June 1, 2018 (Vol. 38, No. 11)
Josh P. Roberts
Using CRISPR-Altered Cells to Fight Disease
The CRISPR revolution—which is bringing even nonexperts the ability to rapidly edit genetic material with pinpoint precision—is beginning to infiltrate the clinic. Standing on the shoulders of gene therapy, cell therapy, and bone marrow transplantation, several groups in the United States and Europe are now gearing up to introduce CRISPR-altered cells into humans to fight life-threatening diseases. Many experts spoke about their findings and experiences at the 4th Annual Precision CRISPR Congress held in Boston earlier this year.
CRISPR (clustered regularly interspaced short palindromic repeats) typically uses a version of the Cas9 nuclease to cut double-stranded DNA at a specified point in the genome, after which the cell’s endogenous machinery will repair the break. Under the right circumstances, the cell can be made to incorporate a donor DNA sequence into the break site, thus allowing for gene insertion, modification, or knockout. Unlike random retroviral insertion, CRISPR-Cas9 targets a specific genetic location determined by a homologous RNA (termed a guide RNA or gRNA). And unlike editing with TALENs (transcription activator-like effector nucleases) or ZFNs (zinc finger nucleases), using a gRNA means that no elaborate protein engineering is required to achieve sequence specificity, and is thus easier and faster.
Minimizing Off-Target Effects
But off-target effects (OTEs) of CRISPR-Cas9 editing can’t be merely dismissed out of hand, especially when moving into patients. The main concern with ex vivo editing is cutting at the wrong site, “and you may not even know it happened,” said Mark Behlke, M.D., Ph.D., chief scientific officer of Integrated DNA Technologies (IDT).
While the first line of prevention is bioinformatic screening, “you have to do it because you don’t want to be using a guide that hits at 1,000 places in the genome—it really doesn’t tell you what sites are the riskiest,” noted Dr. Behlke.
Modeling CRISPR is far more complicated than the thermodynamic hybridization rubrics used for designing PCR primers and probes, and we’re not there yet. It’s best to follow up the bioinformatics with an empirical screen. There are several good in vivo and in vitro methods, including GUIDEseq, CIRCLEseq, and SITEseq, that use next-generation sequencing (NGS) as a readout to show where the guide is actually cutting, Dr. Behlke pointed out.
If the purpose is to destroy a gene, there may be several dozen guide sites that will do; if one proves to have too risky an OTE profile, move on to the next one. Yet sometimes, to fix a disease-causing mutation, for example, only one or two sites are available, and so, minimizing risk may require an enzyme with improved specificity. Several studies have been published that show greatly improved off-target profiles.
But, Dr. Behlke explained, these mutant enzymes have lost so much of their on-target activity that they must be overexpressed from plasmids, resulting in longer exposure to and higher concentrations of Cas9. This negates the benefits of these mutants. “Perhaps the single most important factor to increasing off-targeting effects is how long and how much Cas9 you have present,” he said.
By introducing it as part of a ribonuclear protein (RNP) complex instead, “you get very high ‘on’ right at the beginning when you first put the protein in, but then the protein goes away quickly,” Dr. Behlke explained.
He also discussed IDT’s novel HiFi Cas9, discovered by introducing about a quarter of a million mutants into Escherichia coli and simultaneously screening them for on-target cleavage and against off-target cleavage. The enzyme, which Dr. Behlke said is “the only mutant out there that works well as part of an RNP,” maintains the high on-target cutting of wild type but reduces the off-target cutting 20-fold. “This dramatic drop is going to make this much safer for medical treatment,” he declared.
Switch Back to Fetal
Symptoms of the hemoglobinopathies β-thalassemia and sickle cell disease, including anemia, pain, and early death, arise from defective adult β-globin chain, which begins to replace the fetal form around the time of birth. It has been known for decades that the hereditary persistence of fetal hemoglobin (HPFH) will ameliorate the symptoms of hemoglobinopathies, as will a bone marrow transplant from a normal or HPFH donor. Several projects are underway using CRISPR to recapitulate HPFH in hemoglobinopathy patients.
The route CRISPR Therapeutics took was to disrupt the BCL11A gene, which codes for a transcription factor responsible for genetic switching from fetal to adult β-globin production. Building on animal work presented at last year’s congress, T.J. Cradick, Ph.D., head of genome editing at CRISPR Therapeutics, discussed modeling the correction rates necessary to achieve clinical success using patient-derived bone marrow cells.
“North of 90% have productive edits,” he said. “And then if you extrapolate, if our studies are correct when they’re done in humans, not only do you have a high number of cells edited, but you have a sufficient amount of hemoglobin in each cell.” These levels were seen in immunodeficient mice 16 weeks after engraftment, both in the progenitor hematopoietic stem cells (HSCs) “that we believe will be essential for the long-term cure” as well as more differentiated lineages.
Optimization allowed CRISPR Therapeutics to maintain the high level of editing in GxP manufacturing at clinical scale, and regulatory/toxicity studies revealed no issues. The company plans to begin a clinical trial for β-thalassemia in Europe in 2018, and hopes to conduct a trial in the United States in sickle cell patients beginning next year.
Correct the Mutation
Daniel P. Dever, Ph.D., a research instructor in the Stanford University lab of Matthew Porteus, M.D., Ph.D., is pursuing a biologically easier (but technically more difficult) strategy: simply correct the point mutation that’s responsible for generating sickle hemoglobin and let the relevant gene make normal adult hemoglobin.
The lab previously showed that high levels of correction can be achieved using adeno-associated viral (AAV) vector delivery of a homologous donor along with a CRISPR-Cas9 RNP. Treated patient-derived cells were able to engraft in immunodeficient mice and differentiate in significant numbers.
The Porteus group met with the FDA for a pre-IND “where we outlined our manufacturing strategy, clinical protocol, and preclinical data,” recalled Dr. Dever. He believes that the trial, for which the group hopes to file an IND in 2019, will be the first-in-human gene correction for hematopoietic stem cells.
“Our FDA meeting was a chance to participate in standards development, discussing matters such as off-targets. We’re trying to set a standard of using GUIDEseq for bioinformatics and to couple that with the biology,” explained Dr. Dever. “Another thing that came up was immunity … we think we have a leg up because this is all ex vivo.”
Lack of precedent can reach into the seemingly mundane as well. There is currently no guidance, for example, for how pure gRNA should be. “The quality attributes of these things are not defined—nobody has done it before,” said Shantanu Kumar, Ph.D., senior scientist at Juno Therapeutics, a Celgene company.
“That’s why we’re starting from scratch and setting the standards ourselves, basically looking for a vendor who can make this gRNA in a GMP setting and a Cas9 vendor who can make GMP Cas9 with all the documentation,” he explained. Another vendor is needed to mix these at clinical grade and scale to produce “a single-vial system which is made in a GMP environment, done with all the QC testing that can qualify the drug-like product, so that we can use it in a GMP facility for T-cell engineering.”
Dr. Kumar was surprised to find that there are very few, if any, contract manufacturing organizations (CMOs) that meet his criteria: “We are talking to a few CMOs, trying to teach them, give them protocols, and set standards. Basically, we are going to pay some service fees and all the development costs that will occur. It’s a slow process, but we will get there.”
Using CRISPR to Correct Duchenne Muscular Dystrophy Mutations
Is it possible to heal heart muscle? Chengzu Long, Ph.D., assistant professor at the Leon H. Charney Division of Cardiology at the New York University School of Medicine, thinks so. He utilizes CRISPR-Cas9 technology to correct for Duchenne muscular dystrophy (DMD) mutations in mice and in human cardiac muscle cells.
DMD is caused by mutations in the DMD gene, which codes for dystrophin. Humans with this disease suffer a gradual loss of muscle mass, which ultimately leads to premature death due to respiratory or heart failure.
In preclinical research, Dr. Long and colleagues drew blood from DMD patients and reprogrammed the cells into induced pluripotent stem cells. Next, they introduced guide RNAs and Cas9 into the cells before differentiating them into cardiomyocytes. They used PCR to confirm the edits and harvested either treated mixtures of cells or single-cell clones. The team then expanded these cells before functional analysis. A small 3D model of the heart was created using a hydrogel/collagen matrix to measure contractility, and the investigators found that even as little as a 30% correction is enough to counteract the contractile dysfunction of DMD mutant cells.
CRISPR efficiency is variable and edited sequences are heterogeneous, according to Dr. Long. Technologies like Bio-Rad’s droplet digital PCR (ddPCR) can be used to determine the abundance of the edited cells and provide confirmation of the desired edits. ddPCR detects CRISPR edits made via nonhomologous end joining or homology-directed repair, providing precise and sensitive detection and quantification of edits at or below 0.5% frequency, he points out.
There are over 400 genes linked to 800 neuromuscular diseases. Dr. Long’s work on DMD is just scratching the surface of potential targets for CRISPR therapies.