TITLE: Uppsala Fungal Barcoding using ONT: DNA extraction -&gt; Library preparation V1
AUTHORS: Karl Soler

[DESCRIPTION]
Abstract

This protocol was developed by Uppsala Svampklubb (fungal society), in cooperation with Evolutionary Biology Centre (EBC) at Uppsala University, as a cost efficient method to help amateurs with fungal species identification using DNA sequences. 

The advantage of Nanopore technology is that over 2000 samples can be sequenced at the same time, thereby significantly lowering the cost per sample. The more that is sequenced at the same time, the cheaper it becomes per sample. The main challenge is to balance a high enough success rate with a low cost per sample, which is fairly optimized in this protocol, especially for agarics.

The protocol follows the process all the way from collecting fungi in the field to having a library of DNA ready for sequencing. It yields an 85% success rate for mushrooms and toadstools (fairly easy to sequence). Hydnoids, polypores and corticioids usually also works well. Ascomycetes are usually more difficult but works with some tweaking of the protocol.

Time required:
Time consumption is shown on a note in the fist step of each section. The table below summarizes the time consumption of the entire barcoding process; from collecting specimens to analysing and documenting DNA sequences. 

 


[STEPS]
SECTION: Fungal collecting, storing and documentation
1. 
 

Before starting:
In order to maximize the chances that your barcoding is successful there are several things to consider when collecting, storing and documenting your fungal specimens. Collections coupled with a DNA sequence are also more valuable when donated to a herbarium (a voucher), in which case it is important to properly conserve them so that they meet the standards of the herbarium. 

Some things to consider:
When possible collect many fruit bodies in the field. Larger collections are better than small ones since material is constantly being used for research. 
Document the finding. Minimum parameters for herbarium donations are: date, region, locality and coordinates. The locality should be described as detailed as possible in case the coordinates are wrong. Specimens should preferably be determined to at least genus level.
Specimens should be dried in 30-35 °C until they are completely dry. The optimal temperature depends on the type of fungus and other factors. DNA might be damaged if the temperature is too high, or the specimen is dried too quickly, or too slowly. If specimens are dried too slowly they also often get eaten by worms and insects. Room temperature is often sufficient for fungi with lower water content, e.g. many hard wood living fungi such as polypores and corticioids. 
Very large specimens should be sliced into several thinner pieces before drying in order to fit in herbarium sheets.
If the fungus is an epifyte on a fragile plant it is good to use a plant press or similar.
Store the specimens as dark as possible as light exposure destroys DNA over time. Properly dried and stored collections lasts at least a few years before DNA deteriorates to a level that makes sequencing harder.
Keep a database, e.g. an excel sheet, with all your data on the collected specimens. It is practical to assign personal collection numbers to your specimens which can be used to keep track of them when they enter the barcoding stage or are deposited to a herbarium.
Report the finding to an observation database such as iNaturalist or Artportalen (in Sweden).
SECTION: Adapter ligation and clean-up
90. Make up the library to 12 µL at 35-50 fmol for simplex reads. This may require using nuclease-free water and a new 1.5 mL LowBind tube.

SECTION: Adapter ligation and clean-up
89. Quantify 1 µL of the sample using a Qubit fluorometer. Depending on how many PCR plates were pooled into the sample, and the success rate (percentage off cells with successful amplification) of the PCR plates, the Qubit value may end up around 50-100 ng/µL. 

SECTION: Adapter ligation and clean-up
88. Remove and KEEP 15 µL of eluate, which now contains the DNA library, in a 1.5 mL LowBind tube. The beads can be disposed of.

SECTION: Adapter ligation and clean-up
87. Place the tube back on the magnet for at least 1 min, until the eluate is clear. 

SECTION: Adapter ligation and clean-up
86. Remove the tube and resuspend the beads in 15 µL of Elution Buffer (EB). Spin down and incubate for for 10 min at 37 °C. We typically use a dry heat block. Every 2 minutes, flick the tube for 10 s to encourage DNA elution.

SECTION: Adapter ligation and clean-up
85. Spin down the tube and place it back on the magnet. Pipette off any residual supernatant. 
SECTION: Adapter ligation and clean-up
84. Repeat the previous step.   

SECTION: Adapter ligation and clean-up
83. Wash the beads with 125 µL of Short Fragment Buffer (SFB). Flick the the tube to resuspend the beads in the buffer, then briefly spin down. Then return to the magnetic rack and wait for the beads to stick to the magnet. Pipette off the eluate and discard.

Do NOT use ethanol here as it will ruin the sequencing reaction. 


 

SECTION: Adapter ligation and clean-up
82. Spin down the sample and place on the magnetic rack. When the beads are stuck to the magnet and the eluate is clear, pipette off eluate and discard.
SECTION: Adapter ligation and clean-up
81. Resuspend the AMPure XP Beads by vortexing. Then add 20 µL of beads to the reaction and mix by pipetting. Incubate on a Rotator mixer for 10 min at room temp.

SECTION: Adapter ligation and clean-up
80. Thoroughly mix the reaction by gently pipetting at least 10 times and briefly spinning down. Then incubate for 20 min at room temperature.

SECTION: Adapter ligation and clean-up
79. Add 5 µL of the Quick T4 DNA Ligase and pipette mix 10 - 20 times. The LowBind tube should now contain a total of 50 µL.

SECTION: Adapter ligation and clean-up
78. Add 10 µL of the NEBNext Quick Ligation Reaction Buffer (5X) and pipette mix 10 - 20 times.

SECTION: Adapter ligation and clean-up
77. Add 5 µL of the Native Adapter (NA) and pipette mix 10 - 20 times.

SECTION: Adapter ligation and clean-up
76. Add 30 µL of the pooled, barcoded sample to a new 1.5 mL LowBind tube. 


SECTION: Adapter ligation and clean-up
75. - Spin down the Native Adapter (NA), pipette mix and place on ice. 

- Thaw the Elution Buffer (EB) at room temperature and mix by vortexing. Then spin down and place on ice.

- Thaw the Short Fragment Buffer (SFB) at room temperature and mix by vortexing. Then spin down and place on ice.


 

Reagents:
  
  

SECTION: Adapter ligation and clean-up
74. 
 

1. Thaw the NEBNext Quick Ligation Reaction Buffer (5X) and Quick T4 DNA Ligase at room temperature. 

2. Then spin down the two tubes for 5 s. 

3. Ensure that each tube is fully mixed by performing 10 full volume pipette mixes. 

4. When the Quick Ligation Reaction Buffer is sufficiently pipette mixed (no precipitate can be seen), vortex the tube for several seconds to ensure thorough mixing. Do NOT vortex the Quick T4 DNA Ligase.

5. Place on ice.

 

Reagents:
  

SECTION: Native barcode ligation
72. It is a good idea to check if there is sufficient DNA concentration in the eluate using a Qubit fluorometer. In that case use 1 µL of the sample, using more can result in the DNA concentration being too high for the fluorometer to read.

Take forward the sample to the "Adapter ligation and clean-up" section. 
SECTION: Native barcode ligation
71. Place the tube back on the magnetic rack until the eluate is clear. Then remove and KEEP 35 µL of eluate in a new 1.5 mL LowBind tube. 

SECTION: Native barcode ligation
70. Incubate for 10 min at 37 °C. We typically use a dry heat block. Every 2 minutes, flick the tube for 10 s to encourage DNA elution.

SECTION: Native barcode ligation
69. Remove the tube from the rack and resuspend the beads in 35 µL nuclease-free water by gently flicking the tube. 

SECTION: Native barcode ligation
68. Spin down and place the tube back on the magnet. Pipette off residual ethanol. Allow to dry for not more than 30 s.

SECTION: Native barcode ligation
67. Repeat the previous step.   

SECTION: Native barcode ligation
66. Wash the beads with 700 µL of 80% ethanol without disturbing the beads. If the beads are disturbed, wait until they have gathered on the magnet before proceeding. Pipette off the ethanol and discard.

SECTION: Native barcode ligation
65. Spin down the pooled sample and place it on the magnetic rack for 5 min until it the eluate is clear from beads. Pipette of the eluate and discard.

SECTION: Native barcode ligation
64. Prepare 2 mL of fresh 80% ethanol in nuclease-free water for use in step    and forward.

SECTION: Native barcode ligation
63. Resuspend the AMPure XP Beads by vortexing. Then add 0.4X bead solution to the pooled sample and mix by pipetting. Incubate on a rotator mixer for 10 min at room temp. 

SECTION: Native barcode ligation
62. Pool all the barcoded samples in a 1.5 mL LowBind tube. The total pooled volume should now be 
24 µL X number of barcodes/samples. 
SECTION: Native barcode ligation
61. Add 4 µL of EDTA (with the blue cap) to each tube. Mix thoroughly by pipetting and spin down briefly. 

=> EDTA is added to stop the barcode reaction.

SECTION: Native barcode ligation
60. Incubate for 20 min at room temp. 

SECTION: Native barcode ligation
59. Add 10 µL of Blunt/TA Ligase Master Mix. Pipette mix 10-20 times. Spin down briefly.

SECTION: Native barcode ligation
58. Add 2.5 µL of the corresponding barcode to each tube. Pipette mix 10-20 times. 

SECTION: Native barcode ligation
57. Add 7.5 µL of End-prepped DNA from step   to each corresponding tube.   

SECTION: Native barcode ligation
56. If not already done: Prepare a clean 0.2 mL thin-walled PCR tube for every sample and label it with the barcode number. 
SECTION: Native barcode ligation
55. Thaw the number of native barcodes (NB01-24) required for your samples (one per sample) at room temp. Individually mix them by pipetting, spin down and place them on ice. The barcode number for each sample (PCR plate) should be carefully noted in your excel data sheet. Only use one barcode per sample.

 

SECTION: Native barcode ligation
54. Thaw the EDTA (with the blue cap) at room temp and mix by vortexing. Then spin down and place on ice.

 

SECTION: Native barcode ligation
53. 
 

Thaw the Blunt/TA Master mix at room temp. Spin down the tube for 5 s. Mix it by pipetting 10 times.

Reagent:
  

SECTION: End-prep
51. It is a good idea to check if there is sufficient DNA concentration in each sample using a Qubit fluorometer. 
SECTION: End-prep
50. Pipette 10 µL of the clear eluate into clean 1.5 mL LowBind tubes. OBS: The clear eluate contains DNA and must not be discarded!

SECTION: End-prep
49. Place the tubes back on the magnets until the eluate is clear. 
SECTION: End-prep
48. Remove the tubes from the magnets and resuspend the beads in 10 µL nuclease-free water. Pipette mix until the beads are homogenously suspended. Spin down and incubate for 2 min at room temp.

SECTION: End-prep
47. Repeat the previous step  . Briefly spin down the tubes and put them back on the magnets. Pipette off any residual ethanol. Let the tubes dry for 30 s. Do not wait longer than this since the the beads will crack and clump together!

SECTION: End-prep
46. Carefully wash the beads with 200 µL of fresh 80% ethanol. Then remove the ethanol using a pipette and discard it. If you disturb the beads, wait for them to gather at the magnet again before removing the ethanol.

SECTION: End-prep
45. Spin down the samples and place them on the magnetic rack until the eluate is clear. Pipette off the eluate and discard it. 
SECTION: End-prep
44. Prepare a sufficient volume of 80% freshly prepared ethanol in nuclease-free water for step .

SECTION: End-prep
43. Resuspend the AMPure XP beads by vortexing. Then add 15 µL to each tube and mix by flicking the tube. Incubate on a rotator mixer for 5 min at room temp. If you don't have access to a rotator mixer, hold the tubes on a rack and wave them around in a circular motion.

SECTION: End-prep
42. Transfer each sample into a 1.5 mL LowBind tube. 
SECTION: End-prep
41. Incubate in a thermal cycler at 20 °C for 5 min followed by 65 °C for 5 min. 


 


SECTION: End-prep
40. The total volume in the tubes should now be 15 µL. Mix by pipetting and spin down in a mini-centrifuge. 

SECTION: End-prep
39. Add 1.75 µL of Ultra II End-prep Reaction buffer to the new tubes. Pipette mix 10-20 times.

Then:

Add 0.75 µL of Ultra II End-prep Enzyme Mix to the tubes. Pipette mix 10-20 times.

=> The easiest way to do this is to make a master mix with the Ultra II buffer and enzyme mix. Then adding 2.5 µL to each tube.

SECTION: End-prep
38. Using a maximum of 12.5 µL, transfer 200 fmol of DNA (195 ng for 1.5 kb amplicons)  from the eluate in step   to new 0.2 mL thin-walled PCR tubes. If less than 12.5 µL is needed, make up the rest with nuclease-free water.

=> In this step you need to know approximately how much DNA your cleaned samples contain. 
SECTION: End-prep
37. Thaw the Ultra II reagents on ice. Flick or invert the tubes to ensure that they are well mixed. Do not vortex the Enzyme mix! Spin down the tubes before opening. The Reaction buffer should be brought to room temperature, then pipette mixed several times. Contrary to the enzyme mix, it should then be vortexed for at least 30 seconds before usage. Make sure that potential precipitate in the bottom of the tube is solubilized. 

Reagents:
  


 

SECTION: End-prep
36. Thaw the AMPure XP beads at room temperature.

 

SECTION: End-prep
35. 
 

This section, along with the "Native barcode ligation" and "Adapter ligation and clean-up" sections require reagents from the ONT Native Barcoding Kit 24 V14, and the following protocol is modified from the manuals provided with the kit: Native barcoding protocol

 

SECTION: Post PCR pooling &amp; cleaning
33. Make sure that the eluate contains enough DNA for the End-prep section  . This can be measured using a Qubit fluorometer.

 

SECTION: Post PCR pooling &amp; cleaning
32. Place the tubes back on the magnets for 1 min to separate the beads. Transfer the eluate to a new 1.5 mL LowBind tube. OBS: the eluate contains the DNA and must not be discarded!


 

SECTION: Post PCR pooling &amp; cleaning
31. Remove the tubes from the magnets and add 30 µL of molecular water. Pipette mix at least 10 time, making sure that the beads are homogenously eluted in the water. Incubate for 2 min. 

SECTION: Post PCR pooling &amp; cleaning
30. Repeat the previous step   . In this step it is important to remove all the ethanol. Let the tubes dry for 30 s to evaporate small traces of ethanol. Be careful not to dry the beads any longer since they will crack and clump together making it hard to elute them.

SECTION: Post PCR pooling &amp; cleaning
29. It is now time to wash the beads with 70% ethanol. Use an ethanol volume that is at least the total volume of the sample and the bead solution you removed in the previous step, and a minimum of 200 µL. Pipette the ethanol into the tubes and incubate for 30 s. Then remove and discard the ethanol. 

Important: Freshly prepare the ethanol yourself by mixing >99% ethanol and nuclease-free water in 0.7/0.3 proportions.
SECTION: Post PCR pooling &amp; cleaning
28. Remove the solution using a pipette and discard it. Be careful not to come in contact with the beads. It is not important that 100% of the solution is removed in this step. Instead, be careful not to suck beads into the pipette.

 

SECTION: Post PCR pooling &amp; cleaning
27. Spin down the tube for 2 s in a mini centrifuge. Place the tube on a magnetic rack/magnetic separator (see materials section) and let the beads separate from the solution during 2 min. When all the beads are stuck to the magnet, proceed to the next step. 


 


SECTION: Post PCR pooling &amp; cleaning
26. The magnetic beads should have been brought to room temperature and are ready to use. Shake the magnetic bead bottle to resuspend any magnetic particles that may have settled. Then add bead solution to the LowBind tube with amplicons. The amount you add should be 1,8X the amount of amplicons you transferred in the previous step. Pipette the mix up and down in the tube 10 times to thoroughly mix it and the incubate for 5 min at room temperature. DNA is now stuck to the beads.

SECTION: Post PCR pooling &amp; cleaning
24. The following instructions are modified from: AMPure XP Protocol. 


In this step you will transfer X µL from the amplicon pool to a new 1.5 mL LowBind tube. The volume you transfer depend on how much DNA there was in your PCR plate. A PCR plate with a low success rate (fewer cells with successful amplification = less total DNA) of around 30% may require 300 µL, while a high success rate may require 100 µL. If you performed gel electrophoresis on the plate you will know the success rate. Otherwise it can often be estimated by the proportions of easy vs. difficult fungi in the plate. Mushrooms, toadstools etc are relatively easy while wood-living fungi, hydnoids and many ascomycetes are examples of more difficult groups, often with lower success rates in this protocol. Estimating which species are easy vs. difficult requires some experience. 

IMPORTANT: Save both the pool tubes and the LowBind tubes containing your pooled amplicons. Make sure to properly label them!


SECTION: Post PCR pooling &amp; cleaning
23. Pool the contents of the 8-strip into a single 1.5 mL LowBind tube with a pipette. The tube will then contain 96 x 3 = 288 µL of amplicon solution, i.e. the contents from an entire PCR plate (unless the plate was not completely filled). After this step you should have an equal amount of 1.5 mL tubes to PCR plates.


 

SECTION: Post PCR pooling &amp; cleaning
22. At this point you should have a number of ready PCR plates with amplicons. The following steps are done for every plate. Quickly centrifuge the PCR plate so that there is enough amplicons in the bottom om the cells. 

Start by pooling 3 µL from each cell on the PCR plate into a new 0.2 mL 8-strip using a multichannel pipette. This can be done without changing the pipette tips between columns since they will be pooled together anyway. After this, every tube in the 8-strip should contain the contents from 12 cells in its corresponding row in the PCR plate (12x3=36 µL). Keep the PCR plate with the remaining amplicons in case something goes wrong later. 


 

SECTION: Post PCR pooling &amp; cleaning
21. The   magnetic beads, which are stored in the fridge, should be brought to room temperature. Initially the beads are in the bottom of the bottle and need to be suspended by shaking it. 


 

SECTION: Post PCR pooling &amp; cleaning
20. 
 

This section, along with the "End-prep", "Native barcode ligation" and "Adapter ligation and clean-up" sections will go smoother if the needed tubes are prepared and labeled beforehand. Keep in mind that one tube for each PCR plate is needed, until the end of the "Native barcode ligation" section when DNA from all the plates is pooled together (right most tubes in the picture below). It is a good idea to label the tubes with plate number and barcode number, especially the latter (barcodes are explained in step  ). 

Tube guide

-> X stands for the number of PCR plates you want to sequence
-> all tubes with contents from a single PCR plate should be labeled with plate number and barcode number
-> When performing Qubit, 2 additional tubes to the samples are always needed for the standards

Post PCR pooling & cleaning
(X) 0.2 mL 8-strips 
(X) 1.5 mL LowBind tubes -> for pooling
(X) 1.5 mL LowBind tubes -> for cleaning
(X) 1.5 mL LowBind tubes -> cleaned
(X+2) Qubit tubes

End-prep
(X) 0.2 mL thin-walled tubes -> for end-prep
(X) 1.5 mL LowBind tubes -> for cleaning
(X) 1.5 mL LowBind tubes -> cleaned
(X+2) Qubit tubes

Native barcode ligation 
(X) 0.2 mL thin-walled tubes -> for barcoding
(1) 1.5 mL LowBind tube -> for pooling & cleaning
(1) 1.5 mL LowBind tube -> cleaned
(3) Qubit tubes

Adapter ligation & clean-up
(1) 1.5mL LowBind tube -> for adapter ligation & cleaning
(1) 1.5 mL LowBind tube -> cleaned library
(3) Qubit tubes
(1) 1.5 mL LowBind tube -> finished library


 

SECTION: PCR Protocol
18. Set the PCR-machine according to the picture below. Annealing temperature should be 63 °C and the number of cycles should be 35. Set the reaction volume to 14 µL. Start the PCR machine. The program will take around 90 min.


 

Optional: If you want to check if the PCR was successful you can perform gel electrophoresis on the entire PCR plate. We strongly recommend this in your early experiments as many things can go wrong during the extraction and PCR process. From performing electrophoresis you will also know the amplification success rate in your plate which will be useful when calculating the amount of DNA you have in later stages.
SECTION: PCR Protocol
17. Carefully place the lids containing the second primer on the PCR plate. The cells now contain 14.3 µL of mastermix, template and primers and the plate is ready for PCR. Before going to the thermocycler (PCR machine), vortex the plate for 5-7 seconds and then centrifuge it up to 4000 rpm.


 

SECTION: PCR Protocol
16. Add 11 µL of the mastermix you prepared in step   to each cell on the plate using a dispenser pipette. Be careful when using the dispenser pipette! Make sure not to get air bubbles in the liquid which will cause the pipette to dispense uneven amounts between cells. The same will happen if one tries to use all the liquid in the pipette before refilling it. Always make sure to refill the pipette before it almost runs out so that this does not happen.

SECTION: PCR Protocol
14. Place your 12 PCR plate lids (8-strip lids) upside down on the workbench. Prepare the next primer in your primer pair (FÖRKLARING) by pipetting 1.4 µL of the primer into the PCR plate lid using a multichannel pipette. This will ensure that primers do not come into contact with each other in the plate cells while you prepare the PCR plate. This is important as primers attach to each other over time rendering them useless.

Important: It is important that the master mix is ready and that you immediately proceed to the next step after performing this one. The small primer drops are prone to evaporate quickly so make sure that they are not left out for too long.

 

SECTION: PCR Protocol
13. Vortex the 8-strips containing your DNA template for a few seconds to ensure even concentration of DNA. Then add 0.5 µL of DNA template to each cell in the PCR plate using a multichannel pipette. Make sure that the position of each specimen stays the same (1A-12H). 

-> We reduced the initial template volume (1.4) to 0.5 in order to reach a 1:50 dilution without having to use an extra set of tubes and pipettes in   
SECTION: PCR Protocol
12. Prepare an empty PCR plate (a plate with 96 cells). Add 1.4 µL of the LR5 reverse primer to each cell using a multichannel pipette. 

-> Be sure to align the primer plate and the PCR plate (columns) to avoid mistakes. As the multichannel pipette can only transfer primers to 8 columns (cells) at a time, pipetting the 12 LR5 reverse primers must be done in two rounds; one with 8 pipette tips and one with 4. One can use the same pipette tips within the same round (same primers) but NOT between rounds (different primers). 

 

SECTION: PCR Protocol
11. For the PCR protocol in this manual we use a PHUSION mastermix:   
and dNTP:s from:
  

Start with preparing the mastermix. The amount of mastermix you will need depends on how many PCR plates you prepare on the occasion. Use the table below to calculate how much of each reagent you will need. We recommend that you prepare a bit more mastermix than is needed on paper since some will get stuck in the tube and pipette. 7% or 9% extra solution have been proven to be sufficient for us. Important: Wait with adding the polymerase to just before you are going to add mastermix to the PCR plate 
(  ). The mix with the other reagents can sit on ice until then.

 

Prepare a tube that can hold the volume of mastermix that you will need. Pipette the reagents into the tube. Gently tilt the tube 180 degrees back and forth to mix the reagents. The mastermix is now ready for use.

 
 

SECTION: PCR Protocol
10. 
 

We will start this section by making a primer plate. The primer plate will cover the primer use for over 10 PCR plates and therefore only needs to be made once in a while. The instructions are based on the ITS1 forward primer (White et al. 1990) and LR5 reverse primer (Hopple & Vilgalys 1994) with 8 and 12 tags respectively. We have, however, also started using the ITS1-F_KYO2 forward primer (Toju et al. 2012) for difficult ascomycetes and the ITS4 reverse primer (White et al. 1990) for old material.

First time using the primers: Start with thawing your newly purchased primers. Add nuclease-free water to each of the primer tubes so that the resulting volume has a concentration of 100 micromolar (µM). As each tube contains a different amount (moles) of primer, you will have to  calculate the volume of water you need to put in each tube individually. See the picture below for an example.

Vortex the tubes for a few seconds and spin down. You may want to repeat this step to make sure that the primer is evenly mixed.

 

Prepare (20) 1.5 mL tubes; 12 for the LR5 reverse primers and 8 for the ITS1 forward primers. Label them according to the tag number. 

Make a 1:20 dilution (5 micromolar (µM)) with nuclease-free water in the new tubes by pipetting a volume of choice from the 100 µM tubes, e.g. 50 µL of primer + 950 µL of water. Your primer setup should look something like the picture below.

Vortex the 1.5 mL tubes for a few seconds and spin down.

 

Prepare an empty PCR plate (a plate with 96 cells). 

Place an 8-strip lid horizontally over one of the rows. Cut an 8-strip lid in half to a 4-strip and place it next to the 8-strip to fill out the row with lids. Label the 12 lids with the LR5 reverse primer number (tag) you will use in each cell. 

On another row, place a new 8-strip lid. Label the 8 lids with the ITS1 forward primer number (tag) you will use in each cell. 

Transfer 150 µL of 5 micromolar (µM) primer solution from the 1.5 mL tubes to the corresponding cell on the primer plate. 

Label the primer plate with primer information, date and concentration. It should now look something like the picture below. 

 
 


SECTION: DNA extraction from dried fungi
8. Add 190 µL of nuclease-free water to the strips in order to dilute the samples. 

-> The 1:50 dilution have proven to be sufficient for most mushrooms and toadstools. However, different genera often require different dilutions depending on the amount of DNA and PCR inhibitors present in the material. Strongly pigmented fungi often require a higher dilution. What method works the best for certain fungal groups is highly experience based. In some cases even a 1:1000 dilution may work better. In this step we have diluted the sample to 1:20. Later, in the PCR protocol, we also reduced the initial template volume from 1.4 to 0.5 in order to achieve a ~1:50 dilution. If you think that a higher dilution (>1:60) is necessary, you will need to use an extra set of tubes and pipette tips in this step (dilute further in new tubes). 
SECTION: DNA extraction from dried fungi
7. Spin down the strips in a mini centrifuge for 10 seconds to ensure that all the tissue and liquid is in the bottom. Make sure that the tissue is in the fluid and that no large air bubbles are left. Heat the tubes with the tissue for 60 min to 90 min at 80 °C on a dry heat block or in a thermocycler. The heat will break down the cell walls, releasing DNA.


 

SECTION: DNA extraction from dried fungi
6. Pipette 10 µL of Mason & Botella (2020) extraction buffer (20 mM Tris (pH 8), 25 mM NaCl, 2.5 mM EDTA, 0.05% (wt/vol) SDS, and 2% (wt/vol) PVP-40) in each tube with tissue using a dispensing pipette or multipipette. This step can be done in the lab both before and after placing the tissue in the tubes. We recommend that it is done beforehand since the tissue is easier to stick in the tubes when there is liquid inside. Static electricity can otherwise make it difficult to handle small pieces of tissue in plastic tubes.

 


 

SECTION: DNA extraction from dried fungi
5. Consider performing step   before this step.

Put a very small piece of tissue in each tube. Usually 0.2 mg or 1-2 mm2 is well enough for this manual and one can often use even smaller amounts. The amount of tissue can be changed for different species if needed but keep in mind that one may also need to change the amount of buffer when doing so in order to maintain optimal dilution. If you are new to this protocol it is a good idea to calibrate your weight/mass perception of different fungal species beforehand using a high resolution lab scale. Later, when doing this at home, you will then be better at estimating amounts. It is often preferable to choose tissue from the fertile part of the fungus, i.e. the hymenium containing spores.

Dried tissue can be broken off from the fungus using e.g. a scalpel or tweezers and a white paper. Choose a piece that is close to the desired size and put it in the tube. Toothpicks are very useful for putting small pieces of tissue into the tubes. Between specimens, wipe the scalpel or tweezer with paper. If you want to be extra careful, dip it in ethanol before wiping. 

 

 

SECTION: DNA extraction from dried fungi
4. Put the specimens in numerical order on your work bench with regards to the pairing of your personal collection numbers with the row-column (number-letter) combination on the 96 tubes. The collection assigned to tube 1A should be placed first in order etc. for efficient DNA extraction. 
SECTION: DNA extraction from dried fungi
3. Assign your fungal specimens to a row and column number in your excel data sheet.

It is very important that you keep track of which collection belongs which tube. Without this information you will later have no way of knowing which DNA sequence belong to which collection.

SECTION: DNA extraction from dried fungi
2. 
 

Start with laying out (96) 0.2 mL-tubes (12X 8-strips) with individual caps on a 96-well PCR plate rack. Label the tubes with their row and column number in the plate, i.e. a combination of columns 1-12 and row A-H (1A-12H). Also label each 8-strip with the pIate number. It is convenient to use two plate racks, one as your current workspace and one with your completed strips. 


 

SECTION: DNA extraction from dried fungi
9. Save point: The tubes with diluted DNA will be used in the following section (PCR protocol) and can be stored in the fridge overnight. For long term storage both the tubes with tissue and dilution can be stored in the freezer without degrading the DNA. The DNA will only degrade after repeated freezing and thawing which should not be a problem here. 
SECTION: PCR Protocol
19. Save point: The primer plate should be stored in the freezer. The PCR plate can after amplification be stored in the fridge for short term storage (a few days) and in the freezer for long term storage. IMPORTANT: The PCR plate can not be stored before amplification as the primers will malfunction.
SECTION: Post PCR pooling &amp; cleaning
34. Save point: Tubes with cleaned amplicons can be stored in the fridge until the start of the next section.
SECTION: Post PCR pooling &amp; cleaning
25. Save point: Store the pool tubes in the fridge until you have successfully completed the protocol. If something goes wrong during the rest of the protocol, the pool tubes are your lifeline and you can start over from this point. When the protocol is completed, they can be stored in the freezer.
SECTION: End-prep
52. Save point: End-prepped DNA can be stored in the fridge until the start of the next section.
SECTION: Native barcode ligation
73. Save point: If the protocol is too long to be completed in one go, the barcoded sample can be stored at 4 °C overnight.
SECTION: Adapter ligation and clean-up
91. Save point: The prepared library is used for loading on the the flowcell of the nanopore device. Store the library at 4 °C for short term or repeated use. For long term (more than 3 months) or single use store the library at -80 °C in LowBind tubes.
SECTION: PCR Protocol
15. Add the Phusion DNA polymerase to the mastermix. It is now ready to be added to the PCR plate.