Massively Parallel Reporter Assays in Cultured Mammalian Cells

1. Sequence Design and Synthesis

1. Begin MPRA with the design and synthesis of sequences to be assayed for regulatory activity. For compatibility with the pMPRA vector series, design each sequence using the following template: 5’-ACTGGCCGCTTCACTG-var-GGTACCTCTAGA-tag-AGATCGGAAGAGCGTCG-3’ where var denotes the sequence to be assayed and tag denotes one or more identifying tags.
2. The two variable regions (var and tag) are separated by a pair of KpnI (GGTACC) and XbaI (TCTAGA) restriction sites to facilitate directional ligation of a reporter gene fragment between them. In addition, two distinct SfiI (GGCCNNNNNGGCC) sites will be added by PCR to allow directional ligation into the pMPRA backbone. The variable regions must not contain additional copies of any of these restriction sites. If necessary, one or more of these restriction enzymes can be replaced, as long as the replacements 1) allow efficient cutting close to the ends of DNA molecules, and 2) do not cut elsewhere in the vector sequence. See Discussion for additional details.
3. Obtain the required primers listed in Table 1 from a commercial vendor.

Table 1. Primer sequences. [index] denotes a 6 to 8 nt index sequence used for multiplexed sequencing. Obtain at least 8 TagSeq-P2 primers with different indices. All of the primers should be purified by HPLC or PAGE.

4. Oligonucleotide libraries can be obtained from a commercial vendor or core facility, or generated by using a programmable microarray-based oligonucleotide synthesizer as described by the manufacturer. Resuspend the OLS products in 100 µl TE 0.1 buffer.
5. Run the oligonucleotide libraries on a 10% TBE-Urea denaturing polyacrylamide gel. Load approximately 1 pmol per lane and stain with ssDNA-sensitive fluorescent stain to assess their quality (Figure 2A).
6. If a discrete band corresponding to the full-length oligonucleotides can be visualized (Figure 2A, lanes 1 and 2), excise the corresponding acrylamide gel slice and elute oligonucleotides into 100 µl TE 0.1 overnight at room temperature with shaking. Otherwise, proceed using the raw OLS suspension.
7. Amplify and add SfiI tails to the oligonucleotide library using emulsion PCR ⁹.
8. Set up 50 µl PCR reaction mixtures as described in Table 2. Then combine with 300 µl oil and surfactant mixture from the Micellula DNA Emulsion and Purification kit and vortex for 5 min at 4 °C.

Table 2. Emulsion PCR reaction mix (water phase).

9. Use the concentration that gives the maximal yield of amplification products without the appearance of artifacts (see section 1.12).
10. Dispense the resulting emulsion into PCR tubes and perform 20 cycles of PCR (2 min at 95 °C, 20 x[20 sec at 95 °C, 20 sec at 55 °C, 30 sec at 72 °C], 3 min at 95 °C).
11. Pool and break each 350 µl emulsion PCR reaction by adding 1 ml isobutanol and briefly vortexing, and then purify the amplification product using a DNA purification column.
12. Run an aliquot on a 2% or 4% agarose gel to verify artifact-free amplification (Figure 2B).

Figure 2. Preparation of oligonucleotide synthesis libraries. A) Three different raw oligonucleotide synthesis libraries (OLS) run on denaturing 10% TBE-Urea polyacrylamide gels. Bands corresponding to full length oligonucleotides (*) can be visualized and excised from libraries 1 and 2. Library 3 contains contaminants that interfere with PAGE purification. If this is the case, proceed directly to PCR amplification. B) Products of open and emulsion PCR amplification of the same oligonucleotide library run on an agarose gel. PCR amplification of complex oligonucleotide libraries frequently creates chimeric products and other artifacts that may appear as higher and lower bands. Emulsion PCR can minimize these artifacts.

2. Library Construction

1. Prepare a linearized plasmid backbone by digesting vector pMPRA1 with SfiI at 50 °C for 2 hr, run on a 1% agarose gel, excise the backbone band (in this case, 2.5 kb) and purify it using a gel-purification spin column.
2. Digest the emulsion PCR products from step 1.4 with SfiI at 50 °C for 2 hr and purify using DNA purification columns.
3. To ligate the promoter-tag library into the linearized vector backbone, set up reactions containing 100 ng of the digested PCR product, 50 ng of linearized vector backbone and T4 DNA ligase. Incubate overnight at 16 °C and then heat at 65 °C for 20 min to inactivate the ligase.
4. Transform E. coli with the ligation reaction. To preserve library complexity, aim to obtain at least 10x more cfu than there are distinct promoter-tag combinations in the designed library.
5. When the total number of tags is approximately 100,000, the target cfu can typically be achieved by performing 6-8 parallel transformations per library.
6. For each transformation, combine 50 µl electrocompetent E. coli cells with 2 µl of ligation mix on ice. Transform by electroporation, recover the cells in 800 µl SOC medium by shaking at 37 °C for 1 hr, and then combine cells from the parallel transformations.
7. To evaluate the transformation efficiency, plate serial dilutions from an alinquot of recovered cells on LB agar plates with 50-100 µg/ml carbenicillin and incubate overnight at 37 °C. Estimate total cfu as (observed cfu) × (dilution factor) × (V – v) / v, where V is the total volume of recovered cells and v is the volume of the aliquot taken for the serial dilutions.
8. At the same time, use the remainder of the recovered cells to inoculate 200 ml LB supplemented with 100 µg/ml carbenicillin. Grow cells at 37 °C overnight in a shaker incubator and then isolate the plasmid DNA following standard procedures.
9. Digest an aliquot of the isolated plasmid library with SfiI at 50 °C for 2 hr and run on a 1% agarose gel to confirm the presence of inserts.
10. Linearize the plasmid library by cutting between the promoter variants and tags using KpnI and XbaI. To maximize the digestion efficiency, perform serial digestions:
11. First, digest with KpnI at 37 °C for 1 hr and purify using magnetic beads. Second, digest with XbaI with addition of 1 U Shrimp Alkaline Phosphatase at 37 °C for 2 hr, heat inactivate at 65 °C for 5 min, and then purify using magnetic beads.
12. Run an aliquot on a 1% agarose gel to verify complete linearization. If uncut plasmid is visible, the linearized fragment should be gel purified.
13. To generate an MPRA library suitable for transfection into mammalian cells, ligate an ORF with KpnI/XbaI-compatible ends into the linearized intermediate library.
14. To prepare a compatible luc2 ORF fragment from pMPRAdonor1, digest this plasmid with KpnI and XbaI at 37 °C for 1 hr and run on a 1% agarose gel. Excise the ORF fragment (1.7 kb in this case) and purify using a gel purification spin column.
15. Clone the ORF fragment into the linearized intermediate library as described in steps 2.3-2.8.
16. Digest an aliquot (corresponding to 1-2 µg) of the MPRA library with KpnI at 37 °C for 1 hr and run on a 1% agarose gel. If the digested library runs as a single band, proceed to section 3. If additional bands are observed (typically corresponding to carryover of intermediate constructs), the library can be further purified as follows:
17. Digest 3-5 µg of the contaminated library with KpnI at 37 °C for 1 hr and run on a 0.8% agarose gel overnight at 4 °C.
18. Excise the correct library band, purify the DNA using a gel purification spin column and perform self-ligation using high-concentration T4 DNA ligase at 37 °C for 1 hr. Then repeat the transformation and final library DNA isolation as described in step 2.8.

3. Transfection, Perturbation, and RNA Isolation

1. For each independent transfection, culture the required number of cells (as determined by the MPRA library complexity, see Discussion) in appropriate medium. For example, culture HEK293T/17 cells in DMEM supplemented with 10% FBS and L-glutamine/penicillin/streptomycin. Culture cells for at least two independent transfections for each library and experimental condition.
2. Transfect the cultured cells with MPRA plasmids. The transfection method and conditions must be optimized for each cell type. For each transfected sample, retain an aliquot (50-100 ng) of the plasmid DNA as a matched control.
3. For example, transfect 0.5 x 10⁷ HEK293T/17 cells grown to ~50% confluence in a 10 cm culture dish with 10 µg plasmid DNA in 1 ml Opti-MEM I Reduced Serum Medium using 30 µl Lipofectamine LTX and 10 µl Plus Reagent. Remove the transfection mixture after 5 hours and allow the cells to recover for 24-48 hr.
4. Optionally, perform any perturbation required to activate context- or signal-dependent regulatory sequences in the designed library.
5. Harvest the cells and isolate poly(A)+ mRNA using standard oligo(dT) cellulose columns or beads using their manufacturer’s instructions.
6. Ensure that the maximum binding capacity of the columns or beads exceed the total amount of mRNA expected from the harvested cells. For example, the expected yield from section 3.3 is approximately 0.5-2.5 µg mRNA.

4. Tag-Seq

1. To eliminate carryover of vector DNA from the transfected cell lysates, treat each 20 µl mRNA sample with 1 µl Turbo DNase (2 U) and 2.3 µl 10x Turbo DNase buffer at 37 °C for 1 hr, add 2.4 µl Turbo DNase Inactivation Reagent at RT for 5 min with mixing, centrifuge at 10,000 g for 90 sec, and then transfer the solution to a fresh tube.
2. Verify purity by performing PCR as described in section 4.6 on 60-100 ng of each mRNA sample, and then running the products on an agarose gel. If specific amplicons are visible (0.25 kb if using pMPRA1 with luc2), column purify the treated mRNA and then repeat the DNase treatment.
3. To generate Tag-Seq sequencing libraries, convert reporter mRNA to cDNA and add sequencing adapters by PCR.
4. Set up mRNA/RT primer mixtures as described in Table 3. Incubate at 65 °C for 5 min, then place on ice. In parallel, set up cDNA synthesis reactions as described in Table 4.

Table 3. RNA/Reverse transcription primer mix for cDNA synthesis.

Table 4. cDNA synthesis reaction mix.

5. Gently combine mRNA/RT primer mixes with cDNA synthesis mixes. Incubate at 50 °C for 50 min, 85 °C for 5 min and then place on ice. Finally, incubate with 2 U of Ribonuclease H at 37 °C for 20 min.
6. Set up PCR reactions as described in Table 5 with 4-6 µl cDNA reaction mixture or 50 ng reporter plasmid as templates. Then perform PCR amplification (95 °C for 2 min, 26 x [95 °C for 30 sec, 55 °C for 30 sec, 72 °C for 30 sec], 72 °C for 3 min).

Table 5. Tag-Seq PCR reaction mix.

7. Run the PCR products through a 2% agarose gel, excise bands corresponding to the Tag-Seq library amplicons (0.25 kb if using pMPRA1 with luc2) and purify these using gel purification spin columns.
8. Pool, denature and sequence the purified Tag-Seq amplicons directly with an Illumina sequencing instrument.
9. Filter low-quality reads by removing all that a) contain one or more position with a Phred quality score less than 30 within the sequenced tag or b) does not exactly match a designed tag. Count the number of times each remaining tag appears in each library. Normalize the tag counts to TPM (tags per million sequenced tags) and then compute the ratio of mRNA-derived tag counts over plasmid-derived tag counts for each pair of sequencing libraries. If multiple different tags were linked to each sequence variant, use their median ratios for downstream analysis.