Written by Dr. Marija Cvetanovic Edited by Dr. Ronald Buijsen
Researchers from the University of California show they can “edit” the Frataxin gene in human cells from Friedreich’s Ataxia and transplant them into mice. This lays the groundwork for this method to be tested for safety.
Friedreich’s ataxia is a progressive, neurodegenerative movement disorder. It is often associated with heart issues and diabetes. Symptoms first start to appear in patients when they are around 10 to 15 years old. Friedreich’s ataxia has the prevalence of approximately 1 in 40,000 people and is inherited in a recessive manner. This means that patients with Friedreich’s ataxia inherited a disease gene from both the father and mother. Friedreich’s ataxia is caused by an overexpansion of the GAA repeat in the Frataxin gene, all these extra repeats causes less Frataxin protein to be made.
Human hematopoietic stem and progenitor cells (HSPCs) are the stem cells that give make to other types of blood cells. You can find HSPCs in the blood all around the body.
HSPCs are ideal candidates for use in stem cell therapy because of a few reasons. First, you can easily get them out of the body through a blood donation (at least easier than some other types of cells!). Second, they can self-renew, meaning they will make more of themselves. Third, other folks have researched this type of cell before, so we know they are fairly safe. Researchers wanted to test if these cells could be used to help treat Friedreich’s ataxia.
Stem cells are cells that provide new cells during growth, and replace cells that are damaged or lost during life. They have the following two important properties that enable them to do this:
The ability to develop (differentiate) into many other, different cell types, for example brain cells, heart cells or liver cells.
The capacity to replicate (self-renewal) and generate more of these important cells.
Mammals have two types of stem cells; embryonic stem cells (ESCs) and adult stem cells. ESCs are derived from an early-stage embryo, while adult stem cells are found throughout the body after development. The main difference between these cells is their ability in the number and type of different cell types they can become. Embryonic stem cells are pluripotent, meaning that they can become all cell types. Adult stem cells are multipotent and can only develop into a limited set of cell types.
Induced pluripotent stem cells
In 2006, the lab of Shinya Yamanaka showed that mouse skin cells could be converted into stem cells by using four specific factors, the Yamanaka factors (1). These cells were named “induced pluripotent stem cells (iPSC). For this important finding Yamanaka was awarded the 2012 Nobel Prize together with Sir John Gurdon. One year later a similar strategy was used to successfully reprogram human skin cells into human iPSCs (2). Nowadays, these iPSCs are an important tool in biomedical research and used for disease modelling, to study disease mechanisms, for drug development and cell replacement studies. To model a disease, a skin biopsy or a urine sample from a patient can be used to generate patient-specific iPSCs. Another option is to modify iPSCs using CRISPR/Cas9. Using defined protocols, these iPSCs can be converted into, for example, brain cells or even mini brains (organoids), which can be used to study a disease. Furthermore, when differences (phenotypes) are found in these cultures, this can be used to screen for drugs that reverse this particular aspect of the disease.
Stem cell-based therapies for SCA
A last aspect is that these iPSCs can be used to replace damaged or lost cells. Since the first stem cell-based clinical trials to replace brain cells that are lost in Parkinson Disease, stem cell replacement therapy has evolved and numerous clinical trials were initiated. In the spinocerebellar ataxia (SCA) field, the first preclinical experiment of embryonic transplants into a mouse model of SCA1 showed a positive effect on animal behaviour and brain pathology (4). Although these first preclinical experiments in SCA disease models are positive, an iPSC-based therapy for SCAs is far from a clinical application.
The CRISPR gene-editing toolbox expanded with the addition of prime editing. Prime editing has astounding potential for both basic biology research and for treating genetic diseases by theoretically correcting ~89% of known disease-causing mutations.
What is prime editing?
Prime editing is coined as a “search-and-replace” editing technique that builds on the “search-and-cut” CRISPR technology. Like CRISPR, prime editing utilizes the Cas9 enzyme targeted to a specific location in the genome by a guide RNA (gRNA). With a few ingenious modifications, including an enzyme called a reverse transcriptase (RT) fused to Cas9, prime editors can be targeted to nearly anywhere in the genome where the RT writes in new DNA letters provided by a template on the gRNA.
How is prime editing different from CRISPR?
Scientists are excited about prime editing because it has several advantages and overcomes many of the limitations of previous CRISPR systems. CRISPR Cas9, an endonuclease, cuts—like scissors—both DNA strands to inactivate a gene or to insert a new sequence of donor DNA. Unlike CRISPR edits, the prime editing Cas9, a nickase, cuts a single DNA strand and does not rely on the cell’s error-prone repair machinery, thereby minimizing any resulting deleterious scars left on the DNA. It has a broader range of targets because it is not limited by the location of short DNA sequences required for Cas9 binding to DNA. The versatility and flexibility of the system allows for more control to inactivate genes as well as to insert, remove, and change DNA letters, and, combine different edits simultaneously—analogous to a typewriter. Importantly, the edits are precise with relatively infrequent unwanted edits. Initial indications showed fewer off-target edits in the genome, possibly because more steps are required for a successful edit to occur. In some cases, it may be more efficient than CRISPR, depending on the targeted cell type, such as in a non-dividing cell like a neuron in the brain. However, with all these advantages, CRISPR still remains the tool of choice for making large DNA deletions and insertions because the prime editing system is limited by the RT and template RNA length.
How could prime editing help ataxia patients?
Prime editing offers enormous possibility for correcting heritable ataxia mutations accurately and safely. In dominantly inherited SCAs, like SCA1 or SCA2, prime editing could shorten the pathogenic repeat expansion allele to the normal length, or inactivate the pathogenic allele without creating unwanted, deleterious mutations. It also provides researchers with a powerful tool to study disease-causing genes in cells and animal models in new ways to advance our knowledge about the underlying mechanisms in ataxia.
What barriers are there to using prime editing as a treatment?
Prime editing will require rigorous testing in cells and animals before moving into humans in a clinical trial. Optimizing delivery and efficiency in target cells and tissues, and minimizing side-effects will be the key barriers to overcome.
To read the original Nature article describing prime editing, it can be found from the Liu lab here.
A common nuisance for bacteria is the bacteriophage: a virus that uses the internal machinery of a bacteria to replicate its own genetic material. Bacteriophages do this by latching onto bacteria and injecting their DNA into the cell. As the cell grows and divides, the bacteriophage’s hope is that their genetic material is replicated alongside the bacteria’s own genome. Unfortunately for bacteriophages, many bacteria have evolved a method to fight off their attacks. After recognizing a viral infection, the bacteria integrate portions of the injected viral DNA into their own genome. The area where these viral DNA segments end up is known as the CRISPR sequence (short forclustered regularly interspaced short palindromic repeat). The viral DNA segments that were integrated into the CRISPR sequence are then replicated and attached to a bacterial protein called Cas9 (CRISPR-associated protein 9). These CRISPR-Cas9 pairs patrol the cell, acting as the bacteria’s antiviral immune system. If the same viral infection happens again, the DNA in one of the CRISP-Cas9 pairs will match part of the injected viral DNA and bind to it. Once bound, Cas9 cuts the viral DNA, which is then destroyed.
Recently, scientists have found a way to harness this system for manipulating genes (a process broadly called genetic engineering). By making an artificial CRISPR sequence, attaching that sequence to Cas9, then introducing the man-made CRISPR-Cas9 into a cell, it becomes possible to make a targeted cut in any gene. Making a CRISPR-Cas9 pair that targets one specific gene is as simple as making a CRISPR sequence that matches that gene.
Unlike in bacteria, most organisms repair rather than simply destroy cut DNA. This leaves the targeted genetic sequence available for further manipulation, including the introduction of a short mutation or even the insertion of a whole new DNA sequence. In essence, using the CRISPR-Cas9 system, scientists are now able to edit genes in a simple, targeted way.
CRISPR-Cas9 has become quite popular as a genetic tool in research settings: as of now, the genomes of anything from worms and fruit flies to mice and monkeys have been altered using this technique. While its use in humans is still in its early stages – the first patient treated using CRISPR began therapy earlier this year – is plausible that CRISPR-Cas9 could prove useful in altering the genomes of patients with genetic disorders (like, for instance, the SCAs). For patients, this might sound like a miracle cure. However, it is important to note that several concerns remain as to the ethics of human genetic engineering – the concept of “designer babies” being one of them.