Fragment Library Preparation

Transcription

1 Fragment Library Preparation 5500 Series SOLiD Systems QUICK REFERENCE Note: For safety and biohazard guidelines, refer to the Safety section in the Fragment Library Preparation: 5500 Series SOLiD Systems User Guide (Part no ). For every chemical, read the SDS and follow the handling instructions. Wear appropriate protective eyewear, clothing, and gloves. Prepare a single fragment library 1 Quantitate the DNA For accuracy, determine sample DNA concentration using a double-stranded DNA-specific fluorescence assay. Use the Invitrogen Qubit dsdna HS Assay Kit (Invitrogen Part no. Q32851 or Q32854) to measure dsdna concentrations from 10 pg/µl to 100 ng/µl. For samples outside this range, use the Qubit dsdna BR Assay Kit for higher concentrations of DNA (Invitrogen Part no. Q32850 or Q32853) or the Invitrogen Quant-iT PicoGreen dsdna Assay Kit for lower concentrations (Invitrogen Part no. P7589). 2 Shear the DNA with the Covaris S220 System Note: For shearing conditions with the Covaris S2 System, see Shear the DNA with the Covaris S2 System on page 14. a. Mix in a 1.5-mL LoBind Tube: Amount DNA 10 ng 5 µg 1 Low TE Buffer Variable µl Shear Buffer 1.2 µl Total 120 µl b. Load the DNA into the Covaris S220 System: 1. Prepare the Covaris S2 tank: Fill the tank to level 12. Ensure that the circulated water chiller is set to 2 5 C. Supplement the chiller not the actual water bath with 20% ethylene glycol. Ensure that the water bath temperature is 5 10 C. 2. Place a Covaris microtube into the loading station and transfer the mixture to the Covaris microtube.

2 2 Shear the DNA with the Covaris S220 System (continued) c. Shear the DNA using the appropriate program on the Covaris S220 System: IMPORTANT! Ensure that the bath temperature limit is set at 15 C, and keep the bath temperature to 10 C. Condition Setting Number of cycles 6 Bath temperature 5 C Bath temperature limit 15 C Mode Water quality testing function Frequency sweeping d. Place the Covaris microtube into the loading station, then transfer the DNA solution to a new 1.5-mL LoBind Tube. Off Duty Factor 10% Peak Incident Power Cycles/burst 100 Time 175 Watts 60 seconds 3 End-polish the DNA a. Combine in a new 1.5-mL LoBind Tube: Amount Sheared DNA 120 µl 5 Reaction Buffer 40 µl 10 mm dntp 8.0 µl End Polishing E1 8.0 µl End Polishing E2 10 µl Nuclease-Free Water 14 µl Total 200 µl b. Vortex the mixture for 5 seconds, pulse-spin, then incubate at room temperature (20 25 C) for 30 minutes. 2

3 4 Size-select the DNA by Agencourt AMPure XP Bead Reagent a. Resuspend the Agencourt AMPure XP Reagent beads and allow the mixture to come to room temperature. b. Prepare 1 ml of 70% ethanol. c. Bind the DNA to the resuspended, ambient Agencourt AMPure XP Reagent: 1. Prepare the bead suspension in the sample reaction: Volume Sheared DNA 200 µl Agencourt AMPure XP Reagent 100 µl Total 300 µl Equal to 0.5 volume of end-polished DNA. 2. Vortex the beads for 10 seconds, then pulse-spin. 3. Incubate the mixture at room temperature (20 25 C) for 5 minutes. 4. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution clears. The solution is clear of brown tint when viewed at an angle, as shown below: d. Carefully transfer the supernatant, which contains the DNA of the desired size, to a new 1.5- ml LoBind Tube. Discard the pellet. e. Bind the size-selected DNA in the supernatant to the Agencourt AMPure XP Reagent: 1. Combine: Volume Supernatant ~300 µl Agencourt AMPure XP Reagent 60 µl Total ~360 µl Equal to 0.3 volume of the end-polish reaction volume of 200 µl. 2. Vortex the beads for 10 seconds, then pulse-spin. 3. Incubate the mixture at room temperature (20 25 C) for 5 minutes. 4. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution clears, then remove and discard the supernatant. Save the pellet, which contains the DNA. 3

4 4 5 Size-select the DNA by Agencourt AMPure XP Bead Reagent (continued) Quantitate the sizeselected DNA f. Wash the DNA-bead complex 3 times. For each wash: 1. Add 200 µl of freshly prepared 70% ethanol to the tube, mix by inverting the tube a few times, then pulse-spin. 2. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution clears, then remove and discard the supernatant. g. Remove the tube from the DynaMag -2 magnetic rack, pulse-spin the tube, return the tube to the magnetic rack; then remove and discard the supernatant with a 20-µL pipettor. h. Open the tube, then dry the beads at room temperature (20 25 C) for 5 10 minutes. i. Elute the DNA: 1. Remove the tube from the DynaMag -2 magnetic rack, then add 36 µl Low TE Buffer directly to the pellet to disperse the beads. 2. Pipette the suspension up and down to mix. 3. Vortex the beads for 10 seconds, then pulse-spin. 4. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution clears. 5. Transfer the supernatant containing the size-selected DNA to a new 1.5-mL LoBind Tube. Quantitate the DNA by using the Quant-iT dsdna HS Assay Kit, the NanoDrop ND-1000 Spectrophotometer, or the Agilent Technologies 2100 Bioanalyzer. IMPORTANT! Average yield of size-selected DNA is 30% of input quantity. If the yield is substantially <20%, troubleshoot the low yield, then repeat the procedure from Shear the DNA with the Covaris S220 System on page 1. 6 Add a da-tail to the sizeselected DNA a. Combine in a 1.5-mL LoBind Tube: Amount Size-selected DNA 34 µl 5 Reaction Buffer 10 µl 10 mm datp 1.0 µl A-Tailing Enzyme I 5.0 µl Total 50 µl b. Incubate the mixture at 68 C for 30 minutes, then cool to room temperature. Note: While the reaction is incubating, calculate the amount of adaptors needed for ligation (see Ligate adaptors to the DNA on page 5). 4

5 7 Ligate adaptors to the DNA IMPORTANT! Do not use P1 and P2 Adaptors that are designed for fragment library preparation and sequencing on the SOLiD 4 System. These adaptors are not compatible with reverse-read sequencing on the 5500 Series SOLiD Sequencers. Use on P1-T and Barcode-T-0XX Adaptors that are designed for the 5500 Series SOLiD Sequencers. If the input DNA before shearing is: <100 ng: Use 0.06 µl of each adaptor or an equivalent amount of adaptor after dilution. For example, use 0.6 µl of a 10-fold dilution of an adaptor or 1.2 µl of a 20-fold dilution. 100 ng: Calculate the amount of adaptor needed, Y, for the reaction based on the amount of DNA from the last purification step. If DNA fragments were sheared using the standard protocol for fragment library preparation, the average insert size should be approximately 165 bp before adaptor ligation. IMPORTANT! Do not use P1 and P2 Adaptors that are designed for fragment library preparation and sequencing on the SOLiD 4 System. These adaptors are not compatible with reverse-read sequencing on the 5500 Series SOLiD Sequencers. Use on P1-T and Barcode-T-0XX Adaptors that are designed for the 5500 Series SOLiD Sequencers. a. Calculate the amount of adaptors needed: μg-to-pmol conversion factor = 10 6 pg 1 μg 1 pmol 660 pg (μg-to-pmol Y μl adaptor needed = # μg DNA 10 conversion factor) 1 Average insert size 1 μl adaptor needed 50 pmol b. In a new 1.5-mL LoBind Tube, combine: to prepare the ligase master mix: Volume 5 Reaction Buffer 3.0 µl P1-T Adaptor, 50 µm Barcode-T-001, 50 µm c. Combine the 15 µl of ligase mixture to the 46 µl A-tailing reaction. d. Vortex the mixture for 5 seconds, pulse-spin, then incubate the mixture in a thermocycler with the lid heater on: IMPORTANT! Incubation nick translates the DNA. YµL YµL T4 DNA Ligase, 5 U/µL 6.5 µl 10 mm dntp 1.2 µl Nuclease-free Water Variable µl Total 15 µl Stage Temp Time Holding 20 C 30 min Holding 72 C 20 min Holding 4 C 5

6 7 8 9 Ligate adaptors to the DNA (continued) Quantitate the ligated DNA (Optional) Amplify the library e. For speed and high yield, purify the DNA with up to two rounds of purification with the Agencourt AMPure XP Reagent. In the first round, use 0.6 per sample of the Agencourt AMPure XP Reagent (see Purify the DNA using Agencourt AMPure XP Reagent on page 14). In the second round, use 1.5 per sample volume of the reagent. As an alternative to bead-based purification, for convenience, purify the DNA with the SOLiD Micro Column Purification Kit (see Purify the DNA with the SOLiD Library Micro Column Purification Kit on page 15). Quantitate the DNA by using the Quant-iT dsdna HS Assay Kit, the NanoDrop ND-1000 Spectrophotometer, the Agilent Technologies 2100 Bioanalyzer, and/or by qpcr. IMPORTANT! The current protocol is optimized for maximum yield from input DNA. In most cases, library amplification is not needed. If library amplification is needed, minimize the number of cycles, based on the amount of starting input DNA. Use minimal cycling to avoid over-amplification and production of redundant molecules. a. In a 0.2-mL PCR tube, prepare the PCR mixture: Volume Adaptor-ligated, purified DNA 20 µl Platinum PCR Amplification Mix 100 µl Library PCR Primer 1, 50 µm 2.5 µl Library PCR Primer 2, 50 µm 2.5 µl Total 125 µl <20 µl is acceptable. Do not adjust the PCR volume. b. Vortex the mixture for 5 seconds, pulse-spin the mixture. c. Determine the number of PCR cycles: Starting amount of DNA Number of cycles ng 10 cycles 100 ng 1 µg 6 8 cycles 1 2 µg 4 6 cycles 2 5 µg 0 3 cycles d. Run the PCR: Stage Step Temp Time Holding Denature 95 C 5min Cycling Denature 95 C 15 sec Anneal 62 C 15 sec Extend 70 C 1 min Holding Extend 70 C 5min Holding 4 C e. Purify the DNA using the Agencourt AMPure XP Reagent (see Purify the DNA using Agencourt AMPure XP Reagent on page 14). 6

7 10 Quantitate the DNA Quantitate the DNA by using the Quant-iT dsdna HS Assay Kit, the NanoDrop ND-1000 Spectrophotometer, the Agilent Technologies 2100 Bioanalyzer, and/or by qpcr. 11 Check the size distribution of the library Use 1 µl of sample in the Agilent Technologies 2100 Bioanalyzer. If you see the expected size distribution, proceed directly to emulsion PCR [refer to the SOLiD EZ Bead Emulsifier Getting Started Guide (Part no )]. If you do not see the expected size distribution, troubleshoot or contact your Life Technologies Applications Specialist. Prepare multiple fragment libraries This procedure is for N number of libraries, where N 12. Prepare pooled, barcoded libraries for multiplexed sequencing or prepare multiple libraries in parallel that are not pooled. 1 Quantitate the DNA For accuracy, determine sample DNA concentration using a double-stranded DNA-specific fluorescence assay. Use the HS Assay Kit (Invitrogen Part no. Q32851 or Q32854) to measure dsdna concentrations from 10 pg/µl to 100 ng/µl. For samples outside this range, use the dsdna BR for higher concentrations of DNA (Invitrogen Part no. Q32850 or Q32853) or PicoGreen dsdna Assay Kit for lower concentrations (Invitrogen Part no. P7589). 2 Shear the DNA with the Covaris S220 System Note: For shearing conditions with the Covaris S2 System, see Shear the DNA with the Covaris S2 System on page 14. a. For each library, mix in a 1.5-mL LoBind Tube: Amount DNA 10 ng 5 µg 1 Low TE Buffer Variable µl Shear Buffer 1.2 µl Total 120 µl b. Prepare the Covaris S220 tank: Fill the tank to level 12 and set the chiller temperature to between 2 to 5 C to ensure that the temperature reading in the water bath is 5 C. c. Place a Covaris microtube into the loading station and transfer the mixture to the Covaris microtube. d. Shear the DNA using the appropriate program on the Covaris S220 System: Condition Setting Number of cycles 6 Bath temperature 5 C Bath temperature limit 10 C Mode Water quality testing function Frequency sweeping e. Place the Covaris microtube into the loading station, then transfer the DNA solution to a new 1.5-mL LoBind Tube. Off Duty Factor 10% Peak Incident Power Cycles/burst 100 Time 175 Watts 60 seconds 7

8 3 End-polish the DNA a. Combine in a new 1.5-mL LoBind Tube to prepare the end-polishing master mix: Amount per library Master mix for N libraries 5 Reaction Buffer 40 µl 40 µl (1.1 N) 10 mm dntp 8.0 µl 8.0 µl (1.1 N) End Polishing E1 8.0 µl 8.0 µl (1.1 N) End Polishing E2 10 µl 10 µl (1.1 N) Nuclease-Free Water 14 µl 14 µl (1.1 N) Total 80µL 80µL (1.1 N) 4 Size-select the DNA by Agencourt AMPure XP Bead Reagent b. Label 1.5-mL LoBind Tubes, one tube for each library. c. Transfer 120 µl of each sheared DNA from Shear the DNA with the Covaris S220 System on page 7 to the appropriately labelled tube. d. Pipet 80 µl of the master mix from step a into each labeled tube. While pipetting the master mix, take care not to cross-contaminate libraries. e. Vortex each reaction for 5 seconds, pulse-spin, then incubate at room temperature (20 25 C) for 30 minutes. a. Resuspend the Agencourt AMPure XP Reagent beads and allow the mixture to come to room temperature. b. Prepare 1 ml N of 70% ethanol. c. Size-select the DNA with resuspended, ambient Agencourt AMPure XP Reagent: 1. Prepare the bead suspension in the sample reaction for each library: Volume Sheared DNA 200 µl Agencourt AMPure XP Reagent 100 µl Total 300 µl Equal to 0.5 volume of end-polished DNA. 2. Vortex the beads for 10 seconds, then pulse-spin. 3. Incubate the mixture at room temperature (20 25 C) for 5 minutes 4. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution clears. The solution is clear of brown tint when viewed at an angle, as shown below: 8

9 4 Size-select the DNA by Agencourt AMPure XP Bead Reagent (continued) d. Carefully transfer the supernatant, which contains the DNA of the desired size, to a new 1.5- ml LoBind Tube. Discard the pellet. e. Bind the size-selected DNA in the supernatant to the Agencourt AMPure XP Reagent: 1. Combine: Volume Supernatant ~300 µl Agencourt AMPure XP bead suspension 60 µl Total ~360 µl Equal to 0.3 volume of the end-polish reaction volume of 200 µl. 5 Quantitate the sizeselected DNA 2. Vortex the beads for 10 seconds, then pulse-spin. 3. Incubate the mixture at room temperature (20 25 C) for 5 minutes. 4. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution clears, then remove and discard the supernatant. Save the pellet, which contains the DNA. f. Wash the DNA-bead complex 3 times. For each wash: 1. Add 200 µl of freshly prepared 70% ethanol to the tube, mix by inverting the tube a few times, then pulse-spin. 2. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution clears, then remove and discard the supernatant. g. Remove the tube from the DynaMag -2 magnetic rack, pulse-spin the tube, return the tube to the magnetic rack; then remove and discard the supernatant with a 20-µL pipettor. h. Open the tube, then dry the beads at room temperature (20 25 C) for 5 10 minutes. i. Elute the DNA: 1. Remove the tube from the DynaMag -2 magnetic rack, then add 36 µl Low TE Buffer directly to the pellet to disperse the beads. 2. Pipette the suspension up and down to mix. 3. Vortex the beads for 10 seconds, then pulse-spin. 4. Place the tube in a magnetic rack for at least 1 minute until the solution clears. 5. Transfer the supernatant containing the size-selected DNA to a new 1.5-mL LoBind Tube. Quantitate the DNA by using the Quant-iT dsdna HS Assay Kit, the NanoDrop ND-1000 Spectrophotometer, or the Agilent Technologies 2100 Bioanalyzer. IMPORTANT! The average yield of size-selected DNA is 30% of input quantity. If the yield is substantially <20%, troubleshoot the low yield, then repeat the procedure from Shear the DNA with the Covaris S220 System on page 7. 9

10 6 Add a da-tail to the sizeselected DNA a. In a new 1.5-mL LoBind Tube, combine to prepare the master mix to add an A tail to each library: Amount per library Master mix for N libraries 5 Reaction Buffer 10 µl 10 µl (1.1 N) 10 mm datp 1.0 µl 1.0 µl (1.1 N) A-Tailing Enzyme I 5.0 µl 5.0 µl (1.1 N) Total 16µL 16µL (1.1 N) 7 Ligate adaptors to the DNA b. Label 1.5-mL LoBind Tubes, one tube for each library. c. Transfer 34 µl of each size-selected DNA to the appropriately labeled tube. d. Pipet 16 µl of the master mix from step a into each labeled tube. While pipetting the master mix, take care not to cross-contaminate libraries. e. Incubate each reaction at 68 C for 30 minutes, then cool to room temperature. Note: While the reaction is incubating, calculate the amount of adaptors needed for ligation (see Ligate adaptors to the DNA ). IMPORTANT! If you are preparing barcoded libraries for multiplexed sequencing, for each sequencing run, use at least one of the following full sets of four barcodes: Barcodes 1 4, 5 8, 9 12, 13 16, 17 20, 21 24, 25 28, 29 32, 33 36, 37 40, 41 44, 45 48, 49 52, 53 56, 57 60, 61 64, 65 68, 69 72, 73 76, 77 80, 81 84, 85 88, 89 92, or Use only one of the barcoded-t-0xx adaptors for each ligation reaction, unless < 4 libraries are being barcoded. Use the barcodes according to these conditions: If <4 libraries are prepared for sequencing, then use multiple barcodes per library in equal ratios. For example, for 2 libraries, use 2 barcodes for each library. For 3 libraries, use 4 barcode adaptors for each library for a total of 12 barcodes. If 4 libraries are prepared for sequencing and libraries are split into sets of 4 to use full sets of barcodes, then use one set of barcodes for the remaining libraries (1,2,or 3 libraries). There is no need to use multiple barcodes per library in equal ratios. If the input DNA before shearing is: <100 ng: Use 0.06 µl of each adaptor or an equivalent amount of adaptor after dilution. For example, use 0.6 µl of a 10-fold dilution of an adaptor or 1.2 µl of a 20-fold dilution. 100 ng: Calculate the amount of adaptor needed, Y, for the reaction based on the amount of DNA from the last purification step. If DNA fragments were sheared using the standard protocol for fragment library preparation, the average insert size should be approximately 165 bp before adaptor ligation. IMPORTANT! Do not use P1 and P2 Adaptors that are designed for fragment library preparation and sequencing on the SOLiD 4 System. These adaptors are not compatible with reverse-read sequencing on the 5500 Series SOLiD Sequencers. Use on P1-T and Barcode-T-0XX Adaptors that are designed for the 5500 Series SOLiD Sequencers. a. Calculate the amount of adaptors needed: μg-to-pmol conversion factor = 10 6 pg 1 μg 1 pmol 660 pg (μg-to-pmol Y μl adaptor needed = # μg DNA 10 conversion factor) 1 Average insert size 1 μl adaptor needed 50 pmol 10

11 7 Ligate adaptors to the DNA (continued) b. In a new 1.5-mL LoBind Tube, combine: to prepare the ligase master mix: Volume per library Master mix for N libraries 5 Reaction Buffer 3.0 µl 3.0 µl (1.1 N) P1-T Adaptor, 50 µm YµL YµL (1.1 N) Barcode-T-0XX, 50 µm YµL Y µl (1.1 N) T4 DNA Ligase, 5 U/µL 6.5 µl 6.5 µl (1.1 N) 10 mm dntp 1.2 µl 1.2 µl (1.1 N) Nuclease-free Water Variable Variable Total 15 µl 15 µl (1.1 N) c. Add 15 µl of the ligation master mix from step b above to each reaction from Add a da-tail to the size-selected DNA on page 10 for a total of 61 µl total volume in each tube. d. Vortex each reaction for 5 seconds, pulse-spin, then incubate each reaction in a thermocycler with the lid heater on: IMPORTANT! Incubation nick translates the DNA. Stage Temp Time Holding 20 C 30 min Holding 72 C 20 min Holding 4 C e. For speed and high yield, purify the DNA with up to two rounds of purification with the Agencourt AMPure XP Reagent. In the first round, use 0.6 per sample of the Agencourt AMPure XP Reagent (see Purify the DNA using Agencourt AMPure XP Reagent on page 14). In the second round, use 1.5 per sample volume of the reagent. As an alternative to bead-based purification, for convenience, purify the DNA with the SOLiD Micro Column Purification Kit (see Purify the DNA with the SOLiD Library Micro Column Purification Kit on page 15). 8 9 Quantitate the ligated DNA (Optional) Amplify the libraries Quantitate the DNA by using the Quant-iT dsdna HS Assay Kit, the NanoDrop ND-1000 Spectrophotometer, the Agilent Technologies 2100 Bioanalyzer, and/or by qpcr. IMPORTANT! The current protocol is optimized for maximum yield from input DNA. In most cases, library amplification is not needed. If library amplification is needed, minimize the number of cycles, based on the amount of starting input DNA. Use minimal cycling to avoid over-amplification and production of redundant molecules. a. Transfer 20.0 µl of each purified library from step e in Ligate adaptors to the DNA. to a 0.2-mL PCR tube. Less than 20µL volume is acceptable. Do not adjust the volume to 20 µl. 11

12 9 (Optional) Amplify the libraries (continued) b. In a new 1.5-mL LoBind Tube, combine for a PCR master mix: Volume per amplification Master mix for N libraries Platinum PCR Amplification Mix 100 µl 100 µl (1.1 N) Library PCR Primer 1, 50 µm 2.5 µl 2.5 µl (1.1 N) Library PCR Primer 2, 50 µm 2.5 µl 2.5 µl (1.1 N) Total 105 µl 105 µl (1.1 N) c. Pipette 105 µl of the master mix from step b above into each labeled tube. While pipetting the master mix, take care not to cross-contaminate libraries. d. Vortex the mixture for 5 seconds, then pulse-spin. e. Determine the number of PCR cycles: Starting amount of DNA Number of cycles ng 10 cycles 100 ng 1 µg 6 8 cycles 1 2 µg 4 6 cycles 2 5 µg 0 3 cycles f. Run the PCR: Stage Step Temp Time Holding Denature 95 C 5min Cycling Denature 95 C 15 sec Anneal 62 C 15 sec Extend 70 C 1 min Holding Extend 70 C 5min Holding 4 C g. Purify the DNA using the Agencourt AMPure XP Reagent (see Purify the DNA using Agencourt AMPure XP Reagent on page 14). 10 Quantitate the DNA Quantitate the DNA by using the Quant-iT dsdna HS Assay Kit, the NanoDrop ND-1000 Spectrophotometer, the Agilent Technologies 2100 Bioanalyzer, and/or by qpcr. 11 Check the size distribution of the libraries Use 1 µl of sample in the Agilent Technologies 2100 Bioanalyzer. If you see the expected size distribution, proceed to (Optional) Pool equal molar barcoded libraries of similar size. If you do not see the expected size distribution, troubleshoot, or contact your Life Technologies Applications Specialist. 12

13 12 (Optional) Pool equal molar barcoded libraries of similar size IMPORTANT! If you are preparing barcoded libraries for multiplexed sequencing, for each sequencing run, use at least one of the following full sets of four barcodes: Barcodes 1 4, 5 8, 9 12, 13 16, 17 20, 21 24, 25 28, 29 32, 33 36, 37 40, 41 44, 45 48, 49 52, 53 56, 57 60, 61 64, 65 68, 69 72, 73 76, 77 80, 81 84, 85 88, 89 92, or Use only one of the barcoded-t-0xx adaptors for each ligation reaction, unless < 4 libraries are being barcoded. Use the barcodes according to these conditions: If <4 libraries are prepared for sequencing, then use multiple barcodes per library in equal ratios. For example, for 2 libraries, use 2 barcodes for each library. For 3 libraries, use 4 barcode adaptors for each library for a total of 12 barcodes. If 4 libraries are prepared for sequencing and libraries are split into sets of 4 to use full sets of barcodes, then use one set of barcodes for the remaining libraries (1,2,or 3 libraries). There is no need to use multiple barcodes per library in equal ratios. a. Quantitate the libraries to be pooled by qpcr [refer to the Applied Biosystems SOLiD Library TaqMan Quantitation Kit protocol (Invitrogen Part no. A12120)]. b. Mix together equal molar amounts of each barcoded library of similar size in an appropriately sized LoBind Tube. Vortex the tube. 13

14 Supplementary procedures Shear the DNA with the Covaris S2 System 1. For each library, mix in a 1.5-mL LoBind Tube: Amount DNA 10 ng 5 µg 1 Low TE Buffer Variable µl Shear Buffer 1.2 µl Total 120 µl 2. Prepare the Covaris S2 tank: Fill the tank to level 12. Ensure that the circulated water chiller is set to 2 5 C. Supplement the chiller not the actual water bath with 20% ethylene glycol. Ensure that the water bath temperature is 5 10 C. 3. Place a Covaris microtube into the loading station and transfer the mixture to the Covaris microtube. 4. Shear the DNA using the appropriate program on the Covaris S2 System: IMPORTANT! Ensure that the bath temperature limit is set at 15 C, and keep the bath temperature to 10 C. Condition Setting Number of cycles 6 Bath temperature 5 C Bath temperature limit 15 C Mode Water quality testing function Frequency sweeping Off Duty cycle 10% Intensity 5 Cycles/burst 100 Time 60 seconds 5. Place the Covaris microtube into the loading station, then transfer the 110 µl of sheared DNA to a new 1.5-mL sample tube provided in the Library Builder Fragment Core Kit for SOLiD 4.0. Purify the DNA using Agencourt AMPure XP Reagent 1. Resuspend the Agencourt AMPure XP Reagent beads, and allow the mixture to come to room temperature. 2. Prepare 70% ethanol. 3. Bind the DNA to the Agencourt AMPure XP Reagent: a. Prepare the bead suspension in the sample reaction according the recommended volumes of sample, Nuclease-Free Water, and Agencourt AMPure XP Reagent. b. Vortex the beads for 10 seconds, then pulse-spin. c. Incubate the mixture at room temperature (20 25 C) for 5 minutes. 14

15 d. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution is clear of brown tint when viewed at an angle; then, remove and discard the supernatant. 4. Wash the DNA 3 times. For each wash: a. Add 200 µl of freshly prepared 70% ethanol to the tube, mix by inverting the tube a few times, then pulse-spin. b. Place the tube in a DynaMag -2 magnetic rack for at least 1 minute until the solution clears, then remove and discard the supernatant. 5. Remove the tube from the DynaMag -2 magnetic rack, pulse-spin the tube, return the tube to the magnetic rack; then remove and discard the supernatant with a 20-µL pipettor. 6. Open the tube, then dry the beads at room temperature (20 25 C) for 5 10 minutes. 7. Elute the DNA: a. Remove the tube from the DynaMag -2 magnetic rack, then add the recommended volume of buffer directly to the pellet to disperse the beads. b. Pipet the suspension up and down to mix. c. Vortex the beads for 10 seconds, then pulse-spin. d. Place the tube in a magnetic rack for at least 1 minute until the solution clears. e. Transfer the supernatant containing the size-selected DNA to a new 1.5-mL LoBind Tube. Purify the DNA with the SOLiD Library Micro Column Purification Kit 1. Pre-spin before use an empty PureLink Micro columns in collection tubes at 10,000 g for 1 minute. 2. Load the DNA onto the PureLink Micro columns: a. To 1 volume of sample, add 4 volumes of Binding Buffer (B2-L) with 40% isopropanol. b. Apply all of the mixture to the PureLink Micro column(s) in collection tube(s). c. Spin the column(s) at 10,000 g for 1 minute at room temperature, then discard the flow-through. dsdna is bound to the column. 3. Wash the column(s): a. Return the PureLink Micro column(s) to the same collection tube(s). b. Add 650 µl of Wash Buffer (W1) with 80% ethanol to wash the column(s). c. Spin the column(s) at 10,000 g for 1 minute at room temperature, then discard the flow-through. d. Spin the column(s) at 14,000 g at room temperature to remove residual wash buffer and dry the silica, then discard the flowthrough and collection tube. 4. Elute the DNA: a. Transfer the column(s) to clean 1.5-mL LoBind tube(s). b. Add 22 µl of Elution Buffer (E1) to the center of the column(s) to elute the DNA, then let the column(s) stand for 1 minute. c. Spin the column(s) at 14,000 g for 1 minute at room temperature. d. (Optional) Add the eluate from the last spin back to the column(s), then let the column(s) stand for 1 minute, then spin the column(s) at 14,000 g for 1 minute at room temperature. 5. If necessary, pool the eluted DNA into one 1.5-mL LoBind Tube. 15

16 For Research Use Only. Not intended for any animal or human therapeutic or diagnostic use. NOTICE TO PURCHASER: PLEASE REFER TO THE APPLIED BIOSYSTEMS 5500 SERIES SOLiD TM FRAGMENT LIBRARY PREPARATIONS USER GUIDE FOR LIMITED LABEL LICENSE OR DISCLAIMER INFORMATION. The trademarks mentioned herein are the property of Life Technologies Corporation or their respective owners. Bioanalyzer is a trademark of Agilent Technologies, Inc. AMPure is a registered trademark of Beckman Coulter, Inc. Covaris is a registered trademark of Covaris, Inc. NanoDrop is a registered trademark of NanoDrop Technologies. TaqMan is a registered trademark of Roche Molecular Systems, Inc. Copyright 2011, Life Technologies Corporation. All rights reserved. Part Number Rev. A 03/2011 Headquarters 5791 Van Allen Way Carlsbad, CA USA Phone Technical Resources and Support For the latest technical resources and support information for all locations, please refer to our Web site at