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

Podsumowanie

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.

Streszczenie

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.

Wprowadzenie

Herbarium collections are a potentially valuable source of both species and genomic diversity for studies including phylogenetics^1,2,3, population genetics^4,5, 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 "treasure chest"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 preservation^9,10, 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→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 depth^12,15, 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.

Protokół

1. Prior to Start

1. 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.
2. Add 10 µL of β-mercaptoethanol per 5 mL of CTAB buffer.
   NOTE: this can be prepared in batches of 50 mL and stored at room temperature for 3–4 weeks.
   1. Heat the CTAB solution in a 65 °C water bath.
3. Chill mortars and pestles in -20 °C for at least 20 min.
4. 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.
5. Chill isopropanol on ice or in a -20 °C freezer.
6. 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).
7. Prepare 80% ethanol.
8. Select herbarium specimens for extraction and retrieve ~1 cm², or 10 mg, of tissue per specimen, preferably leaf material.

2. DNA Extraction

1. Grind ~1 cm² of preselected herbarium tissue using prechilled mortars and pestles. Add liquid nitrogen and 30–50 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.
2. Transfer the frozen powder into two 2 mL tubes (add no more than half of the tube's volume). Add 600 µL 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.
3. Incubate the samples in a 65 °C water bath for 1–1.5 h, vortexing every 15 min.
4. Centrifuge samples at 10,000 x g for 5 min. Transfer the supernatant to a fresh set of labeled tubes (~500 µL). Discard the pellet using a standard non-chlorinated disposal procedure.
5. Add 4 µL of RNase-A (10 mg/mL) to each tube and mix by inverting or pipetting. Incubate the samples at 37 °C in a heat block or water bath for 15 min.
6. Add an equal volume (~500 µL) 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 µL). Discard the organic layer into a chlorinated waste container.
   NOTE: Step 2.6 may be repeated if large quantities of secondary compounds are expected in plant material.
7. Add an equal volume (~400 µL) 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 µL). Discard the organic layer into a chlorinated waste container.
8. Add an equal volume (~300 µL) of prechilled isopropanol and 12 µL of 2.5 M sodium acetate to each tube. Incubate samples at -20 °C for 30–60 min.
   NOTE: The incubation times can be extended (up to overnight incubation), but DNA quality will decrease the longer the samples incubate.
9. 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 300–500 µL). 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.
10. 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 °C). Make sure that the samples do not over-dry, as this can decrease the final yield of DNA.
11. Suspend the isolated DNA in 50 µL of 1x TE. Store in the -20 °C freezer for long term storage or 4 °C for use in the following week.

3. Quality Control (QC)

1. Run an agarose gel for quality check.
   1. 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.
   2. 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.
   3. Mix 3 µL of sample, 2 µL of reagent grade water, and 1 µL of 6x loading dye. Load the samples in the gel matrix, noting their order.
   4. Run the samples for 60–70 min at 60–70 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.
2. Run a dsDNA quantification analysis (see Table of Materials) to determine the quantity of double stranded DNA.
   1. Use 2 µL 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.

4. DNA Shearing

NOTE: This is an optimized version of a commercial double-stranded fragmentase protocol (see Table of Materials).

1. Place dsDNA fragmentation enzyme on ice after vortexing for 3 s.
2. In a sterile 0.2 mL polymerase chain reaction (PCR) tube, mix 1–16 µL isolated DNA with 2 µL of accompanying fragmentation reaction buffer. Bring the total volume to 18 µL by adding nuclease free water. Add 2 µL 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).
3. Incubate the samples at 37 °C for 8.5 min. Then add 5 µL 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.

5. Bead Clean-up

1. Homogenize the SPRI beads by vortexing.
2. Bring the total volume of the sheared DNA to 50 µL by adding 25 µL of nuclease free water. Add 45 µL of room temperature SPRI beads (90% volume) to 50 µL 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.
3. 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.
4. Add 200 µL 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.
5. Remove the tubes from the magnet. Elute the DNA from the beads into 55 µL 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).
6. Pull off 52 µL 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.

6. Library Preparation

NOTE: This is a modified version of a commercially available library kit (see Table of Materials protocol).

1. End Prep
   1. Add 3 µL of endonuclease and phosphate tailing enzymes, and 7 µL of accompanying reaction buffer to 50 µL of cleaned, sheared DNA. Mix thoroughly by pipetting up and down. Spin the tubes to remove bubbles.
      NOTE: The total volume should be 60 µL.
   2. Place the samples in a thermocycler with the following program: 30 min at 20 °C, 30 min at 65 °C, then hold at 4 °C.
      NOTE: Heated lid was set to ≥75 °C
2. Adaptor Ligation
   1. Dilute the adaptor 25–50 fold (working adaptor concentration of 0.6–0.3 µM). Add 30 µL of ligation master mix, 1 µL of ligation enhancer, and 2.5 µL of adaptor for high-throughput short read sequencing to the tubes.
      NOTE: The total volume should be 93.5 µL.
   2. Mix thoroughly by pipetting up and down. Spin the tubes to remove bubbles. Incubate the tubes at 20 °C for 15 min.
   3. Add 3 µL 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 µL. Mix thoroughly and incubate at 37 °C for 15 min using a thermocycler.
      NOTE: The lid should be set to ≥47 °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.
3. Cleanup to Remove Enzymes and Small Fragments
   1. Homogenize magnetic beads by vortexing.
   2. Add 78 µL 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.
   3. 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.
   4. Add 200 µL 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.
   5. Remove the tubes from the magnet. Elute the DNA target from the beads by adding 17 µL of 0.1x TE and mix thoroughly by pipetting up and down.
   6. Incubate at room temperature for 5 min. Place tubes on the magnetic stand and wait for the solution to turn clear (~2 min).
   7. Pull off 15 µL of the supernatant.
4. PCR amplification
   1. Add 25 µL of high fidelity PCR master mix, 5 µL of high-throughput short read sequencing library prep 5' primer, and 5 µL of high-throughput short read sequencing library prep 3' primer, to 15 µL of the cleaned adaptor-ligated DNA.
      NOTE: Total volume should be 50 µL.
   2. 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.

Cycle Step	Temp.	Time	Cycles
Initial Denaturation	98 °C	30 s	1
Denaturation	98 °C	10 s	12
Annealing/Extension	65 °C	75 s	12
Final Extension	65 °C	5 min	1
Hold	4 °C

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.

5. Size Selection for Desired Library Size
   NOTE: This bead step will remove fragments both above and below the target range.
   1. Homogenize SPRI beads by vortexing.
   2. Add 25 µL (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 400–600 base pair range. The supernatant contains smaller fragments.
   3. Add 6 µL 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.
   4. 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.
   5. Add 200 µL 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.
   6. Remove the tubes from the magnet. Elute the DNA target from the beads into 33 µL of 0.1x TE and mix thoroughly by pipetting up and down.
   7. 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 µL of the supernatant and transfer to 2 mL tubes (see Table of Materials) for storage.
      NOTE: The libraries can be kept at -20 ˚C for long-term storage.
6. Quality control
   1. 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.
7. 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.
   1. Dilute the universal reamplification primers 10-fold using 0.1x TE.
   2. Add 25 µL of high fidelity PCR master mix, 5 µL of diluted universal reamplification primer 1 (AATGATACGGCGACCACCGA), and 5 µL of diluted universal reamplification primer 2 (CAAGCAGAAGACGGCATACGA) to 15 µL of low concentration libraries. NOTE: Total volume should be at 50 µL.
   3. 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.
   4. Bead clean-up
   5. Homogenize SPRI beads by vortexing. Add 45 µL of room temperature SPRI beads (90% volume) and mix thoroughly by pipetting up and down.
   6. Let the samples incubate for 5 min. Put the tubes on a magnetic plate for 5 min.
   7. 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.
   8. Add 200 µL 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.
   9. 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.
   10. Remove the tubes from the magnet. Elute the DNA target from the beads into 33 µL of 0.1x TE and mix thoroughly by pipetting up and down.
   11. Incubate at room temperature for 5 min. Place tubes on the magnetic stand and wait for the solution to turn clear (~2 min).
   12. Pull off 30 µL of the supernatant. The libraries can be stored at -20 °C for long-term storage.
8. Quality control
   1. 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.

Wyniki

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...

Dyskusje

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.

Materiały

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