Identification of Functionally-Relevant Lentivirus Integration Sites in an Insertional Mutagenesis Cell Library

Summary

Insertional mutagenesis is an essential tool in forward genetics for identifying functional genomic elements. Here, we describe the Insertion-based Screen for functional Elements and Transcripts (InSET), a method for detecting lentivirus integration sites within a lentivirus-based insertional mutagenesis cell library.

Abstract

The extent of functional sequences within the human genome is a pivotal yet debated topic in biology. Although high-throughput reverse genetic screens have made strides in exploring this, they often limit their scope to known genomic elements and may introduce non-specific effects. This underscores the urgent need for novel functional genomics tools that enable a deeper, unbiased understanding of genome functionality. This protocol introduces the Insertion-based Screen for functional Elements and Transcripts (InSET), a method for identifying lentivirus integration sites within a lentivirus-based insertional mutagenesis cell library. InSET facilitates the capture of genome-wide lentiviral integration sites, with next-generation sequencing used to detect and quantify flanking sequences. InSET's design enables the analysis of integration site abundance variations in phenotypic screens on a large scale, establishing it as a robust tool for forward genetics and for identifying functional genomic elements. A key benefit of InSET is its capacity to reveal previously unidentified genomic elements, including novel functional exons of both protein-coding and non-coding RNAs, independent of prior annotation. Overall, InSET holds significant value in studying the intricate complexity of the human genome and transcriptome, where many genomic elements await functional characterization.

Introduction

Functional genomics remains a crucial and challenging field, even in the post-genome era, with a significant portion of genomic regions still lacking functional characterization. A key reason for these challenges lies in the intricate nature of the human genome and transcriptome, which are characterized by pervasive transcription and universal alternative splicing in both protein-coding and non-coding regions^1,2,3,4,5. This complexity has been extensively highlighted through RNA-targeted enrichment studies^6,7,8,9. It is increasingly recognized that numerous novel genomic elements remain unmapped and functionally unannotated^10,11,12in addition to the well-annotated segments of the human genome.

In recent years, advancements in reverse-genetics tools have been seen, such as RNA interference (RNAi) and CRISPR/Cas systems^{13,14,15,16,17,18,19,20,21}. However, these methods have notable disadvantages, including non-specific effects and high costs associated with generating complex libraries of short hairpin RNAs (shRNAs) or single guide RNAs (sgRNAs)^22,23,24,25. Additionally, these methods predominantly focus on annotated genomic elements. In contrast, forward genetics has the advantage of identifying novel, unannotated functional genomic elements. Given the complexity of the human genome and transcriptome, as well as the vast number of novel genomic elements, this capability of forward genetics is of significant importance.

Forward genetics cell libraries are frequently generated through random retroviral integration, making the efficient detection of genome-wide integration sites crucial for a forward genetics assay. Detecting these integration sites typically involves genome-walking tools²⁶. While there have been significant advancements in these tools, few are optimized for next-generation sequencing (NGS) applications and subsequently applied in insertional mutagenesis studies. Insertional mutagenesis in cell lines often utilizes methods like inverse PCR (I-PCR) or linear amplification-mediated PCR (LAM-PCR) to detect integration sites^27,28,29. These methods, however, usually require restriction enzyme digestion or ligation steps, which can reduce efficiency and introduce biases in the results.

To enhance the efficient detection of genome-wide integration sites in forward genetics screens, a method called the Insertion-based Screen for functional Elements and Transcripts (InSET) method was introduced here. By integrating genome-walking with NGS, InSET enables the high-throughput identification of lentivirus integration sites in lentivirus-based insertional mutagenesis libraries. Compared to existing methods^27,28,29, InSET is relatively straightforward and simple, eliminating the restriction enzyme digestion or ligation steps, making it more convenient and accessible. Furthermore, although the existing forward genetics screen in mammalian cells is limited to very few haploid or near-haploid cell lines^{27,28,29,30,31}, InSET has been proven to work in aneuploid cell lines³².

This study provides a detailed, step-by-step protocol for implementing the InSET method. Additionally, it demonstrates the application of InSET in insertional mutagenesis studies. InSET serves as a powerful tool for identifying novel exons within genomic elements, encompassing both protein-coding and non-coding RNAs in human cell lines.

Protocol

Figure 1 provides a schematic overview of the InSET method. For optimal coverage, prepare five parallel libraries from 1 µg of genomic DNA, then pool them for next-generation sequencing. This procedure has been successfully completed continuously in previous tests, though users may introduce pause points based on their discretion. Detailed information on the reagents and equipment used in this study can be found in the Table of Materials.

1. Capture of lentivirus integration sites in the genome

1. Use the Genomic DNA isolated from lentivirus-based insertional mutagenesis cell library (1 µg), as described in a previous study³², as the template for a linear PCR in a 50 µL reaction containing 1x Taq Buffer, 2.5 U Taq DNA Polymerase, 4 µL of dNTP Mixture (2.5 mM each), and 0.3 µM of biotinylated primer 3-LTR_prime (Table 1) in a PCR tube.
   NOTE: The 3-LTR_prime primer anneals to the LTR region of the lentivirus sequence. If it anneals to the 5' LTR, the amplified fragment includes the flanking genomic sequence. If it anneals to the 3' LTR, the amplified fragment contains an internal lentivirus sequence, which cannot be aligned to the host genome and will, therefore, be filtered out during downstream sequencing data analysis. This primer can also be modified to adapt to other types of vectors used to generate an insertional mutagenesis cell library.
2. Perform 50 cycles of linear PCR in a standard PCR thermocycler, adding 2.5 U of Taq DNA Polymerase at the start and again immediately after 25 cycles. PCR conditions are as follows: 94 °C for 5 min; 50 cycles of 94 °C for 30 s, 55 °C for 30 s and 72 °C for 30 s; 72 °C for 5 min.
   NOTE: To maintain the activity of the Taq DNA Polymerase during prolonged reactions, it is recommended to add the enzyme at both the beginning and the midpoint of the reaction.
3. Transfer the PCR products to a new, clean 1.5 mL centrifuge tube.
4. Mix the PCR products thoroughly with 10 µL of biotin-streptavidin magnetic beads.
5. Incubate the mixture for 2 hours at room temperature, shaking at 400 rpm in a metal bath.
   NOTE: Take out the tube to mix well with the pipette every 15 min.
6. Use a magnetic stand to collect the beads, then carefully remove the supernatant.
7. Wash the beads with 300 µL of Binding/Wash Buffer (10 mM of Tris-HCl (pH 7.5), 1 mM of EDTA, 1 M of NaCl, 0.1% Tween-20) 6 times and discard the supernatant.
8. Wash with 300 µL (65 °C) DNase/RNase-free distilled water twice and discard the supernatant.
9. For the second strand synthesis, resuspend the beads in a 24 µL reaction volume containing 1x Klenow Fragment Buffer, 2 µL of dNTP Mixture (2.5 mM each), and 6.25 µM of P5_N6 primer (Table 1) in a PCR tube.
   NOTE: Prepare a 100 µM stock solution of P5_N6 primer. The P5_N6 primer is composed of a fixed P5 adaptor sequence at the 5' end and a random hexamer sequence at the 3' end. The random hexamer can bind to a diverse range of target sequences, making it suitable for amplifying various DNA fragments generated through linear PCR.
10. Pre-incubate the mixture at 15 °C for 20 min in a PCR thermocycler.
11. Add 2 U of Klenow Fragment (Large Fragment E.coli DNA Polymerase).
12. Incubate the mixtures in a PCR thermocycler at the following conditions: slowly ramp at 0.5 °C/min from 15 °C to 25 °C followed by 30 min incubation; then slowly ramp at 0.5 °C/min to 37 °C followed by 1 h incubation.
13. Transfer the PCR products into a new, clean 1.5 mL centrifuge tube.
14. Use a magnetic stand to capture the beads, then discard the supernatant.
15. Wash the beads gently with 300 µL of 4 °C DNase/RNase-free distilled water. Collect the beads and discard the supernatant.

2. Construction of NGS library with nested PCR

1. Resuspend the beads from step 1.15 in a 25 µL PCR reaction volume containing 1x Taq Buffer, 1.25 U Taq DNA Polymerase, 0.5 µM of primer P5 (Table 1), 2 µL of dNTP Mixture (2.5 mM each), and 0.5 µM of primer 3-LTR_Nest (Table 1) in a PCR tube.
2. Perform PCR reactions in a PCR thermocycler under the following conditions: 94 °C for 3 min; 20 cycles of 94 °C for 30 s, 55 °C for 30 s and 72 °C for 30 s; 72 °C for 7 min.
3. Use 5 µL of the PCR products from step 2.2 as the template for the next round of PCR in 50 µL volume containing 1x Taq Buffer, 2.5 U Taq DNA Polymerase, 4 µL of dNTP Mixture (2.5 mM each), 0.4 µM of primer Illumina_P5 (Table 1) and 0.4 µM of primer Illumina_P7-3LTR (Table 1).
4. Perform PCR reactions in a PCR thermocycler under the following conditions: 94 °C for 3 min; 15 cycles of 94 °C for 30 s, 55 °C for 30 s, and 72 °C for 30 s; 72 °C for 7 min.
5. Transfer the PCR products to a clean 1.5 mL centrifuge tube.
   NOTE: The protocol can be paused at this step, and the samples can be stored at -20 °C.
6. Purify the PCR products using 1.2x volumes of DNA clean beads, bringing the final volume to 21 µL.
   NOTE: Take 10 µL of PCR products from each of the 5 libraries prepared in parallel, pool them together, and purify to a final volume of 21 µL. The purification protocol for the PCR products is adapted from the manufacturer's instructions for the DNA clean beads (see Table of Materials).
   1. Equilibrate the beads from 2-8 °C to room temperature for 30 min.
   2. Mix the beads thoroughly by inverting or vortexing.
   3. Pipette 1.2x volumes of the beads into a clean 1.5 mL centrifuge tube.
   4. Add the PCR products into the beads and mix thoroughly by pipetting up and down 10 times.
   5. Incubate at room temperature for 5 min to allow DNA binding to the beads.
   6. Place the samples on a magnetic stand and remove the supernatant once the solution is clear (about 5 min).
   7. Add 200 µL of 80% ethanol to wash the beads, incubate at room temperature for 30 s, and remove the supernatant.
      NOTE: The 80% ethanol should be prepared freshly.
   8. Repeat step 2.6.7 once.
   9. Air dry the beads at room temperature with the lid open for about 5 min.
   10. Remove the tube from the magnetic stand, add 22 µL of DNase/RNase-free distilled water, mix thoroughly by pipetting up and down 10 times, and let it sit at room temperature for 2 min.
   11. Place the tube on the magnetic stand until the solution is clear (about 2 min), then transfer 21 µL of supernatant to a clean 1.5 mL centrifuge tube.
       NOTE: The protocol can be paused at this step, and the samples can be stored at -20 °C.
7. Use 1 µL of purified PCR products to measure the concentration using the Fluorometer with the commercially available dsDNA fluorescence detection kit (see Table of Materials).
   NOTE: The concentration measurement protocol follows the manufacturer's instructions for the dsDNA detection kit.
   1. Prepare the Working Solution preparation by mixing the Reagent and Buffer (provided in the kit) at a ratio of 1:200.
   2. Prepare the assay standards by mixing 10 µL of Standard #1 and 10 µL of Standard #2 (provided in the kit), each with 190 µL of Working Solution. Gently vortex the mixture for 2-3 s.
   3. Prepare the assay sample by mixing 1 µL of the purified PCR products with 190 µL of Working Solution. Gently vortex the mixture for 2-3 s.
   4. Incubate the assay standards and assay samples for 2 min at room temperature, keeping them protected from light.
   5. Turn on the Fluorometer.
   6. Select the dsDNA High Sensitivity program from the main menu.
   7. For calibration, insert Standard #1 into the machine, press the Read standard button to measure the first standard, then insert Standard #2 and press the Read standard button to measure the second standard.
   8. For sample measurement, press the Run samples button to go to the sample measurement page. Insert each tube containing the assay sample, and press the Read tube button to obtain the concentration.
      NOTE: The protocol can be paused at this step, and the samples can be stored at -20 °C.
8. Perform NGS on Illumina platforms using paired-end 150 bp (PE150) strategy on a 30 giga-base (GB) scale.
   NOTE: This step is usually outsourced to the sequencing companies.

3. Data analysis

NOTE: The analysis in human cell lines was used as an example in this section.

1. Mapping of lentivirus-integration sites
   1. Filter out sequencing reads with more than 50% low-quality bases (Phred score <20), more than 15 'N' bases, or shorter than 150 bases using fastp tool³³ (see Table of Materials).
   2. Select only paired-end reads with the read 2 starting with LTR tag "GCCTTGTGTGTGGTAGATCCACAG
      ATCAAGGATATCTTGTCTTCGTTGGGAGTGA​AT
      TAGCCCTTCCA".
      NOTE: This sequence depends on the sequence of the vector used to generate the insertional mutagenesis cell library.
   3. Trim the LTR tag sequence from the reads selected from step 3.1.2.
   4. Align the trimmed reads to the GRCh37/hg19 using BWA-MEM ³⁴ (see Table of Materials) with default settings.
   5. Retain only read pairs in which both read 1 and read 2 are uniquely mapped to the genome with correct configuration and spacing, indicated by FLAG 147 and FLAG 163.
   6. Define the insertion position as the first aligned base of the trimmed read 2.
   7. Normalize the read counts at each unique insertion site to the total number of uniquely aligned reads in each sample.
2. Identification of insertions affecting cellular fitness (IACFs)
   NOTE: In the statistical analysis, it is advisable to conduct pairwise comparisons among the three biological replicates of both control and treated samples to identify significantly enriched or depleted positions. The number of replicates mentioned above is minimal. It is recommended to use as many replicates as practically possible to ensure the reliability and accuracy of the results.
   1. Convert the normalized read counts of each position to log₁₀ values.
   2. Set the log₁₀ values of positions with zero reads as −5.
   3. Perform two-sided paired Student's t-test on the log₁₀ values.
   4. Adjust the p-values for multiple comparisons using the Benjamini-Hochberg method within the R environment.
   5. Select positions with a False Discovery Rate (FDR) threshold of 25% as IACFs.
3. Identification of clusters affecting cellular fitness (CACFs)
   NOTE: Three biological replicates of control and treated samples were recommended for this analysis.
   1. Identify the significantly enriched or depleted insertion clusters between control and treated samples using SICER³⁵ (see Table of Materials) with an FDR threshold of 1%. The command for SICER analysis is "sh $SICER/SICER.sh $InputDir $In $Control $OutputDir hg19 1 10 1 0.8 30 0.01".
      NOTE: The meanings of the parameters in the command above are as follows: ["InputDir"] ["treated sample insertion sites bed file"] ["control sample insertion sites bed file"] ["OutputDir"] ["species"] ["redundancy threshold"] ["window size (bp)"] ["fragment size"] ["effective genome fraction"] ["gap size (bp)"] ["E-value"].
   2. Obtain cluster regions shared by at least two biological replicates with the "intersect" function of the BEDTools suite³⁶ (see Table of Materials) and denote these cluster regions as CACFs.

Results

Quality control assessment of an NGS library, typically performed by the sequencing provider, serves as the standard quality check before sequencing. To demonstrate the impact of library quality, two samples were compared. The high-quality library, labeled s2-20-15, was prepared using the standard nested PCR protocol, consisting of two sequential amplification steps: the first with 20 cycles, followed by the second with 15 cycles. In contrast, the low-quality library, labeled s1-30, was prepared using a single-step PCR w...

Discussion

In forward genetics analysis, genome-walking is essential for profiling integration sites in insertional mutagenesis cell libraries. Classical genome-walking methods like I-PCR and LAM-PCR often require restriction enzyme digestion and ligation, which can reduce library complexity and introduce biases due to enzyme site dependency^27,28,29. The InSET method overcomes these limitations by eliminating the restriction dige...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.5 M EDTA	Invitrogen	AM9260G	To make Binding/Wash Buffer
1 M Tris-HCl (Ph 8.0)	Invitrogen	15568025	To make Binding/Wash Buffer
10× Klenow Fragment Buffer	Takara	2140A
10× Taq Buffer	Tiangen	ET101-01-01
5 M NaCl	Invitrogen	AM9659	To make Binding/Wash Buffer
BeaverBeads Streptavidin	Beaver	22307-1	1 μm
BEDTools		Software, v2	https://bedtools.readthedocs.io/en/latest/
BWA-MEM			Software, v0.7.12
dNTP Mixture	Takara	4030	2.5 mM each
Equalbit dsDNA HS Assay Kit	Vazyme	EQ111-02
fastp			Software
HiSeq X Ten	Illumina	SY-412-1001	Illumina platform, outsourced to Novogene Corporation (Beijing)
Klenow Fragment	Takara	2140A
NovaSeq 6000	Illumina	20012850	Illumina platform, outsourced to Novogene Corporation (Beijing)
Qubit 3.0 Fluorometer	ThermoFisher Scientific	Q33216
SICER			Software, v1.1
Taq DNA Polymerase	Tiangen	ET101-01-01
Tween-20	HuShi	30189328	To make Binding/Wash Buffer
UltraPure Distilled Water	Invitrogen	10977015
VAHTS DNA Clean Beads	Vazyme	N411-01