Strand Specific 96-well Library Construction for Illumina Sequencing … · 2018-03-22 · Strand...

Strand Specific 96-well Library Construction for Illumina Sequencing

Saved as: LIBPR.0081 Supersedes: Version 1

Version: 2 Page 1 of 42

Non Controlled Version *Note: Controlled Versions of this document are subjected to change without notice


I. Purpose

To provide specific guidelines for 96-well, strand specific (coding strand only) RNA-seq (WTSS) library construction from cDNA template for Illumina Paired-End Sequencing.

II. Scope

All procedures are applicable to the BCGSC Library TechD and the Library Core groups.

III. Policy

All production procedures shall be documented and controlled by approved systems.

IV. Responsibility

It is the responsibility of all personnel performing this procedure to follow the current protocol. It is the responsibility of the Group Leaders to ensure personnel are trained in all aspects of this protocol. It is the responsibility of Quality Assurance Management to audit this procedure for compliance and maintain control of this procedure.

V. References

Reference Title Reference Number

Sample Preparation for Paired-End Sample Prep Kit from Illumina Version 1.1 (from Prep Kit)

VI. Related Documents

Document Title Document Number

DNA Transfer to Covaris Tube Rack and Sheared DNA Transfer to 96-well Plate Using the Biomek FX

LIBPR.0045

96-well DNA Purification Using Ampure Magentic Beads and Biomek FX

LIBPR.0047

96-well DNA Quantification using PicoGreen and VICTOR3V LIBPR.0048 Operation of Covaris E-Series LIBPR.0041 Operation and Maintenance of the Agilent 2100 Bioanalyzer for DNA samples

LIBPR.0017

Illumina Concentration Checked LIBPR.0030 Operation and Maintenance of the Caliper Labchip GX for DNA LIBPR.0051





Document Title Document Number

samples using the High Sensitivity Assay Operation of the Invitrogen Egel iBase Power System LIBPR_WORKINST.0012

VII. Safety

All Laboratory Safety procedures will be complied with during this procedure. The required personal protective equipment includes a laboratory coat and gloves. See the material safety data sheet (MSDS) for additional information.

VIII. Materials and Equipment

Name Supplier Number: # Model or Catalogue #

NEB Paired-End Sample Prep Kit NEB E6000B-25 �

Fisherbrand Textured Nitrile gloves – various sizes

Fisher 270-058-53 �

Ice bucket – Green Fisher 11-676-36 �

Wet ice In house N/A N/A N/A

AB1000 96-well 200µl PCR plate Fisher AB1000 �

Gilson P2 pipetman Mandel GF-44801 �


Gilson P20 pipetman Mandel GF23600 �



Neptune barrier tips 10 µl Intersciences LPBT10 �




Galaxy mini-centrifuge VWR 37000-700 �

VX-100 Vortex Mixer Rose Scientific S-0100 � AMPure XP, 450mL Beads Agencourt 000132 �

Black ink permanent marker pen VWR 52877-310 �

Clear Tape Sealer Qiagen 19570 �

Eppendorf BenchTop Refrigerated Centrifuge 5810R Eppendorf 5810 R �

Bench Coat (Bench Protection Paper) Fisher 12-007-186 �

Small Autoclave waste bags 10”X15” Fisher 01-826-4 �

MinElute PCR Purification Kit (50) Qiagent 28004 �

DNA Away VWR 53509-506 �

Mussel Glycogen (20mg) Roche Scientific 10 901 393 001

�

3 M Sodium Acetate Sigma EC 211-162-9 �

Anhydrous Ethyl Alcohol (100% Ethanol) Commercial Alcohol

People Soft ID: 23878

�





Mylar PET film, clear (40”x10’x 0.003”) McMaster-Carr 8567K32

�

Plastic wrap In house N/A

�

Agilent DNA 1000 Series II Kit Agilent 5067-1504 �

DNA 1000 Gel Matrix Agilent 5067-1504 �

DNA 1000 Dye Concentrate Agilent 5067-1504 �

DNA 1000 Marker Agilent 5067-1504 �

DNA 1000 Ladder Agilent 5067-1504 �

Agilent DNA 1000 Chips Agilent 6064-8230 �

Agilent Chip Priming Station Agilent Chip Priming Station

� �

IKA Works Vortexer Agilent MS2S9-Agilent-5065-4428

�

22R Microfuge Centrifuge Beckman 22R Centrifuge � Agilent Electrode Cleaner

Agilent 6064-8230 �

Peltier Thermal Cycler MJ Research PTC-225 �

Power Supply, LVC2kW, 48VDCV Tyco Electronics RM200HA100 �

Spin-X Filter Tube Fisher CS008160 �

VICTOR3V Perkin Elmer 1420-040 �

Uracil N-Glycosylase Applied Biosystems N808-0096 �

P2-20 Rainin Lite Manual 12-channel Rainin L12-20 �

�

P20-200 Rainin Lite Manual 12-channel Rainin L12-200

P200 Barrier Rainin tips Rainin RT-L200F

P20 Barrier Rainin tips Rainin RT-L10F

Power PAC BioRad Power PAC 200 �

50 X TAE In House N/A �

1 X TAE In House N/A N/A N/A

100 bp Ladder Invitrogen 15628-019 N/A N/A

IX. Procedure

1. Introduction and Upstream Set Up

1.1. Put on a clean pair of gloves and a lab coat. Wipe down the assigned specific workstation,

pipetman, and small equipment. Lay down new bench coat and retrieve ice and all required reagents.

1.2. It is important to wear gloves when handling sample plates, reagents and equipment, and to treat everything with clean PCR techniques.

1.3. General guidelines:

A. Never re-use plate sealing tape.





B. To avoid cross-well contamination, reaction plates should never be vortexed. C. Plate mixing steps are only to be done using Biomek FX or manual 12-channel

pipette. D. All buffers should be well mixed before addition to the brew. All brews should be

well mixed in tubes before dispensing into reaction plates to assure equal

distribution of all components and thus uniformity of enzymatic reactions across a

plate.

1.4. Up to 3 plates per week can be processed at a time by one technician using Biomek FX.

This protocol also allows for processing of partial plates (3 rows of samples or less) using manual Rainin 12-channel pipette. Plate type to be used in this protocol is AB1000.

1.5. Plates can be stored at -20°C overnight after every step except after “A” addition and after

dUTP strand digestion.

Note: “A” addition and adapter ligation reactions should be performed on the same day.

Similarly, dUTP digestion and PCR should be performed on the same day as well.

1.6. Store “A” tailed reactions and dUTP digested product at 4°C (or on ice) until ready to

proceed with same day XP bead clean-up.

1.7. All pre-PCR work should be done on the 5th Floor. Post PCR work is to be performed on the 6th floor.

1.8. Single use aliquots of PE Illumina adapter once taken out of the freezer should be kept on

ice and never refrozen.

1.9. Indexing PCR primers are allowed to be thawed only up to 3 times. Each time the plate is taken out of the freezer and thawed it should be marked even if it was not used (including when plates are taken out during freezer maintenance). Ideally Indexing PCR primer plates should be single use.

1.10. The input material for this procedure is double stranded, strand specific cDNA generated

using SOP: “Strand Specific_ 96well cDNA Synthesis”. 1.10.1. Prior to starting this procedure the quality and quantity of cDNA template should be

assessed using HS Agilent Assay at least for controls and 24 random samples from each plate. For cDNA quality control acceptance criteria refer to “Strand Specific_ 96well cDNA Synthesis” document.





1.11. The minimum input material for this pipeline is 15ng of strand specific cDNA as measured by Qubit or Picogreen assays after Covaris E-series shearing and one XP beads clean up. It is the minimum expected yield of strand specific cDNA from 2ug of human or mouse total RNA of RIN 7.0 or above.

2. Covaris E-series shearing

2.1. Adjust the volume of cDNA to 60uL/well using Qiagen EB buffer:

2.1.1. Open project “FG Indexing” and select program:“P165B_384Axygen_to_AB1000” 2.1.2. Add 25µL of Quiagen EB buffer to every well (after cDNA synthesism, Ampure XP

bead clean up, and QC you should have 35µL remaining/per well). 2.1.3. Follow the displayed deck layout for set up and click “OK”. 2.1.4. When the program is complete, seal the plates with clear tape sealer and quick spin

in an Eppendorf centrifuge using Program 2 (2000g for 1 minute at 4ºC). Keep the plates on ice.

2.2. For Covaris E-series shearing set-up refer to protocols:

a) LIBPR.0045 DNA Transfer to Covaris Tube Rack and Sheared DNA Transfer to 96-

well Plate Using the Biomek FX b) LIBPR.0041 Operation and Maintenance of Covaris E-series

2.3. Shearing conditions for strand specific PET library construction using Covaris E-series are:

Duty cycle: 20% Intensity: 5 Time: 55 seconds shearing Cycles per burst: 200

Note: Samples in Covaris tubes can be left in the 4°C-6°C water bath overnight. In order

to save time the technician can start Covaris-E series shearing at the end of the

working day and retrieve the completed plate the following morning.

2.4. After the Covaris shearing, please run 11 samples on the Agilent High Sensitivity

DNA chip to make sure the samples have been sheared to the correct size. See

Operation and Maintenance of the Agilent 2100 Bioanalyzer for DNA samples-

LIBPR.0017

2.5. LIMS tracking:





2.5.1. Use a handheld scanner. Enter your user name and password, select sequence database, Lib Construction department, 6th floor printers (GE), and click “log in” button.

2.5.2. Select the Lib Construction tab. Save or retrieve plate set. 2.5.3. From the Protocol drop-down menu, select “96Well_Covaris_Shear_DNA” and

click “Continue Prep”. 2.5.4. Scan the equipment used for shearing, enter freezer rack location, and select IDX

pipeline. In the Comments field enter the shearing time and click on “Completed Covaris shear DNA”.

3. Ampure Magnetic XP Beads Cleanup after Covaris shearing

3.1. After shearing, DNA material requires XP bead clean before entering end-repair. The

DNA volume coming out of Covaris shearing is 60uL.

3.2. To clean up DNA after Covaris shearing:

3.2.1. When processing full plates, follow protocol LIBPR.0047 – 96-well DNA Purification Using Ampure Magnetic Beads and Biomek FX.

3.2.2. When processing 3 rows of samples or less, both the protocols:

a) LIBPR.0047 96-well DNA Purification Using Ampure (or XP Beads) Magnetic

Beads and Biomek FX or

b) LIBPR.0073 Manual Bead Clean using Ampure XP Beads can also be used in this step.

3.3. LIMS protocol:

3.3.1. Use a handheld scanner. Enter your user name and password, select sequence

database, Lib Construction department, 6th floor printers (GE), and click “log in” button.

3.3.2. Select the Lib Construction tab. Retrieve plate set. 3.3.3. From the Protocol drop-down menu, select “96Well_Cleanup_DNA” and click

“Continue Prep”. 3.3.4. Click on “Completed Cleanup DNA”.

4. PicoGreen DNA quant





4.1. When processing full plates, quant DNA following the protocol: LIBPR.0048 – 96-well

DNA Quantification using PicoGreen and VICTOR3V.

4.2. When processing 3 rows of samples or less, quant DNA using Qubit.

4.3. Using excel sheet, calculate total ng in each well.

4.4. Both the quality and the quantity of the final library product are directly linked to the cDNA input amount. Less than 15ng of strand specific cDNA entering this pipeline may result in failures and/or low data quality. For comparison:

4.4.1. 2µg of human or mouse total RNA (RIN 7.0 or above) typically results in about 15-

25ng or more of strand specific cDNA as measured by Qubit after Covaris shearing and one Ampure XP bead cleanup.

4.4.2. 4µg of human or mouse total RNA typically results in about 40-50ng or more of strand specific cDNA as measured by Qubit after Covaris shearing and one Ampure XP bead clean up.

5. End-Repair and Phosphorylation

Note: Reaction brew must be made in the PCR Clean Room laminar flowhood on the 5th

floor, room 510.

5.1. Retrieve and thaw 5x Ligase Buffer and 10mM dNTPs at room temperature then place

them on ice. Leave enzymes in the freezer until you are ready to add them to the brew.

5.2. Reaction set up using LIMS:

5.2.1. In alDente login window type in your user name and password, select Database: sequence, Department: Lib Construction, and Printer group: 6th Floor Printers (GE). Click “Log In” button.

5.2.2. Under Mix Standard Solutions select “LibConst_End_Repair_Brew”.

5.2.2.1. If processing full plates, leave “1” in the first box and choose 96 for number of samples. Click “Mix Standard Solution”. If processing more than one plate, under Parameters, enter the number of plates, leave the default settings of 96 for both # of BrewSourceWells and # of Wells. Click “Re-calculate Standard Solution”.





5.2.2.2. If processing a partial plate, enter number of samples in the first box and “1” in the second. Click “Mix Standard Solution”.

Figure 1. Mix Standard Solution

5.2.2.3. If processing a partial plate, under Parameters, enter 12 for the # of

BrewSourceWells, and enter the number of wells being processed. Click “Re-calculate Standard Solution”.

Figure 2. Re-calculate Standard Solution

5.2.3. Enter solution numbers into the brew calculator. 5.2.4. Select Type: Reagent, select Grp: Lib Construction, and Barcode Label: 1D Large

Solution/Box/Kit Labels. Double check that everything is entered correctly, then click “Save Standard Mixture”.

5.2.5. Retrieve both the brew barcode and reagent check list label. Place both in your lab notebook.

5.2.6. In the case where LIMS is unavailable, prepare the brew using the “LibConst_ End

Repair and Phosophorylation_Brew” excel worksheet located in: R:\Functional_Genomics\Library_Core\Library Production\Plate based library construction\LIMS Backup Brew

5.3. Prepare brew in a clean 15mL tube according to the printed calculator label (or if LIMS is down excel sheet), checking off reagents using a blue pen as they are added. 5.3.1. Mix each reagent (buffers by pulse-vortexing and enzymes by gentle flicking of the

tube) and spin down in a minifudge before addition to the brew.





5.3.2. Mix the brew very well by gentle, repeated pulse-spin vortexing. Spin down in a minifuge and store the brew on ice.

5.4. If processing full plates, on ice, using Gilson Pipetman dispense the following volume/well

into one 96-well AB-1000 Brew Source Plate: 5.4.1. For one plate: 20µL/well 5.4.2. For two reaction plates: 35µL/well 5.4.3. For 3 reaction plates: 50µL/well

5.5. If processing a partial plate, on ice, using Gilson Pipetman, based on the number of rows,

dispense the correct amount of the End Repair brew into one row of AB1000 plate. 5.5.1. For one row of samples: 20µL/well (15µL+5µL dead volume) 5.5.2. For 2 rows of samples: 35µL/well (15µL*2 +5µL dead volume) 5.5.3. For 3 rows of samples: 50µL/well … ect.

5.6. Cover brew source plate with plate seal and quick spin at 4ºC, 2000g for 1 minute. Keep it

on ice.

5.7. If processing a partial plate, on ice, using Rainin P2-20 Manual Multichannel transfer 15µL of the brew into each row of DNA samples. After addition mix each row 15 times. Use a circular mixing motion: draw volume from the bottom of the well and dispense higher up at the liquid level. During mixing monitor the volume in each tip to make sure that all wells within the row are getting mixed. Change tips between rows. Mark each row on the side of the plate so that no rows are skipped or repeated. Seal the plate and spin it down at 4°C, 2000g for 1min. Go to step 5.14.

5.8. If processing full plates, log into Biomek FX-5. Under Project “FG Indexing”, open

program: “Lib_Construction_Reaction_Setup_AB1000”.

5.9. Wipe the Biomek and bench with ddH2O followed by 70% EtOH.

5.10. Click on the green arrow “Run” button in the top menu bar. The program will prompt you to enter the number of reaction plates. Select “N” for “AdapterLigation”, select “N” for dA_Addition; select “Y” for EndRepair. Enter number of plates being processed.

Note: Reaction brews vary in viscosity. In order to assure the accuracy of transferred

volume it is very important to select the correct Biomek technique, in this case “Y”

(yes) for End Repair.





5.11. Use P20 Beckman FX barrier tips. The End Repair brew plate is the “Source”. “Dest” is the DNA plate (or plates if processing more than one). Make sure to remove all the covers, both from plates and tip boxes.

5.12. Confirm that the location of the plates and tip boxes on the Biomek deck matches the

software deck layout on the computer screen and click OK to start the program.

Figure 3. Deck layout to dispense 15µL of End Repair brew into the plates containing DNA.

5.13. After the program is completed, seal the plates and quick spin in an Eppendorf centrifuge

on Program 2 (2000g at 4ºC for 1 minute). Discard tips and recycle the boxes.

5.14. Check the brew source plate for the remaining dead volume. There should be about 5µL or less of the brew left in every well. Record in your lab notebook if there are any variations in the left over volume and notify your supervisor. Discard the source brew plate.

5.15. Incubate End Repair reaction plates at room temperature for 30 minutes.

5.16. LIMS: Retrieve the plate set number. From the Protocol drop-down menu, select “Index_End_Repair” and click “Continue Prep”. Scan the barcode of the brew solution. The brew volume is defaulted to 15µL. Enter the Pre-PCR Intermediates -20ºC freezer rack location. Click on “Completed Scan Index 96_ER Brew-Add 15µL to DNA”.

5.17. This is a safe stopping point. After 30 minutes room temperature incubation, plates can be stored at -20°C.





6. Ampure Magnetic Bead Clean Up after End Repair

Note: Use Biomek FX-5 on the 5th

floor for Pre-PCR work. Do not use the 6th

floor Biomek.

6.1. The DNA volume going into this step is 50µL/well.

6.2. To clean up DNA after end repair reaction:

6.2.1. When processing full plates, follow protocol LIBPR.0047 – 96-well DNA

Purification Using Ampure Magnetic Beads and Biomek FX. 6.2.2. When processing 3 rows of samples or less, both the protocols:

a) LIBPR.0047 96-well DNA Purification Using Ampure (or XP Beads)

Magnetic Beads and Biomek FX or


6.3. LIMS: Retrieve the plate set number. From the Protocol drop-down menu, select

“Index_Cleanup_ER” and click “Continue Prep”. Scan the Ampure magnetic bead solution number. The bead volume is defaulted to 100µL. Enter the Pre-PCR Intermediates -20ºC freezer rack location. Make sure IDX pipeline is selected. Click on “Completed Transfer to 96-well ABGene”. This will generate a new plate set number. Record the new plate set number in your lab notebook and retrieve the barcode from the barcode printer.

6.4. This is a safe stopping point. Plates can be stored at -20°C.

7. Addition of an ‘A’ Base to the 3’ End of the cDNA Fragments

Note: Step 6 needs to be started in the morning as “A” addition and Adapter Ligation MUST

be performed on the same day.

Note: “A” addition reaction brew must be made in the PCR Clean Room laminar flowhood

on the 5th

floor, room 510.

7.1. Retrieve and thaw 10x Klenow Buffer and 10mM dATP at room temperature then place

them on ice. Leave enzymes in the freezer until you are ready to add them to the brew.






7.2.1. Under Mix Standard Solutions select “LibConst_A_addition_Brew”.

7.2.1.1. If processing full plates, leave “1” in the first box and choose 96 for number of samples. Click “Mix Standard Solution”. If processing more than one plate, under Parameters, enter the number of plates, leave the default settings of 96 for both # of BrewSourceWells and # of Wells. Click “Re-calculate Standard Solution”.

7.2.1.2. If processing a partial plate, enter number of samples in the first box and “1” in the second. Click “Mix Standard Solution”. If processing a partial plate, under Parameters, enter 12 for the # of BrewSourceWells, and enter the number of wells being processed. Click “Re-calculate Standard Solution”.

7.2.2. Enter solution numbers into the brew calculator. 7.2.3. Select Type: Reagent, select Grp: Lib Construction, and Barcode Label: 1D Large

Solution/Box/Kit Labels. Click “Save Standard Mixture”. 7.2.4. Retrieve both the brew barcode and reagent check list label. Place both in your lab

book.

7.3. Reaction set up using spreadsheets: In the case where LIMS is unavailable, prepare the brew using “LibConst_A_addition_Brew” excel worksheet located in: R:\Functional_Genomics\Library_Core\Library Production\Plate based library construction\LIMS Backup Brew Calculators

7.4. Prepare brew in a clean 15mL tube according to the printed calculator label (or if LIMS is down excel sheet), checking off reagents using a blue pen as they are added. 7.4.1. Mix each reagent (buffers by pulse-vortexing and enzymes by gentle flicking of the

tube) and spin down in a minifudge before addition to the brew. 7.4.2. Mix the brew very well by gentle, repeated pulse-spin vortexing. Spin down in a

minifuge and store the brew on ice.

7.5. If processing full plates, on ice, using Gilson Pipetman dispense the following volume/well into one 96-well AB-1000 Brew Source Plate: 7.5.1. For one plate: 20µL/well 7.5.2. For two reaction plates: 35µL/well





7.5.3. For 3 reaction plates: 50µL/well

7.6. If processing a partial plate, on ice, using Gilson Pipetman, based on the number of rows, dispense the correct amount of the End_Repair brew into one row of AB1000 plate. 7.6.1. For one row of samples: 20µL/well (15µL+5µL dead volume) 7.6.2. For 2 rows of samples: 35µL/well (15µL*2 +5µL dead volume) 7.6.3. For 3 rows of samples: 50µL/well … ect.

7.7. Cover brew source plate with plate seal and quick spin at 4ºC, 2000g for 1 minute. Keep it on ice.

7.8. If processing a partial plate, on ice, using P2-20 Manual Rainin Multichannel transfer 15µL of the brew into each row of DNA samples (previously end-repaired and XP bead cleaned). After addition mix each row 15 times. Use a circular mixing motion: draw volume from the bottom of the well and dispense higher up at the liquid level. Change tips between rows. Mark each row on the side of the plate so that no rows are skipped or repeated. Seal the plate with PCR cover and spin it down at 4°C, 2000g for 1min. Go to step 7.15.

7.9. If processing full plates, use the Biomek FX and P20 Beckman FX barrier tips to add 15µL

of the “A” addition brew to the DNA plates.



floor

Biomek.

7.10. Log into Biomek FX-5. Under Project “FG Indexing”, open program: “Lib_Construction_Reaction_Setup_AB1000”.

7.11. Wipe the Biomek and bench with ddH2O followed by 70% EtOH.

7.12. Click on the green arrow “Run” button in the top menu bar. The program will prompt you

to enter the number of reaction plates. Select “N” for “AdapterLigation”, select “Y” for dA_Addition, select “N” for EndRepair. Enter number of plates.

Note: Reaction brews vary in viscosity and in order to assure an accurate volume transfer

it is very important to select here the correct Biomek technique, in this case “Y”

(yes) for dA_Addition.

7.13. “A” addition Brew plate is the “Source”. “Dest” is the DNA plate (or plates if processing

more than one). Use P20 Beckman FX barrier tips. Confirm that your placement of plates and tip boxes on the Biomek deck matches the software deck layout on the





computer screen. Make sure to remove all of the covers, both from tip boxes and plates. Click OK to start the program.

Figure 4. Deck layout to dispense 15µL of dA Addition brew into the plates containing End Repaired DNA.

7.14. After the program is completed, cover reactions with PCR cover and quick spin in an

Eppendorf centrifuge on Program 2 (2000g at 4ºC for 1 minute). Discard tips and recycle the boxes.

7.15. Check the source brew plate for the uniformity of the remaining dead volume. There should be about 5µL or less left in every well. Record in your lab notebook if there are any variations in the left over volume and notify your supervisor. Discard the source brew plate.

7.16. Use PCR program “37” to incubate reaction plates at 37°C for 30 minutes. Before starting

double check the conditions of the program to make sure that it is correct.


“Index_Add_A-Base_To_ER_DNA” and click “Continue Prep”. Scan the brew solution number. The brew volume is defaulted to 15µL. Enter the Pre-PCR Intermediates -20ºC freezer rack location. Click on “Completed Add A-Base Mix to ER DNA plates”.

7.18. After 30 min incubation, proceed to the next step. This is NOT a safe stopping point.

8. Ampure Magnetic Bead Clean Up after A-Tailing



floor Biomek






8.2. To clean up DNA after “A” addition reaction:







“Index_Cleanup_A-Base”. Scan the Ampure Magnetic bead solution number. The bead volume is defaulted to 100µL. Enter the Pre-PCR Intermediates -20ºC freezer rack location. Make sure IDX pipeline is selected. Click on “Completed Transfer to 96-well ABGene”. This will generate a new plate set number. Record the plate set number in your lab notebook and retrieve the barcode from the barcode printer.

8.4. Proceed to the next step. This is NOT a safe stopping point.

9. PicoGreen DNA quant after “A” base addition.

9.1. When processing full plates, quant DNA following the protocol: LIBPR.0048 – 96-well

DNA Quantification using PicoGreen and VICTOR3V.

9.2. When processing 3 rows of samples or less, you can quant DNA using Qubit.

9.3. Using excel spreadsheet, calculate total ng and the % yield for every well (the starting amount is the DNA quant after shearing and one bead clean). The yield for every well should not be below 50% (the expected yield is 60%). Calculate the average yield and standard deviation. The standard deviation should not be more than 10%. Notify your supervisor of any variations from the expected yield.

9.4. Calculate the total number of molecules of template (pmoles) for every well (size 300bp, 35µL volume). Calculate the amount of Adapter (in pmoles) for each well by multiplying





total pmoles of template in each well times 10. Based on that calculation decide which stock of PE adapter to use, either 1µM or 10µM stock. The maximum volume of adapter per well is 6µL so 1µM stock can be used up to 6 pmoles/well. Above 6 pmoles/well you have to use 10µM stock. Most of the time you will be in range of 1µM adapter.

9.5. The minimum amount of Adapter per well for this pipeline is 1 pmoles/well. If any wells,

based on the calculation, call for less than 1 pmoles of Adapter, in the spreadsheet change the amount of PE Adapter for those wells to 1 pmoles and then calculate the average volume of adapter/well.

9.6. Show your results, and confirm your calculations with your supervisor. If everything is

correct, record the average volume and stock concentration of adapter in your notebook (you will need to input this value into the LIMS calculator).

9.7. For each plate, email the electronic copy of the excel file (saved as .xls) containing well notation (by row), library name, library pla, and total ng/well entering adapter ligation (as per Picogreen or Qubit quant) to your supervisors so that they can upload “Amount_for_Ligation_ng” attribute into LIMS. Include in the file the tra number of the plate to which the attribute should be attached which is the “Index_Cleanup_A-Base” plate.

Note: While uploading cDNA amount entering adapter ligation into LIMS double check that

the library names match the attributes. You should see both the library name and the

corresponding pla next to the attributes.

9.8. LIMS: Retrieve plate set. From the Protocol dropdown menu, select

“96well_PicoGreen_Quant”. Enter the storage location and the file name in the Picogreen_Pre_Ligation attribute field.

10. Ligate Illumina PE Adapter to DNA Fragments

Note: Adapter Ligation brew must be made in the PCR Clean Room laminar flowhood on

the 5th

floor, room 510.

10.1. Retrieve and thaw 5X Ligase Buffer and PE Adapter at room temperature then place them

on ice. (Once thawed, do not keep PE Adapter at room temperature! Transfer to ice.). Note that different adapter concentrations (1µM or 10µM) may be used for special samples, please consult with supervisor. Leave enzymes in the freezer until you ready to add them to the brew.





Note: Thaw PE Adapter in the Tissue Culture Room laminar flowhood on the 5th

floor, room 511. After it is thawed immediately place it on ice.


10.2.1. Click on “Solutions” tab. Under Mix Standard Solutions select “LibConst_Adapter_Ligation_Brew” and enter number of plates and 96 for number of samples. Click “Mix Standard Solution”.

10.2.2. Enter the volume of adapter per well (in uL). Click “Recalculate” button. The ‘BrewSourceWells’ field represents the dead volume and is to be left unchanged at 12. Enter solution numbers into the brew calculator.

10.2.3. Select Type: Reagent, select Grp: Lib Construction, and Barcode Label: 1D Large Solution/Box/Kit Labels. Click “Save Standard Mixture”.


10.3. Reaction set up using spreadsheets:

10.3.1. In the case where LIMS is unavailable, prepare the brew using

LibConst_Adapter_Ligation_Brew excel worksheet located in: R:\Functional_Genomics\Library_Core\Library Production\Plate based library construction\LIMS Backup Brew Calculators

10.4. Prepare the brew in a clean 15mL tube according to the printed calculator label (or if

LIMS is down use the excel sheet), checking off reagents using a blue pen as they are added.

10.4.1. Mix each reagent (buffers by pulse-vortexing and enzymes by gentle flicking of the

tube) and spin down in a minifudge before addition to the brew. 10.4.2. On ice, add T4 DNA Quick Ligase to the brew last. To minimize adapter-adapter

ligation, after addition of the Ligase enzyme it is important to aliquot the brew and subsequently add the brew to the DNA as soon as possible.

10.4.3. Mix the brew very well by gentle, repeated pulse-spin vortexing. Spin down in a minifuge and keep the brew on ice. Immediately proceed to the next step.

10.5. When processing 3 rows of samples or less:

10.5.1. On ice, using Gilson Distiman, based on the number of rows, dispense the correct

amount of the Adapter Ligation brew into one row of AB1000 plate.





10.5.1.1. For one row of samples: 30µL/well (25µL+5µL dead volume) 10.5.1.2. For 2 rows of samples: 55µL/well (25µL*2 +5µL dead volume) 10.5.1.3. For 3 rows of samples: 80µL/well … ect.

10.5.2. Cover brew source plate with plate seal and quick spin at 4ºC, 2000g for 1 minute.

Keep it on ice. 10.5.3. If processing a partial plate, on ice, using P20-200 Manual Rainin Multichannel

transfer 25uL of the brew into each row of DNA samples. After addition mix each row 15 times. Use a circular mixing motion: draw volume from the bottom of the well and dispense higher up at the liquid level. During mixing monitor the volume in each tip to make sure that all wells within a row are getting mixed. Change tips between rows. Mark each row on the side of the plate so that no rows are skipped or repeated. Seal the plate with PCR cover and spin it down at 4°C, 2000g for 1min. Go to step 10.9.

10.6. When processing full plates, on ice, using Gilson Distriman, dispense the following

volume/well into one 96-well AB-1000 Brew Source Plate: 10.6.1. For one plate: 30µL/well. 10.6.2. For two reaction plates: 25µL * 2 plates + 5µL dead volume = 55µL/well of brew

source plate. 10.6.3. For 3 reaction plates: 25µL * 3 plates + 5µL dead volume = 80µL/well of brew

source plate.

10.7. For processing full plates, use the Biomek FX and P20 Beckman FX barrier tips to add 25µL of the Adapter Ligation brew to the A tailed DNA:



floor

Biomek.

10.7.1. Wipe the Biomek and bench with ddH2O followed by 70% EtOH. 10.7.2. Under Project “FG Indexing”, open program:

“Lib_Construction_Reaction_Setup_AB1000.” 10.7.3. Click on the green arrow “Run” button in the top menu bar. The program will

prompt you to enter the number of reaction plates. Select “Y” for “AdapterLigation”, select “N” for dA_Addition, select “N” for EndRepair. Enter number of plates.

Note: Reaction brews vary in viscosity and in order to assure an accurate volume

transfer it is very important to select here the correct Biomek technique, in this

case “Y” (yes) for AdapterLigation.





10.7.4. The Adapter Ligation brew plate is the “Source”. “Dest” is the DNA plate (or

plates if processing more than one). Use P20 Beckman FX barrier tips. Confirm that your placement of plates and tip boxes on the Biomek deck matches the software deck layout on the computer screen. Make sure to remove all of the covers, both from tip boxes and plates. Click OK to start the program.

Figure 5. Deck layout to dispense 15µL of Adapter Ligation brew into the plates containing A tailed DNA.

10.8. After the program is completed, cover reactions and quick spin in an Eppendorf centrifuge

on Program 2 (2000g at 4ºC for 1 minute). Discard tips and recycle the boxes.

10.9. Check the source brew plate for the uniformity of the remaining dead volume. There should be about 5uL left in every well. Record in your lab notebook if there are any variations in the left over volume and notify your supervisor. Discard the source brew plate.

10.10. Incubate Adapter Ligation reaction plates at room temperature OVERNIGHT.

10.11. LIMS: Retrieve the plate set number. From the Protocol drop-down menu, select “Index_Adapter_Ligation” and click “Continue Prep”. Scan the brew solution number, enter the brew volume used, and enter the Pre-PCR Intermediates -20ºC freezer rack location. Make sure IDX pipeline is selected. Click on “Completed Aliquot to 96-well ABGene”. This will generate a new plate set number. Record the plate set number in your lab notebook and retrieve the barcode from the barcode printer.

11. Ampure Magnetic Bead Clean Up after Adapter Ligation







floor Biomek.


11.2. To clean up DNA after PE Adapter Ligation reaction:

11.2.1. When processing full plates, follow protocol LIBPR.0047 – 96-well DNA Purification Using Ampure Magnetic Beads and Biomek FX. The volume of reaction is 60µL so make sure to change the waste volume to 160µL.






“Index_Cleanup_Adapter_Ligation” and click “Continue Prep”. Scan the Ampure magnetic bead solution number. The volume is defaulted to 100µL. Enter the Pre-PCR Intermediates -20ºC freezer rack location. Make sure IDX pipeline is selected. Click on “Completed Transfer to 96-well ABGene”.

11.4. Proceed to the next step or store the plate at -20°C. This is a safe stopping point.

12. dUTP Strand Digestion Reaction with UNG enzyme

Note: dUTP Strand Digestion and PCR reaction MUST be performed on the same day.

12.1. Retrieve UNG (Uracil-N-Glycosylase) enzyme from -20°C freezer. Thaw it on ice.

12.2. When processing a full plate, on ice, using Gilson pipetman, aliquot 48µL of the UNG

enzyme into one row (12 wells) of AB-1000 plate. Cover the plate with plastic seal and spin it down at 4°C, 2000g, for 1min. Keep the plate on ice.

12.3. If processing 3 rows of samples or less, aliquot the appropriate volume of UNG enzyme

plus dead volume into one row of AB1000 plate, based on the number of rows:





12.3.1. For one row: 10µL per well 12.3.2. For two rows: (5µL*2 rows) + 5µL dead volume per well = 15µL/well. 12.3.3. Seal the plate with plastic seal and spin it down at 4°C, 2000g, for 1min. Keep it on

ice.

12.4. Using Manual 12-channel Rainin P2-20 pipette, add 5uL of the UNG enzyme to each row of previously adaptered and Ampure XP bead cleaned DNA. Mix 4 times after each addition. Change tips between rows and place a check mark on the side of the plate after each row addition to make sure no rows are skipped or repeated.

12.5. If processing 3 rows of samples or less, Use Manual 12-channel pipette set to 30uL and

using fresh tips mix each row 10X. While mixing, monitor the volume to make sure that all wells within a row are being mixed. Make sure no rows are skipped. Seal the plate with PCR cover and spin down at 4°C, 2000g, for 1min.

12.6. If processing full plates, use Biomek FX to mix dUTP strand digestion reaction: 12.6.1. Under Project “FG Indexing”, open program:

“Sample_Mix_P20B_or_P165B_NewTips_AB1000_4PlatesMax.” 12.6.2. Click on the green arrow “RUN” button. 12.6.3. Use P165 Beckman FX barrier tips. Select 10 mixes, with 30µL volume. Make

sure that your placement of plates and tip boxes on the Biomek deck matches the software deck layout on the computer screen.

12.6.4. Make sure to remove all of the covers, both from tip boxes and plates. Click OK to start the program.

12.6.5. After the program is completed, seal the plates very well with PCR cover and quick spin in an Eppendorf centrifuge on Program 2 (2000g at 4ºC for 1 minute). Discard tips and recycle the boxes.

12.7. Use a rubber pad on top of the reaction plate. Run PCR program “UNG”:

37°C for 30 minutes 95°C for 15min 4°C forever

12.8. LIMS: Retrieve the plate set number. From the Protocol drop-down menu, select “Index_Uracil_N_Glycosylase_Digestion” and click “Continue Prep”. Scan in UNG enzyme sol number. Enter -20ºC freezer rack location. Click on “Completed step”.





12.9. After the PCR is completed, spin the plate at 4°C, 2000g or 1min then proceed to the next step. PCR amplification should be performed on the same day.

13. Ampure Magnetic Bead Clean Up after dUTP strand digestion.



floor Biomek.


13.2. To clean up DNA after dUTP strand digestion:






13.3. LIMS: Retrieve the plate set number. From the Protocol drop-down menu, select “Index_Cleanup_UN_Glycosylase_Rxn” and click “Continue Prep”. Scan the Ampure magnetic bead solution number. The volume is defaulted to 100µL. Enter the Pre-PCR Intermediates -20ºC freezer rack location. Make sure IDX pipeline is selected. Click on “Completed Transfer to 96-well ABGene”.

13.4. Proceed to the next step. PCR amplification should be performed on the same day.

14. PCR amplification

Note: PCR Brew must be made in the PCR Clean Room laminar flowhood on the 5th

floor,

room 510.

14.1. Retrieve and thaw at room temperature: 5X Phusion HF Buffer, 10mM dNTPs, DMSO, PE PCR primer 1.0 (one aliquot of 250uL per plate), and Indexing primer plate. Once thawed, immediately place all reagents on ice. Leave enzymes in the freezer until you ready to add them to the brew.





Note: Thaw PE PCR primer 1.0 in the Tissue Culture Room laminar flowhood on the 5th

floor, room 511.

14.1.1. Make sure that the Indexing primer plate contains enough volume for the amount of

plates you are processing: 14.1.1.1. For one plate the minimum Index primer plate volume is 10µL/well. 14.1.1.2. For 2 plates the minimum Index primer volume is 15µL/well 14.1.1.3. For 3 plates the minimum Index primer plate volume is 20µL/well.

14.1.2. Once thawed, mark off the Indexing primer plate to keep track of freeze-thaw

cycles. Spin the plate at 4°C, 2000g, for 1min and keep it on ice.

Note: For strand specific library construction the Indexing primer plate can only be

used for a maximum of 3 times. Mark off each time the plate is thawed even if

it is not used. Once the plate is marked off 3 times, after use, it should be

discarded and it should not be placed back into the freezer.

14.2. Reaction set up using LIMS: 14.2.1. Use a handheld scanner. Enter your user name and password, select sequence

database, Lib Construction department, 6th floor printers (GE), and click “log in” button.

14.2.2. Click on the “Solutions” (flask) icon. Under Mix Standard Solutions select “LibConst_IndexingPCR_Brew”.

14.2.3. If processing a full plate or multiple full plates, enter the number of plates and 96 for number of samples. Click “Mix Standard Solution”.

14.2.4. If processing a partial plate:

14.2.4.1. Enter the number of samples in the first box, and “1” in the second box. Click “Mix Standard Solution”.

14.2.4.2. For partial plates, under “Parameters”, enter the correct number of wells and change the “# of BrewSourceWells” to 12. Click “Re-calculate Standard Solution” button

14.2.5. Under “Mixture”, scan in the corresponding solution barcodes. 14.2.6. Select Type: Reagent, select Grp: Lib Construction, and Barcode Label: 1D Large

Solution/Box/Kit Labels. Click “Save Standard Mixture”.






14.3. If LIMS is down, set up the PCR brew using Excel spreadsheet:

“LibConst_IndexingPCR_Brew” located in R:\Functional_Genomics\Library_Core\Library Production\Plate based library construction\LIMS Backup Brew Calculators

14.3.1. If processing full plates, enter the correct number of plates. 14.3.2. If processing a partial plate enter the correct number of wells and change the “# of

BrewSourceWells” to 12.

14.4. Prepare PCR brew in 15mL tube according to printed calculator label (or if LIMS is down according the excel sheet), checking off reagents using a blue pen as they are added. Mix each reagent before addition to the brew. Mix the brew very well by gentle repeated pulse-vortexing. Store the brew on ice.

14.5. When processing a full plate or multiple plates, on ice, use a Gilson Distriman to dispense:

14.5.1. For one plate: 32µL/well of PCR brew into one AB1000 plate (27µL+5µL dead

volume). 14.5.2. For two plates: 59µL/well of PCR brew into one AB1000 plate (27µL*2 +5µL

dead volume). 14.5.3. For 3 plates: 86µL/well of PCR brew into one AB1000 plate (27µL * 3 plates +

5µL dead volume). 14.5.4. Cover brew source plate with plastic seal and quick spin 1min (4°C, 2000g). Keep

the plate on ice. 14.5.5. Proceed to step 13.7 for PCR set up using Biomek FX.

14.6. When processing 3 rows of samples or less:

14.6.1. On ice, use a Gilson Distiman to dispense 27µL the the PCR brew into the

appropriate rows of AB1000 plate. 14.6.2. Cover the plate with plastic seal and quick spin both the plate and Indexing Primer

plate for 1min (4°C, 2000g). Keep plates on ice. 14.6.3. On ice, using Rainin P2-20 Manual 12-channel pipette, add 4µL of the

corresponding Indexing primers into PCR brew well. Mix each row 4 times. Use fresh tips for each row. Mark each row after addition to make sure that no rows are skipped or repeated.

14.6.4. On ice, using Rainin P2-20 Manual multichannel, add 19µL of the DNA into each corresponding PCR reaction well. Mix each row 15 times. While mixing, monitor





the volume in each tip to make sure that all wells within a row are mixed. Use fresh tips for each row. Mark each row after addition to make sure that no rows are repeated or skipped.

14.6.5. Seal the plate well with PCR cover and quick spin at 4°C (2000g, 1min). 14.6.6. Proceed to PCR cycling step 13.8.

14.7. When processing full plates;

14.7.1. Log into Biomek FX-5. Under Project “FG Indexing”, open protocol: “Lib_Construction_iPCR_Setup.” Click green “Run” button and follow the displayed deck layout for setup.

14.7.2. Enter the number of DNA plates being processed. Make sure to enter the correct

number of plates as the volume of Index primers added to the PCR brew plate depends on the number of plates.

14.7.3. Confirm that “Y” is selected to add 96 Indexed Primers to PCR brew source plate. 14.7.4. First, you will be shown the deck layout for adding Indexing primers to PCR brew

mix (see below).

Figure 6. Deck layout to add 96 Index primers to PCR brew and mix.

Protocol: “Lib Construction _iPCR_Setup.”

14.7.5. Use Beckman P20 and P165 Barrier tips. Mark the front of P165 tip box to

maintain the same orientation in the next step. 14.7.6. Spin down both the Indexing primer plate and PCR source brew plate at 4°C for

1min, 2000g. Place them on the Biomek FX deck in their appropriate locations. 14.7.7. Make sure that all of the covers are off from tip boxes and plates. Confirm that all

of the items are placed in their correct locations based on the deck layout displayed on the computer screen and click “Ok”.





14.7.8. The program, based on the number of plates, will add and mix Indexing primers with the PCR source brew.

14.7.9. After the program stops, cover the PCR Brew plate with plastic seal and spin it down at 4°C, 2000g, for 1min.

14.7.10. Confirm that you have marked the Indexing primer plate for the number of thaw cycles:

14.7.10.1. If the Indexing primer plate has been thawed less than 3 times, seal it with

foil tape and place it back into its designated -20°C storage location. 14.7.10.2. If it was already thawed 3 times discard the plate.

14.7.11. Remove and discard P20 barrier tips. 14.7.12. The next step is to aliquot PCR Brew + Indexing Primers into empty PCR Plates

(see below). Follow the deck layout as displayed on the computer screen. Leave the P165Barrier tips from previous step on the Tip loader ALP (make that the same box orientation is maintained). “Dest” are empty AB1000 plates.

14.7.13. Make sure that all items are placed in their correct location and that that all of the covers are removed. Click “ok” to proceed.

Figure 7. Aliquot combined PCR Brew mix and Indexing primers into empty PCR plates.

14.7.14. The last step is to add the DNA template to PCR reaction plates. 14.7.15. Follow the deck layout. Use P20 Beckman FX barrier tips. “Source” is the DNA

template and “Dest” are the aliquoted PCR brew plates.





Figure 8. Deck layout for adding 19uL of DNA template to aliquoted PCR plates containing brew & Indexing

primers

14.7.16. After the program is completed cover all Dest plates with MicroAmp clear

adhesive PCR seal tape. Discard tips and recycle the boxes.

14.7.17. Seal Source DNA Plates and store in -20°C. Make sure to scan them in LIMS to their appropriate RAC barcode.

14.8. Quick spin PCR reaction plates using Program 2 (2000g at 4ºC for 1 minute).

14.9. Over amplification of Adaptered cDNA results in smaller than expected final average

library size. The desired region of 280-400bp is shown below:

Figure 9. Desired region 280-400bp





In this example, 33 ng of cDNA sample entered Adapter Ligation. Blue trace represents product of 10 PCR cycles using half of the template for PCR. Red trace represents product of 13 PCR cycles using the other half of the template for PCR.

14.10. There are 2 PCR program options on the thermocycler for DNA amplification depending

on the amount of cDNA entering PE Adapter Ligation:

14.10.1. For samples with 10ng or less entering Ligation: 13 cycles of PCR is needed 14.10.2. For samples with 10ng-40ng entering Ligation: 10 PCR cycles is sufficient (this

represents the majority of cases) 14.10.3. For samples above 40 ng entering Ligation less than 10 PCR cycles is

recommended (7 PCR cycles should be sufficient).

Please confirm with your Supervisor which PCR cycle to use for the plates that you

are working on.

Run PCR program TSPET10 on the thermocycler for DNA amplification. Use a rubber pad on top of the reaction plate.

TSPET10 PCR parameters:

1. 98˚C 1 min 2. 98˚C 15 sec 3. 65˚C 30 sec 4. 72˚C 30 sec

Go to step 2, 9 more times. 72˚C 5min

5. 4˚C ∞ Run PCR program TSPET13 on the thermocycler for DNA amplification. Use a rubber pad on top of the reaction plate

TSPET10 PCR parameters:

1. 98˚C 1 min 2. 98˚C 15 sec 3. 65˚C 30 sec 4. 72˚C 30 sec

Go to step 2, 12 more times. 5. 72˚C 5min 6. 4˚C ∞





14.11. When the PCR program is completed, remove reaction plates from the tetrad and quick

spin in an Eppendorf centrifuge on Program 2 (2000g at 4ºC for 1 minute).

14.12. LIMS: Retrieve the plate set number. From the Protocol drop-down menu, select “Index_PCR_Indexing” and click “Continue Prep”. Scan the PCR brew solution number and enter 31uL for brew volume. Enter the PPGP -20ºC freezer rack location. Make sure IDX pipeline is selected. Click on “Completed Aliquot to 96-well ABGene”. Record the new plate set number and retrieve the barcode from the barcode printer. Make sure to scan in the Index Primer plate (25µM) solution number. The volume is defaulted to 2µL so change it to 4uL. Enter the PPGP -20ºC freezer rack location. Click on “Completed Scan Indexing Primer Plate”.

15. Caliper QC of Indexed Libraries

15.1 The PCR plate will be QC’d using the Caliper. Please refer to LIBPR.0051 Operation

and Maintenance of the Caliper Labchip Gx for DNA samples using the High Sensitivity

Assay. 2µL of the PCR product will be used for Caliper QC. The profiles should show the majority of the PCR product to be between 280-400bp. If the product is smaller it means too many PCR cycles were performed. An example of the expected PCR profile is shown below:

15.2 Show the Caliper QC profiles to your supervisor.





15.3 LIMS: Retrieve the plate set number. From the Protocol drop-down menu, select “Index_QC_ PCR” and click “Continue Prep”. Click on “Completed QC Indexed PCR DNA”. Enter the Caliper Run ID. Click on “Completed Enter Results”.

16. Picogreen Quant of PCR product (optional)

16.1. Consult with your supervisor if PCR product needs to be quantified before size selection

(it may be necessary before Baraccuda size selection).

16.2. Quant PCR product following the protocol: LIBPR.0048 – 96-well DNA Quantification

using PicoGreen and VICTOR3V.

16.3. Calculate total ng for every well.

16.4. LIMS: Retrieve plate set. From the Protocol dropdown menu, select “96well_PicoGreen_Quant”. Enter the storage location and the file name in the Picogreen_PCR product attribute field.

17. Manual Size selection of PCR product

17.1. Manual size selection should be performed using no more than 12 gels per round. The

samples which are going to be pooled into one sequencing line can be size selected on the same gel. Currently, this means 2 samples per gel. Ask your supervisor for size

selection instructions.

17.2. Cut open the Novex 8% TBE gel cassette pouch to remove the gel cassette, and drain

away the gel packaging buffer. Handle the gel cassette by the edges only. Rinse the gel cassette with deionized water.

17.3. Peel off the tape covering the slot on the back of the gel cassette. In one fluid motion, pull the comb out of the cassette.

17.4. Use a 1mL pipette to gently wash the cassette wells with 1X TBE gel running buffer. Repeat twice, and then fill the sample wells with running buffer.





It is important to wash wells thoroughly to remove preservative residue from wells

that may impede sample running efficiently through gel.

17.5. Assemble the gel apparatus as follows:

17.5.1. Lower the Buffer Core into the Lower Buffer Chamber so that the negative electrode fits into the opening in the gold plate on the Lower Buffer Chamber as shown in the figure 10.

Figure 10. Lower Buffer Chamber and Buffer Core

17.5.2. Insert the Gel Tension Wedge into the XCell SureLock cell behind the buffer core.

Make sure the Gel Tension Wedge is in its unlocked position, allowing the wedge to slip easily into the XCell SureLock unit.

17.5.3. Insert the gel cassette into the lower buffer chamber in front of the core, with the well side of the cassette facing in towards the buffer core. The slot on the back of the cassette must face out towards the lower buffer chamber. Place the Buffer Dam behind the core.





Figure 11. Buffer Chamber Side View

17.5.4. Pull forward on the Gel Tension Lever in a direction towards the front of the XCell

SureLock unit until lever comes to a firm stop and the gel/buffer dam appear snug against the buffer core.

Figure 12. Unlock and Locked Position

17.5.5. Fill the Upper Buffer Chamber with 200mL of the 1X TBE running buffer. Ensure

that the Upper Buffer Chamber is not leaking. If the level of the running buffer drops, the apparatus will need to be reassembled.

17.5.6. Fill the Lower Buffer Chamber with approximately 200mL of running buffer through the gap between the Gel Tension Wedge and the back of the Lower Buffer Chamber as shown below:





Figure 13. Lower Buffer Chamber

17.5.7. Align the lid on the Buffer Core. The lid can only be firmly seated if the (-)

electrode is aligned over the banana plug on the right.

Caution: Power must be off before connecting the XCell SureLock Mini Cell to the

power supply. 17.6. Prepare your samples by adding appropriate amount of 10X loading dye. 17.7. Before loading the sample, carefully rinse out the wells with 1X TBE running buffer

using a 1mL pipette tip.

17.8. Load 10µL of the 100bp DNA Ladder (20ng/µL) in the middle of the gel. Make sure there is no residual ladder on the outside of the tip when loading ladder into the well.

17.9. Load all of the PCR product for each sample split into two wells. When size selecting

two samples per gel, each sample is loaded on the opposite side of the ladder well. When running more than one sample on the same gel, make sure to lead them asymmetrically so that even if the gel is flipped during staining you will be able to recognize which sample corresponds to which well. Leave at least one well empty between the ladder and samples and never load samples or the ladder in the corner wells.

17.9.1. Be aware that some samples have a tendency to float out of the well, so as a test,

load a small volume to ensure that the sample doesn’t float out. If floating does

occur add 1µL of glycerol and re-test.





17.10. Immediately after loading the wells, run the gel @ 200 V for 45-50 minutes. The run is

completed when the first dye marker is out of the gel and the second dye marker is still visible toward the bottom of the gel (just below the 3rd vertical line of the cassette).

17.11. Using colored tape (same color as tape attached to the gel plate) attach a label to the gel

apparatus which states library names and gel location (left of ladder vs. right of ladder) start time, finish time, date, and your initials.

18. Gel Scan and Cutting DNA Fraction from PAGE Gel

18.1. Put on a clean pair of gloves. Pre-chill a centrifuge to 4°C. Retrieve fresh ice and all reagents.

18.2. Set up sets of 0.5mL and 2mL tubes for shearing the gel slices: Make a hole through the bottom of 0.5mL tubes with an 18 gauge needle. Place each 0.5mL tube into a 2mL tube.

18.3. Label each 2mL tube on the side with the tra#, well position, date, and initials.

18.4. Cover the Dark Reader screen with a fresh sheet of plastic wrap. Wrap the right-angle

ruler with plastic wrap.

18.5. Prepare fresh stain: 6µL SYBR Green in 60mL 1X TBE. Minimize exposure to light.

18.6. Stop the gel run after about 45-50 minutes. The first dye marker should be out of the gel and the second dye marker should still be visible toward the bottom of the gel. Dismantle the PAGE apparatus.

18.7. For completed runs, remove the gel cassette and transfer the corresponding label (with

well notation) from the gel apparatus.

18.8. Using a post-PCR dedicated tray, processing 4 gels at a time, stain the gel (each in a separate tray) for 3 minutes. Set the timer.

18.9. After 3min, retrieve the gel from the staining solution and place onto the Mylar sheet.

Also transfer the corresponding tape label.

18.10. Log onto the computer and scan image on high sensitivity setting. Save the image in the appropriate network directory and file folder. Name the file with tra#, plate well locations of the two samples_pipeline_PCR_ DateInitials: Tra26984_A11_A12_Strand Specific_PCR_070315mm.gel





18.11. Confirm that the gel was not flipped over during staining (samples should have been

loaded asymmetrically). If it was, open the scanner tray, correct the gel’s orientation, and re-save the file under the same name.

18.12. Print the image and affix it into your lab notebook.

18.13. Lay the Mylar sheet with the gel on top onto the Dark Reader. For every sample cut out

280-400bp size fraction (see image below). The upper cut is slightly above 400bp ladder mark; the bottom cut is about ¼ down from the 300bp (see where the midpoint is between 200bp and 300bp ladder and go half that distance up for the lower cut). Do not cut below 280bp as we want to avoid the possibility of the concatenated 260bp adapter product.

Figure 14. 280-400bp size fraction

280bp

400bp





18.14. Transfer gel fractions for the same sample into one of the 0.5mL tubes prepared and

labeled earlier for shearing. Make sure that the labeling on the tube matches the labeling of the size selected sample.

18.15. Discard stain, spray with water, and wipe down tray. Discard Mylar sheet and remainder

of the gel. Discard used blade in sharps container. Tidy up area. Change gloves.

18.16. Spin tubes at 12,000rpm at room temperature for 5 minutes. The gel slices should shear through the holes and collect into the bottom of the 2mL tubes.

18.17. After shearing the gel fractions, check that all of the gel has cleared the 0.5mL tubes.

Spin it again, if needed. If no gel remains, discard the 0.5mL tubes and add 350 µL of Elution buffer (5:1, LoTE:7.5M Ammonium Acetate) to each gel slurry. Mix well by vortexing. Pulse spin.

18.18. If time permits, incubate for 1 hour at 65°C to elute DNA and proceed to step 18 for Gel extraction.

18.19. If there is insufficient time to continue, incubate overnight at 4°C.

18.20. Clean PAGE apparatus: Run tap water over PAGE apparatus for 2 minutes; wipe down

with 2% micro90; run water over PAGE apparatus for another 2 minutes. Wipe down the PAGE workstation.

18.21. LIMS: Scan the tra# or plate set number. From the Protocol drop-down menu select “Indexing_Size_Select_plate” and click “Continue Prep”. Enter the PPGP -20ºC freezer rack location. Retrieve new plate barcode from the printer.

19. Gel Extraction and Precipitation

19.1. Gel extractions are to be performed 24 samples per technician per round.

19.2. Retrieve the gel slurries from 4°C. Vortex and pulse spin.

19.3. Incubate at 65°C for 15 minutes. Pulse spin.

19.4. Label Spin-X Filter Tubes with tra#, well position, date, and initials.





19.5. Transfer the contents of each 2mL tube into the corresponding Spin-X Filter Tube. Tap

the slurry into the Spin Filter or use a new disposable spatula to aid transfer. Change gloves.

19.6. Spin at 12,000rpm at 4°C for 5 minutes.

19.7. Check each Spin-X Filter Tube and ensure that the entire 350µL of buffer has spun through the filter. Re-spin the tubes if there is still liquid trapped in the gel material.

19.8. Remove and discard the filter column containing the gel material.

19.9. Transfer the eluate to a single sterile 1.5 mL tube (make sure not to use non-stick tubes) and add the following:

Reagent Volume

Eluate 350 µL

3M Sodium Acetate (1/10 volume) 35 µL

Mussel Glycogen (20 mg/mL) 3 µL

100 % Ethanol (2.5x volume) 970 µL

TOTAL VOLUME 1358µµµµL

19.10. Vortex and pulse spin.

19.11. Chill the tube at -20°C for a minimum of 30 minutes.

19.12. Spin at 14,000rpm at 4°C for 45 minutes. Dispose all waste and partially used reagents

aliquots.

19.13. Carefully decant the supernatant into clean microcentrifuge tube. Keep an eye on the pellet so that it doesn’t slide out.

19.14. Wash the pellet with 1 mL cold 70% ethanol. Spin at 14,000 rpm at 4°C for 3 minutes.

Carefully decant the supernatant into a new microcentrifuge tube as a backup.

19.15. Dab the tube rims on a Kimwipe to remove ethanol. Pulse-spin the tubes and using P10 pipetman carefully remove any residual ethanol.

19.16. Mark the outside bottom of the tube to better locate the pellets.





19.17. Air-dry the pellet at room temperature until translucent. Do not over-dry the pellet.

19.18. Resuspend the pellet in a total volume of 12µL of Qiagen Elution Buffer (EB).

19.19. LIMS Retrieve the plate set number. From the Protocol drop-down menu select

“Index_Gel_ Extraction” and click “Continue Prep”. Enter the PPGP -20ºC freezer rack location. Click on “Completed Aliquot to 1.5mL Tube”. Record the new plate set number in your lab notebook and retrieve the 2D barcode from the barcode

20. QC of Gel Purified product and dilutions to 8nM.

20.1. Use 1µL of each sample for Agilent DNA 1000 Series II assay. Refer to protocol LIBPR.0017 – Operation and Maintenance of the Agilent 2100 Bioanalyzer for DNA samples.

20.2. Use 1µL of each sample for Qubit quant. To minimize photo-bleaching perform Qubit

assay in sets of no more than 24.

20.3. Discuss the Agilent and Qubit results with your supervisor before the libraries are submitted. The expected PPGP profile is shown below. There should be no product visible below 250bp. The minimum submission criteria is 10uL of 8nM library with majority of the product centered at 320-330bp.





Figure 15. expected PPGP profile is shown below

20.4. Based on the Agilent profile, determine the average bp size of each library. Use that

value and the corresponding Qubit concentration in ng/uL to determine nM (molar concentration).

20.5. Dilute each library to a concentration of 8nM in buffer EB supplemented with 0.1%

Tween-20.

20.6. After dilution, confirm the concentration using Qubit. To minimize photo-bleaching Qubit samples in sets of no more than 24. Re-calculate the nM concentration for each sample.

20.7. LIMS: Retrieve the plate set number. From the Protocol drop-down menu select

“Index_QC_Gel_Purified_DNA” and click “Continue Prep”. Enter the Agilent Run ID and the Qubit Run ID. Click on “Completed Qubit & Agilent Gel Purified DNA”.





21. Pooling of Strand Specific Libraries for Sequencing

21.1. Ask your supervisor for pooling instructions. Currently, we are pooling two strand

specific libraries per one line. This means 48 pools from each plate. Ask your supervisor for IX library IDs (one for each pool to be made).

21.1.1. Log into alDente and click on the “Rearrays” icon. 21.1.2. Under Rearray Utilities, select “Tube” from the drop down menu and click

“Manually Set Up ReArray”. 21.1.3. Enter the tra# of the source plate (if pooling from a plate) or the pla#s of tubes to be

pooled. Select “Tube” for the Target Plate Size. Click on “Pool Wells”. (Note: Pooling does not empty source plate. Aliquots are taken but volumes are not tracked.)

21.1.4. In the Library field enter the IX library number. In the Plate Format drop-down menu choose “1.5mL tube”. In the Location field, enter the PPGP -20ºC freezer rack location. In the Pipeline drop-down menu, make sure to pick IPE: SLX-

IPET. 21.1.5. Click on the listed source plate number to open the plate well-map. Click on the

desired wells to pool then click inside the empty Target Plate field on the right to create a re-arrayed wells list.





Figure 16, Create Pool Wells Re-array

21.1.6. Double check that you have selected everything correctly then click “Create Pool

Wells Re-array” red button.

21.1.7. Repeat the steps for every pool, each time selecting the right wells for the assigned INX number.

21.1.8. Retrieve 2D tube barcode labels from the barcode printer.

21.2. Place IX barcodes on sterile 1.5mL tubes and following exactly what was recorded in

LIMS pool equal molar amount of each library scheduled for that pool (if both samples have the same molar concentration – pool equal volume). If you received excel sheet with exact pooling instruction refer to that document. Currently, this will be 48 pools/plate; two libraries in each pool.





22. QC Quant Final Product and Sequencing Submission

22.1. After pooling, Qubit quant 1µL of each IX tube to confirm the molar concentration for submission. To minimize photo-bleaching perform Qubit assay 24 samples at a time. Refer to protocol LIBPR.0030 – Illumina Concentration Checked.

22.2. LIMS: From the Protocol drop-down menu select “Index_QC_Qubit_8nM_Final

Product” and click “Continue Prep”. Enter the rack location of the Next PET Run Box with space available. Enter the library concentration (nM) and enter the Qubit Run ID. Make sure the pipeline IPE: SLX-IPET is selected. Click on “Completed Aliquot to 1.5mL Tube”. This protocol places samples in the ‘SLX -Libraries Ready for Sequencing’ view in LIMS for flowcell scheduling.

22.3. Record the plate set number in your lab notebook and retrieve the 2D barcodes from the barcode printer. Based on Qubit, calculate nM concentration. Record the concentration (nM) of each pool on the corresponding newly printed barcode.

22.4. Replace previous barcodes for each pool with these new submission barcodes.

22.5. Scan and place the tubes in the appropriate Next PET Run boxes.

Date post:	29-Mar-2020
Category:	Documents
Upload:	others
View:	1 times
Download:	0 times