For detailed instructions about the TruSeq Custom Amplicon library preparation methods, refer to your reference guide.

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1 1 TruSeq Custom Amplicon Script Welcome Navigation Objectives Welcome to the TruSeq Custom Amplicon course. Click next to begin. Take a moment to familiarize yourself with the navigation for this course. You can access these instructions at any time by pressing the help button in the top right corner of the page. By the end of this course you will be able to: Describe the items necessary to perform the TruSeq Custom Amplicon assay. Describe the steps in workflow and Identify best practices For detailed instructions about the TruSeq Custom Amplicon library preparation methods, refer to your reference guide. Characteristics Workflow User-supplied Equipment TruSeq Custom amplicon is a library preparation assay made by Illumina with the following characteristics: Targeted resequencing of amplicons per sample. Number of samples can be adjusted to match the desired depth of coverage. Each targeted region is amplified in each sample. Assay takes 6 to 8 hours to complete (2-2.5 hours hands-on-time). Designed for 96-samples per run (95 samples + 1 control). The TruSeq Custom Amplicon workflow is summarized in seven steps. Click each step to learn more. To get started, designate an area of your lab as pre-pcr. A separate room is not necessary as long as pre- and post-pcr equipment is kept separate and a proper cleaning schedule is maintained. A separate plate centrifuge for pre- and post-pcr areas is required. Each centrifuge should reach 2400 g s. An oven or 37 degree incubator for plate assembly is required. Do not use a PCR machine for this purpose. It is preferred to place the oven in the pre-pcr area. A plate incubator or heat block in the pre-pcr area is required. A plate shaker. A 96-sample magnetic stand (validated by Illumina is suggested). Thermal cycler, and AMPure XP beads (that are user supplied) are required in the post-pcr area. Division of preand post-pcr Areas Items to Order from Illumina Pre- and post-pcr areas are ideally in two separate rooms. If two separate rooms is not an option, separate benches or areas are acceptable if: Separate equipment is used, and a Strict cleaning schedule is maintained. Three different components make the TruSeq Custom Amplicon Sequencing libraries. Click each component for more information.

2 2 What is Manifest File? The Manifest File is created after the CAT order is placed in DesignStudio. This file is used in Illumina data analysis to align amplicon sequences It contains a list of all probe and target sequences. It also includes possible off-target sequences. The manifest file is downloaded via MyIllumina after your order has shipped. In addition, download the manifest for the CAT order and the control CAT that is included in the library preparation kit. Library Preparation Workflow The TruSeq Custom Amplicon Sequencing Library Preparation process begins with high-quality DNA, as measured by a fluorescent assay. The CAT or Custom Amplicon oligo is added to each sample. The sample is heated and gradually cooled, and each Downstream Locus-Specific Oligo (DLSO) and Upstream Locus-Specific Oligo (ULSO) hybridizes to its target region. Next an extension-ligation reaction takes place. This reaction completes the creation of your amplicon template. The amplicon is purified and amplified with PCR. The i7 and i5 PCR primers are complementary to overhangs on the ULSO and DLSO, respectively. Each primer has a unique index that is sequenced. These indexes provide each sample with a unique identifier that allows for multiplexing on the sequencer. Assay workflow Illustrated is the workflow for the TruSeq Custom Amplicon Library Preparation Kit. Safe stopping points are marked between steps. Click each step in the workflow for more information. Hybridization of the Oligo Pool: Hybridization of the Oligo Pool During this step, a custom pool containing upstream and down stream oligos specific to your targeted regions of interest are hybridized to your genomic DNA samples. Hybridization of the Oligo Pool: When sealing the plates with foil, use a soft rubber roller. Apply the roller in different directions horizontally, vertically and diagonally to seal every well evenly. Use your thumbs to press the seal down around the edge of each well. When the plate is sealed properly, you can see circular indentation around each well. Proper sealing prevents evaporation during hybridization of the CAT oligo pool.

3 3 Removal of Unbound Oligos: Removal of Unbound Oligos In this step, unbound oligos are washed away using a filter capable of size selection. A filter plate assembly is createdand used in subsequent steps. Samples are added to a pre-washed filter plate. Removal of Unbound Oligos: To assemble the Filter Plate unit, first use a midi plate, upon which you place the metal collar. The metal collar can be reused but clean thoroughly with bleach between uses. Next, place the filter plate on the metal ring, and make sure that it is in place. Finally, the lid is placed on the filter plate. Extension Ligation of Bound Oligos In this step, a DNA polymerase extends from the hybridized ULSO, and a DNA ligase subsequently ligates this fragment to the 5' end of the DLSO. The Extension-Ligation Mix (ELM) is added directly onto the filter. Incubate the filter assembly at 37 C for 45 minutes. PCR Amplification PCR Amplification In this step, the extension-ligation products are PCR amplified using primers that add index sequences for sample multiplexing (i5 and i7), as well as common adapters required for cluster generation (P5 and P7). PCR Amplification In this video, we demonstrate how to set up a PCR index primer plate. After thawing the index primers, vortex each index primer briefly. Next, spin the tubes containing PCR primers. Doing so reduces the risk of contamination when uncapping the tubes. Use 1.5 ml Eppendorf tubes as adapters for centrifugation. Remove the tubes from the centrifuge; arrange them in the index primer fixture in the same order as the plate layout set up in Illumina Experiment Manager. Remove the white caps from the i5 index primers and discard the caps. To reduce the risk of index cross-contamination, do no reuse the caps. Use a multichannel pipette and fine tips to pipette the clear i5 primers into each column of the plate. Before each dispensing, inspect the tips to ensure all channels are functioning properly, with approximately the same volume aspirated. It is not necessary to change tips until all columns are finished. Apply fresh white caps to the i5 index primer tubes. Pick up each cap carefully to avoid contamination. Next, remove the orange caps from the i7 index primers and discard the caps. Do not reuse the caps. Use a multichannel pipette and fine tips to pipette the yellow i7 primers into each row of the plate. Before each dispensing, inspect the tips to ensure all channels are functional properly, with approximately the same volume aspirated. Change tips after each column. The yellow dye allows you to track progress easily. Apply fresh orange caps to the i7 index primer tubes. Pick up each cap carefully to avoid contamination.

4 4 PCR Clean-Up: PCR Clean-Up In this step, AMPure XP beads are used to purify the PCR generated libraries from the other reaction components. PCR Clean-Up: In this step, AMPure XP beads are used to purify the PCR-amplified libraries from the other reaction components. We demonstrate only a few critical steps here. Refer to the User Guide for the complete protocol. Use a full pipette tip box to help you track the current column. When transferring PCR samples from the IAP plate to the CLP plate containing AMPure beads, use a multichannel pipette. Always check the tips to make sure that they are securely attached to the pipette before each column transfer. Change tips after each column. After the incubation of the CLP plate on the magnet for two minutes or until the supernatant has cleared, pay attention to the position of each pellet. If you use the magnet plate as recommended in the User Guide, the beads in columns 1, 3, 5, 7, 9 and 11 are pelleted to the right. The beads in columns 2, 4, 6, 8, 10 and 12 are pelleted to the left. Knowing where the bead pellets are will make it easier to remove the supernatant without disturbing the bead pellets in the next step. Insert pipette tips go in at an angle, point away from where the bead pellets are located. Discard the supernatant and tips after each column. After the entire plate is finished, hold up the plate and inspect whether the supernatant in every well has been removed. The pipette technique introduced in this video is also applicable to the library normalization step, although a different type of beads is used in the other procedure. Library Normalization: Library Normalization This process normalizes the quantity of each library to ensure more equal library representation in your pooled sample. Library Normalization: This process normalizes the quantity of each library to ensure more equal library representation in your pooled sample. If performed correctly, this procedure produces cluster density without using qpcr or Bioanalyzer for library quantitation. Successful library normalization requires complete resuspension of the LNB1 normalizer beads, precise pipette technique, and the use of recommended equipment as listed in the User Guide whenever possible. Bring both LNA1 and LNB1 to room temperature before use. LNA1 contains formamide. Invert and inspect the LNA1 reagent tube and verify that any precipitates have dissolved completely. Next, vortex the LNB1 bead reagent vigorous with intermittent inversion. Flick the tube with the tube inverted to make sure that beads are dislodged from the bottom. Use a P1000 pipette set to 1000 to pipette the beads up and down 20 times. Even if you plan to use only partial tube, use a P1000 pipette, set to 1000 to resuspend the beads. Next, transfer 4.4 ml of LNA1 to a fresh 15 ml conical tube. Use a P1000 pipette set to 800 microliters to pipette the beads up and down a few more times and transfer 800 µl of LNB1 to the conical tube. You do not need to stick the pipette into the conical tube. Next, invert the LNA1/LNB1 mixture 20 times to mix the reagents, and pour it into a trough. Use the resulting bead mix immediately before the beads start to settle. LNA1 and LNB1 cannot be stored together or performance of the beads are severely compromised.

5 5 Library Pooling: Library Pooling In preparation for cluster generation and sequencing, equal volumes of each normalized library are combined and later diluted in Hybridization Buffer. Library Pooling: To create a sample pool to load onto the sequencer, first transfer 5 microliters of each sample that is pooled from the SGP plate into a PCR tubestrip. Always check the tips to make sure that all channels are functioning and transfer approximately the same volume. Make sure that all samples are at the bottom of the tube. Combine and transfer the entire content of each well into a tube labeled PAL. Make sure that there is no sample left behind. Spin the PCR tube strip, if necessary. Mix the PAL thoroughly, by pipetting or by vortexing. Follow the reference guide to determine the volume of PAL needed to create a DAL. Best Practices Tips Online Calculator Summary The penultimate step in the library preparation process consists of: Pooling together normalized libraries, or creating the PAL. The number of libraries pooled together depends on the amount of coverage desired. You should now understand the preparation and workflow for the TruSeq Custom Amplicon library preparation assay including:. Setting up your lab Downloading a manifest Preparing for pooling and Preparing libraries Congratulations Congratulations! You have completed this course.