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How can we make CRISPR therapies safer?

Image credit: Arek Socha/Pixabay

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Jamila
Jamila Oct 21, 2020
How can we hit the brakes on CRISPR therapies to ensure the patient's safety?

CRISPR is a revolutionary gene-editing technology. This year, Jennifer Doudna and Emmanuelle Charpentier won the Nobel prize for their outstanding work on CRISPR-Cas9. CRISPR was initially discovered in bacteria, where it serves as a defense mechanism against bacteriophages. Researchers saw how bacteria could use guide RNAs and Cas to cut up specific sequences belonging to bacteriophages. Realizing CRISPR's potential, the researchers developed it into a gene-editing technology.

There have been several CRISPR clinical trials being conducted around the world. Most of them have been ex vivo - meaning that the patient's cells are taken, edited in the lab, and then infused back into the patient. However, a few in vivo clinical trials are starting - meaning that CRISPR will be directly administered into the patient's body. The in vivo trials are being done for Leber congenital amaurosis and now transthyretin amyloidosis.

The problem is CRISPR can have off-target effects. There are worries that if we directly inject CRISPR into our bodies, CRISPR may delete genes or insert sequences into random places. It might do more damage than good. So, I think it's important to have safety measures to deal with the potential issues related to these CRISPR therapies.
  • There are off-target effects, is there anything else that could make CRISPR therapies potentially unsafe?
  • How could we make CRISPR therapies safer (especially the in vivo ones)?
  • Should we stick to ex vivo CRISPR therapies instead of the in vivo CRISPR therapies?
What are your thoughts?

[1]Terns, Michael P., and Rebecca M. Terns. "CRISPR-based adaptive immune systems." Current opinion in microbiology 14.3 (2011): 321-327.

[2]Anderson, Keith R., et al. "CRISPR off-target analysis in genetically engineered rats and mice." Nature methods 15.7 (2018): 512-514.

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Creative contributions

Why gene editing precision and safety is not only determined by off-target effects.

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Manel Lladó Santaeularia
Manel Lladó Santaeularia Nov 18, 2020
This contribution will cover gene editing precision in the target site, which is an often-forgotten issue. Cas9 generates double-strand breaks (DSBs) that can be repaired following different pathways, including non-homologous end joining (NHEJ), homology-directed repair (HDR) and microhomology-mediated end joining (MMEJ).
  • NHEJ is the most common DSB repair pathway in higher eukaryotes, being active in all phases of the cell cycle, but it is most active during the G1 phase . In short, this system involves several DNA repair factors that, after recognition of the DSB, lead to filling of the gap using random nucleotides. This system is very efficient at quickly repairing the DSB but lacks precision, usually resulting in insertion or deletion (INDELs) of random bases around the cleavage site. This feature of NHEJ is exploited in some gene editing approaches in order to alter the reading frame and thus knock out expression of targeted genes.

  • HDR is mostly active in dividing cells, particularly in the G2 and S phases of the cell cycle. It is associated to DNA damage during chromosomal replication. After recognition of the DSB, HDR factors employ the sister chromatid as a template for strand invasion and homologous recombination followed by polymerase-mediated extension of the single-stranded DNA. This leads to precise repair of the DSB. This mechanism can be exploited in dividing cells by delivering a homologous recombination template flanked by homology arms. This allows correction of mutations as well as integration of exogenous sequences in targeted loci. This approach has been used extensively in vivo but can only be used in dividing cells and thus is very inefficient in most differentiated tissues of the organism.

  • MMEJ is considered to be a mix between NHEJ and HDR. It involves proteins from both pathways but is active through all cell cycle phases, which implies it can be independent of both NHEJ and HDR. In short, the DNA repair machinery can detect microhomologous sequences flanking the site of the DSB and facilitate recombination between those sequences, generating a small and precise deletion between the two microhomologous regions. If possible, this mechanism is quite efficient and leads to predictable INDELs.


The combination of the activity of these three repair mechanisms complicates the generation of intended gene editing outcomes, especially when performing in vivo gene editing. Even without considering HDR, in the case of wanting to generate a knock-out, MMEJ and NHEJ will generate several different kinds of INDELs, potentially causing different effects in the various edited cells. This can be studied by next-generation sequencing of DNA extracted from tissues, but unexpected results can always occur. Especially in cases where a precise modification (i.e. deletion of a particular number of bases) is necessary, this could be a very important limitation. While in vitro gene editing allows for in-depth characterization of modified clones in order to confirm that a generated INDEL leads to the desired phenotypical outcome, in vivo gene editing results can only be measured after it has already generated changes in the individual. Co-administration or up/down-regulation of factors involved in a particular repair pathway, with the objective of increasing a particular outcome of DNA repair, is being investigated but is difficult to apply in vivo.

However, in silico models that predict INDEL outcomes have been generated in the last years, which could greatly help improve the on-target precision of gene editing approaches.
Shen et al. described the machine learning model InDelphi, which was trained with over 2.000 different Cas9 gRNAs paired with DNA target sites . This model is able to predict gene editing outcomes taking into consideration the gRNA sequence as well as the sequences found around the cleavage site. It can accurately predict the genotypes and frequencies of 1 to 60-bp deletions and of 1bp insertions (the most common and almost only type of insertions) in human and mouse cell lines. While this model should be adapted to in vivo gene editing, it could be trained for every different tissue, which would account for the activity of the different DNA repair mechanisms in each tissue, as well as for the state of the chromatin, which can greatly impact the accessibility of repair factors to the DNA.
The most important thing about this model is that not only does it allow to predict the gene editing outcomes, but it proved that, in some sequence contexts, certain gene editing outcomes are significantly more frequent than others. This could allow selection of gRNAs that can generate more than 50% of gene editing outcomes with the same genotype, greatly increasing the on-target precision of gene editing outcomes and reducing the potential safety issues arising from unintended outcomes.

[1]Iyama, T. and D.M. Wilson, 3rd, DNA repair mechanisms in dividing and non-dividing cells. DNA Repair (Amst), 2013. 12(8): p. 620-36.

[2]Takata, M., et al., Homologous recombination and non-homologous end-joining pathways of DNA double-strand break repair have overlapping roles in the maintenance of chromosomal integrity in vertebrate cells. EMBO J, 1998. 17(18): p. 5497-508.

[3]Delacote, F. and B.S. Lopez, Importance of the cell cycle phase for the choice of the appropriate DSB repair pathway, for genome stability maintenance: the trans-S double-strand break repair model. Cell Cycle, 2008. 7(1): p. 33-8.

[4]Anguela, X.M., et al., Robust ZFN-mediated genome editing in adult hemophilic mice. Blood, 2013. 122(19): p. 3283-7.

[5]Li, H., et al., In vivo genome editing restores haemostasis in a mouse model of haemophilia. Nature, 2011. 475(7355): p. 217-21.

[6]Sharma, R., et al., In vivo genome editing of the albumin locus as a platform for protein replacement therapy. Blood, 2015. 126(15): p. 1777-84.

[7]McVey, M. and S.E. Lee, MMEJ repair of double-strand breaks (director's cut): deleted sequences and alternative endings. Trends Genet, 2008. 24(11): p. 529-38.

[8]Shen, M.W., et al., Predictable and precise template-free CRISPR editing of pathogenic variants. Nature, 2018. 563(7733): p. 646-651.

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Antonio Carusillo
Antonio Carusillo6 months ago
It is always nice to smel some CRISPR! May I suggest you also to try to integrate this other tool recently described? It is called MHcut (https://www.nature.com/articles/s41467-019-12829-8) and basically it looks for microhomologies around the target site and it helps to predict the outcome. They have used to generate disease-specific delition. And you can of course imagine to use it as a tool to generate indels whose result can be prediced with high precision. Regarding the co-administration of factors to influence the repair outcome I totally agree with you, they are challenging to be applied in vivo and in the clinics. But maybe we can envision different strategies as you may remember from the talk at the ESGCT in Barcelona 🎉
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Juran6 months ago
Hey! Thank you for a nice introduction of a really interesting part in CRISPR-Cas9 therapy that we missed.

As a strong believer in computers and the giant data sets being the future experimental systems, I support the solution that you proposed as a nice alternative to manage CRISPR off-targets.

As I read, two questions emerged:

- what percentage of "beneficial" gene editing outcome do you think is "acceptable"? Except the tissue specificity which you already mentioned, do you think we need an "ethical layer of decision-making" (e.g. 50% of good gene editing outcome is enough for ovarian cancer stage III, but it must be 80% for diabetes type 2, because there are other safer alternatives)?

- is this method already doable while treating states/diseases that include the extraction of the cells from blood, plasma or any other body liquid, modification of cells and reintroducing them back to a body? We could do these kind of in silico safety checks after the modification and before the reintroducement to the body.
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Manel Lladó Santaeularia
Manel Lladó Santaeularia6 months ago
Juran K. Fist of all, thanks for your comment. You raise some very interesting points.

-For the percentage of "beneficial" gene editing, that is a crucial point but also very difficult to establish. Different applications of gene editing could have different tresholds. For example, if you are trying to edit tumor cells, you want to be very sure that the INDELs you are generating are really having an effect on the cells. This would be crucial especially in this case in order to make sure you are eliminating as many cells as possible, considering how easily cancer cells can adapt to mutations. In a scenario where you want to generate a knock-out of a gene for an inherited disease, a high percentage of "beneficial" gene editing would clearly be preferable, but the treshold would probably be set at the minimum gene editing efficiency that allows you to improve the disease phenotype in the patients. Your point on safer alternatives is interesting, although one of the main advantages of gene editing is that it allows us to target diseases that previously we had no alternative how to treat. A paradigmatic example would be allele-specific gene editing to knock-out mutant alleles causing autosomal dominant diseases. Although miRNAs and other kinds of transcriptional regualtion have been tested, allele-specific gene editing seems to be the option with the most potential in , and thus setting this kind of treshold could indeed be quite important.

-This method would be useful also for ex vivo gene editing, where as you mentioned before, cells are extracted and modified after reimplantation. However, ex vivo gene editing protocols already contemplate in-depth characterization of the different clones generated, as well as selection of particular clone/s with the most desirable INDELs before amplification and reimplantation. Interestingly, in silico prediction could allow to choose a gene editing strategy with a higher turnout of cells edited in the desired way, thus probably reducing costs and improving the pipeline for production of edited cells. However, as far as I know this kind of machine learning would have to be adapted to each particular cell type, so I don't know if it would be readily available to use in, for example, hematopoietic stem cells. On the other hand, in vivo gene editing would benefit way more from this approach, since editing outcomes can only be analyzed after the editing has already occured in the tissue, meaning the potential for unintended gene editing outcomes is significantly higher. In that situation, having a high percentage of a particular, desirable outcome could be especially relevant in order to bring those approaches to the clinic.


Using a synthetic material-based delivery system

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Shubhankar Kulkarni
Shubhankar Kulkarni Oct 22, 2020
Different types of synthetic material (in the form of nanoparticles) can be used for in vivo delivery of genome-editing machinery. They are advantageous over viral vectors:
  1. They can be tailored for delivering different forms of the CRISPR–Cas9 system.
  2. Viral vectors are smaller and the gene-editing system needs to be divided and separately packed in two vectors.
  3. Cells may have a preexisting immunity against viral vectors. The immunocompatibility of synthetic materials can be improved by optimizing the size, shape, coating, and surface chemistry and charge.
  4. Retroviral and lentiviral vectors have a large genome of size. However, these vectors may integrate the transgene into the host genome, disrupting functional genes, and increasing off-target gene editing. Synthetic-material-mediated delivery can avoid such problems although random integration has been observed.
  5. Synthesis/ production of the synthetic material and its conversion to nanoparticles is more cost-effective and suitable for large-scale production than viral vectors.

[1]Tong, S., Moyo, B., Lee, C.M. et al. Engineered materials for in vivo delivery of genome-editing machinery. Nat Rev Mater 4, 726–737 (2019). https://doi.org/10.1038/s41578-019-0145-9

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Shubhankar Kulkarni
Shubhankar Kulkarni7 months ago
I read a bit further and found some issues with the synthetic material-delivery systems in the same paper. They have also mentioned the use of viral-non-viral-hybrid delivery methods (a combination of viral and non-viral delivery approaches) - a synthetic material delivers the Cas9 nuclease and a viral vector delivers the gRNAs and the DNA donor template.

This reduces the off-target effect issue of the viral vehicles and the bulkiness of the synthetic materials. Also, although the positively charged surface of the synthetic material allows for the easy entry in the nuclear space and the release of the cargo, it leads to extravasation of the delivery vehicles into the interstitial space from highly permeable vessels. Hence, repeated injections are required to observe the desired amount of the on-target effect.

Although the viral-non-viral-hybrid delivery methods have not been used extensively, they seem to be advantageous than both the viral and synthetic material methods.
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Jamila
Jamila 7 months ago
Shubhankar Kulkarni The delivery method is an essential aspect as it will determine the CRISPR therapy's safety and effectiveness. In many studies, viruses have been used to deliver CRISPR-Cas to the cells. [1]

✔️Viruses are quite efficient – infecting the host genome is a natural ability.
✖️ There could be viral integration into the host genome.
✖️ Immunogenicity and toxicity associated with viruses
✖️ Adeno-associated viruses (AAV) have a limited capacity for gRNAs and template DNA. [1]

Previously, nonviral delivery methods were less efficient than viral delivery methods [2] – but I think the consensus is that they are improving rapidly. So, viruses may not be the best way to deliver CRISPR in the future.

I've never heard of this viral-non-viral-hybrid delivery method you mention – it sounds fascinating that researchers can use viruses to deliver the guide RNAs and synthetic methods to deliver Cas proteins. It seems like the perfect combination that might overcome significant issues, like viral integration, capacity limits, immunogenicity, improve efficiency, etc.

References
1. Wang, Dan, Feng Zhang, and Guangping Gao. "CRISPR-Based therapeutic genome editing: Strategies and in vivo delivery by AAV vectors." Cell 181.1 (2020): 136-150.
2. Nayerossadat, Nouri, Talebi Maedeh, and Palizban Abas Ali. "Viral and nonviral delivery systems for gene delivery." Advanced biomedical research 1 (2012).
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Jamila
Jamila 7 months ago
I also read a paper where researchers developed virus-like nanoparticles (VLN) to deliver CRISPR components. VLN has a core-shell - mesoporous silica nanoparticle (MSN)-based core and a lipid shell. Apparently, this structure allows VLN to remain stable in the blood. This method also showed no signs of inducing cytotoxicity in cells. So, VLN can be used to co-deliver CRISPR and drugs to cancer cells. [1]

Reference
1. Liu, Qi, et al. "Virus-like nanoparticle as a co-delivery system to enhance efficacy of CRISPR/Cas9-based cancer immunotherapy." Biomaterials 258 (2020): 120275.

Controlling CRISPR edited cells via Auxotrophy

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Antonio Carusillo
Antonio Carusillo Nov 27, 2020
The topic of the session is CRISPR safety and as well explained in the contributions, most of the concerns regarding CRISPR and its usage come from the possibility to introduce undesired genetic modifications ( due to off-targets) whose consequences may escalate till a point at which cells may transform and acquire a cancer-like behaviour.

I - and others - have already mentioned strategies to make CRISPR preciser and safer overall. In this contribution, I would like to tackle the same issue but from a different angle.

Can we render CRISPR safer by making the modified cells safer?

This is a question that was explored by Porteus lab which is well known for its work in Sickle Cell Disease treatment based on which they have also opened a company.
The Porteus lab decided to exploit an "old" biological phenomenon because of which an organism -is unable to synthesize a particular organic compound required for its growth. Thus the organism needs to acquire it from the environment.
Porteus lab explored this concept .
In particular, they used CRISPR to target and knock out the uridine monophosphate synthetase enzyme complex, encoded by UMPS. The UMPS it is necessary for the catabolic pathways involved in Nucleic Acid Synthesis, Lipid metabolism and Carbohydrate metabolism.
Porteus lab could show that by using CRISPR to knock-out the UMPS gene, the edited cells become dependent on supplementation of Uridine. Cells whose culturing media was not supplemented with Uridine died while behaved normally - as the untreated - when Uridine was added. Very importantly this was not just proved in a petri dish ( a.k.a in vitro ) but they also engrafted UMPS KO cells in a mice model. If the mice received Uridine with the food, the UMPS KO cells could survive and persist in the mice, contrary withdrawal of Uridine resulted in the loss of the UMPS KO cells.
This is a very elegant strategy proves that it is possible to control cell persistence in vitro and in vivo by supplementing the required organic compound.
Such strategy presents some pros and cons.

The pros:
  • it allows controlling edited cells without the need to integrate bulky construct in the cell. Like a safety-switch as you would do for CART cells for example. So you can envision to perform along with the editing at the desired target also the KO of the UMPS gene. This means that if after re-infusing the edited cells in the patient if some side effects are observed ( maybe as a consequence of undetected off-targets ), withdrawing the organic compound will be enough to get rid of the cells
The cons:
  • if you infuse such cells in the patient and the patient benefits from them, you will need to keep these cells alive. How you do it? By supplying for example Uridine. Unfortunately, even if Uridine has been used in humans for the treatment of hereditary orotic aciduria and fluoropyrimidine toxicity, it is poorly absorbed and broken down in the liver. This can be circumvented by the administration as the prodrug uridine triacetate (UTA; also known as PN401), which has approval from the US Food and Drug Administration (FDA). So, you are maybe solving one problem but you are creating a dependency on the patients on constant Uridine supplement. At least until the edited cells have completed their job
  • to create cells holding both the UMPS gene KO and the modification in the gene of interest you will need to introduce two DNA double-stranded break (DSB) at the same time. This increases the probability to introduce gross DNA rearrangements like translocations or head-to-head chromosomic fusions .
In conclusion, we can see that besides making CRISPR safer we may also think about strategies to control the cells once edited. The auxotrophy approach is novel and elegant, but at the moment it is hard to see how it may be used in a clinical setting. We will probably need to wait until more papers will be based - maybe - on the strategy proposed by the Porteus lab.
In the meantime, can we think about other strategies to control the edited cells in a way that if they start acquiring a cancer-like phenotype we can discard them without endangering the patient's life?

[1]Wiebking, V., Patterson, J.O., Martin, R. et al. Metabolic engineering generates a transgene-free safety switch for cell therapy. Nat Biotechnol (2020). https://doi.org/10.1038/s41587-020-0580-6

[2]CRISPR-Cas9 Causes Chromosomal Instability and Rearrangements in Cancer Cell Lines, Detectable by Cytogenetic Methods Emily Rayner, Mary-Anne Durin, Rachael Thomas, Daniela Moralli, Sean M. O'Cathail, Ian Tomlinson, Catherine M. Green, and Annabelle Lewis The CRISPR Journal 2019 2:6, 406-416

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Jamila
Jamila 5 months ago
Antonio Carusillo Oo I like this idea! Thanks for sharing.

The system is excellent because it would provide a novel safety layer for gene-editing. Also, from the studies you mention, it does seem promising.

However, I wonder about the uridine associated side effects. A study found that long term usage of uridine caused an increase in weight and decreased insulin signalling in mice, making them pre-diabetic. [1] The prolonged use of UTA might not have the same problems, but it should be tested (if it hasn't been already). On the other hand, using a low dosage/having a relatively small number of edited cells in the body might offset any uridine associated side-effects.

Also, uridine is present in various foods like tomatoes, beer, broccoli, and much more. Would having foods that contain small amounts of uridine interfere with the system? Having a small quantity of uridine in food might not make a difference. However, if it did make a difference, then it would need to be carefully assessed - specific dietary requirements might be required with auxotrophy.

Reference
1. Urasaki, Yasuyo, Giuseppe Pizzorno, and Thuc T. Le. "Chronic uridine administration induces fatty liver and pre-diabetic conditions in mice." PLoS One 11.1 (2016): e0146994.

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Shubhankar Kulkarni
Shubhankar Kulkarni5 months ago
Antonio CarusilloNice strategy! How long do the edited cells survive on uridine? Have any studies done a long-term follow-up? - preferably in vivo but in vitro may also provide some insight. The artificial change brought about in the cells may have certain "side effects". I wonder if they have an impact on the lifespan of the cells.

Use Anti-CRISPRs as an antidote to CRISPR.

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Jamila
Jamila Oct 27, 2020
The CRISPR-Cas system is naturally occurring in bacteria. It's a way to fight off invading bacteriophages. Over time, bacteriophages have developed ways to overcome the CRISPR-Cas system. One of these methods includes using anti-CRISPRs (Acr). Acr proteins are produced by bacteriophage to inhibit the CRISPR-Cas system in the bacteria, thus leading to bacteriophage survival. There are many different types of Acr proteins available, and they inhibit the CRISPR-Cas system differently. The three main ways that Acr inhibit CRISPR-Cas are:
  1. They inhibit the guide RNA and Cas complex.
  2. They prevent Cas binding to the DNA.
  3. They prevent DNA cleavage by Cas.
Researchers have tested whether Acr can inhibit CRISPR-Cas when carrying out CRISPR editing. Shin and colleagues conducted CRISPR gene-editing on human cells and then added AcrIIA4. They found that adding Acr after gene-editing decreased the off-target effects of Cas9.

Furthermore, the high activity of CRISPR-Cas9 has been reported to induce toxicity. So, it is crucial to regulate the levels of CRISPR-Cas9. In one study, using AcrII4 and AcrII2 was able to reduce the toxicity of CRISPR-Cas editing in human hematopoietic stem cells.

Acr has been used in vivo too. Researchers delivered CRISPR-Cas9 and AcrIIC3N to adult mice for in vivo editing. The researchers did not find any toxicity associated with CRISPR editing.

Therefore, Acr proteins have the potential to be used in combination with CRISPR editing to make it safer. However, researchers need to conduct more in vivo experiments to confirm whether using acr is safe for humans and whether Acr can improve the safety profile of CRISPR editing, i.e. reduce the off-target effects and associated toxicity.

[1]Marino, Nicole D., et al. "Anti-CRISPR protein applications: natural brakes for CRISPR-Cas technologies." Nature Methods (2020): 1-9.

[2]Shin, Jiyung, et al. "Disabling Cas9 by an anti-CRISPR DNA mimic." Science advances 3.7 (2017): e1701620.

[3]Li, Chang, et al. "HDAd5/35++ adenovirus vector expressing anti-CRISPR peptides decreases CRISPR/Cas9 toxicity in human hematopoietic stem cells." Molecular Therapy-Methods & Clinical Development 9 (2018): 390-401.

[4]Lee, Jooyoung, et al. "Tissue-restricted genome editing in vivo specified by microRNA-repressible anti-CRISPR proteins." RNA 25.11 (2019): 1421-1431.

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Shubhankar Kulkarni
Shubhankar Kulkarni7 months ago
That is a good idea! I could think of a few questions after reading the suggestion:
1. Did the anti-CRISPRs decrease the ON-TARGET efficiency of CRISPR?
2. Adding both CRISPR and anti-CRISPR in a living system is like the movie "Aliens vs predators" - two space species fighting a war on Earth. Will our body suffer any collateral damage?
3. If these therapies become mainstream, can pathogens use either of these systems (CRISPR and anti-CRISPR) to launch an infection?

I know that the field is in its infancy and we may not know the answers but thought that these questions are important. Do we know the answers to any of these?
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Antonio Carusillo
Antonio Carusillo6 months ago
Shubhankar Kulkarni

1- from this more recent paper regarding Anti-CRISPR.Protein ( https://advances.sciencemag.org/content/6/6/eaay0187 ) they show that the fine tuning of such ACr ( either over-expressed or fused directly to the Cas9) has an effect of both the ON target and Off-target. However, the effect on the ON-target depends on the target site ( in same instances the ACr did not have any effect in others the effect was more evident ). While the Off-targets within a certain concentration of the ACr used are dramatically reduced compared to the ON-Target. So you may give up on a bit of ON-target editing in order to achieve a preciser Nuclease. Nevertheless we have to take in consideration that the " target-dependency " issue is always a present factor. So, regardless the strategy you should always evaluate 3 to 5 different guiding RNAs for the same target and seee which one ( in the absence of any "tricks" ) gives you the bast ON-target/ OFF-target ratio and from there you can start using all the sort of strategies that I may explain in upcoming contribution. I just need to find the time to write it ( and to make it short enough ).
2- I do not expect Any collateral damage ( neither toxicity was reported in the paper). The ACr acts on CRISPR system by preventing it to interact with the DNA. The CRISPR system acts on the DNA inducing a DSB ( the real threat, but this is somenthing I will explain in the contribution or contributions ). So no Alien vs Predator scenario. As the " preys " are different 😄
3- Anti-CRISPR are indeed found in bacteriophage ( https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6086933/ ) as system to fight back the CRISPR system ( is an endless Cold War down there ). However, this system was evolved cause bacteria were indeed expressing CRISPR. So from an evolutionary point of view, the ACr was a form of " adaptation". Human therapy - so far - does not include integrating a CRISPR constantly expressed by our cells. So, pathogenes won´t take ( theoretically ) any advantage from evolving anti-CRISPR proteins.
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Jamila
Jamila 6 months ago
Shubhankar Kulkarni Thanks for the questions.

1. In Shin’s study, they found that anti-CRISPRs seemed to reduce the off-target effects of CRISPR editing but didn’t heavily impact the on-target efficiency – this effect was seen in Shin’s study, at least! (https://advances.sciencemag.org/content/advances/3/7/e1701620.full.pdf)

2. Yes, that is very true. More research definitely needs to be conducted on the safety of adding CRISPR-Cas and anti-CRISPRs into the body together. The in vivo studies are pretty much in their infancy. However, in one study by Lee, the group added CRISPR and anti-CRISPRs simultaneously into adult mice – the researchers reported no toxicity at all. (https://pubmed.ncbi.nlm.nih.gov/31439808/) In Lee’s study, the mice were euthanized 7 days after receiving the vector containing CRISPR-Cas and Acr.

The safety of anti-CRISPRs and CRISPR-Cas combination needs to be evaluated for extended periods because, in humans, these CRISPR treatments will be long term, not just a 7-day treatment. In conclusion, more in vivo studies using CRISPR-Cas and anti-CRISPRs need to be carried out to validate the safety further.

3. I’m not sure if pathogens can use CRISPR/anti-CRISPRs to launch an infection, but I would think they wouldn’t be able to do that. The clinical trials using CRISPR in humans have shown to be safe with no immune responses. (https://www.nature.com/articles/d41586-020-00339-3) However, this might be different when using anti-CRISPRs too. I can’t say for sure, so I’d suggest determining the immune response when adding CRISPR and anti-CRISPR components in vivo.

Why CRISPR is already safe but it may still need improvements

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Antonio Carusillo
Antonio Carusillo Oct 31, 2020
This contribution will cover different aspects regarding CRISPR and safety, for some of them I will link some previous answers I discussed on Quora. This not for self-advertisement but to end up writing a 10 pages document. I don’t want just to send you references either, cause for some of the Quora answers I used 5 or 6 each. So, this could be easily a humble mini-review, thus I will do my best to keep it short ( but still comprehensible ). So let’s get started.


The point I will cover here are:
  1. why CRISPR is already safe enough
  2. strategies used to refine CRISPR precision
  3. why although precise, CRISPR has an inner “ defect “
  4. alternative to CRISPR to address the previous point.

If this introduction didn’t scare you off already, let’s get started.

1- CRISPR therapies are already safe enough

The session is about “how to make CRISPR therapies safer “. They are safe enough already. In fact, since the first paper about use CRISPR in mammalian cells ( less than 10 years ago ), we have already different clinical trials based on CRISPR going on, at this very moment .
As a reminder, these are mostly Phase I/II Clinical trials aiming mainly to test the safety aspect of CRISPR treatment, not its efficacy. Worth to be mentioned are two ex-vivo ( meaning that the patient cells are edited in a petri dish before being infused back to the patient ):

- Sickle Cell Disease treatment
- CAR-T cells engineered via CRISPR

Now those are ex-vivo edited cells. After ex-vivo editing, the treated cells undergo a full characterization looking in particular for off-targeting ( I will discuss this matter later ), chromosomal aberration and karyotype. If everything is fine, the cells can be infused to the patient.

However, this year also CRISPR was used for the first time in vivo. Which means that CRISPR was injected directly into the patient in the tissue of interest. In particular, CRISPR was packaged into a non-integrating viral vector ( known as Adeno Associated Virus ) and injected into patients affected by Leber Congenital Amaurosis Type X. A genetic disease which in the worst scenarios can lead to blindness. Contrary to ex-vivo editing here the a priori tests are the ones you can conduct in an animal model, ex-vivo human tissue but at the time of the therapy, there is no way to check the cells before you infuse back into the patient.

It is a one-way ticket. No turning back.

The study named BRILLIANCE has been conducted by Editas and Allergan. No disclosure so far regarding the results.

However, the fact that such a study was conducted kind of gives you the idea that CRISPR was considered safe enough to attempt an in vivo editing. A scenario in which, if things go on you can’t really rewind the tape and start over again.

So this may be already an answer for this thread. But I would like to go a bit more into details with part 2 regarding the refinement strategies you may use to make CRISPR preciser ( and safer, from a certain point of view ).

2- Refinement strategies

Even if CRISPR can be considered safe enough to be used in human clinical trials, there is still room for improvement.
To this end, different strategies are in place to render CRISPR preciser and safer.
When we speak about precision, we refer to the possibility that CRISPR may target undesired locations within the genome, introducing unwanted changes. Such changes are referred to as “ off-target”.

Some current strategies are:

- Short guide-RNAs : the guide-RNA is the component of the CRISPR system which directs the CRISPR effector ( the Cas9 nuclease ) towards its target. Studies have shown that shortening the gRNA may help to reduce the off-target activity of CRISPR. Even if you may lose some on-target efficiency as well.
- Two components dead CRISPR-Fok1 : another smart approach has been to use a catalytically inactive Cas9 which can only bind to the DNA but cannot cleave it. That’s why has been defined as dead-Cas9. Now to drive the cleavage a catalytic Fok1 domain ( the one used in previous gene-editing technologies like TALENs ) has been fused to the dead Cas9. Why? Cause the Fok1 domain can cleave only when paired with another 1 Fok1 domain. So scientists to cut the DNA deliver 2 dead-Cas9s which have to be on the same target at the same moment to cut the DNA. So the chances that at the off-target level, both dead-Cas9 are present at the same time on the same are fewer compared to a 1 component system. This results in fewer off-targets
- High-Fidelity Cas9: the two mentioned strategies are nice concepts, however, you have to give up on ON-target activity ( first case ) or you have to rely on more bulky components ( second case ). The current strategy of choice is the Hi-Fi Cas9 which is an engineered Cas9 nuclease mutated in specific residues to allow the Cas9 to have less off-target activity without losing on-target one. This because the mutations are creating a more “ relaxed “ status of the binding activity of the Cas9. This “relaxed status “ is strict enough for the Cas9 to bind firmly to its on-target but not enough to the off-target. Most of the clinical trials are using Hi-Fi-Cas9.

Those are the strategies that help CRISPR to be precise, there are also strategies to control its expression. But I will skip this as at the moment no therapeutic applications have been based on these. Just for you to know, some strategies allow controlling CRISPR activity in cells ( to decide when to activate it and when to turn it off ). If interested I will provide you with the references to this aspect in the comments.

3- CRISPR and DNA damage

So, as I said before CRISPR is already “safe-enough” to be used in human clinical trials and there are also strategies that may help to make it even safer. However, there is one aspect that still poses concerns ( from time to time ) and can’t be solved due to the way CRISPR it works. CRISPR to work needs to cut the DNA. While we think it is a very cool approach, the cells may think otherwise. A DNA cut known as a DNA Double-Stranded Break (DSBs )is the worst injury the cell can experience as we are damaging – on purpose – the core of the genetic information. DSBs are not well tolerated by the cells which have in place a proper response system called DNA-Damage Response (DDR ) and one of the main actors is P53, the known oncosuppressor. Well, in two papers it was found that CRISPR edited cells were dying out because of the DSB induced. In particular, the DSB was activating the P53 response which was apoptosis cell death. However, some cells survived and the scientists found out that these cells have mutations in P53. Meaning that likely, these cells had already inherent mutations for P53 and when edited they were “selected” for survival. This news resonated in the scientific community to a point that the CRISPR stocks dropped dramatically for a while. However, after these 2 papers no more P53 issues came up. Nevertheless, the warning was there: we should always check on the edited cells for P53 alterations. Something that you may do when editing cells ex-vivo but how can we do it when we go in vivo? As I said no more shreds of evidence were shown and before in-vivo editing a lot of tests are run, but the fact remains: a DSB is not a good thing for the cells.

4- CRISPR “ DSB-free “ alternatives

This brings me to the conclusions. We saw so far that CRISPR is used in clinical trials but that even if we manage to make it precise like a scalpel, we can’t make up for the fact that we are introducing a DSB.
These are were alternatives – still based on CRISPR – come into play.
You can find here the full discussion, while here I will just sum up the highlights.

I am talking about:

Base Editing: By using a nickase Cas9 ( able to introduce only a Single Strand Break ) fused to a cytosine base editors (CBEs) or adenine base editors (ABEs ) it is possible to introduce chemical modifications able to convert a C•G base pair into a T•A base pair, A•T base pair to a G•C base pair, respectively. Such an approach is being tested ( not clinical trials ) to address mutations characterized by single-nucleotide changes ( SNPs ) or to introduce stop codons to generate gene knock out. All of this without introducing a DSB. On this technology, Beam Therapeutics was founded by Dr David Liu.
Prime Editing: it can be considered an evolution of base editing in the terms that without introducing a DSB, howver contrary to base editing where you can only introduce point mutations, with Prime editing at the moment you can introduce up to 100 bp changes without the need to cut both strands the DNA. Dr David Liu also founded Prime Medicine for further translation of it to the clinics.

So is it done?

Not really, base editing and prime editing are still at their very early stages compared to CRISPR. Even if they are already showing in mouse models great promises, like curing Progeria, the use of these tools is hampered by their size. Of course, they can be used in vitro, but as we saw is in vivo that we need it the most. So, current challenges are how to delivery big payloads in vivo and still retain their full efficiency.

[1]Hirakawa MP, Krishnakumar R, Timlin JA, Carney JP, Butler KS. Gene editing and CRISPR in the clinic: current and future perspectives. Biosci Rep. 2020;40(4):BSR20200127. doi:10.1042/BSR20200127

[2]Zhang, J., Li, X., Neises, A. et al. Different Effects of sgRNA Length on CRISPR-mediated Gene Knockout Efficiency. Sci Rep 6, 28566 (2016). https://doi.org/10.1038/srep28566

[3]Havlicek S, Shen Y, Alpagu Y, et al. Re-engineered RNA-Guided FokI-Nucleases for Improved Genome Editing in Human Cells. Mol Ther. 2017;25(2):342-355. doi:10.1016/j.ymthe.2016.11.007

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Jamila
Jamila 6 months ago
Antonio Carusillo Thanks for the insight.

1. I completely agree with you. It must be safe to some extent because we have clinical trials using this technology right now. With ex vivo editing, researchers can easily check whether there are any abnormalities before infusing the cells back into the patient. However, in vivo editing hasn’t got this pre-check – so it seems a lot riskier! (https://www.researchgate.net/publication/282392356_Advances_in_therapeutic_CRISPRCas9_genome_editing) Of course, the researchers can monitor the patients during the trial with blood tests and other health checks, but the pre-check gives extra satisfaction.

2. The way I see it, CRISPR can be made safer with various strategies like the ones you mention. We can optimize the guide RNA and Cas to reduce the risk of off-target effects:
- The size and GC content of the guide RNA can alter the off-target effects.
- Some Cas variants may have fewer off-target effects. For example, Cas9 derived from Staphylococcus aureus (SaCas9) might have less off-target effects compared to Cas9 derived from Streptococcus pyogenes (SpCas9). Furthermore, SaCas9 is smaller than SpCas9, which is better for delivery purposes.
(https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7407193/)

3/4. Yeah, that is a problem. We could either use:
- “DSB free” methods
- regularly check the patient’s cells after in vivo editing

Maybe the P53 issue was limited to those two specific research papers – it could be to do with the methods used in their research paper. So, it might not be a problem.

Perhaps the DSB free CRISPR methods would be safer and better alternatives to traditional CRISPR gene editing. Base editing and prime editing are both outstanding DSB free methods. There is also RNA editing, epigenetic editing, and transcriptional regulation – all of which are also DSB free methods! (https://www.nature.com/articles/s41467-018-04252-2) It seems there is a vast array of CRISPR editing methods that could be used in the future – but of course, these methods are still in their infancy compared to CRISPR gene-editing.

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Antonio Carusillo
Antonio Carusillo6 months ago
Jamila Ahmed 1. Yes exactly what I meant. They took the chance to do it directly in vivo without any pre check beside previous experiments in animal model or in vitro. Meaning that for now CRISPR is considered safe enough to be used in a trial

2. yes diffeent Cas9 species may have different or lower off targets, you may also have different activities and off-target. You may also forced evolution to develop brand new Cas9 variants like the so called XCas9 (https://blog.addgene.org/xcas9-engineering-a-crispr-variant-with-pam-flexibility)

3. Yes RNA editing or Epigenome Editing are DSB free methods but they are also transient. Theoretically Epigenetic modifictions based on DNA methylation should be fixed or pretty stable, but no studies has be done so far showing 3 or 4 years monitoring DNA methylation status. RNA editing per se will be also transieent. So for such applications, you have to consider multiple administrations although you may find out that one or two administrations are enough.

Bioinformatics and sequencing to increase the safety of CRISPR

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J
Juran Oct 31, 2020

Background

In every biological experiment where you need to use certain oligonucleotides, such as the silencing of genes using transient transfection with siRNA or just a regular PCR, you need to make sure that your oligonucleotides are designed specifically enough to target only what you want. To check if your designed sequence is targeting the desired DNA/RNA region, you can use online blasting tools . These tools search the regions in a desired genome that match the sequence you entered. Results you get are the genes that share the sequence with your query sequence, with calculated percentages and probabilities.

With the on-going advancements in sequencing data analysis, the human genome is not so mystic anymore. Since the CRISPR method is usually used to change the genome sequences, no post-transcriptional or translational modifications should be disturbed (but it can also be checked).

The point of this

When designing the short-guide RNAs, if we take into account the available bioinformatics tools and run the sequence on many of them, that should strongly say that there will be no off-targets.

One proof that I can remember is the melting curve in the qPCR experiment, which is used to show how many different products are produced during the reaction. If one melting curve, at a specific temperature (e.g. 67 degrees Celsius) is seen, that means only one product was produced. If there are two or more curves with different temperatures, it means your sequence was not so specific. We can surely find more ways to prove the sgRNAs target specifically enough.

Ethical questions

Also, if a person is having a deadly disease, is it more ethical to let him die or to try to cure him with possible side-effects?
How many drugs on the market do you know that are proven to have 0 side-effects?

[1]https://www.ncbi.nlm.nih.gov/tools/primer-blast/index.cgi

[2]https://genome.ucsc.edu/cgi-bin/hgPcr

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Jamila
Jamila 6 months ago
For CRISPR editing, the guide RNA and target sequence must match, and the right PAM should be located near the target sequence. Bioinformatics and sequencing are quite useful tools that can make CRISPR safer. Specifically, these tools can help design guide RNAs with the lowest risk of off-target effects and detect off-target effects after CRISPR editing. [1]

Here's a list of some bioinformatics tools used in CRISPR:
- Cas-OFFinder
- uCRISPR
- CCTop
- CRISPRtarget
- CHOPCHOP
- E-CRISP [1]

Very interesting ethical questions. All the drugs on the market do have side-effects – but these drugs mainly target proteins, not the DNA itself. DNA-targeting drugs are relatively novel, so I guess there is an added worry that you might make considerable changes to the DNA, which you might not be able to reverse later down the line. So far, the CRISPR therapies have been well-tolerated and "safe" in the clinical trials.

I guess the overall aim is to make the CRISPR therapies as safe as possible by minimizing the risks of severe side effects – but yes, there will always be side effects, just like any drug.

Reference
1. Alkhnbashi, Omer S., et al. "CRISPR-Cas bioinformatics." Methods 172 (2020): 3-11.
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Antonio Carusillo
Antonio Carusillo6 months ago
Juran K. Bioinformatics is at the forefront of predicting and analysing CRISPR mediated off-targets. A possible tool is, for example, you can use CHOPCHOP (https://chopchop.cbu.uib.no/) to design a guide RNA with the highest efficiency and lowest off-target activity. However, these prediction tools have the problem to be not always " extendable " to all the conditions like cell type, nuclease delivery system, experimental time point ( days, weeks, month ). So they can return an "average" but there is no way for you, at the moment, to just pick one guide RNA and be sure that this one will be 100% effective and 100% precise. In the lab where I work, we usually select 3 to 5 gRNAs based on CHOPCHOP prediction and we try them all to see which one - in our conditions - perform the best.
At this point, bioinformatics comes handy for in cellula prediction like you would do using CIRCLE-Seq (https://www.nature.com/articles/nmeth.4278). This kind of analysis gives you the " worst-case scenario". By performing such an assay in vitro, you won´t take into account elements like chromatin status. Chromatin accessibility may affect the way CRISPR cuts at the on and off-target. Such assay assumes that CRISPR can equally access to both on-target and off-targets and so you can detect also off-targets that in vivo won´t be cut.
Now with these 2 first steps, you have selected the guide RNA with the best performance and is time to go in vivo ( human cells, mouse model etc ). After the editing, you can perform further bioinformatics pipelines that will tell you in your final model how many off-targets did you get (https://science.sciencemag.org/content/364/6437/286). Thereby the best recommendation is to apply this 3-layers system where you check: in silico, in vitro and in vivo for the precision of CRISPR. This is what is done in most of the labs willing to devise therapeutics based on gene editing :)
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Shubhankar Kulkarni
Shubhankar Kulkarni6 months ago
Juran K.This is thoughtful progress. I like the idea of utilizing the available technology than creating a new one. Although we can design the CRISP Repeats after carefully screening the genome, the genome is only about 1.2% of the DNA that our cells contain. There is a high probability that the remaining 98.2% "junk" may have the target sequences and those can be modified. Although such changes may usually not have a drastic effect, since these regions do not translate into proteins, the modification using CRISPR can create new start and stop codons in the "junk" part of our DNA leading to their translation into alien proteins. This will be also, probably, considered as an "off-target" effect of CRISPR.

Perhaps drug-induced CRISPR systems could make CRISPR safer.

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Jamila
Jamila Nov 13, 2020
CRISPR, being consistently expressed in our cells, may contribute to off-target effects and toxicity. CRISPR technology should come with an "off button" to make it safer.

Research groups have developed ways to control CRISPR. These include using light or drug-inducible CRISPR systems. The drug methods work in two ways: It can target Cas at the protein level or target CRISPR/Cas at the transcriptional level. Here are some drug-induced CRISPR systems: Tet, CIP/SPLIT systems, intein, HIT, etc.

An example of the tet system
If you use doxycycline, this will either activate or inhibit transcription of the CRISPR-Cas vector. So, you can easily turn CRISPR off or on using this approach.

Researchers compared the off-target effects of wild-type Cas9 to drug-inducible Cas9 (intein system). The researchers looked at three genes and 11 possible off-target sites and found that intein Cas systems had better specificity than wild-type Cas9.

Some of these drug-inducible methods aren't reversible (like the intein system). That would be one thing to look out for when developing a drug-inducible CRISPR system.

[1]Zhang, Jingfang, et al. "Drug inducible CRISPR/Cas systems." Computational and Structural Biotechnology Journal 17 (2019): 1171-1177.

[2]Davis, Kevin M., et al. "Small molecule–triggered Cas9 protein with improved genome-editing specificity." Nature chemical biology 11.5 (2015): 316-318.

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AP
Andrew Pan6 months ago
Hi Jamila

Just wanted to chip in here. While inducible CRISPR systems are fantastic from a research perspective for diseases modeling, for a therapeutic/clinical persspective I do not think it would be feasible. For one: it would necessitate each cell to carry CRISPR, which means that it would need to be imported at some early stage during embryonic development, or every single cell in the body or a signficiant portion of cells of interest will need to safely integrate CRISPR.

Secondly, I am currently working with doxycycline induced CRISPRi systems in iPSCs, and there is some level of basesline gene expression (leakiness) even without the presence of drug. Furthermore, from a therapeutic perspecitve, you would need to use the drug at a low enough dose to be safe but still effective, and delivered to the tissue of interest.
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Jamila
Jamila 6 months ago
Andrew Pan Hi Andrew.
Thanks for the comment 😊

I do wonder if all the cells would need to be targeted for us to see a therapeutic benefit, or would a small but highly targeted portion of cells be enough. As you say, the major hurdle is CRISPR being safely integrated into these cells.

Very nice work! The leakiness would be a problem; the gene expression should be tightly controlled with these drug-induced methods. I do wonder if some drug-inducible methods are better than the others like whether they also have leakiness. I guess there is a lot of refinement still required for drug-induced CRISPR systems, but I do see potential in them.

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Shubhankar Kulkarni
Shubhankar Kulkarni6 months ago
Hi Andrew Pan ! Is the leakiness due to any specific molecule (is it identified?) that is present in the system or any other parallel drug that needs to be introduced for reprogramming, culturing, or any other experimental procedure?

Temporary and tissue-specific expression of Cas9 could be crucial to improve its safety.

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Manel Lladó Santaeularia
Manel Lladó Santaeularia Nov 20, 2020
This contribution will cover different strategies that are being used in order to achieve temporary and tissue-specific Cas9 activity for in vivogene editing. This is important because, normally, we want to use Cas9 to target a single gene inside the cells of that tissue. Because of simple stochastic probability, having Cas9 be present in the cells for a long time increases the probability of Cas9 and its gRNA recognizing and binding to off-target sites, which also increases genomic instability. I will not address drug-induced Cas9 because it has already been very well described in another collaboration. Thus, I’ll focus on Cas9 transient delivery for in vivo gene editing,
The most widely used platform for delivering Cas9 in vivo are viral vectors. Due to their natural properties, viral vectors can only deliver DNA, which can then remain in the target cells as an episome or integrate in the cellular genome. This causes Cas9 expression to be stable. Additionally, the need for nuclear entry of the DNA in order to achieve expression of Cas9 can reduce efficiency of delivery by limiting the levels of Cas9 expression inside transduced cells. Instead, having a restricted time for Cas9 to act in the cell would, in theory, reduce the probability of off-target effects, but could have the disadvantage of lower on-target cleavage efficiency. How do we achieve this transient activity of Cas9? Several methods have been proposed:

-mRNA delivery: Delivering Cas9 mRNA to the target cells is a good strategy to achieve only temporary expression of Cas9. mRNA very rarely integrates in the cell DNA, and does not need to enter the nucleus the generate Cas9 expression. Additionally, expression is robust and peaks in the first hours after delivery and then persists for several days, but eventually disappears when the mRNA is degraded. mRNA degradation can, however, be a limitation for mRNA-based Cas9 delivery, as different tissues present different mRNA degradation systems and thus the half-life of the mRNA could be too short to achieve meaningful levels of Cas9 expression. To solve this, chemical modifications of the mRNA can increase its half-life, although there should be a fine tuning that allows Cas9 protein levels to be high enough to achieve efficient gene editing but decrease rapidly afterwards. However, in vivo mRNA delivery presents important complications that are being addressed by the scientific community: efficiency and specificity of delivery to particular tissues, potential degradation of mRNA by serum or tissue-specific nucleases, as well as the development of an immune response, limit both systemic and local delivery. To address these issues, several non-viral delivery systems have been developed. They consist of lipid or polymer nanoparticles that can carry the long Cas9 mRNA. For lipid nanoparticles, cationic and zwitterionic lipids have been used to deliver Cas9 mRNA together with chemically modified sgRNA to different tissues. However, improvements on the composition of those nanoparticles could increase efficiency and tissue specificity of mRNA delivery after systemic injection. Nanopolymers work in a similar way and could be easier to modify to target them to specific tissues, as has been done to target xenograft tumors.

-Ribonucleoprotein delivery: Cas9 and its gRNA bind to form a stable ribonucleoprotein (RNP) complex that can be introduced in nanoparticles and delivered in vivo. This approach achieves a very strong and quick efficiency of gene editing in the targeted cells, and the RNP gets rapidly degraded. This leads to very low off-target cleavage. However, in vivo RNP delivery also presents several limitations: first, Cas9 proteins are derived from bacterial species that regularly infect humans, and thus a large part of the population presents preexisting antibodies against Cas9. This could drastically reduce the efficiency of RNP delivery. Additionally, after the RNP has entered the target cells, degradation of the RNP by the proteasome can lead to exposure of Cas9 peptides to the immune system through the major histocompatibility complex, thus initiating an immune response against Cas9 that could lead to elimination of the modified cells. Even after considering immunity-related limitations, systemic delivery of RNP nanoparticles presents another very important limitation: differently to mRNA delivery, where translation of the RNP implies signal amplification, RNP delivery depends on getting enough Cas9 inside the cells so that gene editing is efficient. This is problematic due to the cost of production and safe isolation of the RNP complex, as well as the doses needed to achieve efficient systemic delivery to a relevant number of target cells. To improve on this, modification of nanoparticles to render them more tissue-specific and more stable, as well as cheaper to produce, is the next crucial step.
-Self-inactivating Cas9: An interesting method that has been proposed by some authors is the generation of a self-inactivating Cas9 that, after being expressed, would cleave the DNA sequence of its own gene and thus block the expression of more Cas9. The KamiCas9 design targeted Cas9 to the ATG of its own gene, thus leading to disruption of this ATG. This proved reduction of off-target effects related to transient expression of Cas9 delivered with a lentiviral vector. More recently, a similar system has been described to use Cas9 to cleave the AAV genome, leading to reduction of Cas9 expression. However, these systems don’t take into consideration the potential for non-homologous end joining (NHEJ)-mediated integration of fragments of the viral genome into the target site that was shown by Suzuki et al. after cleavage of both the AAV viral vector and the target site. This potential integration represents a major safety concern for the use of this technology.


Since all these different methods have clear advantages and disadvantages, further development is necessary to improve their safety and efficacy. Moreover, none of these methods has proven to be substantially better than the others, and thus different methods may be preferable depending on the target tissue and the therapeutic strategy to be used. The next years will for sure see the rise of several new approaches that may find the key to solving this issue.

[1]Zhang, H.X., Y. Zhang, and H. Yin, Genome Editing with mRNA Encoding ZFN, TALEN, and Cas9. Mol Ther, 2019. 27(4): p. 735-746.

[2]Zuris, J.A., et al., Cationic lipid-mediated delivery of proteins enables efficient protein-based genome editing in vitro and in vivo. Nat Biotechnol, 2015. 33(1): p. 73-80.

[3]Merienne, N., et al., The Self-Inactivating KamiCas9 System for the Editing of CNS Disease Genes. Cell Rep, 2017. 20(12): p. 2980-2991.

[4]Suzuki, K., et al., In vivo genome editing via CRISPR/Cas9 mediated homology-independent targeted integration. Nature, 2016. 540(7631): p. 144-149.

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Jamila
Jamila 6 months ago
Hello, thank you for your contribution.

Non-viral methods can use receptor-mediated delivery methods to deliver CRISPR components too. This is whereby cell-specific ligands are bound to the extracellular vesicles that carry the CRISPR components. The ligand will only bind to cell-specific receptors. Once the ligand and receptor bind, the extracellular vesicle can be internalised via endocytosis. [1]

Doudna reported that liver-specific CRISPR editing could be conducted. Doudna's lab targeted the asialoglycoprotein receptor – a receptor that is nearly exclusive to liver cells. [2] Other researchers have also used receptor-mediated delivery methods with great success. [2-4]

Problems with the receptor-mediated delivery method include:
- Finding a target that is specific for every tissue/cell might be hard to do because the target must be exclusive to that particular tissue/cell type.
- When you block a specific receptor with this delivery method, what are the consequences of this? Are we blocking a receptor that is required for our daily biological functions!? Wouldn't this have harmful side-effects on us? [4]

I never heard of a self-inactivating Cas9 before this. That's so interesting. Thanks for sharing! It would be great to find other ways to develop self-inactivating Cas – methods that hopefully don't cause viral integration.

It would be interesting to see which of the delivery methods you mention are better than the other – I guess with time we will find out.
Being able to target specific tissues with CRISPR-Cas would hopefully reduce the off-target effects elsewhere in the body. [1] Therefore, developing tissue-specific delivery methods is crucial in making CRISPR-Cas technologies safer and more effective.

References
1. Shalaby, Karim, Mustapha Aouida, and Omar El-Agnaf. "Tissue-Specific Delivery of CRISPR Therapeutics: Strategies and Mechanisms of Non-Viral Vectors." International Journal of Molecular Sciences 21.19 (2020): 7353.
2. Rouet, Romain, et al. "Receptor-mediated delivery of CRISPR-Cas9 endonuclease for cell-type-specific gene editing." Journal of the American Chemical Society 140.21 (2018): 6596-6603.
3. Large, Danielle E., et al. "Advances in receptor‐mediated, tumor‐targeted drug delivery." Advanced Therapeutics 2.1 (2019): 1800091.
4. Rouet, Romain, and Daniel Christ. "Efficient Intracellular Delivery of CRISPR-Cas Ribonucleoproteins through Receptor Mediated Endocytosis." ACS chemical biology 14.3 (2019): 554-561.
5. Muro, Silvia. "Challenges in design and characterization of ligand-targeted drug delivery systems." Journal of Controlled Release 164.2 (2012): 125-137.

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AP
Andrew Pan6 months ago
Jamila Another problem I would like to point out with any drug delivery system is that currently, most of the nanoparticle systems in question face significant systemic barriers. For example, much of the injected doses is typically trapped in the liver and spleen (which does incidentally simplify gene therapies targeted to these organs). For example, Alnylams Patisiran LNP (lipid nanoparticle) system works by targeting hepatocytes and is one of the first if not the first LNP based gene therapy vectors available now.

This review by Saltzmans group at Yale explains why receptor-mediated targeting is not necessarily as promising as previously thought (https://www.sciencedirect.com/science/article/pii/S147149141830100X), but in brief, it is because the "attractive forces" between the receptor and its ligand operate on very short range.
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Manel Lladó Santaeularia
Manel Lladó Santaeularia6 months ago
Jamila You raise some very interesting points about receptor-mediated tissue specificity for Cas9 delivery. However, as both you and Andrew have mentioned, there are several limitations that still have to be addressed. I expect that, in the next years, this field will greatly evolve.

However I don't believe there is going to be one single strategy that demonstrates to be exceedingly better than the others for all tissues. It may very well be that different strategies are better for different tissues or different types of deliveries. It's important that, although most tissues can be reached with intravenous administration, this presents important limitations for mRNA delivery, which makes me think that ribonucleoprotein delivery will be more promising. However, this is very dependant on both liver retention (as Andrew very importantly states) and tissue targeting. In some cases, maybe local administration (e.g. electroporation) would be preferable? In those cases, delivery to a sizeable portion of the tissue could be a limitation, as well as potential clinical applicability.

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