by standard DNA regions, facilitates prompt and accurate identification of traditional medicinal plants. The process of DNA barcoding entails six basic steps: 1) processing the medicinal plants, 2) extracting high-quality total DNA from the medicinal plants using centrifugal column method, 3) amplifying target DNA region internal transcribed spacer 2 (ITS2) with universal primers of plants and performing Sanger sequencing, 4) splicing and aligning sequence to obtain the target sequence, 5) matching the barcode sequence against the barcode library for identification, 6) aligning sequence, comparing intraspecific and interspecific variation, constructing phylogenetic neighbor-joining tree. As shown in the results, the universal primer can amplify the target region. Basic Local Alignment Search Tool (BLAST) demonstrates the percentage identified was 100%, and the neighbor-joining tree demonstrates that the splicing sequences were clustered with the A. sinensis OR879715.1 clade, and the clade support value is 100. This protocol provides a reference for applying DNA barcoding technology as an effective method to identify medicinal plants and adulterants.

Author Spotlight: Harnessing DNA Barcode Technology to Enhance the Efficiency of Medicinal Plant Identification

The present protocol describes the use of DNA barcoding technology to authenticate plant-based medicinal materials, and the medicinal plant Angelica sinensis (Oliv.) Diels was identified as an example.

Medicinal plants are valuable resources globally and are used worldwide to maintain health and treat disease; however, the presence of adulteration ...

Gel-seq: A Method for Simultaneous Sequencing Library Preparation of DNA and RNA Using Hydrogel Matrices

The ability to amplify and sequence either DNA or RNA from small starting samples has only been achieved in the last five years. Unfortunately, the standard protocols for generating genomic or transcriptomic libraries are incompatible and researchers must choose whether to sequence DNA or RNA for a particular sample. Gel-seq solves this problem by enabling researchers to simultaneously prepare libraries for both DNA and RNA starting with 100 - 1000 cells using a simple hydrogel device. This paper presents a detailed approach for the fabrication of the device as well as the biological protocol to generate paired libraries. We designed Gel-seq so that it could be easily implemented by other researchers; many genetics labs already have the necessary equipment to reproduce the Gel-seq device fabrication. Our protocol employs commonly-used kits for both whole-transcript amplification (WTA) and library preparation, which are also likely to be familiar to researchers already versed in generating genomic and transcriptomic libraries. Our approach allows researchers to bring to bear the power of both DNA and RNA sequencing on a single sample without splitting and with negligible added cost.

Gel-seq enables researchers to simultaneously prepare libraries for both DNA- and RNA-seq at negligible added cost starting from 100 - 1000 cells using a simple hydrogel device. This paper presents a detailed approach for the fabrication of the device as well as the biological protocol to generate paired libraries.

The ability to amplify and sequence either DNA or RNA from small starting samples has only been achieved in the last five years. Unfortunately, the ...

Bioengineering

Ultra-long Read Sequencing for Whole Genomic DNA Analysis

Third generation single-molecule DNA sequencing technologies offer significantly longer read length that can facilitate the assembly of complex genomes and analysis of complex structural variants. Nanopore platforms perform single-molecule sequencing by directly measuring the current changes mediated by DNA passage through the pores and can generate hundreds of kilobase (kb) reads with minimal capital cost. This platform has been adopted by many researchers for a variety of applications. Achieving longer sequencing read lengths is the most critical factor to leverage the value of nanopore sequencing platforms. To generate ultra-long reads, special consideration is required to avoid DNA breakages and gain efficiency to generate productive sequencing templates. Here, we provide the detailed protocol of ultra-long DNA sequencing including high molecular weight (HMW) DNA extraction from fresh or frozen cells, library construction by mechanical shearing or transposase fragmentation, and sequencing on a nanopore device. From 20-25 &#181;g of HMW DNA, the method can achieve N50 read length of 50-70 kb with mechanical shearing and N50 of 90-100 kb read length with transposase mediated fragmentation. The protocol can be applied to DNA extracted from mammalian cells to perform whole genome sequencing for the detection of structural variants and genome assembly. Additional improvements on the DNA extraction and enzymatic reactions will further increase the read length and expand its utility.

Long-read sequences greatly facilitate the assembly of complex genomes and characterization of structural variation. We describe a method to generate ultra-long sequences by nanopore-based sequencing platforms. The approach adopts an optimized DNA extraction followed by modified library preparations to generate hundreds of kilobase reads with moderate coverage from human cells.

Third generation single-molecule DNA sequencing technologies offer significantly longer read length that can facilitate the assembly of complex genomes ...

Isolation and Transcriptome Analysis of Plant Cell Types

In multicellular organisms, developmental programming and environmental responses can be highly divergent in different cell types or even within cells, which is known as cellular heterogeneity. In recent years, single-cell and cell-type isolation combined with next-generation sequencing (NGS) techniques have become important tools for studying biological processes at single-cell resolution. However, isolating plant cells is relatively more difficult due to the presence of plant cell walls, which limits the application of single-cell approaches in plants. This protocol describes a robust procedure for fluorescence-activated cell sorting (FACS)-based single-cell and cell-type isolation with plant cells, which is suitable for downstream multi-omics analysis and other studies. Using Arabidopsis root fluorescent marker lines, we demonstrate how particular cell types, such as xylem-pole pericycle cells, lateral root initial cells, lateral root cap cells, cortex cells, and endodermal cells, are isolated. Furthermore, an effective downstream transcriptome analysis method using Smart-seq2 is also provided. The cell isolation method and transcriptome analysis techniques can be adapted to other cell types and plant species and have broad application potential in plant science.

The feasibility and effectiveness of high-throughput scRNA-seq methods herald a single-cell era in plant research. Presented here is a robust and complete procedure for isolating specific Arabidopsis thaliana root cell types and subsequent transcriptome library construction and analysis.

In multicellular organisms, developmental programming and environmental responses can be highly divergent in different cell types or even within cells, ...

Robust DNA Isolation and High-throughput Sequencing Library Construction for Herbarium Specimens

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Automated Gel Size Selection to Improve the Quality of Next-generation Sequencing Libraries Prepared from Environmental Water Samples

Purification of High Molecular Weight Genomic DNA from Powdery Mildew for Long-Read Sequencing

Optimization and Comparative Analysis of Plant Organellar DNA Enrichment Methods Suitable for Next-generation Sequencing

Empirical, Metagenomic, and Computational Techniques Illuminate the Mechanisms by which Fungicides Compromise Bee Health

Ultralow Input Genome Sequencing Library Preparation from a Single Tardigrade Specimen

Hybrid De Novo Genome Assembly for the Generation of Complete Genomes of Urinary Bacteria using Short- and Long-read Sequencing Technologies

Application of DNA Barcoding to Identify Medicinal Plants

Gel-seq: A Method for Simultaneous Sequencing Library Preparation of DNA and RNA Using Hydrogel Matrices

Ultra-long Read Sequencing for Whole Genomic DNA Analysis

Isolation and Transcriptome Analysis of Plant Cell Types