TITLE: Fuhrman Lab 515F-926R 16S and 18S rRNA Gene Sequencing Protocol
AUTHORS: David Needham, Erin Fichot, Alma Parada, Yi-Chun Yeh, Jed Fuhrman

[DESCRIPTION]
<div class = "text-blocks"></div>

[STEPS]
?. [PCR]
Use UV-Crosslinker to treat consumables. Press “Time”, then “10.0” to run for 10 minutesPCR strip tubes (cat# )PCR waterAny tubes needed to make a master mix
?. [Assess Amplification]
Preparing gel1.0 g agarose100 mL TAE buffer (or TBE)Swirl to mix agarose and buffer
?. [Assess Amplification]
Microwave 1 minute, swirl. If solution is not clear, microwave in 10 sec increments and swirl. **caution, hot**
?. [PCR]
Wipe down inside of the PCR hood and all pipettes with 10% bleach. Turn the UV light on (15 min).
?. [PCR]
Make a master mix of PCR water and 5’ Master Mix (5% extra to account for pipetting error)Flick-mix, spin tubePipet 22.5 µL into each PCR strip tube
?. [PCR]
Add 1.5 µL of primer mix to each tubeFor primer plate, use a Kimwipe to wipe 10% bleach on aluminum foil coverThen use Kimwipe to wipe 70% EtOH on aluminum foil coverWait for EtOH to evaporate before pipetting through aluminum foil.When done, place new foil directly on top of punctured foil.
?. [PCR]
Add 1 µL of template DNA to appropriate tubesDo not add mock community template in the clean hood, since these are pooled PCR products and could conceivably cause contamination of the hood. Instead, wait until all other sample tubes are closed, take everything out of the hood, add mock community template to appropriate tubes, then proceedAmplify ≥ 1 even and ≥ 1 mock community per sequencing run
?. [PCR]
Flick-mix tubes, then centrifuge briefly.
?. [PCR]
Place tubes in thermocycler:  ABC1Initial Denaturation:
95°C&nbsp; 120s
**Our default PCR setup is 30 cycles with 0.5ng template230 Cycles** of:
95°C&nbsp; 45s
50°C&nbsp; 45s
68°C&nbsp; 90s
3Final Elongation Step:
68°C&nbsp; 300s
4Refrigeration: &nbsp;&nbsp;&nbsp;&nbsp;&nbsp;
4°C&nbsp;&nbsp; forever
ABC1Initial Denaturation:
95°C&nbsp; 120s
**Our default PCR setup is 30 cycles with 0.5ng template230 Cycles** of:
95°C&nbsp; 45s
50°C&nbsp; 45s
68°C&nbsp; 90s
3Final Elongation Step:
68°C&nbsp; 300s
4Refrigeration: &nbsp;&nbsp;&nbsp;&nbsp;&nbsp;
4°C&nbsp;&nbsp; forever
?. [Assess Amplification]
Add 7 µL SYBR-safe dye, vortex.
?. [Assess Amplification]
Prepare casting tray, making sure it is level and that orange gaskets are arranged properly
?. [Assess Amplification]
Pour gel into tray
?. [Assess Amplification]
Add comb(s)
?. [Assess Amplification]
Preparing SamplesCut strip of parafilm.
?. [Assess Amplification]
Pipet 2 µL loading dye onto parafilm, one dot per sample
?. [Assess Amplification]
Pipet 3 µL of sample into loading dye, pipet up and down 2 times.
?. [Assess Amplification]
Loading and running gelPlace solidified gel in proper orientation (samples running down toward you)
?. [Assess Amplification]
Pour used TAE running buffer (or TBE, if that was used to make gel) into chamber, fill until it just covers gel
?. [Assess Amplification]
Remove comb
?. [Assess Amplification]
Pipet 5 µL of 100 bp ladder into first well (left)
?. [Assess Amplification]
Pipet 5 µL of dye/sample into each remaining well
?. [Assess Amplification]
Plug in electrodes to power source
?. [Assess Amplification]
Run for ~100 V, 30-45 minutes.
?. [Assess Amplification]
What to look forAmplification ~563 bpMay see 2 bands if eukaryotic sequences are present (band will be about 200 bp longer)No-template-control PCR blanks, no amplificationPrimer dimer may be present, but should be removed upon clean-up (below)
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
If cleaning only a few samples at a time, use the following protocol (Ampure Beads). Otherwise, clean PCR reactions with SequalPrep Normalization Plate (Invitrogen A10510-01).
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Before beginning:Warm aliquot of beads at RT for 30 minutes.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Spin down the PCR rxns tubes using the microcentrifuge (brief spin)
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Bring up volume of remaining PCR product to 40 µl with TE
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Should have 22 µL PCR product remaining, so add 18 µL TE
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Label another set of PCR tubes for collection of cleaned amplicons
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Label another set of PCR tubes (or a plate) for 1:5 dilution of cleaned amplicons (to be used for quantification by Pico Green assay, see below)
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
To this set of tubes, add 4 µl of TE
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Once beads are warm, add the beads to 0.8 x ratio in PCR tubes
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Vortex strip tubes for several seconds
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Allow the beads and DNA to bind by waiting 5 minutes.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
After 5 minutes, place the tubes in magnetic separator.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Wait 3 minutes for full separation
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Remove and discard the clear buffer
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Add 200 µL of freshly-made 80% ethanol. (Ethanol is hygroscopic such that it will pull water out of the air and be diluted. While is this probably more of a consideration for 100% Ethanol, it is good lab practice to remake fresh 80% Ethanol.)
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Vortex the tube for 5-6 seconds. Not all of the pellet will be dislodged, but that’s ok. Let the ethanol and beads incubate for about 3 minutes.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Again, place the 500 µL tube on the magnet for separation.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Remove the ethanol and keep tubes on magnet.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Repeat the addition of 200 µl of Ethanol.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Remove the ethanol (no incubation necessary).
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Remove the tube and spin for several seconds to collect the remaining ethanol in the bottom of the tube. Put back on magnetic plate
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Remove the remainder of the ethanol
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Allow to dry on magnet with open tube for ~5 minutes. The beads should not crack, you will lose DNA, supposedly.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Add 10 uL of TE Buffer to the beads and pipette several times making sure to break up all of the bead pellet. The TE buffer elutes the DNA from the beads
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Incubate for about 5 minutes, separate on the magnet.
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Collect your DNA by pipetting off about 9.5 µL of the TE (being careful not to remove any of the beads, though, it is apparently ok if you get a tiny amount, but not ideal)
?. [Clean-Up: Agencourt AMPure XP PCR Purification]
Add 1 µL of this to the 4 µL of TE set aside for the 1:5 dilutions to be used for quantificationNo need to dilute blanksStore cleaned PCR products at 4°C if using within a week. Otherwise, freeze at -20°C.
?. [Quantification with Pico-Green dsDNA Quant-iT Assay Kit]
Samples need to be in the range of the standard curve (0-15 ng/µL), otherwise the quantification is not accurateIn general, well-amplified PCR samples are around 10-40 ng/uL, so it is best to run 1:5 dilutions of cleaned PCR products to get them within the range of the standard curve  1:5 dil of 10 ng/µL = 2 ng/µL
 1:5 dil of 40 ng/µL = 8 ng/µL
  Probably best not to dilute no template controls (blanks). One might  get a better idea of if there might be contamination present. Note, no template controls  usually do have measurable amounts of DNA, but we think this is likely from primer dimer or another source. See further details below regarding blanks.
?. [Quantify]
Invitrogen P7589 Kit Components20X TE bufferpico-green dyeDNA lambda standard (100 ng/µL)
?. [Quantify]
Before beginningWalk to the ice machine with ice bucket, retrieve ice.
?. [Quantify]
Locate reagents in fridge and place on ice. Keep Pico Green fluorescent reagent in the dark.
?. [Quantify]
Aseptically remove 96-well plate from bag and place optically-clear strip caps over all wells
?. [Quantify]
Cut plate so that only the number of wells that are needed needed, are used (keeping other capped wells for later).
?. [Quantify]
ProcedureTurn on Stratagene ≥ 20 min before runningOpen softwareSelect "quantification plate"Make sure lamp is warming up (light bulb icon will be yellow or green, not red)
?. [Quantify]
Dilute TE from kit according to the number of samples to be quantified.  ABCDEFGHIJKL1&nbsp;
# samples&nbsp;→
10
20
30
40
50
60
70
80
90
100
2&nbsp;
water (µL)760
2x522.5
2x665
2x807.5
2x950
3x760
3x823.3
3x918.3
4x760
4x831.2
3TE
20X stock (µL)40
55
70
85
100
120
130
145
160
175
4&nbsp;
total volume (mL)800
1100
1400
1700
2000
2400
2600
2900
3200
3500
5&nbsp;
tube size
2 mL
2 mL
2 mL
2 mL
2 mL
5 mL
5 mL
5 mL
5 mL
5 mL
ABCDEFGHIJKL1&nbsp;
# samples&nbsp;→
10
20
30
40
50
60
70
80
90
100
2&nbsp;
water (µL)760
2x522.5
2x665
2x807.5
2x950
3x760
3x823.3
3x918.3
4x760
4x831.2
3TE
20X stock (µL)40
55
70
85
100
120
130
145
160
175
4&nbsp;
total volume (mL)800
1100
1400
1700
2000
2400
2600
2900
3200
3500
5&nbsp;
tube size
2 mL
2 mL
2 mL
2 mL
2 mL
5 mL
5 mL
5 mL
5 mL
5 mL
?. [Quantify]
Dilute pico-green dye from kit according to the number of samples to be quantified  ABCDEFGHIJKL1&nbsp;
# samples&nbsp;→
10
20
30
40
50
60
70
80
90
100
2&nbsp;
1x TE (µL)398
547.25
696.5
845.75
995
2x597
2x646.8
2x721.4
2x796
2x870.6
3pico
200X stock µL)2
2.75
3.5
4.25
5
6
6.5
7.25
8
8.75
4&nbsp;
total volume (µL)400
550
700
850
1000
1200
1300
1450
1600
1750
5&nbsp;
tube size
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
ABCDEFGHIJKL1&nbsp;
# samples&nbsp;→
10
20
30
40
50
60
70
80
90
100
2&nbsp;
1x TE (µL)398
547.25
696.5
845.75
995
2x597
2x646.8
2x721.4
2x796
2x870.6
3pico
200X stock µL)2
2.75
3.5
4.25
5
6
6.5
7.25
8
8.75
4&nbsp;
total volume (µL)400
550
700
850
1000
1200
1300
1450
1600
1750
5&nbsp;
tube size
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
?. [Quantify]
Dilute DNA stockDo serial dilutions, briefly vortexing and spinning down after each dilutionMake in 0.5 mL LoBind tubesKeep on ice  ABCD1&nbsp;
10 ng/µl1 ng/µl0.1 ng/µl21x TE (µL)18
36
18
3volume of stock (µL)2
4
2
4total volume (µL)20
40
20
5stock (ng/µL)100
10
1
ABCD1&nbsp;
10 ng/µl1 ng/µl0.1 ng/µl21x TE (µL)18
36
18
3volume of stock (µL)2
4
2
4total volume (µL)20
40
20
5stock (ng/µL)100
10
1
?. [Quantify]
Make standard curveDo in duplicate (columns 1+2 on plate)Keep on ice until ready to run  ABCDEFGHI196-well row #&nbsp;→
H
G
F
E
D
C
B
A
2[final]
0
0.1
0.5
1
5
7.5
10
15
31x TE (µL)15
14
10
14
10
7.5
14
13.5
4volume of stock (µL)0
1
5
1
5
7.5
1
1.5
5total volume (µL)15
15
15
15
15
15
15
15
6stock [] ng/µL---
0.1
0.1
1
1
1
10
10
ABCDEFGHI196-well row #&nbsp;→
H
G
F
E
D
C
B
A
2[final]
0
0.1
0.5
1
5
7.5
10
15
31x TE (µL)15
14
10
14
10
7.5
14
13.5
4volume of stock (µL)0
1
5
1
5
7.5
1
1.5
5total volume (µL)15
15
15
15
15
15
15
15
6stock [] ng/µL---
0.1
0.1
1
1
1
10
10
?. [Quantify]
Load 14 µL of 1x TE to each well to be used for sample measurement
?. [Quantify]
Load 1 L of sample to appropriate wellEx:  ABCDEFGHI1H
G
F
E
D
C
B
A
&nbsp;
2std-0
std-0.1
std-0.5
std-1
std-5
std-7.5
std-10
std-15
1
3std-0
std-0.1
std-0.5
std-1
std-5
std-7.5
std-10
std-15
2
4sample 1
sample 2
sample 3
sample 4
sample 5
sample 6
sample 7
sample 8
3
5sample 9
sample 10
sample 11
sample 12
sample 13
sample 14
sample 15
sample 16
4
6&nbsp;
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5
7&nbsp;
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6
8&nbsp;
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7
9&nbsp;
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8
10&nbsp;
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9
11&nbsp;
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10
12&nbsp;
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11
13&nbsp;
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&nbsp;
&nbsp;
12
ABCDEFGHI1H
G
F
E
D
C
B
A
&nbsp;
2std-0
std-0.1
std-0.5
std-1
std-5
std-7.5
std-10
std-15
1
3std-0
std-0.1
std-0.5
std-1
std-5
std-7.5
std-10
std-15
2
4sample 1
sample 2
sample 3
sample 4
sample 5
sample 6
sample 7
sample 8
3
5sample 9
sample 10
sample 11
sample 12
sample 13
sample 14
sample 15
sample 16
4
6&nbsp;
&nbsp;
&nbsp;
&nbsp;
&nbsp;
&nbsp;
&nbsp;
&nbsp;
5
7&nbsp;
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6
8&nbsp;
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7
9&nbsp;
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8
10&nbsp;
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9
11&nbsp;
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10
12&nbsp;
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11
13&nbsp;
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&nbsp;
12
?. [Quantify]
Add 15 µL of diluted pico-green dye to each well
?. [Quantify]
Briefly vortex and spin, and keep in the dark for ≥ 5 minutes
?. [Quantify]
Read fluorescence on stratagene (typical SYBR-like fluorescence)Mark wells used for standards and type in actual concentration, click on SYBRMark wells used for samples as “unknown”s, click on SYBRCheck Gain settings—typically use 4 but can use 8 if you need more light“Run”
?. [Quantify]
What to look for after completion:Std curve  Duplicates should be close to each other (can remove if necessary)R2 should be ≥ 0.95Should be linear (can remove higher concentrations if necessary to keep linearity)Samples  Values should fall within the corrected standard curve  If not, they need to be re-diluted and quantified againTo calculate actual concentration, need to multiply quantification by dilution factor  For example, if pico value for 1:5 dilution gave a concentration of 5.7 ng/µL, 5.7 x 5 = actual undiluted concentration of 28.5 ng/µLBlanks typically have between 0 and 0.6 ng/µL in them after cleanup, likely from primer dimer
?. [Dilute]
Based upon quantification, dilute each sample to equimolar concentrations with TE, 1 ng/µL.
?. [Dilute]
Calculate how much TE is needed to add to 1 µL of un-diluted, bead-cleaned PCR product to get a final concentration of 1 ng/µLEx:  ABCDEF1sample
measured 1:5 dilution [], ng/uL
calculated full [], ng/uL
vol un-diluted, bead-cleaned PCR product for dilution (uL)
vol TE for dilution (uL)
dilution [] ng/uL
21
4.6
23.0
1
22.0
1
32
2.1
10.5
1
9.5
1
43
7.9
39.5
1
38.5
1
54
5.2
26.0
1
25
1
6blank
n/d
0.1
5
0
0.1
ABCDEF1sample
measured 1:5 dilution [], ng/uL
calculated full [], ng/uL
vol un-diluted, bead-cleaned PCR product for dilution (uL)
vol TE for dilution (uL)
dilution [] ng/uL
21
4.6
23.0
1
22.0
1
32
2.1
10.5
1
9.5
1
43
7.9
39.5
1
38.5
1
54
5.2
26.0
1
25
1
6blank
n/d
0.1
5
0
0.1
?. [Dilute]
Accurately pipet calculated volume of TE into PCR plate according to calculations
?. [Dilute]
Accurately pipet 1 µL of undiluted, bead-cleaned PCR product into TE
?. [Dilute]
Cap, vortex, and spin 1 ng/µL dilutions
?. [Dilute]
Store cleaned PCR products at 4°C if using within a week. Otherwise, freeze at -20°C.
?. [Pool]
Pool by adding 1 µL of each sample into one 1.5 mL LoBind tube.Don’t forget blanks and mock communitiesEasiest to collect all samples from all users for run into one pool, instead of individual users each creating own cleaned poolCurrent MiSeq requirements at UC-Davis (10/2015)  15 µL5 nM (~1.8 ng/µL)If we pool 1 µL of each 1 ng/µL sample, we need 27 samples to meet this requirement, but since we like to pool enough to send half and keep half, we need ≥ 63 samples to meet this requirement (35 µL total vol)Store pool at 4°C if using within a week. Otherwise, freeze at -20°C.
?. [Clean and Concentrate]
Clean pool with SPRIselect (Beckman Coulter B23317) beads.SPRIselect beads are just like the Ampure beads, but the SPRIselect beads are QC’d for precise size-selectionFollow same protocol as described above for Ampure beads, except  Elute with 35 µL TECollect elution into a well-labeled 0.5 mL LoBind tube  Pipet 15 µL of this into a separate, well-labeled 0.5 mL LoBind tube  This 15 mL tube is the one that will be sent for sequencingCurrent MiSeq requirements at UC-Davis (10/2015)  15 µL5 nM (~1.8 ng/µL)The remaining bead-cleaned pool will be used for:  On-site storageAssessment by:  Quantification (pico-green)Size distribution (bioanalyzer trace)Store 0.5 mL LoBind tube with 15 µL for sequencing at -20°C until sent for sequencingStore remaining cleaned pool (also in 0.5 mL LoBind tube) at 4°C if assessing via pico and bioanalyzer within a week. Otherwise, freeze at -20°C.
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
Sequencing pool needs to be assessed for quantity and size distribution before submitting.Invitrogen Q32854 Kit Components:Qubit dsDNA HS Buffer, RT200X Qubit dsDNA HS Reagent dye, RTHS Standard #1 (0 ng/µL), 4°CHS Standard #2 (10 ng/µL), 4°C
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
Before beginningGet iceLocate standards in fridge and place on iceLocate RT reagents and assay tubes
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
ProcedureLabel assay tube lids, 1 per sample and 2 for standards
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
Prepare assay solution, keep dark, RT  ABCDEFGHIJK1# samples&nbsp;→
1
2
3
4
5
6
7
8
9
10
2Buffer (µL)700
900
2x550
2x650
2x750
2x850
2x950
3x700
3x766.7
3x833.3
3200X dye (µL)3.5
4.5
5.5
6.5
7.5
8.5
9.5
10.5
11.5
12.5
4total volume (µL)703.5
904.5
1105.5
1306.5
1507.5
1708.5
1909.5
2110.5
2311.5
2512.5
5tube size
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
5 mL
5 mL
5 mL
ABCDEFGHIJK1# samples&nbsp;→
1
2
3
4
5
6
7
8
9
10
2Buffer (µL)700
900
2x550
2x650
2x750
2x850
2x950
3x700
3x766.7
3x833.3
3200X dye (µL)3.5
4.5
5.5
6.5
7.5
8.5
9.5
10.5
11.5
12.5
4total volume (µL)703.5
904.5
1105.5
1306.5
1507.5
1708.5
1909.5
2110.5
2311.5
2512.5
5tube size
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
2 mL
5 mL
5 mL
5 mL
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
Add assay solution to assay tubes (keep as dark as possible)Standards: 190 µLSamples: 199 µL
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
Add DNA to assay tubesStandards: 10 µLSamples: 1 µL
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
Vortex, spin, keeping dark
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
After ≥ 5 min, run on Qubit:Select correct assay: “Qubit dsDNA HS”Follow directions on prompt  When measuring samples, change volume of input DNA to 1 µL
  When finished, select “Check Calibration” to record actual fluorescence values for the standards.When measuring samples, change volume of input DNA to 1 µL
?. [Assess Pool: Quantification with Qubit dsDNA HS Assay Kit]
What to look for:Most sequencing centers quantify DNA via QubitCould also quantify with pico-green as outlined above  No need to dilute sample first  63 samples minimum, 1 ng/sample  63 ng in 35 µL TE = 1.8 ng/mL256 samples maximum, 1 ng/sample (currently with our 16x16 barcodes)  256 ng in 35 uL TE = 7.31 ng/uLAs long as the pool concentration is ≥ 1.8 ng/mL (for this primer set), you can send the pool as is.If it is
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
Agilent 5067-4626 Kit ComponentsHS Chip, RTSpin filtersDNA Ladder, 4°CDNA markers, 4°CDNA dye, 4°CDNA gel matrix, 4°C
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
Before beginningEquilibrate gel and dye (or gel-dye mix) to RT in the dark, 30 min.
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
Make sure gel-dye mix is prepared and Vortex dye 10 sec, spinPipet 15 uL dye into gel matrix vial.Vortex 10 secPipet into spin filter (provided)Spin 10 min, RT, 2240 gLabel tube cap with date of preparationGood for 5 chips
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
Test syringe for gel priming station for pressure by holding gloved finger against tip while dispensing 1 mL air
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
Add 400 uL new, clean PCR water to bioanalyzer’s electrode washing chip
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
ProcedurePrime the plate  Set chip priming station to bottom position (“C”), move plunger to 1 mL volume, and insert new HS chipPipet 9 uL gel-dye mix into well marked with black “G”Set timer for 60 seconds, then lock the latch of priming stationPress the plunger until it is locked into the clipWait exactly 60 seconds, then release the plunger with the clip release mechanismVisually inspect that the plunger moves back at least to the 0.3 mL markWait for 5 s, then slowly pull back the plunger to the 1 mL positionOpen the chip priming stationOpen bioanalyzer lid to let electrodes dry
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
Load remaining reagentsPipet 9 µL of gel-dye mix to each of the wells marked with a grey “G”Pipet DNA marker into all remaining wells (ladder and sample wells)  5 µL for ladder well5 µL for sample wells6 µL for sample wells where no sample is to be loadedPipet 1 µL DNA ladder into well marked with ladder symbolPipet 1 µL DNA into appropriate sample well5 µL for ladder well5 µL for sample wells6 µL for sample wells where no sample is to be loadedPipet 1 µL DNA ladder into well marked with ladder symbolPipet 1 µL DNA into appropriate sample well
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
Mix reagents
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
Load into Agilent 2100 BioanalyzerSelect Assay: dsDNA, High Sensitivity DNA.xsyEnter sample informationIf running fewer than 11 samples, indicate how many wells to run (Run samples 1 to 9, for example)  All samples indicated need to be run if user would like to toggle between time and size of fragment assessment.Could manually stop run after sample 9, for example, but then option to alter x-axis to fragment length would not be availablePress Start
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
After chip is run (~30 min)Remove HS chip and throw away (can clean and re-use…)Add 400 uL new, clean PCR water to bioanalyzer’s electrode washing chip
?. [Bioanalyzer Chip: High Sensitivity DNA Kit]
What to look for:Can have sequencing facility run chip for us instead  UC Davis currently charges $184/chip or $53/sample for 1-3 samples, 11/2015We currently pay $49.14/chip (7/2015)Trace can identify primer dimers  Greatly interfere with sequencing since these are preferentially sequencedIf present, do another clean-up before sending pool for sequencingTrace can show if multiple fragment sizes are present  May indicate separate 16S and 18S pools as this primer set can detect bothMay indicate a greater issue with ampliconsTrace indicates amplicon size  Need amplicon size for sequencing formCan better estimate molarity of sample
?. [Submit]
Each sequencing run should include:  ≥ 1 PCR blank (1 per person)≥ 1 even mock community≥ 1 staggered mock communityDilute multiple pools to an equal concentration  If submitting one pool:  Mix multiple pools in the ratio in which you desire sequencing depthEx: Pool 1 has 50 samples and is 1.8 ng/µL; Pool 2 has 100 samples and is 1.8 ng/µL. If you desire each sample to have equal sequencing depth, mix 11.7 µL Pool 1 and 23.3 uL Pool 2.If submitting multiple pools,  Typically charged for pooling: Currently $96 at UCDavis (11/2015)Need to send bioanalyzer trace and quantification information for each pool sentNeed to send:  Submission form (quantification information)Bioanalyzer trace for each pool sentIndices used for demultiplexing
?. [Appendix 1: Primer Strategy]
Primer Strategy from Hilary Morrison (et al. at MBL). Also basically used in http://aem.asm.org/content/suppl/2011/05/13/77.11.3846.DC1/Bartram_ms_AEM_S-corrected.pdf1, though I think they only did reverse indexing with no inline barcode on the forward.The primers are quite ‘universal’; the details:515F (modified from the EMP primer, here’s a reference4)NOTE: Currently used 515F primer is modified to contain a Y in 4th position of the 5’ end2, instead of the published C, this increases the Thaumarchaea hits to from 0.4% to 70%, and the total Archaea from 50% to 80%Matches From the Silva v106 (using 515F with C in 4th position)Arch (0mm): 54.4%Notably, does not hit most of the Thaumarchaea with zero mismatch, the mismatch is 4 bases from the 5’ end.Bact (0mm): 95.4%Euk (0mm): 92.2Arch (1mm): 96.2%Bact (1mm): 98.4%Euk (1mm): 97%926R (perhaps this is a paper where this primer appears, i.e., 3: http://www.biomedcentral.com/1471-2105/12/38/ and referenced in http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3721114/pdf/ismej201344a.pdf, Headwaters are critical reservoirs of microbial diversity for fluvial networks, and Development of a standardized approach for environmental microbiota investigations related to asthma development in childrenfor what that’s worth.Matches from the Silva 115Arch (0mm): 89%Bact (0mm): 92.7%Euk (0mm): 92.2%Arch (1mm): 94.1%Bact (1mm): 97.4%Euk (1mm): 97.5%Indices. This is one aspect that, if ordering new primers, could be changed. Since the indexing read is a dedicated read that does not reduce the data from other reads, having the commonly used 12 base index would only benefit. But practically, if might create a big problem if one was to mix indeces with 6 bases and others with 12 bases in the same run, please contact sequencing facility if this would even work. Generally taken from https://wikis.utexas.edu/display/GSAF/Illumina+-+all+flavors
?. [Appendix 2: Mock Community and Blanks]
Usage: We amplify an ‘even’ and ‘staggered’ mock communities as ‘positive controls’ with each set of PCRs that we run. For example for every 48 PCR samples, we will include 2 mock communities (1 even and 1 staggered) and at least 1 no template control. The lyophilized mock community should be diluted with 125 µL of TE (the final concentration will be about 0.001ng/µL). Add the TE, and then vortex, wait 5 minutes and vortex again (like you would do for a primer). Aliquot this into about 10 aliquots. I have observed some degradation from multiple freeze thaw cycles. We use 1 µL of 0.001ng/µL of mock community for each amplification because this is approximately the concentration that we expect 16S rRNA gene abundances to be found in our environmental samples. This is because we usually load 1ng environmental DNA to each PCR reaction, and 16S is roughly 1/1000 of each genome.  Do not take the mock communities into clean, UV/PCR hood.It would be interesting to add the mock communities to an environmental sample and leave the same environmental sample blank in a separate reaction. If you do this, you could see how much the mock community results are skewed by the presence of other templates, we have not done this would be interested in your experiences if you were to experiment with this.The following table includes the clone names, accession number of the clones, and the SILVA taxonomy for the sequences in the mock community, and the proportion of each of the clones in version 2 of the mock community. For version 1, remove the Thaumarchaea sequences (red text) and re-calculate proportions.  ABCDEFG1clone_names
ARISA
Accession
Silva taxonomy v. 119
% ID to Silva119 rep_seq/99 match
(full/ 515-926/ no primers)
proportion in staggered.v2
proportion ineven.v2
2SAR11
ARISA_667.6
DQ009197.1
Bacteria; Proteobacteria; Alphaproteobacteria; SAR11 clade; Surface 1
99.22/99.76/99.73
31.5
9.1
3OCS155_a
ARISA_435.5
D1009124.1
Bacteria; Actinobacteria; Acidimicrobiia; Acidimicrobiales; OM1 clade; Candidatus Actinomarina
98.36/100/100
15.8
9.1
4OCS155_b
ARISA_419.5
DQ009123.1
Bacteria; Actinobacteria; Acidimicrobiia; Acidimicrobiales; OM1 clade; Candidatus Actinomarina; uncultured bacterium
99.17/99.76/99.73
9.0
&nbsp;
5Tharumarchaea_MGI_a*
thaum_890_2_0.0997
KT036446
Archaea; Thaumarchaeota; Marine Group I; Unknown Order
99.1/99.76/99.73
9.0
9.1
6Prochlorococcus
ARISA_828.8
DQ009356.1
Bacteria; Cyanobacteria; Cyanobacteria; SubsectionI; FamilyI; Prochlorococcus; uncultured bacterium
99.51/99.76/100
6.8
9.1
7SAR86_a
ARISA_402.4
DQ009149.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; SAR86 clade; uncultured bacterium
99.39/99.75/99.73
4.5
9.1
8AEGEAN-169
ARISA_676.9
DQ009262.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rhodospirillales; Rhodospirillaceae; AEGEAN-169 marine group
99.09/100/100
2.3
&nbsp;
9SAR116_a
ARISA_653.1
DQ009264.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rickettsiales; SAR116 clade; uncultured bacterium
100/100/100
2.3
9.1
10Euryarchaea_MGII*
eury25
KT036445
Archaea; Euryarchaeota; Thermoplasmata; Thermoplasmatales; Marine Group II; uncultured archaeon
99.17/99.76/99.73
1.8
9.1
11Flavobacteria
ARISA_726.4
DQ009108.1
Bacteria; Bacteroidetes; Flavobacteriia; Flavobacteriales; Flavobacteriaceae; NS2b marine group
99.03/100/100
1.8
9.1
12Planctomyces**
6013
AF029077
Bacteria; Planctomycetes; Planctomycetacia; Planctomycetales; Planctomycetaceae; Blastopirellula
100/100/100
1.8
9.1
13SAR116_b
ARISA_703.7
DQ009270.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rickettsiales; SAR116 clade
100/100/100
1.8
&nbsp;
14SAR202_a
bats256
AY534095
Bacteria; Chloroflexi; SAR202 clade; uncultured Chloroflexi bacterium
100/100/100
1.8
9.1
15SAR406
ARISA_627.8
DQ009157.1
Bacteria; Deferribacteres; Deferribacteres; Deferribacterales; SAR406 clade(Marine group A); uncultured bacterium
98.47/99.76/99.73
1.4
9.1
16Flavobacteria_Formosa
ARISA_779.2
DQ009099.1
Bacteria; Bacteroidetes; Flavobacteriia; Flavobacteriales; Flavobacteriaceae; Formosa; uncultured bacterium
98.83/99.76/99.73
0.9
&nbsp;
17Flavobacteria_NS9
ARISA_540.1
EF572761.1
Bacteria; Bacteroidetes; Flavobacteriia; Flavobacteriales; NS9 marine group; uncultured bacterium
99.17/99.01/98.91
0.9
&nbsp;
18Pseudospirillum
ARISA_933.7
DQ009153.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; Oceanospirillaceae; Pseudospirillum
100/100/100
0.9
&nbsp;
19SAR86_b
ARISA_634.7
DQ009142.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; SAR86 clade; uncultured bacterium
100/99.76/99.73
0.9
&nbsp;
20SAR92
ARISA_762.8
DQ009136.1
Bacteria; Proteobacteria; Gammaproteobacteria; Alteromonadales; Alteromonadaceae; SAR92 clade; uncultured marine bacterium
100/100/100
0.9
&nbsp;
21Thaumarchaea_MGI_b*
thaum_890_3_0.0100
KT036447
Archaea; Thaumarchaeota; Marine Group I; uncultured marine archaeon
98.03/100/100
0.9
&nbsp;
22Verrucomicrobia
ARISA_738.8
DQ009368.1
Bacteria; Verrucomicrobia; Opitutae; MB11C04 marine group
99.46/100/100
0.9
&nbsp;
23Rhodobacteraceae
ARISA_840
EU804911.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rhodobacterales; Rhodobacteraceae; uncultured
100/100/100
0.7
&nbsp;
24SAR86_d
ARISA_657.6
DQ009141.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; SAR86 clade; uncultured bacterium
100/100/100
0.7
&nbsp;
25Flavobacteria_NS5
ARISA_749.6
DQ009088.1
Bacteria; Bacteroidetes; Flavobacteriia; Flavobacteriales; Flavobacteriaceae; NS5 marine group; uncultured bacterium
98.37/100/100
0.5
&nbsp;
26SAR86_c
ARISA_584
DQ009125.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; SAR86 clade
100/100/100
0.2
&nbsp;
27SAR116_c
ARISA_765.7
DQ009276.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rickettsiales; SAR116 clade; uncultured bacterium
97.82/99.76/99.73
0.1
&nbsp;
28SAR202_b
bats259
AY534094
Bacteria; Chloroflexi; SAR202 clade; uncultured Chloroflexi bacterium
100/100/100
0.1
&nbsp;
full sequence (27f-1492r)/ primed amplicon(515f-926r)*full sequence (20f-1392r)/ primed amplicon(515f-926r)**full sequence (27f-1392r)/ primed amplicon(515f-926r)
ABCDEFG1clone_names
ARISA
Accession
Silva taxonomy v. 119
% ID to Silva119 rep_seq/99 match
(full/ 515-926/ no primers)
proportion in staggered.v2
proportion ineven.v2
2SAR11
ARISA_667.6
DQ009197.1
Bacteria; Proteobacteria; Alphaproteobacteria; SAR11 clade; Surface 1
99.22/99.76/99.73
31.5
9.1
3OCS155_a
ARISA_435.5
D1009124.1
Bacteria; Actinobacteria; Acidimicrobiia; Acidimicrobiales; OM1 clade; Candidatus Actinomarina
98.36/100/100
15.8
9.1
4OCS155_b
ARISA_419.5
DQ009123.1
Bacteria; Actinobacteria; Acidimicrobiia; Acidimicrobiales; OM1 clade; Candidatus Actinomarina; uncultured bacterium
99.17/99.76/99.73
9.0
&nbsp;
5Tharumarchaea_MGI_a*
thaum_890_2_0.0997
KT036446
Archaea; Thaumarchaeota; Marine Group I; Unknown Order
99.1/99.76/99.73
9.0
9.1
6Prochlorococcus
ARISA_828.8
DQ009356.1
Bacteria; Cyanobacteria; Cyanobacteria; SubsectionI; FamilyI; Prochlorococcus; uncultured bacterium
99.51/99.76/100
6.8
9.1
7SAR86_a
ARISA_402.4
DQ009149.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; SAR86 clade; uncultured bacterium
99.39/99.75/99.73
4.5
9.1
8AEGEAN-169
ARISA_676.9
DQ009262.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rhodospirillales; Rhodospirillaceae; AEGEAN-169 marine group
99.09/100/100
2.3
&nbsp;
9SAR116_a
ARISA_653.1
DQ009264.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rickettsiales; SAR116 clade; uncultured bacterium
100/100/100
2.3
9.1
10Euryarchaea_MGII*
eury25
KT036445
Archaea; Euryarchaeota; Thermoplasmata; Thermoplasmatales; Marine Group II; uncultured archaeon
99.17/99.76/99.73
1.8
9.1
11Flavobacteria
ARISA_726.4
DQ009108.1
Bacteria; Bacteroidetes; Flavobacteriia; Flavobacteriales; Flavobacteriaceae; NS2b marine group
99.03/100/100
1.8
9.1
12Planctomyces**
6013
AF029077
Bacteria; Planctomycetes; Planctomycetacia; Planctomycetales; Planctomycetaceae; Blastopirellula
100/100/100
1.8
9.1
13SAR116_b
ARISA_703.7
DQ009270.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rickettsiales; SAR116 clade
100/100/100
1.8
&nbsp;
14SAR202_a
bats256
AY534095
Bacteria; Chloroflexi; SAR202 clade; uncultured Chloroflexi bacterium
100/100/100
1.8
9.1
15SAR406
ARISA_627.8
DQ009157.1
Bacteria; Deferribacteres; Deferribacteres; Deferribacterales; SAR406 clade(Marine group A); uncultured bacterium
98.47/99.76/99.73
1.4
9.1
16Flavobacteria_Formosa
ARISA_779.2
DQ009099.1
Bacteria; Bacteroidetes; Flavobacteriia; Flavobacteriales; Flavobacteriaceae; Formosa; uncultured bacterium
98.83/99.76/99.73
0.9
&nbsp;
17Flavobacteria_NS9
ARISA_540.1
EF572761.1
Bacteria; Bacteroidetes; Flavobacteriia; Flavobacteriales; NS9 marine group; uncultured bacterium
99.17/99.01/98.91
0.9
&nbsp;
18Pseudospirillum
ARISA_933.7
DQ009153.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; Oceanospirillaceae; Pseudospirillum
100/100/100
0.9
&nbsp;
19SAR86_b
ARISA_634.7
DQ009142.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; SAR86 clade; uncultured bacterium
100/99.76/99.73
0.9
&nbsp;
20SAR92
ARISA_762.8
DQ009136.1
Bacteria; Proteobacteria; Gammaproteobacteria; Alteromonadales; Alteromonadaceae; SAR92 clade; uncultured marine bacterium
100/100/100
0.9
&nbsp;
21Thaumarchaea_MGI_b*
thaum_890_3_0.0100
KT036447
Archaea; Thaumarchaeota; Marine Group I; uncultured marine archaeon
98.03/100/100
0.9
&nbsp;
22Verrucomicrobia
ARISA_738.8
DQ009368.1
Bacteria; Verrucomicrobia; Opitutae; MB11C04 marine group
99.46/100/100
0.9
&nbsp;
23Rhodobacteraceae
ARISA_840
EU804911.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rhodobacterales; Rhodobacteraceae; uncultured
100/100/100
0.7
&nbsp;
24SAR86_d
ARISA_657.6
DQ009141.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; SAR86 clade; uncultured bacterium
100/100/100
0.7
&nbsp;
25Flavobacteria_NS5
ARISA_749.6
DQ009088.1
Bacteria; Bacteroidetes; Flavobacteriia; Flavobacteriales; Flavobacteriaceae; NS5 marine group; uncultured bacterium
98.37/100/100
0.5
&nbsp;
26SAR86_c
ARISA_584
DQ009125.1
Bacteria; Proteobacteria; Gammaproteobacteria; Oceanospirillales; SAR86 clade
100/100/100
0.2
&nbsp;
27SAR116_c
ARISA_765.7
DQ009276.1
Bacteria; Proteobacteria; Alphaproteobacteria; Rickettsiales; SAR116 clade; uncultured bacterium
97.82/99.76/99.73
0.1
&nbsp;
28SAR202_b
bats259
AY534094
Bacteria; Chloroflexi; SAR202 clade; uncultured Chloroflexi bacterium
100/100/100
0.1
&nbsp;
?. [Appendix 2: Mock Community, blanks and simple data analyses and WARNING on 18S.]
Data analysis:We add perfect sequences in with the quality filtered reads from the sequencer to easily assess the accuracy of the PCR-to-OTU table procedures. If you just analyze the mock communities by themselves, with the perfect sequences, you might over or underestimate the ability of clustering algorithms ability to correctly split your environmental sequences. For example, we found that pairing the amplified mock community sequences with those of environmental sequences split some of the taxa with certain settings or pipelines, which did not occur when we analyzed them separately.Find 2 files on the Fuhrman lab website that will help with assessment and analysis of the mock communities. These files happen to be formatted for the QIIME pipeline, which you may use, but you may like to instead format them for mothur, or another 16S analysis tool of choice. The two 4 of files are correspond to the version 1 or 2 of the staggered and even communities.  These files have the full length expected 16S sequences for the mock communities, in fasta format, in the proportions that would be if the mock community performed perfectly. You will probably want to trim the sequence files to only include the 16S fragment that your assay amplified. If you are using the 515F/926R (our preferred primers), the following primer removal will work, if you download the cutadapt software: http://cutadapt.readthedocs.io/en/stable/installation.html#remove reverse primer (reverse complement of primer)cutadapt -a AAACTYAAAKRAATTGRCGG in.silico.full.even.fa -o no.rev.in.silico.even.fa -e 0.2 --discard-untrimmed#remove forward primer (5'-3' orientation of primer)cutadapt -g GTGYCAGCMGCCGCGGTAA no.rev.in.silico.even.fa -o no.prime.no.rev.in.silico.even.fa -e 0.2 --discard-untrimmed#remove reverse primer (reverse complement of primer)cutadapt -a AAACTYAAAKRAATTGRCGG in.silico.full.stag.fa -o no.rev.in.silico.stag.fa -e 0.2 --discard-untrimmed#remove forward primer (5'-3' orientation of primer)cutadapt -g GTGYCAGCMGCCGCGGTAA no.rev.in.silico.stag.fa -o no.prime.no.rev.in.silico.stag.fa -e 0.2 --discard-untrimmedNow that you have a file with the trimmed version of your mock community sequences, formatted for QIIME, add them to your sequences read files for clustering (probably at the bottom so they are least like to be seeds for cluster generation). If you trimmed and removed the primers from the files above, one good option is to add them to your sequences after they have been demulitplexed, merged, quality filtered, and primers trimmed. Like this:#simple unix concatenate command to concatenate all of your sample sequence reads, including the amplified mock community reads and the in silico (perfect) mock community readscat seqs.to.cluster.fa no.prime.no.rev.in.silico.even.fa no.prime.no.rev.in.silico.stag.fa > seqs.to.cluster.plus.mock.faThen, follow through your chimera checking, clustering, classification, and OTU table generation pipeline just as you normally would. If you are using QIIME, with the file formats above, the mock communities will be called “in.silico.even” and “in.silico.stag” in your OTU table.WARNING: 18S forward and reverse reads DO NOT overlap, therefore in merging steps common to most rRNA analyses all 18S sequences would be lost if the step requires overlap. Therefore we recommend first trimming for quality control then merging forward and reverse reads separated by a single N and using k-mer base classifier.Here is a sample set of unix commands that our lab uses for this:#!/bin/bash# Program:# This pipeline is used for 16s and 18s sequences analysis# Pipelines and applications used in this script:# Trimmomatic, Usearch, QIIME,seqtk, cutadapt, BLAST# Required files: # R1 and R2 fastq files for each sample separately# Final outputs:# A 16S OTU table, a 18S OTU table, and two chloroplast table showing  how the chloroplast OTUs are reassigned in PhytoRef and NCBI databases# History:# Liv, 8/21/2017: First release# Liv, 8/23/2017: add steps of classifying chloroplast reads using blastn against NCBI and PhytoRef# Note that the way to create a list depends on how you name your samplels *R1.fastq| cut -d '.' -f 1-13>sample.namefor i in `cat sample.name`;do# Do qualify filtering on fwd and rev reads using Trimmomatic java -jar /korriban/liv/Trimmomatic-0.36/trimmomatic-0.36.jar PE -phred33 "$i"_R1.fastq "$i"_R2.fastq R1_pe R1_se R2_pe R2_se SLIDINGWINDOW:4:20 MINLEN:200;# Rename files that pass QC and remove files that don't pass QCmv R1_pe "$i"_R1_trimSW4_20.fastq;mv R2_pe "$i"_R2_trimSW4_20.fastq;rm R1_se;rm R2_se;# Merge paired ends using Usearch (alternative: Vsearch or FLASH) (Note that merged reads are 16s, and non-merged reads are 18s)~/bin/usearch9.2.64_i86linux32 -fastq_mergepairs "$i"_R1_trimSW4_20.fastq -reverse "$i"_R2_trimSW4_20.fastq -fastqout "$i".merged.fastq -fastq_maxdiffs 3 -fastq_minmergelen 300 -fastqout_notmerged_fwd notmerged_"$i".R1_trimSW4_20.fastq -fastqout_notmerged_rev notmerged_"$i".R2_trimSW4_20.fastq########## for 16s reads ########## # Convert to fasta fileconvert_fastaqual_fastq.py -f "$i".merged.fastq -o 16s.fna/ -c fastq_to_fastaqual# Remove primers using cutadaptcutadapt -g ^GTGYCAGCMGCCGCGGTAA -o 16s.fna/test.fwdprimer.removed.fna 16s.fna/"$i".merged.fna --discard-untrimmed;cutadapt -a AAACTYAAAKRAATTGRCGG$ -o 16s.fna/primers_removed_"$i".merged.fna 16s.fna/test.fwdprimer.removed.fna --discard-untrimmed# Do chimera checking by sample and remove chimeric readsidentify_chimeric_seqs.py -m usearch61 -i 16s.fna/primers_removed_"$i".merged.fna -o 16s.fna/usearch61_chimera_checking/ --suppress_usearch61_reffilter_fasta.py -f 16s.fna/primers_removed_"$i".merged.fna -o 16s.fna/primers_removed_nochimera_"$i".merged.fna -s 16s.fna/usearch61_chimera_checking/chimeras.txt -n########## for 18s reads ########## # Trim all the non-merged fwd reads to the same lengthjava -jar /korriban/liv/Trimmomatic-0.36/trimmomatic-0.36.jar SE -phred33 notmerged_"$i".R1_trimSW4_20.fastq notmerged_"$i".R1_trimSW4_20_len190.fastq CROP:190 MINLEN:190;# Trim all the non-merged rev reads to the same lengthjava -jar /korriban/liv/Trimmomatic-0.36/trimmomatic-0.36.jar SE -phred33 notmerged_"$i".R2_trimSW4_20.fastq notmerged_"$i".R2_trimSW4_20_len190.fastq CROP:190 MINLEN:190# Convert the non-merged rev reads to its reverse complement with seqktseqtk seq -r notmerged_"$i".R2_trimSW4_20_len190.fastq > notmerged_"$i".R2_trimSW4_20_len190_rc.fastq# Split fwd and rev fastq files into fasta and quality filesconvert_fastaqual_fastq.py -f notmerged_"$i".R1_trimSW4_20_len190.fastq -o 18s.fna/ -c fastq_to_fastaqualconvert_fastaqual_fastq.py -f notmerged_"$i".R2_trimSW4_20_len190_rc.fastq -o 18s.fna/ -c fastq_to_fastaqual# Add N at the beginning of reverse complementsed -e 's/^/N/' 18s.fna/notmerged_"$i".R2_trimSW4_20_len190_rc.fna | sed -e 's/N>/>/' > 18s.fna/N.added.notmerged_"$i".R2_trimSW4_20_len190_rc.fna# Extract headers from fwd readssed -n '1~2p' 18s.fna/notmerged_"$i".R1_trimSW4_20_len190.fna >18s.fna/R1;# Extract headers from rev readssed -n '1~2p' 18s.fna/notmerged_"$i".R2_trimSW4_20_len190_rc.fna >18s.fna/R2;# Get shared headerscat 18s.fna/R1 18s.fna/R2 | sort | uniq -d | cut -d '>' -f 2 >18s.fna/filter.txt;# filter fasta file filter_fasta.py -f 18s.fna/notmerged_"$i".R1_trimSW4_20_len190.fna -o 18s.fna/temp_R1.fna -s 18s.fna/filter.txt;filter_fasta.py -f 18s.fna/notmerged_"$i".R2_trimSW4_20_len190_rc.fna -o 18s.fna/temp_R2.fna -s 18s.fna/filter.txt;# Remove headers from reverse complementcut -d '>' -f 1 18s.fna/temp_R2.fna > 18s.fna/r2.no.header;# Paste r1 and reverse complement of r2 togetherpaste 18s.fna/temp_R1.fna 18s.fna/r2.no.header -d '' > 18s.fna/"$i".mergedN.18s.fna;# Remove primers using cutadaptcutadapt -g ^GTGYCAGCMGCCGCGGTAA -o 18s.fna/temp.fwdprimer.removed.fna 18s.fna/"$i".mergedN.18s.fna -e 0.2 --discard-untrimmedcutadapt -a AAACTYAAAKRAATTGRCGG$ -o 18s.fna/primers_removed_"$i".mergedN.18s.fna 18s.fna/temp.fwdprimer.removed.fna -e 0.2 --discard-untrimmed# Do chimera checking by sample and remove chimeric readsidentify_chimeric_seqs.py -m usearch61 -i 18s.fna/"$i".mergedN.18s.fna -o 18s.fna/usearch61_chimera_checking/ --suppress_usearch61_reffilter_fasta.py -f 18s.fna/"$i".mergedN.18s.fna -o 18s.fna/nochimera_"$i".mergedN.18s.fna -s 18s.fna/usearch61_chimera_checking/chimeras.txt -ndonemkdir 16s.fna/no.chimeramv 16s.fna/primers_removed_nochimera_* 16s.fna/no.chimera/mkdir 18s.fna/no.chimeramv 18s.fna/primers_removed_nochimera_* 18s.fna/no.chimera/# Combine all the 16S fasta files and in silico 16S mock fasta files into a single fasta filecat 16s.fna/no.chimera/*.fna ../insilico_16s/*fasta>16s.insilico_mock_combined_chimera_removed.fna# Combine all the 18S fasta files and in silico 18S mock fasta files into a single fasta filecat 18s.fna/no.chimera/*.fna ../insilico_18s/*fasta>18s.insilico_mock_combined_chimera_removed.fna# OTU picking using uclustpick_otus.py -i 16s.insilico_mock_combined_chimera_removed.fna -o otu_picking_16s_uclust99/ -s 0.99pick_otus.py -i 18s.insilico_mock_combined_chimera_removed.fna -o otu_picking_18s_uclust99/ -s 0.99# Pick the representative fnapick_rep_set.py -i otu_picking_16s_uclust99/16s.insilico_mock_combined_chimera_removed_otus.txt -f 16s.insilico_mock_combined_chimera_removed.fna -m most_abundant -o otu_picking_16s_uclust99/rep_otus.fastapick_rep_set.py -i otu_picking_18s_uclust99/18s.insilico_mock_combined_chimera_removed_otus.txt -f 18s.insilico_mock_combined_chimera_removed.fna -m most_abundant -o otu_picking_18s_uclust99/rep_otus.fasta# Assign taxonomy (reference database: SILVA (for 16S) and PR2 (for 18S))assign_taxonomy.py -i otu_picking_16s_uclust99/rep_otus.fasta –t /Silva119_release/taxonomy/99/taxonomy_99_7_levels.txt -r /Silva119_release/rep_set/99/Silva_119_rep_set99.fna --similarity 0.97 -o otu_picking_16s_uclust99/silva119_taxonomy/assign_taxonomy.py -i otu_picking_18s_uclust99/rep_otus.fasta –r /pr2_gb203_version_4.5/pr2_gb203_version_4.5.fasta -t /pr2_gb203_version_4.5/pr2_gb203_version_4.5.taxo -o otu_picking_18s_uclust99/pr2_taxonomy/ -m rdp --rdp_max_memory 40000# make otu tablemake_otu_table.py -i otu_picking_16s_uclust99/16s.insilico_mock_combined_chimera_removed_otus.txt -t otu_picking_16s_uclust99/silva119_taxonomy/rep_otus_tax_assignments.txt -o otu_picking_16s_uclust99/16s.insilico_mock.biommake_otu_table.py -i otu_picking_18s_uclust99/18s.insilico_mock_combined_chimera_removed_otus.txt -t otu_picking_18s_uclust99/pr2_taxonomy/rep_otus_tax_assignments.txt -o otu_picking_18s_uclust99/18s.insilico_mock.biom# Convert biom to txtbiom convert -i otu_picking_16s_uclust99/16s.insilico_mock.biom -o otu_picking_16s_uclust99/16s.insilico_mock.txt --header-key taxonomy --to-tsv biom convert -i otu_picking_18s_uclust99/18s.insilico_mock.biom -o otu_picking_18s_uclust99/18s.insilico_mock.txt --header-key taxonomy --to-tsv #####for 16S reads assigned as chloroplast in SILVA database ########## # Get a Chloroplast only OTU table from 16s OTU tablefilter_taxa_from_otu_table.py -i otu_picking_16s_uclust99/16s.insilico_mock.biom -o otu_picking_16s_uclust99/chloroplast_only.biom -p Chloroplast# Extract a Chloroplast only fasta file based the Chloroplast only OTU tablefilter_fasta.py -f otu_picking_16s_uclust99/rep_otus.fasta -o otu_picking_16s_uclust99/chloroplast_rep_otus.fasta -b otu_picking_16s_uclust99/chloroplast_only.biom # A BLAST search against NCBI nucleotide blastn -db /home/korriban/db/blast_nt_genomic_refseq/nt –query chloroplast_rep_otus.fasta -out result3.out -num_threads 120 -max_target_seqs 1 -outfmt "6 qseqid sseqid pident length mismatch gapopen qstart qend sstart send evalue bitscore sseqid sallseqid sgi sacc staxids sscinames scomnames stitle"# Create a custom PhytoRef database makeblastdb -in PhytoRef_with_taxonomy.fasta -out phytoref.db.taxo -dbtype nucl# A BLAST search against PhytoRef databaseblastn -db /korriban/liv/PhytoRef/phytoref.db.taxo -query otu_picking_16s_uclust99/chloroplast_rep_otus.fasta -out 16s_chl_blast_phytoref.out -num_threads 8 -max_target_seqs 1 -outfmt 6
?. [Appendix 2: Mock Community and Blanks]
Background: We generated a mock community that approximates a surface microbial community at the San Pedro Ocean Time-series (SPOT) for several reasons: 1.) examine the extent that PCR bias influences the assessment of a community with a known input, 2.) assist in examining run-to-run variability of sequencing runs, 3.) serve as a control for clustering, classification, or other analytical procedures (e.g., is the data quality high enough for oligotyping? Yes: https://www.nature.com/articles/ismej201729).  The mock community is made from pooled 16S rRNA gene clones generated from SPOT, and 3 clones from the Giovannoni lab’s clone collection (taxa that we did not have: Chloroflexi and Planctomycetes). The clones were re-grown overnight from their -80C freezer stocks, and plasmids purified via Miniprep. We then amplified the plasmids with vector primers (remove any extra E. coli contamination), followed a PCR with 16S rRNA gene primers that amplified nearly the full 16S primer (generally a 8-28F position and 1392-1492R). We then quantified the products and pooled them in equal proportions (even community) or from 0.1%-30% (staggered community), taking into consideration each taxon’s amplicon length and sequence.Originally the even community consisted of 10 clones (even version 1) and the staggered community consisted of 25 clones (staggered version 1). However, we later added 1 Thaumarchaea clone to the even mock community (even version 2) and 2 Thaumarchaea clones to the staggered community (staggered version 2).
?. [Appendix 3: General PCR Considerations]
Generally PCR strips should be performed in the UV, Biosafety hood (we typically use the Stratlinker for plastics and PCR water, and other reagents that can be UV’d). In general we have not been able to trace contamination issues to lab practices, but it is probably a good idea to take some steps to reduce contamination such as cross contamination or from the lab.Some steps to perhaps reduce contaminationWipe down the inside of the Biosafety hood with 10% Bleach. Wipe down the pipettes with 10% Bleach.Leave the UV light on the hood for about 15 minutes.
 Aliquot all PCR reagents and ideally only use them one time, especially the buffer, MgSO4, and water into thin walled PCR strip tubes.You can also treat all plastic ware in the Stratalinker and/or soak in a 10% bath of bleach.Consider opening tubes with DNA individually, being careful where your thumb goes as to avoid cross contamination.Add master mix to PCR tubes before DNA.Treat reagents, that are not harmed by UV, with UV: the water, buffer and magnesium. This can be performed in 300uL strip tubes in the Stratalinker by setting the timer to 9999. This is about 10 minutes. I typically perform this step at the beginning of a days PCR setup.Load all samples + blanks into thermocycler before moving on to the mock communities which are relatively concentrated with 16S PCR product.We used to typically run reactions in triplicate by adding, say 72uL of MM to a tube and then 3uL of DNA and then split into 3 tubes of 25uL each, though we have not confirmed that this makes a difference, if nothing else it gives more PCR product with fewer cycles.After PCR combine triplicate reactions on lab bench. At this point if there is slight cross contamination it is not as big deal of a deal as it was before since now the samples are all barcoded. The worst it could do is mess up your quantifications, which is not ideal (potentially, for example, making your blanks quantify higher than they should).The polymerase master mix indicated below was used for some Fuhrman Lab amplicons, and Alma/David found that the mock communities generated with this polymerase were equivalent to those generated with the 5-Prime Taq master mix indicated in this protocol.Platinum Hifi PCR Master Mix----------------------------1x  Invitrogen Hifi Buffer (11304-029) (10x)--------2.5 µL----1x [final]dNTPs (10mM) (promega cat #: U1515) ------------0.5 µL--200uM [final]Forward Primer (10µM)---------------------------1 µL-----0.4uM [final]Reverse Primer (10µM)---------------------------1 µL-----0.4uM [final]Invitrogen MgSO4 (11304-029)(50mM)--------------1 µL------2mM [final]Invitrogen Platinum Hifi Taq (11304-029)--------0.2 µL--x Unit [final]DNA Template (generally, 1ng/µl)----------------1 µL------1ng [final]Molecular Grade Water (VWR cat#:95043-414)-----17.8 µLTotal Reaction Volume -------------------------25µL
?. [Appendix 4: PCR clean-up notes]
When more high-throughput sequencing is desired, can clean-up PCR reactions with normalization plates. We have successfully used SequalPrep Normalization Plate (Invitrogen A10510-01), but tested when we were doing triplicate PCR reactions/sample.According to protocol, most PCR reactions give 25-100 ng/uL in 20 µL reactions.Add at least 250 ng of PCR product suggested per wellStandard elutions using 20 µL elution volume yields concentrations of 1-2 ng/µL  aka, 25 ng kept
  Theoretically negates the need to quantify and dilute cleaned PCR reactions before pooling.
?. [Appendix 5: Sequencing Submission ]
Current requirements/pricing (11/2015)  ABCD1&nbsp;
&nbsp;
UCDavis
USC
2HiSeq 2500 Rapid Run, PE250
min vol
15 mL
&nbsp;
3&nbsp;
min nM
5 nM, ~1.8 ng/uL for our amplicons
&nbsp;
4&nbsp;
$
$5380/lane
$3700/lane
5&nbsp;
&nbsp;
&nbsp;
&nbsp;
6MiSeq 600 cycles v3 PE300
min vol
15 mL
&nbsp;
7&nbsp;
min nM
5 nM, ~1.8 ng/uL for our amplicons
&nbsp;
8&nbsp;
$
$2669
&nbsp;
9&nbsp;
&nbsp;
&nbsp;
&nbsp;
10MiSeq 500 cycles v2 PE250
min vol
15 mL
&nbsp;
11&nbsp;
min nM
5 nM, ~1.8 ng/uL for our amplicons
&nbsp;
12&nbsp;
$
$1967
&nbsp;
13&nbsp;
&nbsp;
&nbsp;
&nbsp;
14Library Pooling
$
$96, up to 6 samples
&nbsp;
15&nbsp;
&nbsp;
&nbsp;
&nbsp;
16DNA Quant (Qubit)
$
$96, up to 12 samples
&nbsp;
17&nbsp;
&nbsp;
&nbsp;
&nbsp;
18Library Quant (qPCR)
$
$36/sample
&nbsp;
19&nbsp;
&nbsp;
&nbsp;
&nbsp;
20Bioanalyzer HS
$
$53, 1-3 samples
$184, 4-11 samples
&nbsp;
21&nbsp;
&nbsp;
&nbsp;
&nbsp;
22QC: Qubit, bioanalyzer, bead-clean
$
&nbsp;
$120/sample, 1-11
ABCD1&nbsp;
&nbsp;
UCDavis
USC
2HiSeq 2500 Rapid Run, PE250
min vol
15 mL
&nbsp;
3&nbsp;
min nM
5 nM, ~1.8 ng/uL for our amplicons
&nbsp;
4&nbsp;
$
$5380/lane
$3700/lane
5&nbsp;
&nbsp;
&nbsp;
&nbsp;
6MiSeq 600 cycles v3 PE300
min vol
15 mL
&nbsp;
7&nbsp;
min nM
5 nM, ~1.8 ng/uL for our amplicons
&nbsp;
8&nbsp;
$
$2669
&nbsp;
9&nbsp;
&nbsp;
&nbsp;
&nbsp;
10MiSeq 500 cycles v2 PE250
min vol
15 mL
&nbsp;
11&nbsp;
min nM
5 nM, ~1.8 ng/uL for our amplicons
&nbsp;
12&nbsp;
$
$1967
&nbsp;
13&nbsp;
&nbsp;
&nbsp;
&nbsp;
14Library Pooling
$
$96, up to 6 samples
&nbsp;
15&nbsp;
&nbsp;
&nbsp;
&nbsp;
16DNA Quant (Qubit)
$
$96, up to 12 samples
&nbsp;
17&nbsp;
&nbsp;
&nbsp;
&nbsp;
18Library Quant (qPCR)
$
$36/sample
&nbsp;
19&nbsp;
&nbsp;
&nbsp;
&nbsp;
20Bioanalyzer HS
$
$53, 1-3 samples
$184, 4-11 samples
&nbsp;
21&nbsp;
&nbsp;
&nbsp;
&nbsp;
22QC: Qubit, bioanalyzer, bead-clean
$
&nbsp;
$120/sample, 1-11
?. [Appendix 5: Sequencing Submission ]
Current UCDavis form (11/2015)
?. [Primers]
Bartram, A. K., Lynch, M. D. J., Stearns, J. C., Moreno-Hagelsieb, G. & Neufeld, J. D. Generation of Multimillion-Sequence 16S rRNA Gene Libraries from Complex Microbial Communities by Assembling Paired-End Illumina Reads. Appl. Environ. Microbiol. 77, 3846–3852 (2011).Parada, A., Needham, D. M. & Fuhrman, J. A. Every base matters: assessing small subunit rRNA primers for marine microbiomes with mock communities, time-series and global field samples. Environ. Microbiol. (2015). doi:10.1111/1462-2920.13023Quince, C., Lanzen, A., Davenport, R. J. & Turnbaugh, P. J. Removing Noise From Pyrosequenced Amplicons. BMC Bioinformatics 12, 38 (2011).Walters, W. A. et al. PrimerProspector: de novo design and taxonomic analysis of barcoded polymerase chain reaction primers. Bioinformatics 27, 1159–1161 (2011).
?. [PCR]
5 PRIME HotMasterMix  ABCD1&nbsp;
[stock]
vol per rxn (µL)[final]/rxn
2PCR water*
&nbsp;
12.5
&nbsp;
35’ Master Mix
2.5 x
10****
0.5 U taq, 45 mM Kcl, 2.5 mM Mg2+, 200 uM each dNTP
41:1, 515F:926R primer mix**
5 µM each primer1.5
0.3 mM each primer
5DNA***
0.5 ng/µL1
0.5 ng
6total vol
&nbsp;
25
&nbsp;
*VWR cat# 95043-414**Equivalent to adding 0.75 µL of each 10 µM working primer stock; We have tested/used 0.2-0.4 mM successfully***Can be modified according to [DNA] or DNA quality as long as you use the same amount (ng) in each reaction for a particular study. We have tested/used 100 pg – 2 ng. ****For trouble samples, increase master mix to 12 uL, and this seems to improve amplification (0.6 U taq, 54 mM KCl, 3 mM Mg2+, 240 uM each dNTP). Note, we initially used more expensive Taq such as HiFi from NEB, but saw no appreciable difference in terms of sequence abundances in our mock communities and samples.
ABCD1&nbsp;
[stock]
vol per rxn (µL)[final]/rxn
2PCR water*
&nbsp;
12.5
&nbsp;
35’ Master Mix
2.5 x
10****
0.5 U taq, 45 mM Kcl, 2.5 mM Mg2+, 200 uM each dNTP
41:1, 515F:926R primer mix**
5 µM each primer1.5
0.3 mM each primer
5DNA***
0.5 ng/µL1
0.5 ng
6total vol
&nbsp;
25
&nbsp;
?. [Primers]
We have obtained primers from Operon/Eurofins (https://www.eurofinsgenomics.com/en/home/), 50  nmol  scale,  salt-free  purified.            - Reconstitute main stocks in TE, 100 µM, store at -80° C            - Working stocks: dilute in TE, 10 µM, store at -20° C (can be stored at 4° C for several days to avoid freeze/thaw)Suggestion: prepare primer plates containing 1:1 mix of unique combinations of 10 µM 515F:926R barcoded primers    ([final] of each is 5 µM). Cover plate with sterile foil (USA Scientific, 2923-0110)These 16 forward and reverse primers allow for 256 unique combinations for multiplexing, more are possible.