Beginner Wizard

CRISPR-KO (Knockout)

Cas9 creates indels that disrupt the gene. Permanent loss-of-function. Best for genes where complete knockout is tolerated.

CRISPRi (Interference)

dCas9-KRAB represses transcription without cutting DNA. Reversible, tunable. Best for essential genes where KO is lethal.

CRISPRa (Activation)

dCas9-VP64/p65 activates transcription. Gain-of-function. Best for finding genes whose overexpression affects phenotype.

Prime Editing MAVE

Variant-level pooled screening is supported through Prime Editing MAVE. The current configuration supports human GRCh38 SNVs with PE2-style NGG pegRNA design and evaluates functional effects from pegRNA abundance only.

Dropout (Negative Selection)

Guides targeting essential genes are depleted over time. Compare endpoint to T0. Most common for genome-wide viability screens.

Positive Selection

Guides targeting genes that confer resistance are enriched. Only a small fraction survives. e.g., drug resistance screens.

Treatment Response

Compare treated vs. untreated. Guides affecting drug response are enriched or depleted. Requires matched control.

FACS (Fluorescence Sorting)

Sort cells based on fluorescence (e.g., reporter, surface marker). Collect high/low bins. Major bottleneck at sorting step.

What is this?

A CRISPR library is a collection of sgRNAs targeting your genes of interest. Genome-wide libraries target all ~19,000 human genes. Focused libraries target a subset.

Why does it matter?

Library size directly determines how many cells you need. A 76,000-guide library at 500× coverage requires ~175 million cells per replicate.

Bacterial amplification

After cloning, transform and plate enough bacteria to maintain representation. Rule of thumb: library size × desired bacterial coverage (typically 500-1,000×). Pool the amplified colonies and prepare plasmid DNA.

NGS QC of the amplified plasmid library

Sequence the final amplified plasmid pool once, before viral packaging. Check:

• Guide representation (Gini coefficient < 0.1 ideal)
• Missing guides (< 1% missing)
• Abundance distribution (no extreme skew)

For a custom library: use QC to upload FASTQ from the amplified plasmid pool and the guide manifest; confirm representation before producing virus.

Package the sgRNA library into lentivirus

Use a third-generation compatible packaging workflow in HEK293T cells under your institution's approved biosafety procedures. The transfer plasmid is your pooled sgRNA library; the standard helper plasmids are psPAX2 (Addgene #12260) and pMD2.G (Addgene #12259).

Starting DNA ratio

Use transfer plasmid : psPAX2 : pMD2.G = 4 : 3 : 1 by mass as a starting point. For a 10 cm dish with 10 µg total DNA, this is 5.0 µg library plasmid, 3.75 µg psPAX2, and 1.25 µg pMD2.G. Optimize the transfection reagent and DNA amount for your local HEK293T culture conditions.

Workflow

1. Seed HEK293T cells the day before so they are approximately 70-80% confluent at transfection.
2. Transfect the three-plasmid mixture, then replace medium according to the transfection-reagent protocol.
3. Collect viral supernatant at 48 h and, if compatible with cell health, again at 72 h; clarify through a 0.45 µm low-protein-binding filter.
4. Use freshly collected or appropriately stored virus for the titration below. Determine the functional titer in the intended target cell type, not only in HEK293T cells.

Why pilot first?

Before your full screen, perform a small-scale viral titration to determine the viral volume needed to achieve your target MOI.

How?

Transduce cells with different virus volumes. After 48-72h, measure the fraction of infected cells (via antibiotic selection or fluorescent marker).

MOI ≈ -ln(1 - f_infected)

What is MOI?

Multiplicity of Infection = average number of viral integrations per cell. At MOI=0.3, ~25.9% of cells are infected; among infected cells, ~85.7% have exactly one integration and ~14.3% have multiple integrations.

Why low MOI?

Low MOI enriches for single-integrant infected cells, making phenotype attribution to one gene perturbation more reliable. The exact fraction is shown below from the Poisson model.

How many cells do I need?

Cells needed = (library size × target representation) / (P(single integration) × selection survival)

For a 78,000-guide library at 500× with MOI=0.3 and 30% selection survival, you need ~585 million cells per replicate.

After transduction (24-48h), add antibiotic to select for transduced cells. Duration: typically 2-4 days for puromycin.

Important: Perform a kill-curve pilot first. The concentration and duration depend on your cell line.

After selection, expand cells to reach the required number for your screen. Maintain representation at every passage — never split below target coverage × library size.

What is T0?

T0 is the baseline timepoint collected after selection and expansion, before the experimental selection/treatment begins. It represents the starting guide distribution.

Why is it needed?

T0 serves as the reference for comparing endpoint guide abundance. Without T0, you cannot distinguish guide depletion from stochastic loss.

Plasmid reference vs. cellular T0

Some labs use the plasmid library as reference. However, cellular T0 captures the actual post-transduction distribution, which may differ from the plasmid library due to transduction bias.

This is where your phenotype is applied. The type depends on your screen:

• Dropout: Simply grow cells; depleted guides indicate essential genes
• Positive selection: Apply selective pressure; only resistant cells survive
• Treatment: Add drug; compare to untreated control
• FACS: Sort cells by fluorescence; collect high/low bins

Bottleneck warning: This stage can dramatically reduce representation. If survival fraction is low, you need proportionally more starting cells.

During the screen, cells need to be passaged to maintain growth. Each passage is a potential bottleneck.

Rule: Always maintain ≥ target representation × library size cells at every passage. Never split below this threshold.

At the endpoint, harvest all cells and extract genomic DNA. Record the total cell count — this determines your gDNA yield and downstream PCR capacity.

Cells → Genome Equivalents → DNA Mass

Each diploid cell contains 2 genome equivalents (2 × 3.5 pg = 7.0 pg DNA).

For aneuploid lines, adjust the ploidy parameter.

Why is PCR critical?

PCR amplifies the sgRNA cassette from gDNA. If only 100 genome equivalents per guide enter PCR, your effective molecular representation is 100× — regardless of how many cells you started with.

Key formula

PCR representation = (total DNA × genomes per µg) / (library size)

Sequencing cannot fix low PCR representation. If PCR gives 100×, sequencing 10,000 reads/guide gives 100 independent observations, not 10,000.

How many reads?

Target: 200-500 reads per guide per sample. Total reads = guides × reads/guide × samples.

Only ~70% of reads will map to your library (usable read fraction).

Important

More reads cannot compensate for low molecular representation. If PCR input is 100 genomes/guide, effective independent observations = 100, not your read count.

When sequencing is complete, go to Analyze to upload your FASTQ or count table. Each input accepts your own files and provides downloadable examples to test the workflow first.

Click below to run the complete calculation and generate your experimental plan.