General construction of double-indexed fusion primers for pair-end sequencing on MiSeq:

P5 (adapter)         linker index i5   SBS3 (sequencing primer, read 1) template primer fwd
AATGATACGGCGACCACCGA GATCT  xxxxxxx  ACACTCTTTCCCTACACGACGCTCTTCCGATCT ZZZZZZZZZZZZZZZZZZZZZZZ

P7                    linker index i7  SBS12 (sequencing primer, read 2) templ primer reverse

CAAGCAGAAGACGGCATACGA GAT    xxxxxxx  GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT YYYYYYYYYYYYYYYYYYYY

This construction is based on the TruSeq chemistry. At ICBR, they know it but it might be worth emphasizing when you submit your libraries. For more details on preparing a library using these fusion primers go here.

The steps of pair-end sequencing are as follows:

1. SBS3 priming, read 1 sequencing
2. i5 sequencing
3. i7 sequencing
4. SBS12 priming, read 2 sequencing

Therefore, the i7 index is the one that’s read first. We already have 96 versions of the read 2 primer, the one with the i7 index. The i5 index can multiply your multiplexing power.

Issues with base diversity

Illumina machines have trouble detecting signal from individual DNA clusters if all/most clusters have the same base across the lane. This is especially problematic with amplicons, where most sites will have the same letter in most reads. It can result in unclear signal. That is important to avoid especially in the indexes and at the beginning of the read when the instrument is still calibrating itself. There are two strategies to avoid low-quality reads due to homogeneity of DNA:

1. Spiking with PhiX (a phage genome) – ICBR does that automatically, but you need to tell them that you have an amplicon, so they add more. That works, but it wastes sequencing power.
2. Make the sequences mutually off-set, either by adding heterogeneity spacers, or by using sliding template primers. We use the latter. It is possible is the surrounding of the priming site is also fairly conserved. An example of a sliding LR0R primer (fungal LSU) is here:

priming site on template: 5'...GCCACCCGCTGAACTTAAGCATAT..3'
original LR0R:                    ACCCGCTGAACTTAAGC
LR0R(1):                           CCCGCTGAACTTAAGCA

The first PCR (to enrich for your fragment) should be done with the same uniform template primer, but these sliding primers can be incorporates into the fusion primers. They assure that the reads are off-set by one position which creates the desired base diversity at every sequencing step.
It is also very important to make sure that indexes  re ballanced

Issues with read length

Our primers

Here are ALL the barcodes/indexes you can choose from. Although the combination of two indexes should unambiguously identify the original sample, it is not a bad idea to use unique indexes throughout, both for the i5 and i7 segments:

Nested PCR

1. The initial DNA extract template is subjected to amplification with the template-specific primers LR0R and JHY-LSU-369rc for 30 cycles. The number of cycles is lower than usual to prevent various biases resulting from excessive amplification.
2. PCR reactions are not cleaned up, to avoid loss of DNA (we used to do it, but it is probably not necessary).
3. 5μl of each amplicon is run on a gel to assess failures/successes. You may not see very much on your gel! Even faint band should be used. When band is invisible, I wouldn’t use it even though there may still be some amplicon – it was probably not a good unbiased amplification.
4. A 1-μl aliquot of each successful amplicon is used as a subsequent PCR template and subjected to a 20-cycle amplification with the fusion primers.
5. The secondary (final) amplicons are cleaned up with AmPure DNA cleanup kit.
6. The DNA concentration in each cleaned PCR product is measured using the PicoGreen-based Quant-iT kit.
7. Products are pooled in equimolar proportions to create an amplicon library at a final concentration of 10 ng μl−1.
8. The pooled amplicon library is cleaned up using UltraClean PCR clean-up kit and submitted to ICBR.
9. When submitting to ICBR, make sure you specify that we need raw output, non-demultiplexed fastq data.

The LSU fusion primers

LR0R –> JH-LSU-369rc: From 5’ of the first primer site to the 3’ end of the reverse primer: 400bp. The most variable region (positions): 100-240

Primer construction
To prepare an order, simply concatenate. You need one Forward, and as many Reverse as you need barcodes. We order form IDT, which only does synthesis at 100nM for oligos this long. I have negotiated this synthesis for the price of 25nM synthesis, so do purchases through me. Specify that you want them in the “Lab Ready” suspension.
Forward primer:
P5 AATGATACGGCGACCACCGA
Linker GATCT
SBS3 ACACTCTTTCCCTACACGACGCTCTTCCGATCT
LR0R ACCCGCTGAACTTAAGC

Reverse primer:
P7 CAAGCAGAAGACGGCATACGA
Linker GAT
Barcode xxxxxxx
SBS12 GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
JH-LS-369rc CTTCCCTTTCAACAATTTCAC

Barcodes:
We already have the JH-LS-369rc fusion primers with these barcodes :
1 AAGGTGG
8 GCTGAGG
15 ACAGCGG
23 CATGGTG
31 TCGGTTG
35 CGAGATG
41 TGTGCTG
45 CTAGGAG
51 TATGTAG
56 GTGGAAG
60 TAATCAG
64 TCTGGCG
70 CCAGTCG
75 CTGTACG
78 GATGCCG
80 ACGTGGT
85 TCTGTGT
93 TAAGAGT
98 GTTGCGT
103 ATTGGTT
109 CTGGATT
114 TAGGCTT
120 GGTGGAT
127 CAAGTAT
134 TCGTAAT
138 CCGGCAT
142 AATTGCT
146 ACGGTCT
150 GAGGACT
155 TTAGCCT
158 CATTGGA
167 GATCTGA
176 GCTGGTA
181 AGTGACA
189 TTCAGGC
202 GACGGTC
Other available barcodes:
2 CTTGTGG
3 TGAGTGG
4 CAATTGG
5 TTCTTGG
6 GGTATGG
7 GTGCTGG
9 ATCGAGG
10 GAGTAGG
11 TGTTAGG
12 AGGAAGG
13 TCGCAGG
14 AATCAGG
16 TACGCGG
17 ATGTCGG
18 GGATCGG
19 CAGACGG
20 GTCACGG
21 CGTCCGG
22 TTACCGG
24 TAGTGTG
25 AGATGTG
26 CTCTGTG
27 ACTAGTG
28 TGCAGTG
29 ATGCGTG
30 GCACGTG
32 GTAGTTG
33 AGCGTTG
34 CTGATTG
36 ACGTATG
37 TTATATG
38 GTTAATG
39 AACAATG
40 CAGCATG
42 GCCGCTG
43 AATTCTG
44 CCATCTG
46 CCTTGAG
47 GGAAGAG
48 CACAGAG
49 TGGCGAG
50 GATCGAG
52 GCGTTAG
53 ATATTAG
54 TCCATAG
55 AGTCTAG
57 TGCGAAG
58 CTCCAAG
59 ATTGCAG
61 AGCTCAG
62 GCTACAG
63 AAGCCAG
65 GTCGGCG
66 AAGAGCG
67 CTTAGCG
68 CCGCGCG
69 TAACGCG
71 TGGTTCG
72 AGAATCG
73 CATCTCG
74 ATCCTCG
76 GCATACG
77 GGCAACG
79 TCGACCG
81 CTATGGT
82 CGTAGGT
83 GACAGGT
84 AGACGGT
86 ATAGTGT
87 TAGTTGT
88 GCATTGT
89 AGCTTGT
90 CCGATGT
91 AATATGT
92 CTCCTGT
94 CACTAGT
95 GTGAAGT
96 TCCAAGT
97 CGGCAGT
99 CGAGCGT
100 CCTTCGT
101 TGGACGT
102 GAGCCGT
104 TCAGGTT
105 GTGTGTT
106 AGGAGTT
107 CCTCGTT
108 TACCGTT
110 GGCGATT
111 GCTTATT
112 TATAATT
113 AGTCATT
115 CGGTCTT
116 ATCTCTT
117 GCGACTT
118 CTTACTT
119 GTACCTT
121 AACGGAT
122 GAATGAT
123 CGCTGAT
124 TTGAGAT
125 ACAAGAT
126 GCGCGAT
128 GCCGTAT
129 CTGTTAT
130 TGATTAT
131 GAGATAT
132 GTTCTAT
133 AGCAGGA
135 GTCTAAT
136 CATCAAT
137 GGACAAT
139 AGTACAT
140 TACACAT
141 TCTCCAT
143 TTCTGCT
144 GCTAGCT
145 TGAAGCT
147 CGTGTCT
148 TCACTCT
149 GGCCTCT
151 AGATACT
152 CTCAACT
153 TTGCACT
154 AACCACT
156 GGTTCCT
157 ATGACCT
159 GCCTGGA
160 CTGAGGA
161 TAGCGGA
162 ACTCGGA
163 GTACGGA
164 CGCGTGA
165 TTAATGA
166 AGGCTGA
168 CCACTGA
169 CTAGAGA
170 GACGAGA
171 CCGTAGA
172 TTCCAGA
173 AATGCGA
174 TGCTCGA
175 TCTACGA
177 TCGAGTA
178 GTCAGTA
179 CGGCGTA
180 ACAGTTA
182 TATTACA
183 CGGAACA
184 CTTCACA
185 CTGGCCA
186 ACCGCCA
187 TCATCCA
188 TTCACCA
190 GTCGTGC
191 CAGCTGC
192 TGTCTGC
193 ACGGAGC
194 CGTGAGC
195 TTGTAGC
196 GGCTAGC
197 GTTCAGC
198 AGCGCGC
199 GATTCGC
200 CTCTCGC
201 GCACCGC
203 ACCTGTC
204 GGTAGTC
205 AATCGTC

PCR mastermix

1. Use the cycling conditions labeled as M13 (under Lab or Martin) on Hulcr lab thermocycler.
2. For extra care (ie publications), prepare a replicate PCR for each extraction.

Running the gel

1. Prepare a 1% agarose gel (careful, it will be fragile). Make sure the gel is level when pouring!
2. Use extra reagents, but make sure the volume will fit in the comb you use for making the wells:

1. When loading, arrange samples from similar morphotypes together. If replicate PCRs were prepared, do not group them directly next to corresponding replicate sample.
2. Place both a 100bp and 1000 bp ladder on either side of every 5-10 reactions ran. This will aid in reading the gel, and will help to show if the DNA is moving at different speeds in different parts of the gel.
3. Run for up to 3 hours at 100-110V. Check every 45min-1 hour and take photo. The longer you run the gel, the more differentiated the bands will become, making it easier to read.

Reading the gel:

1. Use a photo editor (ie photoshop) to adjust the contrast/brightness so that all bands can be seen and differentiated between samples.
2. Use a line tool in the photo editor to connect bands in ladders with the same lengths. Use the slope of these lines to gage the distances traveled between samples.
3. Note any differences in fragment location and presence/absence between samples. Same species should be basically identical in their arrangement. Samples that appear different should be sequenced at a variable locus for confirmation.

Currently, GENEWIZ offers the best rate for sequencing 48 or fewer reactions with clean up. If you are submitting 48 or less, please follow the GENEWIZ protocol below, using 8 strip cap tubes. If you have greater than 48 samples, please submit using a prepaid plate through Eurofins, following the Eurofins protocol below.

You may prefer to use one company over the other for a multitude of reasons. For submitting larger numbers of samples to GENEWIZ, the protocol is roughly the same, but you use semi skirted plates and strip caps. For submitting fewer samples to Eurofins, we have barcodes (used for loose 1.5mL vials, which they use for orders of less than 24 samples)and packaging material available. Please follow the guidelines for sample submission on the Eurofins Genomics website.

GENEWIZ

In general, we will be using the “Custom” option for GENEWIZ. This entails sending 10uL of diluted un-purified PCR product per sample, with 5uL of 5uM primer per sample. For most cases, we will opt to have clean up performed at the facility.

Primers: Send 5uL of 5uM primer per sample to be sequenced. To make a solution of 5uM in 5uL, mix 2.5uL of your 10x primer stock with 2.5uL PCR water. Multiply these values by your number of samples + 2 and send in one Eppendorf tube per primer. Label with the primer name and concentration (5uM).

Template: Send in 10uL of your un-purified PCR product per well. This can be diluted with water, 0.5uL PCR product mixed with 0.5uL water tends to work well for most. If your band is strong and you know your concentration is high, you can increase the dilution to conserve PCR product. If it is weak or you know the concentration is low, you can send in less dilute product.

Submitting samples:

1) Create or log in to your GENEWIZ account. If this is your first time creating it, use 11575668 as the PI/Institution ID. Contact Zach with your account email to be added to the Lab Group to share payment info and sequence results.

2) Create a sequencing order from your account page. Choose the options for “Same Day” and “Upload Excel Form.” Choose and download the “Custom Form.”

3) Fill out and upload the Excel form by typing in your sample names, primer names, and Primer Concentrations. Select your DNA Type, DNA Length, and Special Protocols from their drop down menus. You can skip DNA Concentration, GENEWIZ Primer, and Notes.

4) Select payment method. You may use your own P-card if you have one. If not, access to our Lab Group should allow you to use Zach’s.

5) Select the Newins-Ziegler Hall Mailroom as your sample pickup location.

6) Submit your request, here you can print off the form you will include with your samples. This will also give you vial labels if you are sending in samples in strip tubes, label these values on the sides. Plates do not need labels as the excel file assigns well positions. Use 8 strip domed caps to seal vials and plates. You can wrap in parafilm for extra security.

7) Place samples in ziploc sample bag with primers and the provided order form. Drop off your sample in the GENEWIZ box in the NZH mail room before 4pm on the day of submission for pickup. If you cannot make it by this time, send an email to DNASeq@genewiz.com on the day you drop it off to notify them so it will be picked up.

Pricing: For custom orders the cost is $4 per sample. Clean up is an additional $1. You can perform clean up yourself, however this saves us little money given the cost of ExoSAP.

Electrophoresis Gel Procedure

Materials:
TAE buffer (In transparent plastic bottle)
Erlenmeyer flask with cap (glass petri dish)
Weigh paper
PCR Grade Agarose (1% agarose in buffer)
Electrophoresis machine
2 gel combs

Procedure:
• Weigh 0.7 grams of agarose using the blue plastic spoon in the agarose jar, and put this in the flask.
• Measure 70 mL of TAE buffer and pour into the flask
• Heat the flask with cap in the microwave at 2 minutes until you hear a tinkling sound – at this point take it out and swirl. Use the green PPE glove.
• Repeat microwaving in increments until you hear the tinkling sounds. Keep repeating until the agarose is completely dissolved (it will not look like there are clear granules in the liquid).
• Let the flask cool after microwaving (you may run water over the flask to cool it faster). It should cool to about 50 degrees C before you pour it. The flask shouldn’t be painful to the touch.
• Ensure that the gel casting tray is clean and dry. It may be cleaned with DI water if dirty. Never use bleach to clean the acrylic.
• Ensure that the gel container is oriented so that the current will run from black to red.
• Check that the gel container is level using the rounded level.
• Ensure that the gel casting tray is oriented so that the seals are against the walls of the gel container.
• Pour the gel from the flask into the casting tray. Use a pipette tip to remove any bubbles.
• Include 2 gel combs in the notches in the tray. Note: the combs have different volume sizes and amount of wells on each side, so be sure to pick the well volume and number of wells you want.
• The gel takes 1.5 to 2 hours to fully solidify. It can be checked intermittently during this time, and will turn opaque when solid and dry.
• Turn the gel casting tray so that the well contents will flow from black to red. If oriented from red to black, the bands will just fall off of the gel into the buffer. Pour buffer so that it is touching the electrodes.
• Once buffer is covering everything, the combs may be taken out. Be careful not to rip the gel. Pull the combs directly upwards.

Filling Wells

Materials:
SYBR green (In freezer)
Loading dye
Ladder 100 BP (fridge)
Approximately 6 inches of parafilm
10 uL pipette and tips

Procedure:
• Beforehand, mix 300 uL SYBR green with 600 uL loading dye and keep in freezer.
• Centrifuge the PCR products and then put in a frozen rack. Centrifuge SYBR green mixture and put in the larger frozen rack wrapped in aluminum foil. It is light sensitive.
• Add SYBR green mixture in dots on the parafilm, with 2 uL of the mixture per dot. You will need the amount of dots for your number of samples plus 1 for the ladder. (8 dots for 5 beetles, a negative and positive control, and the ladder).
• Add 6-8 uL of ladder in the first dot only, and pull the liquid up and down in the pipette tip to mix. Note: if electrophoresing two rows of wells, it can be helpful to put a ladder in the first well of the second row as well as the first.
• Add 6 uL of each PCR product in each of the other dots and mix up and down in the pipette tip (It is important at this point to make sure you know what order your samples are in).
• Set the pipette to 9.5 uL and pull the first dot up with the pipette. Be careful not to introduce air bubbles. Add the liquid to the first well as vertically as possible, and being careful not to touch the gel. Don’t press the pipette button down the second half of the way so that air bubbles are not introduced. It is better to leave a very small amount of PCR product in the tip than to introduce bubbles.
• Repeat this with each of the dots into each of the wells.
• Slide the lid onto the gel container.
• Plug the red cord into the red outlet, and the black cord into the black outlet on the machine.
• Turn the machine on, and put on 100 volts for 45 minutes.
• Take a photo of the gel within 10 minutes after it finishes.

Gel Photos Procedure:
• Take the gel casting tray out of the gel container and place it into the tray of the Enduro machine. Make sure everything on the machine is closed, because it emits UV light to take the photo.
• Open the Enduro GDS software on the computer, and click capture image, then click illuminate. This should make the gel with bands visible. You can adjust the illumination to make the bands more visible. Once done adjusting, you can take a photo.
• Take the gel casting tray out of the machine, making sure the UV is off first.
• Gel can be thrown away, and the gel casting tray can be cleaned with DI water. Wipe the Enduro tray with DI water as well.

Preparing Gel

For the small tray, you need 60mL of 1% gel (0.6g of agarose to 60 ml of TAE buffer). Boil over in microwave (about 2 mins). Cool down under cold water, pour into the rig with combs, wait for about 15 min.

For the medium tray, you need 160mL of 1% gel (1.6g of agarose to 160mL of TAE buffer). Boil over in microwave (about 2 mins). Cool down under cold water, pour into the rig with gel combs, wait for about 15 min.

Preparing SYBR Green

For loading directly to the wells in the gel with your sample

From www.lonza.com: SYBR® Green I Stain can be added directly to the loading buffer at a final concentration of 1:1000. First prepare a 1:100 dilution of SYBR® Green I Stain in high-quality anhydrous DMSO. The 1:100 dilution can be stored in the freezer and reused. Add 1 μl of this dilution to 9 μl-10 μl of your sample before loading.

SYBR Green & Dye mastermix

For one sample, combine 0.5uL of 100x SYBR (labeled 1:100X) and 1uL of 6X Loading Dye. Mix that with 5uL of PCR product. Or, more typically, make and store a mastermix:

For mastermix, combine 300uL of 100x SYBR and 600uL of Dye.

Loading the Gel

Prepare ~1.5uL droplets of SYBR & Dye mastermix on parafilm, there are aliquots of this mix prepared by the lab manager in the shared reagent box in the door of the small freezer. Add 5uL of PCR product, mix by sucking up & down with the pipette. Load into slot on gel.

For GreenTaq PCR product (product is green): add 5uL of sample to respective droplets of only SYBR Green, mix by sucking up&down with the pipette. No other loading dye needed.

Run on about 110 Volts for 45 minutes for running about halfway down the gel. Lower time for a shorter distance. If there is not enough liquid in the gel tray, add TAE buffer, just enough to cover the gel.

A cheap and possibly mutagenic alternative to SYBR Green is 1:1000X Ethidium Bromide added directly to the agarose after cooling. WEAR GLOVES!
Load DNA ladder in the last well.

3D Printed gel combs

We have designs for 3D printed gel combs compatible with our gel trays. These combs are designed to match our multichannel pipettes and rest either on top of, or in the slots of, the gel trays. The designs have three pieces: (1) the comb itself, slotted to fit into one of the holder types and with single and dual lane options; (2) an on top of tray holder which allows you to place the comb at any distance down the tray, and (3) an in line holder which allows you to use the standard distance slots in the tray. Links to the 3D printable files can be found below. You may print these through the UF libraries for a low cost as student, faculty, or staff.

PLEASE DO NOT EDIT THESE DESIGNS. To prepare one for printing, simply select the piece you would like to print, choose “Export,” and export just that piece as an .stl file. This can then be uploaded to the printing lab. Do not try to download all three pieces as one file, each file must be a single connected piece. You can submit multiple files at once to have them printed at the same time.

Medium gel design

Diluting primers

Reconstituting lyophilized primers to create 100X stock:
Resuspend to 100 uM/L concentration: for 28.5 nM of lyophilized primer, add 285 ul water (10 times more, in uL).

Dilute to make working solution :
To 10uL of the working stock above, add 90 uL of water, to make 10 uM/L (or 10 pM/ul) solution.

Working concentration: Use 1uL of the working solution above in 25uL reaction (10/25=0.4pM per rxn).

Primers we use

R= A G (purine) D= G A T

Buying primers

ITS1 example CType: DNA Oligos – modified & non-modified
Oligo Name: …
Synthesis Scale: 0.025 µmol
Purification: Desalt
Sequence (5′ to 3′): …
Format: Lab-ready
Number of Tubes per Oligo: 1
Concentration (µM): NA

454 primers – NO LONGER IN USE

From IDT: to get reaction-ready concentration, get 1nm in 100uL (concentration: 10)
Old FLX chemistry Fungi.

Old FLX chemistry bacteria and archaea.

Barcodes on 515R primer ordered on Feb 06 2012 (order “515F-806R-454barcoded”). R primer mixed with each barcoded F primer.

A1 GTAGTGTCTAGC
A2 GTCGCTGTCTTC
A3 GTGAGGTCGCTA
A4 GTTGACGACAGC
A5 GTCACGACTATT
A6 GTCTGACAGTTG
A7 GTGTGCTATCAG
A8 TACAGATGGCTC
A9 TACTTACTGCAG
A10 GGATCGCAGATC
A11 GTAGACTGCGTG
A12 GTCAACGCGATG
B1 GTCTCTCTACGC
B2 GTGTCTACATTG
B3 GTATGTTGCTCA
B4 GTCTCATGTAGG
B5 GTGTACCTATCA
B6 TAGCTGAGTCCA
B7 GGCGACATGTAC
B8 GTAGATGCTTCG
B9 GGTGCGTGTATG
B10 GCTTACATCGAG
B11 GTATCCATGCGA
B12 TAACTCTGATGC
C1 TACTAATCTGCG
C2 TAGCCTCTCTGC
C3 GCTTGCGAGACA
C4 GTCATATCGTAC
C5 GTACGGCATACG
C6 GTATGCGCTGTA
C7 GTCTATCGGAGT
C8 GTGGCGATACAC
C9 TACACGATCTAC
C10 TACTGGACGCGA
C11 TAGCTCGTAACT
C12 GGCAGTGTATCG
D1 GTAGAGCTGTTC
D2 GCTTCATAGTGT
D3 TAACAGTCGCTG
D4 TACGTGTACGTG
D5 TAGCATCGTGGT
D6 GTACAAGAGTGA
D7 TACGCGCTGAGA
D8 TAGATCCTCGAT
D9 GCTGTGTAGGAC
D10 GGTCGTAGCGTA
D11 GTCTGGATAGCG
D12 GTGTGTGTCAGG
E1 TACAGTCTCATG
E2 TACTTCGCTCGC
E3 TAGGTATCTCAC
E4 GTCGTAGCCAGA
E5 GCTGTAGTATGC
E6 GGTCACTGACAG
E7 GTGATAGTGCCG
E8 GTATATCCGCAG
E9 GTCATTCACGAG
E10 GTCTTCGTCGCT
E11 GTGTTGCAGCAT
E12 TACATCACCACA
F1 TAGACTGTACTC
F2 GTAGCGCGAGTT
F3 GTCCATAGCTAG
F4 TAGTCGTCTAGT
F5 GGCTATGACATC
F6 GTGACCTGATGT
F7 TAGTGCTGCGTA
F8 GGTATACGCAGC
F9 GTAGCTGACGCA
F10 GTCGACTCCTCT
F11 GTGACTGCGGAT
F12 GTTCGCGTATAG
G1 TACGATGACCAC
G2 TAGATAGCAGGA
G3 TAGTGTGCTTCA
G4 GTTAGAGCACTC
G5 TACCGCTAGTAG
G6 TAGAGAGAGTGG
G7 TAAGCGCAGCAC
G8 TACTACATGGTC
G9 TAGCGACATCTG
G10 GGACGTCACAGT
G11 GTACTCTAGACT
G12 TACACACATGGC
H1 TACTGCGACAGT
H2 TAGCGGATCACG
H3 GTATGACTGGCT
H4 GTCTACACACAT
H5 GTGCACATTATC
H6 GGCGTACTGATG
H7 GTAGCAACGTCT
H8 TAGCACACCTAT
H9 TACGGTATGTCT
H10 GTTGTATACTCG
H11 GTCGTGTGTCAA
H12 GTGCAATCGACG

The complete order, including the barcodes above:

Titanium Lib-a chemistry, NOT YET USED, need to confirm correctness. Fungal LSU primers. Four-bp multiplex identifiers (MIDs). Only one-directional sequencing needed, from the more variable 3′ end, so only B adapters have multiple MIDs attached.

Again – Titanium Lib-A chemistry, not yet tested, Bacterial primers.

Picogreen on the RT PCR Machine

Per Chris Dervinis, Sr Biologist and Forest Genomics Lab Manager, SFRC, Univ. of FL :

“We use the sybr green filters for the assay, which is not mentioned in the protocol.  One thing to keep in mind is that the linear range for picogreen in these smaller assay is lower than it is with a plate reader and larger volume size, so you will need to be sure that your samples are under 50ng/ul or you will need to dilute for an accurate approximation.”

Plate Setup

1.Set up the standards using the Lambda DNA – Component C (100 ng/µl) with a total volume of 5 µl

For example: Make a 10 ng/µl Standard from the Lambda DNA

Add TE to the plate, according to the scheme below:

Add DNA Standard (10 ng/ μL – kit supplies DNA standard 100 ng/ μL) and the samples, according to the scheme bellow:

1.Add 25 µl of Picogreen solution into each well

1. Picogreen dilution: (1 Component A: 200 TE)

NOTE: Protect Picogreen solution from light (photo-sensitive)

1.Add the PCR flat caps over each column with the Strip Cap Tool (avoid touching the top of caps with your hands because the reading must pass through the caps)

1. Vortex and spin down

RT PCR Set up

Lamp Warm up takes 20 mins so have the lamp warming up before starting plate setup.

1.Open the realplex program (password is “e”). If the program is not recognizing the thermocycler, make sure the USB  connection to the computer is in the right port (both the port and the cord are marked with marker). All other connections can be checked based on the arrangement in the manual (on computer desktop).

1. Choose “Quantitative Plate Read”
2. Set the plate grid
   1. Highlight wells that are for standard and click on “Standard” (right hand side)
   2. Highlight wells that are for samples and click on “Unknown” (right hand side)
   3. Highlight ALL wells that will be read and click SYBR
   4. Set the values for the Standards (for example: 50, 40, 30, 25, 20, 15, 10, 0)
      1. Set the 2 standard as replicates (right hand side) so the 2 readings will be averaged for 1 point on the standard curve
      2. Change the quantity from “copies” to “nanogram” (right hand side)
         1. It will ask for the conversion; Click “Do Not Convert”
         2. Click Run
         3. Click Pre-Read
         4. To analyze the data, click “Analysis” on the top right hand side, click “Results” tab to view Standard curve, Text Report, etc.

Save and export data to powerpoint or excel

IMPORTANT: We now are performing clean up with our sequencing servicers GeneWiz and Eurofins. THis removes the need for clean up, as it is cheaper for them to do it. Please see the “Sequencing” protocol for more information on submitting your un-purified PCR samples for sequencing.

Use the following only with a single band PCR. With more bands, the only option is gel extraction & purification.

Exo-Sap-IT

Keep Exosap on ice and don’t keep out of freezer longer than necessary. This product doesn’t freeze so it is always ready to use straight out of the freezer.  Keep tubes in ice block until ready to put into thermocycler.

1)Per sample for bidirectional sequencing, use an ice block and in each PCR tube mix:

• 0.5uL PCR grade water
• 2uL PCR product

2) After all tubes are prepped, get tube of Exosap out of freezer and promptly add 1 uL to each tube. Immediately return the tube of Exosap to the freezer.

3) After spinning down, run ExoSap program on thermocycler (37C for 15 min, and 80C for 15 min). Depending on the thermocycler, you may need to change the volume of product in the program settings (or when prompted) to 3 uL.

4) When the cycle is finished, spin down again and put half the contents of each PCR tube (1.75 uL) into a 1.5 mL tube, and the remaining half into a second 1.5 mL tube. These will be your forward and reverse submissions for sequencing.

5) Label tubes with the appropriate database contig ID number (format ####-####) (not just the sequencing ID number) and update information in database.

Why the “contig ID” as the sequencing sample identifier?

Genious can easily assemble your forward and reverse sequences into contigs automatically and in batches. It is a one-button operation, and you don’t have to mess around with individual read editing.

What you need to do is to submit your sample names to ICBR in the right format: The name of each contig needs to be “seq_ID”-“PCR_ID”. This is already generated for you in the Isolations form “Sequencing”. Simply grab the contig_ID column that’s there automatically pre-filled, and put that on the spreadsheet that you are submitting to ICBR. Each forward sequence will have a unique seq_ID, but the PCR ID will be shared with the reverse sequence. That’s it.

Then, when you import your reads to Geneious, select the entire batch, go to Align/Assemble -> De Novo Assemble -> Assemble by 2nd position, separated by hyphen. Push Start .

You don’t have to spend an afternoon making arbitrary guesses as to which base is the right call. You also don;t have to trim the ends of the reads – the assembler does that automatically.

Gel extraction and clean-up

Our choice: E.Z.N.A. Gel Extraction Kit, Omega Bio-tek

• 1) cut out gel slice, put in a tube, and weigh it (actual weight minus one empty tube (0.90 g))
• 2) add equal volume of Binding Buffer (x g gel = x ml Buffer)
• 3) incubate at 55-60 C for 7 min (or until gel dissolves)
• 4) add 700uL to spin column, centrifuge at 10,000g for 1 min
• (5) keep adding 700uL of solution to the spin column and spinning until all added)
• 6) add 300uL of fresh Binding Buffer to the column, spin at 10,000 for 1 min
• 7) add 700uL of ‘Wash Buffer, centrifuge at 10,000g for 1 min, discard flow-through
• 8) same thing again: add 700uL of Wash Buffer, centrifuge, discard liquid
• 9) discard liquid, centrifuge the empty column for 2 min at max speed
• 10) put column in clean tube, add 30uL of Elution Buffer
• 11) let soak for 1 minute, then centrifuge at max speed for 1 min
• (12) (if maximum yield is more important than concentration, add another 30uL of Elution Buffer, centrifuge at max speed.)

Notes: When DNA yield is important, use fresh running TAE buffer. Old buffers have high pH and decrease yield.

If Buffer turns orange or red during incubation, pH is too high!

Useful overview of available kits here.

Genewiz folks recommend Qiagen kit, but that is expensive and people report low yields. If you need to load and separate the whole PCR reaction, make a thick gel.

MOBIO UltraClean PCR Cleanup

Protocol modified by Fierer Lab

• 1. Shake to mix the SpinBind before use. Add 5 volumes of SpinBind to your PCR reaction. (ex. 500uL to 100uL of PCR reaction)
• 2. Mix well by pipetting. If an oil overlay was used, you will now have two layers. The top layer is oil.
• 3. Transfer PCR/SpinBind mixture to a Spin Filter column tube, while avoiding the transfer of oil.
• 4. Centrifuge 10-30 seconds at a minimum 10,000 x g ~ 13,000rpm in a tabletop microcentrifuge.
• 5. Remove the Spin Filter basket and discard liquid flow through by decanting then replace basket back in to the same tube.
• 6. Add 300 ul SpinClean Buffer to the Spin Filter.
• 7. Centrifuge 10-30 seconds at a min of 10,000 x g
• 8. Remove the Spin Filter basket and discard liquid flow through by decanting then replace basket back in to the same tube.
• 9. Centrifuge 90 seconds at a min of 10,000 x g
• 10. Transfer Spin Filter to a clean 2ml Collection Tube
• 11. Add 50uL Elution Buffer (10mM Tris) solution provided or sterile water directly onto the center of the white Spin Filter membrane. The choice of Tris or water will not affect yield. DNA is more stable for storage in Tris. Let sit on filter for 5 minutes.
• 12. Centrifuge for 90 seconds at a min of 10,000 x g
• 13. Discard Spin Filter basket from the inside of the Spin Filter. Purified DNA is now in the 2mL Collection tube. The DNA will be free of all reaction components such as primer or linkers, enzyme , salt and dNTPs. Store DNA at -20 degrees Celsius. DNA is now ready to use.

AMPure XP PCR Cleanup with Ring Magnet Plate

Manufacturer says to store AMPure refrigerated (do not freeze) and use within 12 months .

Our Eppendorf skirted plates are NOT compatible with the magnetic ring plate. All other brands of 96 well plates we have on hand at the time of writing (VWR, Fisherbrand, Applied Biosystems) are compatible.

Magnet is very strong: 

• Must be kept well away from computers, thermal cyclers, centrifuge, cell phones and other electrical equipment, credit cards, etc. 
• Must be kept away from people with pacemakers and defibrillator implants.

Do not autoclave magnet plate – looses magnetic strength if heated above 80 C.

1. Gently shake the AMPure bottle to resuspend any magnetic beads that may have settled (color should be uniform)
2. In a 96 well PCR plate (note compatible brands above) add 1.8 uL AMPure  for every 1 uL of PCR product (e.g. add 45 uL Ampure to a 25 uL PCR reaction =70 uL total volume) Keep PCR plate off magnetic plate until step 4.
3. Mix AMPure and PCR product by pipetting 10x, then incubate at room temperature for 5 min.
4. Following incubation, place PCR plate into the magnetic ring plate, let sit for 2 minutes or until solution is clear.
5. Keeping the plate magnetized, remove and discard the clear solution via pipetting (do not disturb the ring of beads around the sides).
6. Add 200 uL 70% ethanol, let sit for 30 seconds, then remove and discard all ethanol without disturbing the beads. Repeat this step for a total of two washes.
7. Let stand for about 5 minutes for any remaining traces of ethanol to evaporate. Avoid allowing the ring of beads to dry for significantly longer, which will create a crackled appearance and inhibit elution.
8. Remove from magnetic plate and add a minimum of 40 uL of elution buffer (=whatever you are resuspending your product in – molecular grade water, TE, etc). Mix by pipetting 10x. All the beads do not need to go back into solution for full elution.
9. Place the plate back on the magnet and let sit for 1 minute to draw the beads back out of suspension.
10. Without disturbing the ring of beads, transfer your samples to a new plate/tubes for storage.

PCR Procedure

Materials:
DNA Extractions
Forward and reverse primers
Premix taq from freezer
Empty Eppendorf tube
Eppendorf tube of PCR water
Positive control (Specimen already known to be high quality)
Two 96-well plates
Eppendorf vial rack

Preliminary Procedure:
• Primers need to be diluted from 10 micromolar to 1 micromolar to be stocked in the freezer for 1-2 months.
• Put your sample information in the database to make new sample numbers for the PCR.
• Label the strip tubes with the new sample numbers using an alcohol resistant marker and put them into a PCR plate. Add extra labels at the end for positive and negative controls.
• Look in “DNA methods” section of lab website for cycling methodology.
• Look in the database to find primers in the freezer.
• Use the PCR excel chart to determine the amount of reagents you need to use in the master mix (the final volume of the master mix should equal 25 uL).
• UV the PCR hood for 15 minutes before starting to do the PCR.

Master Mix Procedure:
o Add x uL of Taq to the empty Eppendorf tube
o Add x uL of each primer to the tube, pipetting up and down to mix
o Add x uL of PCR water to the tube
o Vortex the master mix for 5 seconds

PCR Procedure:
• Vortex the extractions for 1 second, and then centrifuge to eliminate any bubbles.
• Make sure that the DNA extractions are in the same order that the empty PCR tubes are in.
• Pipette 23 or 24 uL of master mix into each PCR tube using the same tip.
• Pipette 1 or 2 uL of each DNA extraction into the PCR tubes. Use a different pipette tip each time, and mix the liquid up and down with the pipette. Try not to introduce bubbles. Close each cap when done pipetting so that none are skipped or none have double the correct amount added.
• Add 1 or 2 uL of PCR water only to the negative control tube.
• Close the tops of all the vials and make sure that all of them are closed. Sometimes they do not close all the way even if they look closed.
• Tap the plate to the table to bring drops in the tubes down.
• Centrifuge strip tubes for 3 seconds to make sure there are no bubbles.
• Put the tubes into the smaller holes of the thermocycler. Make sure the caps are completely closed. Add other tubes of the same kind to balance it.
• Run the program called for in the PCR protocol, and set a timer to be able to take the PCR products out and put them in the freezer.
• Put the extractions and any unused reagents back in the freezer in your box.
• The PCR products should be used as soon as possible.
• Clean everything with ethanol and UV the hood.
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Ensure you enter all PCR runs into the PCR form on the isolations database. You can use the form linked here to properly plan your PCR master mix and database entries. Use the database entry rows at the top to fill in your samples for one run, including a positive control (known working template for these conditions), an extraction negative control (negative control from extraction as template), and a PCR negative control (no template). Then put in the number of reactions in the master mix form, this will give you the necessary amount of reagents to make master mix for your reactions plus two additional.

It is recommended you input your PCR database entries before beginning mixing and your PCR run, this will reserve the spots so that the PCR IDs do not get taken while you are working.

PCR Form Template Excel

Extract-N-Amp PCR ready premix

Typical master mix:

• PremixTAQ 12.5uL, found in freezer door. Fisher Catalog: RR003A
• Primer F (Conc. 10uM) 1uL
• Primer R (Conc. 10uM) 1uL
• PCR H2O, aliquots made by lab manager and stored in shared reagent box in door of small freezer. 9.5uL
• DMSO, aliquots made by lab manager and stored in shared reagent box in door of small freezer. 1uL

Please keep aliquots you open and use in your box after initial use.

Primer stock (100uM) locations can be found the Primer_locations table in the isolations database, these are in the ThermoFisher -80C freezer and will need to be diluted into a 10uM aliquot.

Multiply volumes by number of samples + 2 to make a small amount extra (form above does this). Use 25uL master mix and 1uL template per sample. PCR cycle will vary with primers used.

Common primers and cycle combinations used in our lab. Number in cycle title is usually annealing temperature.

• ITS1F/ITS4 – PCR_55_safe
• LR0R/LR3 – PCR_55_safe
• Bt2a/Bt2b – PCR_52

Another master mix:

• XNA template 4uL (if less, add XNA solutions up to 4 uL!)
• primer F 0.5 uL (ideally 0.4uM)
• primer R 0.5 uL (ideally 0.4uM)
• XNA PCR ready premix 10 uL
• water 5 uL (or add up to 20 uL total)

Typical cycling conditions:

• 1) 94C, 3 min
• 2) 94C, 45s
• 3) xC, 45s
• 4) 72C, 1:30min
• 5) go to 2 34x

NEB Q5 High Fidelity polymerase

High fidelity polymerases have 3′->5′ exonuclease activity, and can chew up primers. Solutions:

1) increase primer concentration

2) protect primers’ 3′ end by phosphorothioate (2 or 3 last bases, not just one).

Non-specific annealing solutions:

1) Optimize annealing temperature.

2) minimize the time that the enzyme has to amplify annealed primers by working on ice, preheating your thermocycler to96C, and using a Hot-Start option (probably not necessary for Q5 which is not active at room temp.)

Typica

Clonetech LA Taq and Buffer

Start cycling with at least 94C for 1 min, can go up to 98C each cycle.

Please record all extractions correctly in the database. See database protocols for isolations for more information.

Sigma Extract-N-Amp

Spin-down protocol (for beetles)

Keep BSA and the Ex-n-A solution cold, on ice or in a pre-frozen rack.

With BSA as a binding agent

• beetle body parts (legs work well from specimens killed with 95% EtOH)
• 40uL of Extraction Solution per sample (in yellow tray in the freezer)
• run protocol Ex-n-Amp on thermocycler (96C for 30 minutes)
• add equal volume(to extract solution) of 3% BSA (will bind extra stuff)— (40uL of Extraction Solution:40uL of BSA)
  • vortex thoroughly
• spin down, store the upper half (30uL) as your final sample
• use the 0.5-1.0uL of this supernatant for PCR

BSA from Fermentas, #00066587, 20mg/mL =~2%

General Ex-N-Amp protocol for fungi:

1. Prepare strip tubes with 40uL of Sigma Aldrich extraction solution. This can be found in aliquots in the shared box in the door of the small freezer.
2. Scrape approximately 10uL of hyphae from fungal colony using a sterile pipette tip (20uL works best and is not used for much else) or scalpel.
3. Add fungal material to extraction solution and vortex.
4. Run Ex-N-A protocol on a thermocycler (96C for 30 minutes).
5. Spin down with high rpm.
6. Pipet off the top 25uL: that’s your extract. It is often safe to dilute this a bit if you will need more than 25 uL, but this is not standard procedure.

Note: If BSA is needed, we have a stock of 2% molecular grade BSA in our freezer. This should be made into aliquots before using for samples. This is not standard, and only used in cases where it increases DNA amplification success.

OLD: Spin-down protocol (for fungi) 

Keep BSA and the Ex-n-A solution cold, on ice or in a pre-frozen rack.

With BSA as a binding agent

• 10uL of original sample
• 20uL of Extraction Solution per sample (in yellow tray in the freezer)
• run protocol Ex-n-Amp on thermocycler (96C for 10 minutes)
• add equal volume(to extract solution) of 3% BSA (will bind extra stuff)— (20uL of Extraction Solution:20uL of BSA)
  • shake thoroughly
• spin it down, store the upper half (20uL) as your final sample
• use the 0.5-1.0uL of this supernatant for PCR

BSA from Fermentas, #00066587, 20mg/mL =~2%

Manufacturer’s protocol – DON’T USE, the Dilution Solution inhibits PCR

With genuine Sigma components. Sigma manual here.

1. 20uL of Extraction Solution per sample
2. 96C for 10 minutes
3. add equal volume of Dilution Solution
4. ready for PCR. Allegedly stable for several months (says Sigma).

PowerLyzer

PowerLyzer Ultra Clean kit from MoBio, catalog # 12255-50

Prepare:

12 samples per one round

ice bucket, or pre-frozen tray in a fridge

tube heater (dry bath) at 65C

number all your tubes on top (four per sample, Bead tubes, Spin Filters, and the final PCR tubes)

Steps

1. spin down sample for 1 min at max speed
2. take 50uL of sediment and add it to bead tubes (with MicroBead solution and MD1)
3. heat at 65C for 10 min
4. while waiting, pre-aliquote solutions into tubes:

1 beads + 300uL MicroBead + 50uL MD1

2 100uL MD2

3 900uL: MD3

5. shake on MoBio vortex adaptor at full speed for 2 minutes

6. continue according to MoBio manual

…

19. transfer yield into labeled 250uL PCR tubes. This is our final sample.

Alternative strategies for small volumes of DNA

• Direct PCR may work too, but not if lots of DNA is present. For direct PCR add 10 – 15 minutes a 95C at the beginning of the PCR.
• Potassium xanthogenate method, Peter Young in York
• Forensic kits, e.g., punch cards

CTAB DNA Extraction