Focus Feature on Gene Editing: Catching up on CRISPR
A roundup of recent CRISPR gene editing R&D news, plus a Q&A and two commentaries from experts in the field
Focus Feature: CRISPR Gene Editing
Catching up on CRISPR
A roundup of recent CRISPR R&D news, beginning with a COVID-19 test from UCSF and Mammoth
By Jeffrey Bouley
Scientists at the University of California, San Francisco (UCSF)—along with researchers at Mammoth Biosciences—have developed what they say is an inexpensive new test that can rapidly diagnose COVID-19 infections.
The partners say that the new test—officially named the SARS-CoV-2 DETECTR—is easy to implement and to interpret, and requires no specialized equipment, which could make the test more widely available than the current crop of COVID-19 test kits, if it were to receive regulatory approval—UCSF researchers are clinically validating the test in an effort to fast-track the approval through the emergency use authorization path.
“The introduction and availability of CRISPR technology will accelerate deployment of the next generation of tests to diagnose COVID-19 infection,” said Dr. Charles Chiu, a professor of laboratory medicine at UCSF and co-lead developer of the new test, which is described in a paper published April 16, 2020, in the journal Nature Biotechnology.
The new SARS-CoV-2 DETECTR assay is among the first to use CRISPR gene-targeting technology to test for the presence of the novel coronavirus. Since CRISPR can be modified to target any genetic sequence, the test kit’s developers “programmed” it to home in on two target regions in the genome of the novel coronavirus. One of these sequences is common to all “SARS-like” coronaviruses, while the other is unique to SARS-CoV-2, which causes COVID-19. Testing for the presence of both sequences ensures that the new DETECTR tool can distinguish between SARS-CoV-2 and closely related viruses.
Much like the diagnostic kits currently in use, the new test can detect the novel coronavirus in samples obtained from respiratory swabs. However, the new test is able to provide a diagnosis much more quickly. While the widely used tests based on polymerase chain reaction (PCR) techniques take about four hours to produce a result from a respiratory sample, the new DETECTR test takes only 45 minutes, rapidly accelerating the pace of diagnosis.
Another key advantage of the new DETECTR test is that it can be performed in virtually any lab, using off-the-shelf reagents and common equipment. This stands in stark contrast to PCR-based tests, which require expensive, specialized equipment.
CRISPR plays key role in new brain cancer model
Sticking with university-based research and the UC system in California—this time in San Diego—we have news from early in the year from researchers at the UC San Diego School of Medicine. Using genetically engineered human pluripotent stem cells, they created a new type of cancer model to study in vivo how glioblastoma, the most common and aggressive form of brain cancer, develops and changes over time.
“We have developed stem cell models that are CRISPR-engineered to have tumor-associated driver mutations in glioblastoma, which harbor essentially all features of patient-derived tumors, including extrachromosomal DNA amplification,” said co-senior author Dr. Frank B. Funari, a professor in the Department of Pathology at the medical school and head of the Laboratory of Tumor Biology in the San Diego branch of the Ludwig Institute for Cancer Research.
“These models, or avatars as we call them, enable us to study human tumor development over long periods in vivo, which has not been feasible with patient-derived tissue samples which already harbor other genetic changes.”
Reporting in the Jan. 28 issue of Nature Communications, researchers used CRISPR editing to make precise mutations in an otherwise “normal” genome to create the genetic conditions that enable tumor development. The resulting avatars are unique in that they behave like a grade 4 glioma—a fast-growing type of tumor that starts in the glial cells of the brain.
“The addition of single-cell RNA sequencing and computational tools enabled efficient analysis of big data to truly evaluate the surprising intra-tumor heterogeneity present in our avatars which replicates what is seen in patients samples,” noted co-senior author Dr. Gene W. Yeo, a professor in the Department of Cellular and Molecular Medicine and the Institute for Genomic Medicine at UC San Diego and faculty member in the Sanford Consortium for Regenerative Medicine.
Existing mouse models work for testing drugs for specific mutations, but do not account for the diverse ways that tumors can develop. Human tissue samples do not allow for standardization in testing. This new avatar modeling system, according to the authors, provides a platform for standardized studies on tumor biology and evolution.
“We can now test which mutations predicted by cancer genome projects are truly tumor-driving, and how they become invasive,” remarked Yeo. “More importantly, these cancer avatars provide systematic, well-controlled opportunities for drug discovery.”
First trial for a CRISPR bacteriophage therapy
Moving on to North Carolina and the more commercial side of CRISPR R&D, Research Triangle Park-based Locus Biosciences announced earlier this year that it was enrolling patients for a Phase 1b clinical trial evaluating LBP-EC01, a CRISPR/Cas3-enhanced bacteriophage (crPhage) product that will target Escherichia coli bacteria causing urinary tract infections (UTIs). As the world’s first controlled clinical trial for a recombinant bacteriophage therapy, this trial represents a significant milestone for the field, the company noted.
LBP-EC01 is a bacteriophage cocktail that has been engineered with a CRISPR/Cas3 construct targeting the E. coli genome. The product works through a unique dual mechanism of action utilizing both the natural lytic activity of the bacteriophage along with the DNA-targeting activity of CRISPR/Cas3. This dual mechanism reportedly makes LBP-EC01 significantly more effective at killing E. coli cells than corresponding natural bacteriophages, as shown both in laboratory tests and in small animal models of urinary tract infection.
“This trial represents a major step toward proving that CRISPR recombinant phage can reach into the human body and precisely remove a specific pathogen,” said Paul Garofolo, CEO of Locus.
Commentary: The many faces of CRISPR/Cas
By Trevor Collingwood of ERS Genomics
CRISPR/Cas genome editing is fundamentally impacting many branches of life science, including gene and cell-based therapy, drug discovery, diagnostics, industrial biology, plant agriculture, livestock and even veterinary medicine. As of late 2019, two classes, six types and 33 subtypes of naturally occurring prokaryotic CRISPR/Cas systems have been described,1 but the so-called CRISPR/Cas9 system has so far proved the most amenable to repurposing for genome editing in eukaryotic systems.
Cas9 is an endonuclease, and by complexing it with a guide RNA that contains approximately 20 nucleotides of homology to a bespoke DNA target, Cas9 can be directed to bind and cleave genomic DNA in cells in a sequence-specific manner. Cleavage generates a DNA double strand break (DSB). A DSB is a key initiating step in genome editing; one that is typically followed by the natural DNA repair processes of non-homologous end joining (NHEJ), which is relatively imprecise, and/or highly precise homology-directed repair (HDR), in which the desired genetic change is copied into the target locus from an accompanying DNA donor template.
This type of controlled “cut and repair” approach has been the mainstay of genome editing for 20 years.
Not making the cut
Creation of a targeted DSB brings with it a risk of off-target DNA cleavage and adverse genetic changes. Variants of Cas9 that bind DNA but cut only one DNA strand (nCas), or do not cleave at all (dCas), have found utility in facilitating other types of genome editing that do not require a DSB. For example, a process called base editing typically uses nCas9 or dCas9 fused to a deaminase effector domain to achieve targeted conversion of a single base to a different base, namely C to T, or A to G, without invoking a DSB.
Much effort has focused on optimizing base editing systems to improve both the efficiency of base modification, as well as overcoming the challenge of selectively modifying a specified base without also modifying its immediate neighbors. Additionally, low-level genome-wide base conversion by the deaminase fusions has been observed that is independent of Cas9 and so likely due to the intrinsic DNA affinity of the deaminase. Recently, a series of engineered deaminase variants was described that exhibit up to a 100-fold reduction in such genome-wide promiscuity while retaining high on-target efficacy.2
In a separate elegant approach called prime editing, dCas9 or nCas9 is fused to a reverse transcriptase. The fusion is able to use a 3’ extension of its own guide RNA as a template from which to reverse transcribe the desired genetic change and insert it directly into the target locus—again, without need for a DSB.3 Both approaches offer intriguing alternatives to DSB-mediated site-specific genome editing, but they too will require further optimization to prove their utility and safety.
More shots on goal
Ideally, a DSB is created as close as possible to the intended site of editing. This is because the efficiency of targeted genome editing at any site decreases markedly with increasing distance from the DSB. Cas9 and other CRISPR nucleases, such as Cas12, are limited in where they can bind because they require a protospacer adjacent motif (PAM)—a conserved sequence of nucleotides that must be present next to the intended target sequence. For the most commonly used Cas9, that which is derived from Streptococcus pyogenes (SpCas9), the PAM sequence is NGG. This is one of the shortest and therefore most frequently targetable PAM motifs, but its required presence still imparts constraint on where editing can occur.
To overcome this barrier, use of directed evolution or structure-guided approaches has been partially successful in relaxing the PAM constraint for SpCas9. A recent report described new variants of SpCas9. One variant in particular (SpRY) was capable of targeting NRN (where R can be A or G) and, to a lesser extent, NYN PAMs (where Y can be C or T). In effect, this enables efficient editing at NNN—essentially removing the PAM requirement altogether. Relaxing the PAM constraint does, though, risk opening the door to increased off-target nuclease activity, and indeed this was seen with SpRY. However, nearly all off-target activity was eliminated upon incorporation of additional “high fidelity” amino acid substitutions previously shown to reduce off-target activity in wild type and other SpCas9 variants.4 Thus, SpRY and similar SpCas9 variants from this group and others may offer an important solution to high-resolution and high-specificity genome editing using only a small selection of Cas9 variants.
Applying the brakes
Efficient DNA cleavage by CRISPR/Cas in eukaryotic cells brings opportunities for therapeutic applications, whether ex-vivo cell-based approaches or direct in-vivo gene editing. The safety threshold for clinical use of CRISPR is understandably high, and many approaches have focused on maximizing on-target nuclease activity while minimizing the possibility for off-target DNA cleavage and the creation of unpredictable and unwanted mutations. Such strategies include optimizing (i) guide RNA and Cas9 nuclease structure, (ii) methods to deliver them to the cells, and (iii) controlling the longevity of the active nuclease complex inside the cell, where idleness is the root of mischief—once the intended edit is made, residual active nuclease left unchecked can do harm.
A particularly intriguing approach to controlling CRISPR systems comes from nature itself and the recently identified naturally occurring inhibitors of CRISPR/Cas, termed anti-CRISPR (Acr) proteins. Acrs derive from mobile genetic elements, such as phage and plasmids, and are themselves used by invaders of prokaryotes as a countermeasure to the latter’s CRISPR/Cas defensive systems. To date, 45 non-homologous Acr proteins have been discovered, many with overlapping activity against different Cas9 and Cas12 orthologs.5 They act by direct interaction with these Cas proteins to disrupt DNA binding, catalytic function or other attributes, and have therefore been considered as potential “kill switches” for CRISPR/Cas.
Acrs could be employed after the intended edit has occurred—perhaps following a successful in-vivo or cell-based therapeutic genome-editing event, or even to halt a runaway gene drive from propagating undesirable traits throughout a population. As an example, Li et al. showed that toxicity related to prolonged expression and activity of SpCas9 in human CD34+ cells was ameliorated by sequential expression of AcrIIA2 together with AcrIIA4 after the editing event.6
Methods for controlled induction of Acr expression itself are also being actively studied as a means to achieve rapid and dynamic control of CRISPR/Cas. Such approaches include (i) transcriptional control of Acrs by driving their expression by promoters that are active at a specific stage of the cell cycle or in response to cellular events; (ii) post-transcriptional regulation of Acrs by tissue-specific microRNAs; (iii) optogenetic control of Acrs, whereby a light-sensitive protein domain inserted into the Acr permits the chimeric Acr to bind to and inhibit Cas9 in the absence of light, but upon photoexcitation, the Acr chimera is destabilized and the Cas9 released; and (iv) ligand-induced expression of Acr by fusing Acr with a destabilizing domain that maintains structural integrity of the fusion in the presence of a ligand, but degrades the Acr fusion once the ligand is withdrawn.3
An orthogonal approach to modulating CRISPR/Cas is to use small-molecule inhibitors of nuclease function. Maji et al. developed a screening platform by which they were able to identify a small-molecule inhibitor of the binding interaction between guide RNA-bound SpCas9 and its NGG PAM sequence, with subsequent attenuation of DNA cleavage.7 The strength of inhibition was very sensitive to structural changes affecting the Cas9/PAM interaction, suggesting that for different Cas9 variants or orthologs, different inhibitors would likely be needed. Nevertheless, small-molecule inhibitors offer the advantages of being cell permeable, reversible, proteolytically stable, non-immunogenic and non-cytotoxic, as well as exhibiting fast kinetics.
Limited only by imagination
Safe, efficient and sophisticated targeted modification of genomic DNA in a broad array of cell types has been the primary goal for genome editing to date. However, CRISPR/Cas now comes in many forms and is proving itself a fascinating tool not only for basic research, but also for new diagnostic and detection applications, as evidenced by the plethora of recent submissions to the Biorxiv preprint server for the use of Cas12a or the RNA-cleaving Cas13 in the detection or even treatment of SARS-CoV-2 infection.
It has been nearly eight years since the seminal paper by Jinek et al. set the CRISPR/Cas genome editing world alight.8 How this new tool has evolved since then has been astounding. CRISPR/Cas9 is now in the clinic, and CRISPR-modified food products are making their way to market. The coming years will, without doubt, expand on the many faces of CRISPR/Cas and bring forth tools and applications hitherto unconceived.
Dr. Trevor Collingwood brings over a decade of experience in the field of genome editing and its applications to his role heading up business development at ERS Genomics. He has held key roles in R&D, business development, marketing and alliance management for early-stage development of genome-editing technologies at leading companies in the field.
1. Makarova, K.S., et al., Evolutionary classification of CRISPR-Cas systems: a burst of class 2 and derived variants. Nat Rev Microbiol, 2020. 18(2): p. 67-83.
2. Doman, J.L., et al., Evaluation and minimization of Cas9-independent off-target DNA editing by cytosine base editors. Nat Biotechnol, 2020.
3. Anzalone, A.V., et al., Search-and-replace genome editing without double-strand breaks or donor DNA. Nature, 2019. 576(7785): p. 149-157.
4. Walton, R.T., et al., Unconstrained genome targeting with near-PAMless engineered CRISPR-Cas9 variants. Science, 2020.
5. Marino, N.D., et al., Anti-CRISPR protein applications: natural brakes for CRISPR-Cas technologies. Nat Methods, 2020.
6. Li, C., et al., HDAd5/35(++) Adenovirus Vector Expressing Anti-CRISPR Peptides Decreases CRISPR/Cas9 Toxicity in Human Hematopoietic Stem Cells. Mol Ther Methods Clin Dev, 2018. 9: p. 390-401.
7. Maji, B., et al., A High-Throughput Platform to Identify Small-Molecule Inhibitors of CRISPR-Cas9. Cell, 2019. 177(4): p. 1067-1079 e19.
8. Jinek, M., et al., A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science, 2012. 337(6096): p. 816-21.
Q&A: Easi-CRISPR expands the limits on gene editing
By DDNews Staff
When generating a genetically engineered rodent model, various gene-editing techniques are available for consideration. CRISPR/Cas 9 is increasingly popular for its speed, but there is a limit to the size and complexity of the genetic modifications CRISPR can handle efficiently and accurately. The Easi-CRISPR technology expands the boundaries and opens the door to larger genetic modifications than possible with traditional CRISPR.
DDNews recently talked with Jochen Welcker, director of molecular biology and scientific development at Taconic Biosciences, about the Easi-CRISPR technology Taconic licenses from the University of Nebraska, exploring what makes it different, the advantages it offers and what’s next for this gene-editing technique.
DDNews: Can you give a brief rundown of how the process of Easi-CRISPR differs from other CRISPR approaches?
Welcker: Genetic modifications, such as deletions, insertions or replacements, became much more feasible using CRISPR technology. However, some caveats exist for standard CRISPR approaches. Insertions and replacements require an HDR (homology directed repair)-template. Single-stranded DNA (ssDNA) is preferred since it is not prone to unintended, random insertions—however, chemical synthesis of ssDNA molecules is limited to up to 200 nucleotides. For HDR-templates larger than 200 nucleotides, double-stranded DNA (dsDNA) molecules must be used; however, dsDNA is highly prone to random insertions and has a low HDR-frequency. Therefore, long ssDNA molecules are required to perform insertions or replacements in a controlled fashion. This is where Easi-CRISPR comes into play. Easi-CRISPR uses long ssDNA (instead of dsDNA) as the HDR-template, enabling large insertions or replacements while avoiding random insertions. Since long ssDNA cannot be generated by chemical synthesis, Easi-CRISPR is more labor-intensive than standard CRISPR approaches.
DDNews: What are the benefits of Easi-CRISPR? What kind of gene editing does it enable beyond that of other CRISPR approaches?
Welcker: The size of genetic information that can be built into a given genomic locus is even smaller than the 200 nucleotides that standard CRISPR approaches use for HDR templates. Homology arms flanking the genetic element must be introduced, and each homology arm comprises approximately 40 nucleotides. That leaves only 120 of the 200 nucleotides in the HDR-template available to encompass a novel genetic sequence. This is sufficient for introducing point mutations or small protein tags, but not for larger modifications. Since Easi-CRISPR utilizes much longer template molecules which can be several kilobases in length, it enables the insertion of entire human or reporter genes as well as genomic replacements in vivo.
DDNews: Given that Easi-CRISPR allows for larger insertions than standard CRISPR, does it come with a greater risk of off-target or downstream effects?
Welcker: No, it does not. All parameters related to the guide RNA—e.g., cutting efficiency and the number of potential off targets—are identical for both CRISPR approaches. Easi-CRISPR comes with a lower risk of downstream effects because unintended random insertions should not occur. The quality of ssDNA templates, which is dependent on the production method, is of utmost importance with Easi-CRISPR. ssDNA can be obtained through a range of methods, such as in-vitro transcription to generate RNA followed by reverse transcription, enzymatic digestion of dsDNA by strandase, and asymmetric PCR.
In-vitro transcription and reverse transcription, as well as asymmetric PCR, harbor the risk of introducing mutations into the ssDNA. Strandase digest of dsDNA is never complete, therefore ssDNA generated this way contains a high fraction of contaminating dsDNA. In our hands, the most reliable method resulting in high-fidelity ssDNA with little to no dsDNA contaminants is the nicking of ssDNA out of plasmid templates. This method relies on the use of nicking endonucleases and does not rely on enzymatic replication or purification.
Nevertheless, stringent quality control—for example, quality control of the resulting genetically modified animals—is mandatory. Unintended mutations (arising from the ssDNA template itself or resulting from errors during the integration into the genome) cannot be entirely excluded. Also, even though random integration of ssDNA occurs with a much lower frequency than that of dsDNA, this might also happen with ssDNA.
DDNews: What are the indications for which this technology is particularly well suited?
Welcker: Easi-CRISPR is ideally suited to the insertion of commonly used sequences like reporter genes (e.g., green fluorescent protein), recombinases (e.g., Cre or Flp) or protein tags. Genomic replacements can also be achieved effectively with Easi-CRISPR, unless they exceed a certain size. Since the length of ssDNA is still limited in size, Easi-CRISPR is not suited to large gene replacements or genomic modifications.
DDNews: What do you consider to be the most interesting or promising aspect of Easi-CRISPR?
Welcker: Easi-CRISPR enables genomic modifications in vivo, which otherwise would only be possible by targeting embryonic stem (ES) cells—a time-consuming procedure requiring solid ES cell lines and access to a cell culture laboratory. Instead, Easi-CRISPR introduces targeted modifications directly into mouse or rat embryos via pronuclear injections, omitting the cell culture and significantly reducing the timeline, costs and effort to generate animal models. Thus, Easi-CRISPR gives scientists faster access to animal models for the most current research topics. A timely example is the rapid generation of a mouse model expressing the human ACE2-receptor under control of endogenous mouse regulatory elements to study SARS-CoV2 infections.
DDNews: The gene editing field has obviously seen explosive growth since the widespread adoption of CRISPR; do you expect to see that growth continue as Easi-CRISPR gains traction?
Welcker: The advantages of speed and simplicity made possible by Easi-CRISPR expanded the toolbox available for more advanced genome modifications. Easi-CRISPR enables the use of genome editing for specific scientific questions which were only achievable by ES cell targeting previously. As of now, Easi-CRISPR is still limited regarding template size and complexity. However, technical improvements in the future may enable using Easi-CRISPR for large replacements of complex genome modifications. Easi-CRISPR pushes the limitations of gene editing and opens the door not only to generating animal models where no ES cell lines are available yet, but could eventually also lead the way to new gene therapy strategies in humans as well.
Jochen Welcker joined Taconic Biosciences in 2012 after completing his postdoc at the Max-Delbrück-Center for Molecular Medicine, during which he worked on the genetic aspects of mouse development. He has more than a decade of experience in designing, generating and analyzing genetically engineered mice.
Commentary: Using gene editing to produce more effective cell lines for research and vaccine production
Dr. Elizabeth Turner Gillies, Scientist for ATCC
For decades, most vaccines have been made in chicken eggs, which act as a simple tissue culture system for viral production. Vaccine makers simply inject the virus and, after the appropriate incubation period, purify replicated viral particles. The process is simple, relatively inexpensive and works quite well for a number of routine vaccines such as influenza.
While this approach usually meets normal vaccine demand, it can fall short during a public health emergency, as vaccine production in eggs can be difficult to initiate and scale up from a small R&D laboratory operation to an industrial manufacturing campaign.
Procuring millions of extra eggs for industrial-scale vaccine production is challenging. Retasking the stocks devoted to influenza production, for example, would deplete stocks to produce that critical vaccine. In addition, the eggs must be incubated in enormous, temperature- and humidity-controlled warehouses. As with egg stocks, warehouses would also need to be reassigned, reducing incubation space for other vital vaccines.
Producing vaccines in a mammalian tissue culture system offers a practical solution for these challenges. While this approach can be a bit more expensive in the short-term, working with a bioreactor rather than a warehouse full of eggs can accelerate the response.
In addition, strategically editing cell line genomes can make these systems even more powerful, boosting research and improving vaccine production.
Editing Vaccine Cell Lines
Vaccine production relies predominantly on four cell lines: Vero, WI-38, MDCK and MRC-5. While researchers have made incremental improvements over the years—much in the same way farmers might improve their stock through selective breeding—by adjusting the lines through different bioreactors, media conditions, scale-up processes and clonal selection, these decades-old lines have remained mostly unchanged.
While entirely new cell lines can be developed, stringent regulatory guidelines could significantly delay vaccine development. Rather than attempting to reinvent the wheel with a new line, CRISPR/Cas9 gene editing can provide a more straightforward method to improve existing cells.
The antiviral interferon response system is one promising pathway. Because they are partially interferon-production deficient, Vero cells are commonly used to produce many viruses. These deficient cells cannot signal the presence of an ongoing viral infection to neighboring cells or activate anti-viral interferon response genes, which modulate protein synthesis, DNA replication, RNA production and other antiviral mechanisms.
These mechanisms decrease the number of viral particles that can be generated in target cells, which is great news for cell survival, but not for virologists and manufacturers trying to scale up vaccine production.
Culturing Different Viruses
Editing genes that drive the host-cell innate immune response could potentially increase production capacity across many viruses. Beyond that, additional gene modifications could increase production even more for specific viruses. However, changes that increase production of one virus might decrease production of another, necessitating new cell line development for each microbe.
Modifying apoptotic or cell stress genes to improve cellular health, or ribosomal regulators to increase output of viral proteins, are other strategies that could improve viral production. However, modifying the host-cell antiviral interferon signaling pathway is a good first step towards improving historical cell lines’ viral production capacity, regardless of virus.
CRISPR/Cas9 gene editing is an attractive, technologically attainable strategy to optimize viral vaccine production; however, it should be followed by rigorous validation studies to ensure traceability and effectiveness. Extensively characterizing and authenticating cell lines, and validating cellular genotype, phenotype and biofunction, is essential before committing a gene-edited line to vaccine production.
STAT1 knockout cell lines offer significant advantages over traditional lines to increase viral production. With legacy cells, researchers had to spend significant time ensuring the target virus would grow in the first place. Only after that problem had been solved could they move on to the scientific questions they were trying to answer. By contrast, STAT1 knockout enhances infection, simplifying this process. CRISPR-edited cell lines will help them cut to the chase.
Ultimately, genome-edited Vero and MDCK cell lines give scientists new tools to accelerate research and vaccine production. In addition, optimizing cell models can make it easier to adapt vaccine production to cells, rather than eggs, increasing overall flexibility and the ability to scale up in a crisis.
Dr. Elizabeth Turner Gillies is a scientist in cell biology research and development at ATCC. She currently works on CRISPR/Cas9 engineered advanced cell models.
Analysis of datasets leads to largest genetic screen resource for cancer research
A comprehensive map of genes necessary for cancer survival is one step closer, thanks to work at the end of 2019 to validate the two largest CRISPR/Cas9 genetic screens in 725 cancer models across 25 different cancer types. Scientists at the Wellcome Sanger Institute and the Broad Institute of MIT and Harvard compared the consistency of the two datasets, independently verifying the methodology and findings.
The results mean that the two datasets can be integrated to form the largest genetic screen of cancer cell lines to date, which will provide the basis for the Cancer Dependency Map in around 1,000 cancer models. The scale of this combined dataset will help to speed up the discovery and development of new cancer drugs.
The Cancer Dependency Map (Cancer DepMap) initiative aims to create a detailed rulebook of precision cancer treatments for patients. Two key elements of the Cancer DepMap are the mapping of the genes critical for the survival of cancer cells and analytics of the resulting datasets. Despite recent advances in cancer research, making precision medicine widely available to cancer patients requires many new drug targets.
Said Dr. Francesco Iorio of the Wellcome Sanger Institute and Open Targets: “It is worth remembering that when these datasets were originally produced we were dealing with a new, unproven technology. This study is important because it demonstrates the validity of the experimental methods and the consistency of the data that they produce. It also means that two large cancer dependency datasets are compatible. By joining them together, we will have access to much greater statistical power to narrow down the list of targets for the next generation of cancer treatments.”