In Vivo High-Throughput Screen Finds Lipid Nanoparticles (LNPs) for Cell-Specific Delivery of Modified mRNA for CRISPR 

  • Multiplexed Screening of Hundreds of LNPs Compositions In Vivo 
  • Barcodes and Sequencing Identify Cell-Type Tropisms
  • In Vivo CRISPR Gene-Editing in Endothelial Cells Provides Proof-of-Concept 

3D illustration of an empty LNP (aka liposome).

In some of our previous blogs, we have featured a variety of publications on lipid nanoparticles (LNPs; aka liposomes) for delivery of therapeutic nucleic acids such as siRNA or mRNA. These and other studies generally involve the empirical screening of a limited number of LNP formulations in vitro, in order to identify the best performing LNP composition. By contrast, in what I view as a “breakthrough article,” Sago et al. have recently reported a high-throughput screening methodology that can identify LNPs with novel cellular tropisms in vivo. They demonstrate this by utilizing LNPs for delivery of functional mRNAs to endothelial cells, a therapeutically important cell-type group that is distinct from hepatocytes, which generally take up LNPs.

Background  

The biggest challenge for realizing the therapeutic potential of nucleic acids is safe and effective delivery vehicles for human use. Among nucleic acid compositions comprised of lipids and lipid analogs, LNPs have been extensively studied in vitro and in vivo. Nonetheless, rational design of these vehicles is lacking. Consequently, empirical screening is used, but due to its relatively slow and laborious nature, this approach limits the number of formulations tested, i.e. screened.

Chen et al. have previously investigated this problematic bottleneck by developing a 24-channel microfluidic method that allows for parallel preparation of short-interfering RNA (siRNA)-containing LNPs (aka lipoplexes) for a large number of components such as cholesterol, poly(ethylene glycol)-conjugated lipids, and phospholipids. This method allows researchers to obtain formulations with the desired biophysical and pharmacokinetic attributes, as shown here.

Adapted with permission from Chen et al. J. Am. Chem. Soc. 134, 6948–6951. Copyright 2012 American Chemical Society.

Chen et al.’s approach has now been extended and enhanced by Sago et al. as a system named Fast Identification of Nanoparticle Delivery (FIND). This method measures cytosolic mRNA delivery by hundreds of LNPs in vivo to any combination of cell types. In the following sections, I will briefly outline various applications of FIND, and how this system was able to quantify how >250 LNPs functionally delivered mRNA to multiple cell types in vivo. Through these processes, two formulations that deliver RNA to endothelial cells were identified. These are especially interesting, as dysfunctional endothelium causes more diseases than any other cell type.

FIND Barcoding, Multiplexing, and Screening Validation

Using the high-throughput microfluidics of Chen et al. mentioned above, Sago et al. co-formulated LNPs with Cre mRNA and a unique DNA barcode. Design details for these DNA barcodes are described elsewhere by Dahlman et al. In brief, as shown here, LNP-1 with composition 1 carried DNA barcode 1 and Cre mRNA; LNP-2 with composition 2 carried DNA barcode 2 and Cre mRNA; etc. up to the Nth particle, LNP-N with composition N that carried DNA barcode N and Cre mRNA. Note that, in the exemplary 58-nt barcode structure shown here, N represents A/G/C/T and red indicates an internucleotide phosphorothioate (PS) linkage. 

Adapted from Sago et al. Proc. Natl. Acad. Sci. 115, E9944-E9952 (2018) with permission from the National Academy of Sciences.

LNPs that met size and stability criteria were screened in vitro, which led to 112 LNPs for further evaluation. Of these, 71 were pooled together and administered to previously described Cre-reporter mice, wherein Cre-reporter cells fluoresce if Cre mRNA is translated into Cre protein, which then translocates to the nucleus and edits target DNA. Cell types of interest were separated by fluorescence-activated cell sorting (FACS) in order to generate an in vivo delivery heatmap, as shown here. Unformulated (aka naked) DNA barcode, which should not be delivered as efficiently as DNA barcodes in LNPs, was included as a negative control. Mice were i.v. injected with the pooled LNPs at a 1.5 mg/kg total mRNA dose (average dose 0.021 mg/kg per particle). Three days later, lung and kidney endothelial cells were FACS-selected.

Adapted from Sago et al. Proc. Natl. Acad. Sci. 115, E9944-E9952 (2018) with permission from the National Academy of Sciences.

Due to the technical complexity of these and many additional experiments, interested readers can consult Sago et al. for details. The remainder of this blog will focus on proof-of-concept results for CRISPR-mediated gene editing using LNP delivery of Cas9 mRNA and single-guide RNA (sgRNA) in endothelial cells. 

FIND-Derived LNPs for In Vivo Gene-Editing in Endothelial Cells

5moU 5’-triphosphate. Taken from TriLink BioTechnologies

After completing the aforementioned in vitro and in vivo validation experiments, Sago et al. further optimized two selected LNPs named 7C2 and 7C3. Mice constitutively expressing SpCas9 were injected three times with 7C2 or 7C3 carrying 1.5 mg/kg of two chemically modified sgRNAs each, with three PS linkages on the 5’ and 3’ ends (custom synthesis by TriLink BioTechnologies) targeting ICAM2 (sgICAM2ab) (1:1 mass ratio). Separately, C57BL/6J mice were injected two times with 2 mg/kg with 7C2 and 7C3 formulated to carry both SpCas9 mRNA and sgICAM2ab at a 3:1 mass ratio. I am very pleased to say that sgICAM2ab and SpCas9 mRNA were obtained from TriLink, with the SpCas9 mRNA being comprised of CleanCap™, as well as 5moU-modified ribonucleotides shown here.

Five days after the last injection, tissues of interest were isolated, and ICAM2 protein expression was measured concurrently as ∼20,000 cells were sorted. Sago et al. state two interesting observations: first, in contrast to accompanying small RNA delivery experiments, 7C3 outperformed 7C2, and second, splenic endothelial cells were targeted efficiently (Panel B). Given that this degree of selectivity has not yet been reported to date, Sago et al. repeated the experiment using additional groups of mice over the course of several months. The 7C3 delivered Cre mRNA to splenic endothelial cells in every experiment (Panel C).

Adapted from Sago et al. Proc. Natl. Acad. Sci. 115, E9944-E9952 (2018) with permission from the National Academy of Sciences.

According to Sago et al., the differences in delivery between 7C2 and 7C3 provide interesting preliminary evidence that suggests changing the composition of helper lipids added to a given ionizable amine can alter tropism, i.e. the type of tissue/cells that take up LNPs loaded with nucleic acids.

Concluding Comments

FIND is a high-throughput method for quantification of functional mRNA delivery mediated by LNPs. Sago et al. state that the distinction between biodistribution and functional delivery is significant: >96% of delivered RNA does not escape the endosome, and endosomal escape may vary with cell type or disease state, making it hard to predict functional delivery using biodistribution. Notably, Cre mRNA must be translated in the cytoplasm, translocate into the nucleus, and edit target DNA. 

Although the aforementioned studies focused on endothelial cells, Sago et al. state that “FIND is agnostic to cell type; [they] envision FIND studies that identify LNPs targeting other cells.” This agnosticism also applies to the type of nucleic acid being delivered, as depicted here for an antisense oligonucleotide (ASO), siRNA, mRNA, and sgRNA.

Taken with permission from Greineder et al. Bioconjugate Chem. 29, 56–66. Copyright 2017 American Chemical Society.

Given that on- and off-target cells can be isolated from the same mouse, FIND may also be used to identify (or iteratively evolve) LNPs with high therapeutic windows, i.e. effective doses vs. toxic doses. To further increase the ratio of splenic endothelial cell to hepatocyte delivery, one may decorate LNPs with reported endothelial-cell-targeting ligands.

Rather than paraphrase Sago et al., here is their long-term vision for the future for FIND:

“More generally, this ability to study how hundreds of LNPs target combinations of cells in vivo may elucidate relationships between LNP structure and in vivo functional delivery. In this case, FIND enabled us to quickly identify two lead LNPs. Here we delivered sgRNAs targeting the inflammation-related gene ICAM-2. However, the long-term utility of FIND will be defined by its ability to identify nanoparticles for in vivo gene editing more efficiently than the current gold standard, which is in vitro screening.”

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I agree with this opinion, and also believe that FIND represents yet another example of what I refer to as ‘seek, and ye shall find’ in a previous blog.

As usual, your comments are welcomed.

Tagging Modified mRNA with TAG (Transglycosylation at Guanosine)

  • Modified mRNA (Mod-RNA) for Therapeutics Drives Development of New Technologies
  • Among These Is Tagging Mod-RNA for Visualization In Vivo
  • Tagging Mod-RNA with TAG Earns Neal Devaraj the 2018 Blavatnik National Award in Chemistry

Back in 2013, I wrote a blog titled Modified mRNA Mania, which drew attention to the then nascent development of an entirely new field of therapeutics, based on the use of biosynthetic modified mRNA (mod-RNA). I also wrote a more recent blog about this still trending field, titled Deluge of mRNA Delivery Publications, featuring a chart of supporting publication data to emphasize the exponentially growing efforts to improve delivery.

Prof. Neal Devaraj. Credit Neal Devaral. With permission

The present blog in this ongoing series on mod-RNA focuses on in vivo visualization of mod-RNA by use of a very clever enzyme-mediated labeling method. Importantly, this labeling procedure can be applied to any biosynthetic mod-RNA of interest. The method was invented by Neal Devaraj, an Associate Professor of Chemistry and Biochemistry at the University of California, San Diego. Because of his work, Prof. Devaraj recently received the 2018 Blavatnik National Award in Chemistry, which you can read about at the end of this blog.

Backstory for Tagging Mod-RNA

In vivo expression of therapeutic proteins encoded in exogenous mRNA provides several advantages compared to delivery of the corresponding complementary DNA. Delivering mRNA is easier because it only needs to enter the cytoplasm, rather than the nucleus, to be functional (depicted here in scheme B). This avoids complications from transcription regulation machinery. Additionally, mRNA does not permanently alter the genome like genomically integrated DNA does (depicted here in scheme A), therefore avoiding permanent and potentially lethal changes.

Taken from Avci-Adali et al. J. Biol. Eng. 2014, 8, 8. Copyright © BioMed Central Ltd. With permission.

In order to overcome the intrinsically unstable and transient nature of natural RNA, the incorporation of modified nucleotides has been—and continues to be—explored to harness RNA as a therapeutic agent. Multiple studies have focused on providing an increase in serum stability, a less active immune response, and an increase in translational capacity, as reviewed elsewhere.

5-moUTP lithium salt. Taken from TriLink BioTechnologies

Many types of modified nucleobases can be incorporated into mRNA transcripts by substituting natural nucleotide triphosphates with one or more modified nucleotide triphosphates during enzymatic synthesis with an RNA polymerase. TriLink BioTechnologies R&D group has synthesized and screened numerous modified nucleotide triphosphates, and 5-methoxyuridine (5moU) has been shown to be especially useful. Interested readers can learn more about mod-RNA at this link to a video presentation in 2018 by Dr. Anton McCaffrey, Senior Director of Emerging Science and Innovation at TriLink.

In order to fully exploit the therapeutic potential of mod-RNAs, novel strategies for the safe and effective “decoration” or tagging of in vitro transcribed RNA, as well as functional moieties or fluorophores to allow for visualization, must be developed. A convenient bioconjugation method that is capable of appending targeting molecules or fluorophores to therapeutic Mod-RNAs would enable new modalities of highly specific uptake of RNA through endocytic pathways, as well as visualization.

Tagging Mod-RNA with TAG

To address the need for a specific and convenient bioconjugation method for mod-RNA, Devaraj and his students envisioned decorating mod-RNA by use of an RNA modifying enzyme. In their 2018 publication in Molecular Pharmaceutics, it was noted that more than 100 RNA post-transcriptional modifications have been reported to date, with a large majority of these modifications occurring on transfer RNAs (tRNAs). Among these, bacterial tRNA guanine transglycosylases (TGTs) have been extensively studied. As depicted here, during the transglycosylation reaction, the N−C glycosidic linkage of a key guanine at the wobble position of the anticodon loop is cleaved, and a 7-deazaguanine derivative named preQ1 (R = H) is substituted.

Taken from Devaraj and coworkers J. Am. Chem. Soc. 2015, 137, 40, 12756-12759. Copyright © American Chemical Society. With permission

Importantly, the RNA-bound TGT crystal structure previously reported by others suggested to Devaraj that there might be enough room to chemically modify the natural preQ1 (R = H) substrate and thus repurpose the enzyme to covalently append a larger R group, such as a fluorophore or affinity tag. This concept was initially reduced to practice in a preliminary communication, wherein the minimal 17-nucleotide hairpin sequence shown above, which was named TAG, was genetically engineered into the 3′-UTR of a full mRNA transcript coding for the red fluorescent protein mCherry (mCherry-TAG). When treated with bacterial TGT and a preQ1 derivative with R = dye, fluorescence-based gel analysis confirmed the formation of the expected mCherry-TAG-dye conjugate.

In a recent Molecular Pharmaceutics publication, Devaraj and his students extended this E. coli TGT activity to recognize TAG in mCherry mod-RNA comprised of the chemically modified nucleobases 5-methylcytosine (5mC) or pseudouracil (Ψ), as well as doubly substituted 5mC + Ψ mod-RNA. The approach depicted here is a two-step process involving RNA-TAG covalent conjugation followed by tetrazine biorthogonal labeling.

Taken from Devaraj and coworkers Mol. Pharmaceutics 2018, 15, 3, 737-742. Copyright © American Chemical Society. With permission

In vitro transcription (IVT) reactions for mCherry-TAG mod-mRNA transcripts were performed with both a partial (25%) and a complete (100%) replacement of the natural bases with 5mC and Ψ by use of the corresponding modified triphosphates, which I’m pleased to say were obtained from TriLink. A cap analog was added to the 5’ end and, after IVT, the 3’ end was polyadenylated to furnish mature mRNAs capable of being translated into mCherry-TAG upon transfection into mammalian cells.

Although mRNA-TAG has been established to efficiently and directly incorporate small molecule moieties, Devaraj and coworkers state that they envisioned a two-step labeling approach of mod-mRNA such that the degree of labeling could be more concretely proven. Additionally, this methodology would also allow a diverse array of targeting molecules, irrespective of size or class, to be easily conjugated and evaluated without the need for multiple syntheses of preQ1 derivatives or the need to quantify each substrate’s efficiency of incorporation.

As depicted here, appending a bioorthogonal tetrazine moiety to mod-mRNA transcripts (red) can be followed by coupling a trans-cyclooctene (TCO)-functionalized fluorescent probe or affinity agent (blue) using previously established, robust tetrazine-ligation chemistry, which is reviewed elsewhere.

Taken from Chaudhuri et al. Bioconjugate Chem. 2017, 28, 4, 918-922. Copyright © American Chemical Society. With permission

The mCherry mod-mRNAs were treated with the TGT enzyme at 37 °C for 4 h to first access tetrazine-labeled mCherry mod-mRNAs. The purified tetrazine-conjugated mod-RNA was subsequently conjugated to a TCO-functionalized sulfo-Cy5 fluorescent probe via tetrazine ligation by incubating for 4 h at 37 °C. The purified RNAs were subjected to polyacrylamide gel electrophoresis and imaged. As shown here, the degree of labeling (DOL) was determined to be ~95% for unmodified transcript, ~84% for 5mC transcript, ~66% for Ψ modified transcript, and ~61% for both Ψ and 5mC modified transcript.

Taken from Devaraj and coworkers Mol. Pharmaceutics 2018, 15, 3, 737-742. Copyright © American Chemical Society. With permission

Devaraj and coworkers hypothesize that the slightly lowered efficiency in labeling of the all 5mC-mod-mRNAs in comparison to unmodified mRNA could be due to variations in the secondary structure of the RNA-TAG hairpin, which may hamper enzyme substrate recognition. More interestingly, and with reference to the TAG hairpin shown above, they further speculate that the decrease in DOL for the Ψ containing mod-mRNA transcripts most likely arises from disruption of key interactions between the TGT enzyme and flanking uridine residues on either side of the exchanged guanine when substituted for the modified base Ψ.

Conclusion

Devaraj and coworkers concluded that they have developed a relatively simple methodology to covalently label mod-mRNAs with a modular two-step approach that can incorporate small molecules such as imaging agents, affinity handles, targeting agents, and drug conjugates using RNA-TAG. They also concluded that this approach provides the possibility of diverse decoration of therapeutically relevant mod-RNAs, without the limitation of length characteristic of synthetic RNA strategies. Finally, they expressed belief that “the RNA-TAG technology could greatly expand the arsenal of therapeutic RNAs by way of conjugation to a variety of functional decorations relevant to further tuning the emerging modality of RNA as a therapeutic class.”

I fully agree with these conclusions, and welcome your comments, as usual.

Footnote

This past June, The Blavatnik Family Foundation and the New York Academy of Sciences announced the 2018 Laureates of the Blavatnik National Awards for Young Scientists, who will each receive $250,000: the largest unrestricted scientific prize offered to America’s most promising faculty-level scientific researchers 42 years of age and younger. Nominated by 146 research institutions across 42 states, the 286 nominees were narrowed to a pool of 31 Blavatnik National Finalists. From this pool of Finalists, a distinguished scientific jury chose three outstanding Laureates, one in each of the Awards’ scientific disciplinary categories: Life Sciences, Physical Sciences & Engineering, and Chemistry.

Life Sciences: Janelle Ayres, PhD, of the Salk Institute for Biological Studies, for her pioneering research in immunology and the study of how bacteria interact with humans. Dr. Ayres’s work is revolutionizing our understanding of host-pathogen interactions and has the potential to solve one of the greatest current public health threats: anti-microbial resistance.

Physical Sciences & Engineering: Sergei V. Kalinin, PhD, of Oak Ridge National Laboratory, for creating novel techniques to study, measure, and control the functionality of nanomaterials at the atomic and nanoscale levels. Dr. Kalinin’s work manipulating individual atoms has the potential to enable scientists to create new classes of materials by assembling matter, atom-by-atom.

Chemistry: Neal K. Devaraj, PhD, of the University of California, San Diego, for his transformative work on the synthesis of artificial cells and membranes, thus creating an exciting new field of research that aims to address one of the great challenges in synthetic biology. Dr. Devaraj has made several game-changing discoveries, and has pioneered the development of new methods for labeling biological molecules, which have already been adopted by researchers globally.

Here is a link to all of Neal Devaraj’s publications currently indexed in PubMed.

Deluge of mRNA Delivery Publications

  • Strong Interest in mRNA Therapeutics Drives Increased Numbers of Delivery Publications
  • Novel Charge-Altering Releasable Transporters (CARTs) Undergo “Self-Immolation”
  • CARTs Outperform Widely Used Lipofectamine In Vitro and Enable In Vivo Delivery

Devotees of this blog may recall my past post in 2013 titled Modified mRNA Mania, which intentionally used the word “mania” to provoke reading about the trending topic on base-modified mRNA as therapeutic agents. My metrics for this mania were a flurry of scientific publications, patent applications staking out intellectual property, and massive investments by venture capitalists and established pharma companies in mRNA therapeutics startups.

As with antisense, siRNA, and antagomir RNA drugs, efficient delivery is widely recognized as a critical technical challenge to overcome. And, not surprisingly, past lipid-based approaches of various sorts are being reinvestigated for repurposing for mRNA delivery.

The focus of the present blog is a new strategy for mRNA delivery developed by a team of collaborators at Stanford University. Although I’ve chosen to highlight this report by McKinlay et al. in prestigious Proc. Natl. Acad. Sci., a search of PubMed for publications indexed to “mRNA delivery” in the title and/or abstract for the period 2005 to 2017 gave articles that can be perused at this link. The graph shown below supports my characterization of this level of activity as “deluge”-like in that there are more than 100 publications, mostly in the last few years, with 40 to 50 more during 2018, by my estimate.

Challenges for mRNA Delivery

Simply stated, the key challenge associated with the use of therapeutic mRNA is an inability to efficiently deliver functionally intact mRNA into cells. Like all nucleic acid-based drugs, mRNA is a macromolecular polyanion and thus it does not readily cross nonpolar cellular and tissue barriers. Moreover, it is also susceptible to rapid degradation by nucleases and ideally it should be protected during the delivery process, even though some success has been reported using intradermal injection of “naked” unmodified mRNA. Finally, after cell entry, rapid release of mRNA in the cytosol and appropriate association with the protein synthesis apparatus is required for translation.

Each of these is a potential point of failure for functional mRNA delivery. In addition to the challenges associated with complexation, protection, delivery, and release, an ideal delivery system would also need to be synthetically accessible, readily tuned for optimal efficacy, and safe.

Charge-Altering Releasable Transporters (CARTs)

McKinlay et al. have successfully addressed each of the challenges mentioned above by developing a highly effective mRNA delivery system comprising charge-altering releasable transporters (CARTs). Since a picture is worth a thousand words, I’ve reproduced here the diagram used by McKinlay et al. to describe their multistep approach with CARTs, namely complexation (1), intracellular delivery (2), and cytosolic release (3) of mRNA transcripts, resulting in induction of protein expression (4).

Taken from McKinlay et al. Proc. Natl. Acad. Sci (2017)

Readers interested in the clever chemistry that underlies CARTs should consult the publication by McKinlay et al. for details. In brief, these dynamic materials, specifically oligo(carbonate-b-α-amino ester)s (1) shown below function initially as polycations that noncovalently complex, protect, and deliver polyanionic mRNA and then subsequently lose their cationic charge through a controlled degradation to a neutral small molecule (2). The proposed mechanism for this degradation mechanism, which McKinlay et al. refer to as “self-immolative,” is pH-dependent.

Proposed rearrangement mechanism for n-mer oligo(α-amino ester)s 1 through tandem five-membered (5) then six-membered (6) transition states to afford an n-2-mer and diketopiperazine 2. Taken from McKinlay et al. Proc. Natl. Acad. Sci (2017)

As exemplified below, CARTs for cellular uptake were synthesized with hydrophobic blocks (n = 15) and cationic blocks (n = 12) such that 11b in physiological phosphate buffered saline (PBS) at pH 7.4 undergoes degradation to form 11c and small molecule 2.

Taken from McKinlay et al. Proc. Natl. Acad. Sci (2017)

These researchers hypothesize that this charge alteration reduces or eliminates the electrostatic anion-binding ability of the originally cationic material, thereby facilitating endosomal escape and enabling free mRNA release into the cytosol for translation. Readers interested in learning more about the complexities of endosomal escape can consult a (free, via Google) book chapter by Uyechi-O’Brien and Szoka titled Mechanisms for Cationic Lipids published in 2003, and a 2012 review by Nguyen and Szoka rhetorically titled Nucleic Acid Delivery: The Missing Pieces of the Puzzle?

Regardless of the actual mechanistic details for CARTs, McKinlay et al. demonstrate the efficacy of these materials to complex, deliver, and release mRNA in various lines of cultured cells including primary mesenchymal stem cells and in animal models, via both intramuscular (i.m.) injection and intravenous (i.v.) administration, resulting in robust gene expression. I’ll briefly outline these findings in what follows; however, the full paper and its supplemental material should be consulted for details.

Incidentally, I’m pleased to add that these CARTs were used to deliver the following base-modified [5-methylcytidine (5meC ) and pseudouridine (Ψ)] reporter mRNAs and dye-labeled mRNA obtained from TriLink BioTechnologies: Enhanced Green Fluorescent Protein (EGFP) mRNA, Firefly Luciferase (Fluc) mRNA, and Cyanine 5 (Cy5)-labeled EGFP mRNA.

Mechanism of Uptake and Release

Using a Cy5-labeled EGFP mRNA it was determined that the mechanism of cell entry for CART mRNA polyplexes is predominantly endocytic by comparing cellular uptake at 4 °C, a condition known to inhibit endocytotic processes, to normal uptake at 37 °C. Consistent with the expected endocytotic mechanism for ∼250-nm particles, HeLa cells displayed a significant (85%) reduction in Cy5 fluorescence at 4 °C.

Cellular uptake and mRNA translation following treatment with CART/mRNA polyplexes were then directly compared with polyplexes formed with non-immolative oligomers. By delivering a mixture of EGFP mRNA and Cy5-labeled EGFP mRNA, analysis of mRNA internalization and expression can be decoupled and simultaneously quantified: Cy5 fluorescence indicates internalized mRNA, irrespective of localization, and EGFP fluorescence denotes cytosolic release and subsequent expression of mRNA.

TriLink Cy5-labeled EGFP mRNA is transcribed with Cy5-UTP and an analog of UTP at a ratio which results in mRNA that is easily visualized and can still be translated in cell culture. Translation efficiency correlates inversely with Cyanine 5-UTP substitution.

This method was used in conjunction with confocal microscopy to compare cellular uptake and mRNA expression of two oligomers, namely, CART D13:A11 (7) and non-immolative, guanidinium-containing D13:G12 (13). Detection included dansylated transporter, Cy5-mRNA, and tetramethylrhodamine (TRITC)-Dextran4400, a stain for endosomal compartments. When cells were imaged 4 h after treatment with CART 7/Cy5-mRNA complexes diffuse fluorescence was observed for both the Cy5 and dansyl fluorophores, indicating that those materials successfully escaped the endosome and dissociated from the polyplexes (i).

Confocal microscopy of HeLa cells treated with Cy5-mRNA complexes using CART 7 or non-immolative oligomer 13 after 4 h. Cells were cotreated TRITC-Dextran4400. Scale bar, 10 μm. Taken from McKinlay et al. Proc. Natl. Acad. Sci (2017)

The two observed puncta in the dansyl signal (ii) was attributed to some intracellular aggregation of the dansyl-labeled lipidated oligocarbonate blocks, resulting from self-immolative degradation of the cationic segments of CART 7. Diffuse fluorescence from (TRITC)-Dextran4400 was also observed and attributed to endosomal rupture and release of the entrapped dextran.

However, when cells are treated with non-immolative 13/Cy5-mRNA complexes, both the Cy5 and dansyl fluorescence remain punctate and colocalized (iii). These signals also strongly overlap with punctate TRITC-Dextran4400, indicative of endosomal entrapment.

Taken together, according to McKinlay et al., these data strongly suggest that the charge altering behavior of CART 7 enables endosomal rupture and mRNA release, contributing to the high performance of these materials for mRNA delivery.

Applications and Animal Experiments

Oligo(carbonate-b-α-amino ester) D13:A11 7 was evaluated in applications to explore the versatility of CART-mediated mRNA delivery. EGFP mRNA expression following delivery by CART 7 was assayed in a panel of cell lines and compared to widely used Lipofectamine 2000 (Lipo). HeLa cells, murine macrophage (J774), human embryonic kidney (HEK-293), CHO, and human hepatocellular carcinoma (HepG2) cells all showed that the percentage of cells expressing EGFP using the CART 7 was >90%, whereas treatment with Lipo induced expression in only 22–55% of the cells. Importantly, in addition to these various immortalized cell lines, mRNA expression was also observed in primary CD1 mouse-derived mesenchymal stem cells (MSCs) with high transfection efficiency.

In vivo bioluminescence imaging (BLI) enables localization and quantification of expression following mRNA delivery in living animals. To assess the efficacy of CART/mRNA complexes following local (i.m.) or systemic (i.v.) routes of administration, CART 7-complexed Fluc mRNA (7.5 μg ) in PBS (75 μL) was given to anesthetized BALB/c mice in the right thigh muscle. As a direct control, naked mRNA was similarly injected in the opposite flank. D-luciferin was systemically administered i.p. at 15 min before imaging for each time point, and luciferase expression was evaluated over 48 h, starting at 1 h after the administration of mRNA complexes.

As shown here, when Fluc mRNA was delivered with polyplexes derived from 7 into the muscle, high levels of luciferase activity were observed at the site of injection. This expression peaked at 4 h and was still observable after 24 h but barely so after 48 h (see publication for percentages). In contrast, i.m. injection of naked mRNA afforded only low levels of luciferase expression, as measured by photon flux, in all five mice (see publication for percentages).

Representative BLI images following i.m. injection of naked mRNA (left flank) or CART/mRNA complexes (right flank).Taken from McKinlay et al. Proc. Natl. Acad. Sci (2017)

Following i.v. injections, the localization of mRNA polyplexes in tissues along the reticuloendothelial system pictured here provides many opportunities in inducing immunotherapeutic responses. According to McKinlay et al., spleen localization is “particularly exciting for future studies involving mRNA-based immunotherapy due to large numbers of dendritic and immune cells in that tissue.” Liver localization was also apparent in these animals, and expression in this tissue “may have applicability for treatment of hereditary monogenic hepatic diseases requiring protein augmentation or replacement such as hereditary tyrosinemia type I, Crigler–Najjar syndrome type 1, alpha-1-antityrpsin deficiency, Wilson disease, and hemophilia A and B, or acquired liver diseases such as viral hepatitis A–E and hepatocellular carcinoma.”

Overview of the reticuloendothelial system. ©Frazier et al. (1996)

Future Perspectives

Rather than paraphrase the future perspectives envisaged by McKinlay et al., here are those views, which to me seem warranted by the promising results summarized above:

“The effectiveness of mRNA delivery using these CARTs represents a strategy for mRNA delivery that results in functional protein expression in both cells and animals. The success of these materials will enable widespread exploration into their utilization for vaccination, protein replacement therapy, and genome editing, while augmenting our mechanistic understanding of the molecular requirements for mRNA delivery.”

As usual, your comments are welcomed.

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Using Nucleotide Analogs for RNA Visualization and Gene Expression Kinetics

  • 5-Azidoalkyluridine in RNA for Click Chemistry in Living Cells for Visualization
  • 4-Thiouridine in RNA in Living Cells for Kinetics by Sequencing

Homeostasis is defined as the stable state (or equilibrium) of an organism and of its internal environment. Maintenance and regulation of homeostasis involves regulated expression of genetic information, and disruption of this process can result in human diseases. The transient existence and function of RNA in the framework of the central dogma of molecular biology necessitates tightly regulated molecular events. In turn, these events control the relative kinetics of RNA transcription, processing, and degradation. To understand the molecular basis for gene regulation, we first must understand the relative kinetics of RNAs biogenesis and degradation in a systematic and transcript-specific manner.

Taken from essayhomeworkhelp.org

Readers interested in learning about the details concerning the biogenesis and degradation of RNA can consult a comprehensive survey and summary by Jackowiak et al., which also serves as a source of references for numerous subtopics. This “chemistry-centric” blog will first focus on fluorescence-based visualization of RNA using click chemistry, which I’ve blogged about previously. Following that, there will be a synopsis of how the chemical properties of the nucleotide analog 4-thiouridine enable high-throughput, sequencing-based RNA, counting for determination of gene expression kinetics.

RNA Visualization

RNA visualization methods commonly rely on metabolic labeling of RNA with ribonucleoside or ribonucleotide analogs, such as 5-bromouridine (BrU) or BrU-5’-triphosphate, followed by immunostaining with fluorescent antibody for BrU, as described here. However, these methods involve laborious assay setups and are not applicable to all cell types and tissue samples due to limited permeability of the antibodies. Endogenous RNA has also been visualized by using fluorescently-modified antisense oligonucleotide probes, molecular beacons, and, more recently, aptamer-binding fluorophores. These methods have varying benefits and limitations, which have been reviewed by Mannack et al. in a freely accessible publication in 2016 titled Current techniques for visualizing RNA in cells that is definitely worth reading as an introduction to this field.

Taken from Mannack et al. F1000Res (2016)

Sawant et al. describe the development of a versatile toolbox composed of azide-modified uridine triphosphates, which facilitates the direct incorporation of azide functionality into RNA transcripts by transcription reaction, as depicted here. The azide-modified RNA is readily functionalized with biophysical probes by various types of reactions, including strain-promoted azide-alkyne cycloaddition (SPAAC) and azide-phosphine Staudinger chemical ligation. Importantly, Sawant et al. show the specific incorporation of azide groups into cellular RNA transcripts by endogenous RNA polymerases for the first time. The azide-labeled cellular RNA transcripts are conveniently visualized in fixed and live cells by fluorescence microscopy upon click reaction with fluorescent alkynes.

Taken from Sawant et al. Nucleic Acids Res (2015)

Interested readers can consult Sawant et al. for extensive details regarding synthesis of the azido-functionalized nucleoside triphosphates and fluorescently labeled reagents that have ring-strained reactive carbon-carbon triple bonds, such as the Cy3-labeled cyclooctyne example shown here.

Taken from Sawant et al. Nucleic Acids Res (2015)

The utility of the SPAAC reaction in the detection of newly transcribing RNA in living cells was first observed by treating the cells with 5-azidomethyluridine-5’-triphosphate (AMUTP) for 1h. Subsequently, the cells were treated with a Cy3-labeled cyclooctyne probe for 30min and washed. The resultant Cy3 SPAAC RNA product was observed by confocal microscopy. Cell viability tests were done to confirm the low toxicity of the probe in these staining reaction conditions. According to Sawant et al., this RNA labeling method is advantageous, as the selective incorporation of azide groups into RNA using AMUTP and the SPAAC dye-visualization procedure provide an alternative route to image actively transcribing RNA in living cells.

Gene Expression Dynamics

Metabolic RNA labeling approaches that employ nucleotide-analogs enable tracking of RNA species over time without interfering with cellular integrity. Among these, 4-thiouridine (s4U) represents the most widely used nucleotide-analog to study the dynamics of RNA expression. This is because it is readily imported into metazoan cells by equilibrate nucleoside transporters, and it provides unique physicochemical properties for thiol(SH)-specific reactivity.

Herzog et al. recently reported SH-linked alkylation for the metabolic sequencing (SLAM-seq) of RNA to precisely locate s4U at single-nucleotide-resolution by reverse-transcription-dependent thymine-to-cytosine(T>C)-conversions in a high-throughput sequencing-compatible manner. For SLAM-seq, Herzog et al. employed the SH-reactive compound iodoacetamide, which covalently attaches a carboxyamidomethyl-group to s4U by nucleophilic substitution, as depicted here.

Taken from Herzog et al. Nature Methods (2017)

Quantitative s4U-alkylation was confirmed by a shift in the characteristic absorbance spectrum of 4-thiouracil from 335 nm to 297 nm. Under optimal reaction conditions, absorbance at 335 nm decreased 50-fold compared to untreated 4-thiouracil, resulting in ≥98% alkylation within 15 min. Quantitative identification of s4U by sequencing presumes that reverse transcriptase (RT) passes alkylated s4U-residues without drop-off. Herzog et al. therefore studied the effect of s4U-alkylation on RT-processivity in primer extension assays, but did not observe a significant effect of s4U alkylation on RT processivity when compared to a non-s4U-containing oligo with identical sequence.

To evaluate the effect of s4U-alkylation on RT-directed nucleotide incorporation, these investigators isolated the full-length products of primer extension reactions, PCR-amplified the cDNA, and subjected the libraries to high-throughput-sequencing. While the presence of s4U led to a constant ~10% T>C-conversions in the absence of alkylation (presumably due to base-pairing variations of s4U-tautomers), s4U-alkylation increased T>C-conversions 8.5-fold, resulting in a >94% T>C conversion rate. Importantly, they showed that iodoacetamide-treatment leaves conversion rates of any given non-thiol-containing nucleotide unaltered. Interested readers can consult the publication for details on how sequencing data are processed to give T>C conversion results.

The overall workflow of SLAM-seq is depicted below. To directly measure mRNA transcript stabilities, Herzog et al. subjected mouse embryonic stem cells (mESCs) to s4U metabolic RNA labeling for 24h, followed by washout and chase using uridine. They also prepared total RNA at various time points along the chase, which was then subjected to alkylation and Quant-seq. Inspection of candidate genes revealed constant steady-state expression across the time-course, while T>C-conversion-containing reads decreased over time in a transcript-specific manner. After suitable calculations, normalized T>C-conversion rates fit well to single-exponential decay kinetics, enabling the determination of transcript half-life.

Workflow of SLAM-seq. Working time for alkylation and Quant-seq library preparation are indicated. Taken from Herzog et al. Nature Methods (2017)

Herzog et al. ranked the 6,665 transcripts for which half-life was determined at high accuracy according to their relative stability. They also performed gene-ontology (GO) enrichment analysis for the 666 most or least stable mRNAs, as shown here. Transcripts with short half-life significantly enriched for regulators of Pol II-dependent transcription, while stable mRNAs associated with the GO-terms translation, respiratory electron transport, and oxidative phosphorylation. Together with gene set enrichment analyses, SLAM-seq measurements confirmed that transcripts encoding proteins with house-keeping function tend to decay at low rates, perhaps reflecting the evolutionary adaptation to energy constraints. In contrast, transcripts with a regulatory role tend to decay faster, most certainly because control over the persistence of genetic information facilitates adaptation to environmental changes.

Taken from Herzog et al. Nature Methods (2017)

Herzog et al. point out a number of limitations and caveats for SLAM-seq RNA kinetics. Because s4U-uptake can vary between cell types, careful assessment of cell-type-specific toxicity is imperative in order to meet s4U-labeling conditions that do not affect gene expression or cell viability. Additionally, the ability to determine de novo synthesized transcripts will depend on (1) the cellular s4U uptake kinetics, (2) the overall transcriptional activity of the cell type and (3) the library sequencing depth. Hence, these parameters need to be taken into account when designing a SLAM-seq experiment, particularly when employing short s4U pulse labeling, where sequencing depth demands adjustments to the given cellular parameters.

Notwithstanding these issues and reflecting on what SLAM-seq has revealed about gene expression kinetics, I was struck by the remarkable effects derived from simply substituting a sulfur atom for oxygen in uridine. Chemistry is amazing!

As usual, your comments are welcomed.

Footnote

Reading about s4U piqued my curiosity about publications in which s4U, as the nucleoside or triphosphate obtained from TriLink, has been used. I searched Google Scholar for such publications, and found a relatively large number and variety of applications, which you can peruse here. My favorite was a protocol by Timothy W. Nilsen in RNA: A Laboratory Manual titled Detecting RNA–protein interactions by photocross-linking using RNA molecules containing uridine analogs.

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In Search of RNA Epigenetics: A Grand Challenge

  • Methylated riboA and riboC are the most commonly detected nucleobases in epigenetics research
  • Powerful new analytical methods are key tools for progress
  • Promising PacBio sequencing and novel “Pan Probes” reported   

In a Grand Challenge Commentary published in Nature Chemical Biology in 2010, Prof. Chuan He at the University of Chicago opined that “[p]ost-transcriptional RNA modifications can be dynamic and might have functions beyond fine-tuning the structure and function of RNA. Understanding these RNA modification pathways and their functions may allow researchers to identify new layers of gene regulation at the RNA level.”

Like other scientists who get hooked by certain Grand Challenges, I became fascinated by this possibility of yet “new layers” of genetic regulation involving RNA, either as conventional messenger RNA (mRNA) or more recently recognized long noncoding RNA (lncRNA). Part of my intellectual stimulation was related to the fact that some of my past postings have dealt with both lncRNA as well as recent advances in DNA epigenetics, so the notion of RNA epigenetics seemed to tie these together.

After doing my homework on recent publications related to possible RNA epigenetics, it became apparent that this posting could be logically divided into commentary on the following three major questions: what are prevalent epigenetic RNA modifications, what might these do, and where is the field going? Future directions were addressed by interviews with two leading investigators: Prof. Chuan He, who is mentioned above, and Prof. Tao, who has been involved in cutting edge methods development.

RNA Epigenetic Modifications

More than 100 types of RNA modifications are found throughout virtually all forms of life. These are most prevalent in ribosomal RNA (rRNA) and transfer RNA (tRNA), and are associated with fine tuning the structure and function of rRNA and tRNA. Comments here will instead focus on mRNA and lncRNA in mammals, wherein the most abundant—and far less understood—modifications are N6-methyladenosine (m6A) and 5-methylcytidine (m5C).

structures

Three Approaches to Sequencing m6A-Modified RNA

Discovered in cancer cells in the 1970s, m6A is the most abundant modification in eukaryotic mRNA and lncRNA. It is found at 3-5 sites on average in mammalian mRNA, and up to 15 sites in some viral RNA. In addition to this relatively low density, specific loci in a given mRNA were a mixture of unmodified- and methylated-A residues, thus making it very difficult to detect, locate, and quantify m6A patterns. Fortunately, that has changed dramatically with the advent of various high-throughput “deep sequencing” technologies, as well as other advances.

(1.) Antibody-based m6A-seq 

An impressive breakthrough publication in Nature in 2012 by a group of investigators in Israel reported novel methodology called m6A-seq for determining the positions of m6A at a transcriptome-wide level. This approach, which is a variant of methylated DNA immunoprecipitation (MeDIP or mDIP), combines the high specificity of an anti-m6A antibody with Illumina’s massively parallel sequencing of randomly fragmented transcripts following immunoprecipitation. These researchers summarize their salient findings as follows.

“We identify over 12,000 m6A sites characterized by a typical consensus in the transcripts of more than 7,000 human genes. Sites preferentially appear in two distinct landmarks—around stop codons and within long internal exons—and are highly conserved between human and mouse. Although most sites are well preserved across normal and cancerous tissues and in response to various stimuli, a subset of stimulus-dependent, dynamically modulated sites is identified. Silencing the m6A methyltransferase significantly affects gene expression and alternative splicing patterns, resulting in modulation of the p53 (also known as TP53) signaling pathway and apoptosis. Our findings therefore suggest that RNA decoration by m6A has a fundamental role in regulation of gene expression.”

Moreover, their concluding sentence refers back to He’s aforementioned Grand Challenge Commentary about RNA epigenetics in 2010, just two years earlier.

“The m6A methylome opens new avenues for correlating the methylation layer with other processing levels. In many ways, this approach is a forerunner, providing a reference and paving the way for the uncovering of other RNA modifications, which together constitute a new realm of biological regulation, recently termed RNA epigenetics.”

(2.) Promising PacBio Single-Molecule Real-Time (SMRT) Sequencing of m6A

In a previous post, I praised PacBio (Pacific Biosciences) for persevering in development of its SMRT sequencing technology that uniquely enables, among other things, direct sequencing of various types of modified DNA bases via differentiating the kinetics of incorporating labeled nucleotides. Attempts to extend the SMRT approach to sequencing m6A have been recently reported by PacBio in collaboration with Prof. Pan (see below) and others in J. Nanobiotechnology in April 2013. Using model synthetic RNA templates and HIV reverse transcriptase (HIV-RT) they demonstrated adequate discrimination of m6A from A, however, “real’ RNA samples having complex ensembles of tertiary structures proved to be problematic. Alternative engineered RTs that are more processive and accommodative of labeled nucleotides were said to be under investigation in order to provide longer read lengths and appropriate incorporation kinetics.

The authors are optimistic in being able to solve these technical problems, and concluded their report by stating:

  “[w]e anticipate that the application of our method may enable the identification of the location of many modified bases in mRNA and provide detailed information about the nature and the dynamic RNA refolding in retroviral/retro-transposon reverse transcription and in 3’-5’ exosome degradation of mRNA.”

Let’s hope that this is achieved soon!

(3.) Nanopore Sequencing of m6A?

It’s too early to be sure, but continued incremental advances in possible approaches to nanopore sequencing suggest applicability to m6A. As pictured below, Bayley and coworkers describe a method that uses ionic current measurement to resolve ribonucleoside monophosphates or diphosphates (rNDPs) in α-hemolysin protein nanopores containing amino-cyclodextrin adapters.

Taken from Bayley and coworkers in Nano Lett. (2013)

Taken from Bayley and coworkers in Nano Lett. (2013)

The accuracy of base identification is further investigated through the use of a guanidino-modified adapter. On the basis of these findings, an exosequencing approach for single-stranded RNA (ssRNA) is envisioned in which a processive exoribonuclease (polynucleotide phosphorylase, PNPase) presents sequentially cleaved rNDPs to a nanopore. Although extension of this concept to include m6A has yet to be demonstrated, earlier feasibility studies by Ayub & Bayley have shown discrimination of m6A (and other modified bases) from unmodified ribobases.

Two Probe-Based Methods for Detecting Specific m6A Sites

1.) “Pan Probes”

As the saying goes, “what goes around comes around”, and in this instance its repurposing 2’-O-methyl (2’OMe) modified RNA/DNA/RNA oligos. This general class of chemically synthesized chimeric “gapmers” was originally used for RNase H-mediated cleavage of mRNA in antisense studies. Very recently, however, Pan and coworkers have cleverly adapted these probes—which I like to alliteratively refer to as “Pan Probes”—to m6A detection in mRNA and lncRNA.

For details see SCARLET workflow; taken from Pan and coworkers RNA (2013)

Pan Probes are comprised of “7-4-7 gapmers” having seven 2’OMe RNA nucleotides flanking four DNA nucleotides, the latter of which straddle known (or suspected) m6A sites, as depicted in the cartoon shown. The indicated series of steps, which involve site-specific cleavage and radioactive-labeling followed by ligation-assisted extraction and thin-layer chromatography, is thankfully called SCARLET by these investigators.

SCARLET was used by Pan and coworkers to determine the m6A status at several sites in two human lncRNAs and three human mRNAs, and found that the m6A fraction varied between 6% and 80% among these sites. However, they also found that many m6A candidate sites in these RNAs were not modified. Obviously, while much more work needs to be done to collect data for deciphering dynamic patterns and implications of m6A RNA epigenetic modifications, these investigators note that SCARLET is, in principle, applicable to m5C, pseudouridine, and other types of epigenetic RNA modifications.

Readers interested in designing and investigating their own Pan Probes can obtain these 7-4-7 gapmers by using TriLink’s OligoBuilder® and simply selecting “PO 2’OMe RNA” from the Primary Backbone dropdown menu, typing the first 7 bases in the Sequence box, selecting the 4 DNA bases from the Chimeric Bases menu and then typing the remaining 7 2’OMe RNA bases.

(2.) Probes for High-Resolution Melting

In a new approach very recently reported by Golovina et al. at Lomonosov Moscow State University, the presence of m6A in a specific position of mRNA or lncRNA molecule is detected using a variant of high-resolution melting (HRM) analysis applicable to, for example, single-nucleotide genotyping. The authors suggest that this method lends itself to screening many samples in a high-throughput assay following initial identification of loci by sequencing (see above). The method uses two labeled probes—one with 5’-FAM and another with 3’-BHQ1 (both available from Trilink’s OligoBuilder®)—that hybridize to a particular query position in a total RNA sample, as shown below for a 23S rRNA model system. The presence of m6A lowers the melting temperature (Tm), relative to A, with a magnitude that is sequence-context dependent.

Taken from Golovina et al. Nucleic Acids Res. (2013).

Taken from Golovina et al. Nucleic Acids Res. (2013).

The authors studied various probe-target constructs, and recommend 12–13-nt-long probes containing a quencher, and >20-nt long probes containing a fluorophore.  They also could advise that the quencher-containing oligonucleotide hybridizes to RNA such that m6A be directly opposite the 3′-terminal nucleotide carrying the quencher. The authors point out that relatively low-abundant, non-ribosomal targets need partial enrichment by, for example, simple molecular weight-based purification or commercially available kits. In this regard, they estimate that, if a particular type of mRNA was present at 10,000 copies per mammalian cell, 107 cells would be required to analyze m6A by this HRM method.

m5C Analysis by Sequencing of Bisulfite-Converted RNA

Selective reaction of bisulfite with C but not m5C in RNA, analogous to that long used for DNA, provides the basis for determining C-methylation status by sequencing. As detailed by Squires et al. in Nucleic Acids Res. in 2013, bisulfite-converted RNA can be sequenced by either of two methods: conversion to cDNA, cloning, and conventional sequencing, or conversion to a next-generation sequencing library. These authors described their salient findings as follows.

“We confirmed 21 of the 28 previously known m5C sites in human tRNAs and identified 234 novel tRNA candidate sites, mostly in anticipated structural positions. Surprisingly, we discovered 10,275 sites in mRNAs and other non-coding RNAs. We observed that distribution of modified cytosines between RNA types was not random; within mRNAs they were enriched in the untranslated regions and near Argonaute binding regions… Our data demonstrates the widespread presence of modified cytosines throughout coding and non-coding sequences in a transcriptome, suggesting a broader role of this modification in the post-transcriptional control of cellular RNA function.”

“Writing, Reading, and Erasing” RNA Epigenetic Modifications

Enzyme-mediated post-transcriptional RNA methylation (aka “writing”) and demethylation (aka “erasing”) are critical processes to identify and fully characterize in order to elucidate RNA epigenetics, and are formally analogous to those operative for DNA epigenetics.

RNA epigenetic “writing” mechanisms have focused on N6-adenosine-methyltransferase 70 kDa subunit, an enzyme that in humans is encoded by the METTL3 gene, and is involved in the posttranscriptional methylation of internal adenosine residues in eukaryotic mRNAs to form m6A. According to Squires et al., two m5C methyltransferases in humans, NSUN2 and TRDMT1, are known to modify specific tRNAs and have roles in the control of cell growth and differentiation.

As for “erasing”, in 2011, He’s lab discovered the first RNA demethylase, abbreviated FTO, for fat mass and obesity-associated protein, which has efficient oxidative demethylation activity targeting m6A in RNA in vitro. They also showed for the first time that this erasure of m6A could significantly affect gene expression regulation. In 2013, He’s lab discovered the second mammalian demethylase for m6A, ALKBH5, which affects mRNA export and RNA metabolism, as well as the assembly of mRNA processing factors, suggesting that reversible m6A modification has fundamental and broad functions in mammalian cells.

So, if Mother Nature evolved these mechanisms for writing and erasing RNA epigenetic modifications, what about the equally important, in between process of “reading” them? He and Pan and collaborators have very recently reported insights to such reading. They showed that m6A is selectively recognized by the human YTH domain family 2 (YTHDF2) “reader” protein to regulate mRNA degradation. They identified over 3,000 cellular RNA targets of YTHDF2, most of which are mRNAs, but also include non-coding RNAs, with a conserved core motif of G(m6A)C. They further establish the role of YTHDF2 in RNA metabolism, showing that binding of YTHDF2 results in the localization of bound mRNA from the translatable pool to mRNA decay sites. The carboxy-terminal domain of YTHDF2 selectively binds to m6A-containing mRNA, whereas the amino-terminal domain is responsible for the localization of the YTHDF2–mRNA complex to cellular RNA decay sites. These findings, they say, indicate that the dynamic m6A modification is recognized by selectively binding proteins to affect the translation status and lifetime of mRNA.

Expert Opinions of the Future for RNA Epigenetics

As I’ve said here before, there is no crystal ball for accurately predicting the future in science, although scientists do enjoy imagining that there is. Opinions of two “hands on” experts in the emerging field of RNA epigenetics are certainly of interest in this regard. Below are some comments offered by the aforementioned Prof. Tao Pan and Prof. Chuan He provided via an email interview in which I posed the question, ‘What do you see as the most important developments for RNA epigenetics?’ These experts have  thrown down the gauntlet, so to speak, by asserting RNA epigenetics as a Grand Challenge.

Prof. Tao Pan

Prof. Tao Pan

“In my opinion, the biggest current challenge for the field is to develop methods that can perturb m6A modification at specific sites to assess m6A function directly in specific genes. RNA interference or overexpression of an mRNA may simply decrease or increase modified and unmodified RNA alike. In a few cases, mutation of a known m6A site in an mRNA resulted in additional modification at a nearby consensus site, so that one cannot simply assume that mutation of a known site would not lead to cryptic sites nearby that may perform the same function. Further, functional understanding of a specific site should also take into account that all currently known m6A sites in mRNA and viral RNA are incompletely modified, so that one may need to explain why cells simultaneously maintain two RNA species that differ only at the site of m6A modification.”   

Prof. Chuan He

Prof. Chuan He

The m6A modification is much more abundant than other RNA modifications in mammalian and plant nuclear RNA and is currently the only known reversible RNA modification. The m6A maps of various organisms/cell types need to be obtained. High-resolution methods to obtain transcriptome-wide, base-resolution maps are important. A future focus should be to connect the reversible m6A methylation with functions, in particular, the studies of the reader proteins that specifically recognize m6A and exert biological regulation. The first example of the YTHDF2 work just published in Nature (above) is a good example. We believe many other reader proteins exist and impact almost all aspects of mRNA metabolisms or functions of lncRNA. 

Besides m6A, there are m5C, pseudoU, 2′-OMe, and potentially other modifications in mRNA and various non-coding RNAs (such as the recently discovered hm6A and f6A). The methods to map these modifications (except m5C) need to be developed and their biological functions need to be elucidated. 

Lastly, potential reversal of rRNA and tRNA modifications needs to be studied. As I stated in the Commentary in 2010, dynamic RNA modifications could impact gene expression regulation resembling well-known dynamic DNA and histone modifications. I think now we have enough convincing data to indicate this is indeed the case. The future is bright.”

Very bright, indeed! Your comments about this posting are welcomed.

Modified mRNA Mania

  • Biosynthetic modified mRNA for gene-based therapy without the gene!
  • AstraZeneca bets up to $420M on Moderna’s “messenger RNA therapeutics”
  • “Me-too” Pharma frenzy to follow?

In a perspective on gene therapy published in Science this year, Inder M. Verma starts by observing that the concept of gene therapy is disarmingly simple. Introduce a healthy gene in a patient and its protein product should alleviate the defect caused by a faulty gene or slow the progression of the disease. He then asks the rhetorical question: ‘why then, over the past three decades, have there been so few clinical successes in treating patients with this approach?’ The answer in part has to do with challenges for cell or tissue-specific delivery, which admittedly is an issue for virtually any type of therapeutic agent. There is also concern for adverse events generally ascribed to unintended vector integration leading to neoplasias. Nevertheless, according to Verma, the present clinical trials pipeline is jammed with more than 1700 (!) clinical trials worldwide, drawing on a wide array of gene therapy approaches for both acquired and inherited diseases.

In view of this scientifically laudable but undeniable—if not frustratingly—slow progress, it’s not surprising that various groups of investigators—and investors—have recently opted to pursue a strategy that eliminates a DNA-encoded gene entirely! Instead, biosynthetic mRNA is delivered in order to directly produce the desired therapeutic protein product—this is now being referred to as “mRNA therapeutics”.

Having said this, let’s consider some pivotal scientific publications, patents, and the emerging commercial landscape for what looks to be a very hot area for research and corporate competition.

Modified mRNA Therapeutic Vaccines

An excellent review published in 2010 by Bringmann et al. entitled RNA Vaccines in Cancer Treatment covers various approaches to using mRNA encoding for tumor-associated antigens to induce specific cytotoxic T lymphocyte and antibody responses. RNA-transfected dendritic cell vaccines have been extensively investigated and are currently in numerous clinical trials (the details for which can be found at the NIH ClinicalTrials.gov website by simply searching RNA vaccines).

Interestingly, clinical feasibility and safety assessment for direct intradermal injection of “naked” unmodified mRNA was reported back in 2008 by Weide et al., who removed metastatic tissue from each of 15 melanoma patients for total RNA extraction, reverse-transcription to cDNA, amplification, cloning, and transcription to produce unlimited amounts of copy mRNA.

Stabilizing unmodified mRNA by packaging in liposomes or forming complexes with cationic polymers has been widely investigated, as well as introducing chemical modifications to mRNA to make it more resistant against degradation and more efficient for translation. The latter includes elongation of the poly-A tail at the 3′-end of the molecule and modifications to the cap structure at the 5′-end. For example, if the original 7-methylguanosine triphosphate is replaced by an Antireverse Cap Analog (ARCA), the efficiency of transcription is strongly enhanced. To provide the immune system with even more potent signals, Scheel et al. modified mRNA with a phosphorothioate backbone in early commercial vaccine development work at CureVac GmbH (Tübingen, Germany) that continues today (see image below).

Effects of mRNA vaccines  (taken from an article in Drug Discovery & Development by Ingmar Hoerr, PhD, CEO and Cofounder of CureVac).

Effects of mRNA vaccines (taken from an article in Drug Discovery & Development by Ingmar Hoerr, PhD, CEO and Cofounder of CureVac).

In summary, in a 2013 review entitled RNA: The new revolution in nucleic acid vaccines, Geall et al. from Novartis Vaccines & Diagnostics (Cambridge, MA, USA) stated that “prospects for success are bright.” They site several reasons for this optimistic outlook including the potential of RNA vaccines to address safety and effectiveness issues sometimes associated with vaccines that are based on live attenuated viruses and recombinant viral vectors. In addition, methods to manufacture RNA vaccines are suitable as generic platforms and for rapid response, both of which will be very important for addressing newly emerging pathogens in a timely fashion. Plasmid DNA is the more widely studied form of nucleic acid vaccine and proof of principle in humans has been demonstrated, although no licensed human products have yet emerged. The RNA vaccine approach, based on mRNA, is gaining increased attention and several vaccines are under investigation for infectious diseases, cancer and allergy.

Modified mRNA for Expressing Clinically Beneficial Proteins

Dr. Katalin Karikó, Adjunct Associate Professor of Neurosurgery and Senior Research Investigator, Department of Neurosurgery, University of Pennsylvania (taken from upenn.edu).

Dr. Katalin Karikó, Adjunct Associate Professor of Neurosurgery and Senior Research Investigator, Department of Neurosurgery, University of Pennsylvania (taken from upenn.edu).

In a landmark publication by Karikó et al. in 2008, it was reasoned that the suitability of mRNA as a direct source of therapeutic proteins in vivo required muting its immunogenicity and boosting its effectiveness. Clues as to how this might be achieved were provided in their earlier work demonstrating the use of base-modified triphosphates to enzymatically synthesize in vitro mRNA having modified nucleosides [such as, pseudouridine (Ψ), 5-methylcytidine (m5C), N6-methyladenosine (m6A), 5-methyluridine (m5U), or 2-thiouridine (s2U)] had greatly diminished immunostimulatory properties. They reasoned that, “if any of the in vitro transcripts containing nucleoside modifications would remain translatable and also avoid immune activation in vivo, such an mRNA could be developed into a new therapeutic tool for both gene replacement and vaccination”.

Using the aforementioned and other base-modified nucleotide triphosphates—all obtained from TriLink BioTechnologiesKarikó et al. found, surprisingly, that mRNA containing pseudouridine had a higher translational capacity than unmodified mRNA when tested in mammalian cells and lysates or administered intravenously into mice at 0.015–0.15 mg/kg doses. The delivered mRNA and the encoded protein could be detected in the spleen at 1, 4, and 24 hours after the injection, and at each time-point there was more of the reporter protein when pseudouridine-containing mRNA was administered. Moreover, even at higher doses, only the unmodified mRNA was immunogenic. [Note: a fascinating follow-on publication provides a non-obvious—at least to me—molecular-level rationale for the surprising enhanced translation of pseudouridine-modified mRNA]. 

uridine

Uridine and pseudouridine differ in bonding to ribose but hydrogen-bond similarly to adenine. Pseudouridine is the most prevalent of the 100+ naturally occurring modified nucleosides found in RNA.

They concluded that, “[t]hese collective findings are important steps in developing the therapeutic potential of mRNA, such as using modified mRNA as an alternative to conventional vaccination and as a means for expressing clinically beneficial proteins in vivo safely and effectively.” Prior to publishing this pivotal report, Katalin Karikó and co-author Drew Weissman filed a patent application in 2006 entitled RNA containing modified nucleosides and methods of use thereof that was issued on October 2, 2012 as US 8,278,036 and is assigned to the University of Pennsylvania.

 

Blood Boosting with Erythropoietin

EPO stimulates the production of red blood cells (taken from proactiveinvestors.com via Bing Images)

EPO stimulates the production of red blood cells (taken from proactiveinvestors.com via Bing Images)

In a very persuasive demonstration of the real possibility of mRNA therapeutics, Karikó et al, reported in 2012 that non-immunogenic pseudouridine-modified mRNA encoding erythropoietin (EPO) was translated in mice and non-human primates. Indeed, a single injection of 100 ng (0.005 mg/kg) of HPLC-purified mRNA complexed to a delivery agent elevated serum EPO levels significantly and levels were maintained for 4 days. In comparison, mRNA containing uridine produced 10–100-fold lower levels of EPO lasting only 1 day. EPO translated from pseudouridine-mRNA was functional and caused a significant increase of both reticulocyte counts and hematocrits. As little as 10 ng mRNA doubled reticulocyte numbers. Weekly injection of 100 ng of EPO mRNA was sufficient to increase the hematocrit from 43 to 57%, which was maintained with continued treatment. Even when a large amount of pseudouridine-mRNA was injected, no inflammatory cytokines were detectable in plasma.

Rhesus macaque (taken from flickr.com via Bing Images)

Rhesus macaque (taken from flickr.com via Bing Images)

Using rhesus macaques (aka rhesus monkeys) they could also detect significantly-increased serum EPO levels following intraperitoneal injection of rhesus EPO mRNA. Other researchers (Kormann et al.) independently used a single injection of modified murine mRNA to produce EPO in mice.

Kick-Start Cardiac Repair with VEGF-A

That’s the catchy title of a News & Views article in the October 2013 issue of Nature Biotechnology with an equally catchy byline that reads “[t]he survival of mice after experimental heart attack is greatly improved by a pulse of RNA therapy.” The featured report by Zangi et al., which is characterized as “a masterpiece of multidisciplinary studies…that will advance our thinking about therapeutic options in the cardiovascular arena,” is indeed impressive. These investigators report that intra-myocardial injections of vascular endothelial growth factor-A (VEGF-A) mRNA modified with 5-methylcytidine, pseudouridine, and 5’ cap structure resulted in expansion and directed differentiation of endogenous heart progenitors in a mouse model of myocardial infarction. They found markedly improved heart function and enhanced long-term survival of recipients. Moreover, “pulse-like” delivery of VEGF-A using modified mRNA was found to be superior to use of DNA vectors in vivo.

A heart attack (myocardial infarction) occurs when one of the heart's coronary arteries is blocked suddenly, usually by a blood clot (thrombus), which typically forms inside a coronary artery that already has been narrowed by atherosclerosis, a condition in which fatty deposits (plaques) build up along the inside walls of blood vessels (taken from drugs.com via Bing Images).

A heart attack (myocardial infarction) occurs when one of the heart’s coronary arteries is blocked suddenly, usually by a blood clot (thrombus), which typically forms inside a coronary artery that already has been narrowed by atherosclerosis, a condition in which fatty deposits (plaques) build up along the inside walls of blood vessels (taken from drugs.com via Bing Images).

Notwithstanding these promising results, the aforementioned News & Views article points out that microgram-scale doses of modified mRNA in mice used by Zangi et al. “would probably correspond to several hundred milligrams…in humans delivered in volumes that might exceed 10 ml per heart. In clinical practice, it would be very difficult to administer such volumes to infarcted hearts.” In my humble opinion, these are legitimate but purely hypothetical issues at this time and, given that it’s very “early days” for therapeutic modified mRNA technologies, it’s not unreasonable to assume that new modifications and/or improved delivery strategies can be developed to enable clinical utility.

From a technical perspective, this work by Zangi et al. involves a form of cell-free reprogramming and, as such, is a good segue into the next section. 

Modified mRNA for Cellular Reprogramming

In 2005, when I first heard of the concept of cellular reprogramming and dedifferentiation—which is to somehow coax a mature, differentiated cell to ‘run in reverse and go backwards biologically’ to a more primitive cell—my immediate impression as a chemist was this was impossible. Surely, I thought, this must violate the Second Law of Thermodynamics or, if not, is completely counterintuitive to how life works. Wow, was I wrong!

Reprogramming of differentiated cells to pluripotency is now firmly established and holds great promise as a tool for studying normal development.  It also offers hope that patient-specific induced pluripotent stem cells (iPSCs) could be used to model disease or to generate clinically useful cell types for autologous therapies aimed at repairing deficits arising from injury, illness, and aging. Induction of pluripotency was originally reported by Takahashi & Yamanaka by enforced retroviral expression of four transcription factors, KLF4, c-MYC, OCT4, and SOX2 (aka “Yamanaka factors”)—collectively abbreviated as KMOS. (TriLink sells these and other factors used to direct cell fate.) Viral integration into the genome initially presented a formidable obstacle to therapeutic use of iPSCs. The search for ways to induce pluripotency without incurring genetic change has thus become the focus of intense research effort.

Consequently, much attention has been given to the 2010 publication by Warren et al. entitled Highly efficient reprogramming to pluripotency and directed differentiation of human cells with synthetic modified mRNA. In this work complete substitution of either 5-methylcytidine for cytidine or pseudouridine for uridine in protein-encoding transcripts markedly improved protein expression, although the most significant improvement was seen when both modifications were used together. Transfection of modified mRNAs encoding the above mentioned Yamanaka factors led to robust expression and correct localization to the nucleus. Expression kinetics showed maximal protein expression 12 to 18 hours after transfection, followed by rapid turnover of these transcription factors. From this it was concluded that daily transfections would be required to maintain high levels of expression of the Yamanaka factors during long-term, multifactor reprogramming regimens.

They went on to demonstrate that repeated administration of modified mRNA encoding these (and other) factors led to reprogramming various types of differentiated human cells to pluripotency with conversion efficiencies and kinetics substantially superior to established viral protocols. Importantly, this simple, non-mutagenic, and highly controllable technology was shown to be applicable to a range of tissue-engineering tasks, exemplified by mRNA-mediated directed differentiation of mRNA-generated iPSCs to terminally differentiated myogenic (e.g. heart muscle) cells.

Modified mRNA reprogramming fibroblasts into induced pluripotent cells for directed differentiation into myofibers, according to Warren et al. in Cell Stem Cell (2010)

Modified mRNA reprogramming fibroblasts into induced pluripotent cells for directed differentiation into myofibers, according to Warren et al. in Cell Stem Cell (2010)

Warren et al. concluded that “we believe that our approach has the potential to become a major enabling technology for cell-based therapies and regenerative medicine.”  According to the Acknowledgements section of this 2010 publication, corresponding author Derrick J. Rossi recently founded a company, ModeRNA [sic] Therapeutics, dedicated to the clinical translation of this technology.” That, we shall see below, has had stunning commercial investment consequences.

By the way, and not surprisingly, Rossi & Warren filed a U.S. patent application in 2012 claiming, among other things, iPSCs induction kits using 5-methylcytidine- and pseudouridine-modified mRNA encoding KMOS human cellular reprogramming factors.

AstraZeneca’s Big Bet on Moderna’s Modified-mRNA Therapeutics

AstraZeneca aims to use Moderna Therapeutics’ modified-messenger RNA technology to develop and commercialize new drugs for cancer and serious cardiovascular, metabolic, and renal diseases, under a multi-year deal that could net Moderna more than $420 million. Moderna is also eligible for royalties on drug sales ranging from high single digits to low double digits per product.

AstraZeneca—ranked 7th in sales in 2010 among the world’s pharmaceutical companies—has the option to select up to 40 drug products for clinical development of what the companies are calling messenger RNA Therapeutics™, which could dramatically reduce the time and expense associated with creating therapeutic proteins using current recombinant technologies, the companies say. Moreover, “where current drug discovery technologies can target only a fraction of the disease-relevant proteins in the human genome, we have the potential to create completely new medicines to treat patients with serious cardiometabolic diseases and cancer,” AstraZeneca CEO Pascal Soriot said in a statement. Mr. Soriot, who had been a senior executive at Roche, very recently joined AstraZeneca, which said it would reorganize R&D and eliminate 1,600 jobs by 2016 as part of a plan to address issues related to failures in clinical trials of several drugs just as big sellers like the antipsychotic Seroquel and the heartburn drug Nexium have lost or are about to lose patent protection.

Moderna, based in Cambridge, Massachusetts, is privately held and was founded in 2010 by Flagship VentureLabs in association with leading scientists from Boston Children’s Hospital and Massachusetts Institute of Technology. Moderna has developed a broad intellectual property estate including 144 patent applications with 6,910 claims ranging from novel nucleotide chemistries to specific drug compositions, according to its website.

DARPA also Bets Big on Moderna’s Modified-mRNA Therapeutics

As the saying goes, “when it rains it pours”, and for Moderna it’s pouring money!

On October 2nd, Moderna announced that the U.S. Defense Advanced Research Projects Agency (DARPA)—whose most successful bets so far have been internet technologies—has awarded the company up to $25 million for R&D using its modified-mRNA therapeutics platform as a “rapid and reliable way to make antibody-producing drugs to protect against a wide range of now and emerging infectious diseases and engineered biological threats.” The statement goes on to say that Moderna’s approach can “tap directly into the body’s natural processes to produce antibodies without exposing people to a weakened or inactivated virus or pathogen, as in the case with the vaccine approaches currently being tested.”

The grant could support research for up to 5 years to advance promising antibody-producing drug candidates into preclinical testing and human clinical trial. The company also received a $700,000 ‘seeding’ grant from DARPA in March to begin work on the project.

If you’re interested in some of the possible ideas associated with the project, go to the 2013 patent application by Moderna entitled Methods of responding to a biothreat, which even envisages a portable, battery operated device for synthesizing modified mRNA. Oh well, never let it be said that DARPA fears a risky bet; on the other hand, since DARPA’s “playing with house money” (aka our taxes!), I suppose it’s easy for them. Let’s hope they/we all win.

Other Commercial Players

In addition to TriLink’s mRNA products, related services, and new cGMP facility, there are other companies to mention here, which I’ll do in alphabetical order.

  • Acuitas Therapeutics has compared the effectiveness of its lipid nanoparticle (LNP) carriers in vivo with the most potent delivery systems reported in the scientific literature, and found that Acuitas LNPs demonstrate much greater luciferase expression in the liver after systemic administration.
  • CureVac is combining both the antigenic and adjuvant properties of mRNA to develop novel and effective mRNA vaccines. CureVac is currently developing therapeutic mRNA vaccines in oncology and therapeutic/prophylactic vaccines for infectious diseases. Information on five of its clinical studies is available at ClinicalTrials.gov.
  • Dendreon has a U.S. patent application for a method to make dendritic cell vaccines from embryonic stem cells that are genetically modified with mRNA encoding tumor antigen. However, no mRNA-searchable items are currently listed on Dendreon’s website.
  • In-Cell-Art is investigating new and improved nanocarriers for mRNA vaccines, and has collaborated with Sanofi Pasteur and CureVac in DARPA-funded studies.
  • Mirus Bio offers a TransIT®-mRNA Transfection Kit for high efficiency, low toxicity, mRNA transfection of mammalian cells, as described by Karikó et al.

Also noteworthy, the 1st International mRNA Health Conference recently held on October 23-24 at the University of Tübingen included talks by numerous key scientists in academia and industry that are well worth looking at in the Conference Program.

In conclusion, I hope that you found this emerging area of modified mRNA therapeutics as interesting and exciting as I did in researching this blog posting, and I welcome your comments.

Postscript

After finishing the above blog, I came across these additional publications on possible mRNA therapies.

Huang and coworkers reported earlier this year that systemic delivery of liposome-protamine-formulated modified mRNA encoding herpes simplex virus 1 thymidine kinase for targeted cancer gene therapy was significantly more effective than plasmid DNA in a therapeutic model of human lung carcinoma in xenograft-bearing nude mice.

Zimmermann et al. reported successful use of mRNA-nucleofection for overexpression of interleukin-10 in murine monocytes/macrophages for anti-inflammatory therapy in a murine model of autoimmune myocarditis. [Note: for a related report on mRNA-engineered mesenchymal stem cells for targeted delivery of interleukin-10 to sites of inflammation see Levy et al.]

Cystic Fibrosis (CF) is the most frequent lethal genetic disease in the Caucasian population. CF is caused by a defective gene coding for the cystic fibrosis transmembrane conductance regulator (CFTR). Bangel-Ruland et al. reported in vitro results indicating that CFTR-mRNA delivery provided a novel alternative for cystic fibrosis “gene therapy”.

The Buzz on the Cut: From Dream to Reality

Targeted Genome Engineering with Zinc-finger Nucleases, TALENs and CRISPR

Targeted genome editing tools such as meganucleases, zinc-finger nucleases, TALENs and CRISPR are among the hottest topics in cell and gene therapy. Dr. Anton McCaffrey, Principal Scientist at TriLink and expert in these areas, gives herein his overview after attending the recent American Society for Gene and Cell Therapy Meeting (May 2013) and the International Society for Stem Cell Research Meeting (June 2013) where there were a number of exciting talks discussing applications of this technology.

amccaffrey


Dr. McCaffrey received his PhD in Biochemistry from the University of Colorado at Boulder in 1999. During his postdoctoral fellowship at Stanford he developed gene therapeutics for hepatitis B and C. He was then Assistant Professor at
University of Iowa where he focused on the role of microRNAs during the pathogenesis of hepatitis C virus and developed RNA interference and zinc-finger nuclease based therapeutics for treatment of hepatitis B virus. Now he manages the RNA Transcription product line at TriLink.

So what is targeted genome engineering with nucleases and why would you want to do this?  The primary goal in cells or animals is to create a specific, localized double-stranded DNA break and then to: 1. correct the sequence of a defective targeted gene, 2. knock in a specific gene mutation to create a disease model or 3. knock out a gene. The basic idea is to rationally design artificial restriction enzymes that recognize a specific location within the DNA genome of a cell or organism and catalyze a double stranded break at this location (Figure 1).

In the first two cases, where the target gene is to be specifically edited, the nuclease(s) are co-transfected with an exogenous donor DNA molecule. This donor DNA contains arms, which share homology with the target loci and will direct homologous recombination at the targeted cut site. The sequence of the donor DNA replaces that of the endogenous locus at one or both alleles. So, for example, a wild-type donor sequence can be used to replace a mutated gene sequence to correct a genetic disease.

If one wishes to inactivate a gene using these technologies, an exogenous donor template is not included. In the absence of a donor, the cell uses non-homologous end joining to repair the double stranded break. At a high frequency, this process introduces deletions and insertions in the gene, which changes the reading frame and inactivates the gene.

Until the advent of these technologies, it was impossible to make transgenic animals other than mice. Using targeting genome engineering is now possible to make transgenic rats, pigs, ferrets and plants. As will be discussed below, advances in messenger RNA (mRNA)-based gene therapy are converging with advances in targeted genome engineering to enable efficient, yet transient expression of designer nucleases without risk of undesired integration of the nuclease expression vector.

Figure 1.  Nuclease Mediated Double Stranded Breaks Stimulate Homologous Gene Replacement or Targeted Gene Inactivation.  If targeted nucleases are co-transfected with a homologous donor DNA fragment, homologous recombination replaces defective DNA with a corrected sequence (left).  In the absence of a DNA donor fragment, non-homologous end joining repairs the break, but with frequent insertions and deletions, thus inactivating the gene.


Figure 1. Nuclease Mediated Double Stranded Breaks Stimulate Homologous Gene Replacement or Targeted Gene Inactivation. If targeted nucleases are co-transfected with a homologous donor DNA fragment, homologous recombination replaces defective DNA with a corrected sequence (left). In the absence of a DNA donor fragment, non-homologous end joining repairs the break, but with frequent insertions and deletions, thus inactivating the gene.

Meganucleases

Techniques for making targeted nucleases are rapidly evolving.  Initial attempts to engineer designer restriction nucleases to target new sequences revolved around changing the specificity of naturally occurring nucleases such as meganucleases.  Meganucleases are restriction enzymes with long recognition sites (12-40 nucleotides). These nucleases could be engineered to recognize related sequences in genomes and cleave them.  However, only a small number of sites could be targeted using this approach (refs A-C).

Zinc-finger nucleases (ZFNs)

Zinc-finger nucleases (ZFNs) were the next major advance in the field. Zinc fingers are the most common DNA binding motif in mammalian transcription factors. These sequence specific binding domains can be engineered to bind to novel DNA sequences. Zinc-fingers can be turned into nucleases by fusing them to non-specific cleavage domains, such as the FokI nuclease. FokI cleaves as a dimer, so pairs of ZFNs are designed to bind to adjacent sites in the genome to allow FokI dimer formation and double stranded DNA cleavage (Figure 2). A number of laboratories published design rules that serve as a starting point to engineer ZFNs with novel DNA binding specificities (refs N-R). In reality, actual binding specificity is context dependent. Several selection protocols in cells also exist for identifying novel ZFNs. ZFNs have been successfully used to modify the genomes of Drosophila, C. elegans, zebrafish and rats (refs D-M). However, identification of functional ZFNs remains challenging and most ZFNs have emerged from a small number of laboratories with specialist skills.

 Figure 2. Zinc-Finger Nucleases Bind as Dimers to Cut Double Stranded DNA. Adapted from Gaj et al.Trends Biotechnol. 2013 May 8. [Epub ahead of print]


Figure 2. Zinc-Finger Nucleases Bind as Dimers to Cut Double Stranded DNA. Adapted from Gaj et al.Trends Biotechnol. 2013 May 8. [Epub ahead of print]

Transcription activator-like effector nuclease (TALENs)

In the last few years, TALENs have emerged as a more generally accessible alternative to ZFNs. Like ZFNs, TALENs utilize a modular DNA binding motif (TALE) that can be modified to introduce new DNA binding specificities. TALENs consist of multiple repeat variable diresidues (RVDs) which each specify binding to a single nucleotide (Refs S-U).  TALEN arrays are made by stringing together RVDs in a specific order to provide specificity and binding affinity to novel DNA sequences. Commonly, engineered TALE sequences are fused to non-specific cleavage domains such as FokI. As with ZFNs, TALENs function as pairs bound to adjacent DNA sequences. Unlike ZFNs, TALENs are not as prone to sequence context effects, which greatly complicate the de novo design of ZFNs. This has made them much more accessible to the general scientific community. A number of groups have published TALEN assembly protocols that allow assembly of these repetitive sequences, including one popular open source assembly method is known as Golden Gate (Refs V-Y).

 Figure 3. TALENs Bind as Dimers to Cut Double Stranded DNA. Adapted from Gaj et al.Trends Biotechnol. 2013 May 8. [Epub ahead of print]


Figure 3. TALENs Bind as Dimers to Cut Double Stranded DNA. Adapted from Gaj et al.Trends Biotechnol. 2013 May 8. [Epub ahead of print]

Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)

The newest kid on the genome-engineering block is CRISPR. CRISPR is a bacterial immune system in which bacteria sample the DNA of pathogens, integrate foreign DNA into their genome in specialized repeat structures, and then use these sequences to produce Guide RNAs that direct cutting of homologous pathogenic DNA sequences. To some degree this is reminiscent of RNA interference in mammals. Once the target site has been delineated by the RNA guide sequence, Cas proteins (CRISPR-associated proteins) do the cutting. A number of groups have adapted this system to create RNA directed genome engineering tools (refs Z-CC) (Figure 4). This new system has generated considerable interest since recognition of the target DNA sequence to be cut is RNA mediated rather than protein mediated. DNA cleavage is carried out by the expressed Cas9 protein. With ZFNs and TALENs, if you wish to target a new site, you have to identify and synthesize two new proteins. With CRISPR, you use the same Cas9 protein each time and just alter the sequence of the guide RNA. Stay tuned for head to head comparisons of the efficiency and specificity of ZFNs, TALENs and CRISPR that are about to be published.

 Figure 4. CRISPR is an RNA Guided Genome Engineering System.  Figure adapted from DiCarlo et al. Nucleic Acids Research, 2013, Vol. 41, No. 7.


Figure 4. CRISPR is an RNA Guided Genome Engineering System. Figure adapted from DiCarlo et al. Nucleic Acids Research, 2013, Vol. 41, No. 7.

Modified mRNA for Transient Expression in Genome Engineering

In each of the three systems described above, one needs to express one or two proteins inside cells or an organism.  Plasmids and viral vectors have been used to achieve this, but these carry a risk. Double stranded DNA breaks catalyze insertion of DNA at the cut site.  At some substantial frequency, the protein expression vectors can integrate at the cut site. These vectors necessarily carry eukaryotic promoters, which can lead to continuous expression of the nuclease or the expression of previously silent sequences. For clinical applications this can be a major issue. One also needs to consider off-target cleavage of by engineered nucleases. Since ZFNs have been around longer than TALENs or CRISPR, the most data exists for ZFNs. It is clear that ZFNs can cut at pseudo-sites that resemble the chosen target site.For this reason, transient expression of nucleases is desirable. Many in the ZFN and TALEN field have moved to expression of these nucleases from synthetic mRNAs because they are transient and have no risk of insertion.  Synthetic mRNAs, which mimic fully processed, capped and polyadenylated mRNAs, can be produced in large quantities by in vitro transcription. Transfected mRNAs made with adenine, cytosine, guanine and uracil are recognized as pathogens by innate immune sensors such as Toll-like receptors, RIG-I and PKR. Kariko et al. showed that mRNAs could be made much less immunogenic and non-toxic by substitution of cytosine and uridine with 5-methylcytosine and pseudouridine (ref DD). Custom syntheses of milligram to gram amounts of 5-methylcytosine and pseudouridine modified mRNAs can be ordered from TriLink BioTechnologies. Cas9 mRNA is also available as a catalog item.

Conclusions

In recent years, designer genome engineering has gone from dream to reality. New editing systems are taking this from the realm of a few elite laboratories and companies to democratizing it for the masses. Concurrent advances in mRNA gene therapy are providing safe and effective delivery systems for expressing the necessary components in cells and animals. There is now huge interest in using targeted genome engineering in patient derived somatic cells and stem cells. Rather than simply knocking genes in or knocking them out, we may now be able to actually correct monogenic genetic disorders.  Clinical trials are currently under way to determine if ZFNs can be used to inactivate the CCR5 HIV co-receptor to make patient T-cells immune to HIV. These technologies will also enable facile creation of disease models in species other than mice. The future is bright for targeted genome engineering. That’s the buzz on the cut.

A sincere thanks to Anton McCaffrey for providing this update on truly exciting trends in nucleic acid-based technologie. As always, I welcome comments and discussions.

References

A. I-SceI-induced gene replacement at a natural locus in embryonic stem cells. Cohen-Tannoudji M, Robine S, Choulika A, et al. Mol Cell Biol 1998;18:1444-8.

B. The yeast I-Sce I meganuclease induces site-directed chromosomal recombination in mammalian cells. Choulika A, Perrin A, Dujon B, Nicolas JF. C R Acad Sci III 1994;317:1013-9.

C. Induction of homologous recombination in mammalian chromosomes by using the I-SceI system of Saccharomyces cerevisiae. Choulika A, Perrin A, Dujon B, Nicolas JF. Mol Cell Biol 1995;15:1968-73.

ZFNs modifying different organisms

D. Efficient gene targeting in Drosophila with zinc-finger nucleases. Beumer K, Bhattacharyya G, Bibikova M, Trautman JK, Carroll D. Genetics 2006;172:2391-403.

E. Efficient gene targeting in Drosophila by direct embryo injection with zinc-finger nucleases. Beumer KJ, Trautman JK, Bozas A, et al. Proc Natl Acad Sci U S A 2008;105:19821-6.

F. Targeted chromosomal cleavage and mutagenesis in Drosophila using zinc-finger nucleases. Bibikova M, Golic M, Golic KG, Carroll D. Genetics 2002;161:1169-75.

G. Genetic Analysis of Zinc-finger Nuclease-induced Gene Targeting in Drosophila. Bozas A, Beumer KJ, Trautman JK, Carroll D. Genetics 2009.

H. Induction and repair of zinc-finger nuclease-targeted double-strand breaks in Caenorhabditis elegans somatic cells. Morton J, Davis MW, Jorgensen EM, Carroll D. Proc Natl Acad Sci U S A 2006;103:16370-5.

I. Heritable targeted gene disruption in zebrafish using designed zinc-finger nucleases. Doyon Y, McCammon JM, Miller JC, et al. Nat Biotechnol 2008;26:702-8.

J. Zinc finger-based knockout punches for zebrafish genes. Ekker SC. Zebrafish 2008;5:121-3.

K. Rapid mutation of endogenous zebrafish genes using zinc finger nucleases made by Oligomerized Pool ENgineering (OPEN). Foley JE, Yeh JR, Maeder ML, et al. PLoS ONE 2009;4:e4348.

L. Targeted gene inactivation in zebrafish using engineered zinc-finger nucleases. Meng X, Noyes MB, Zhu LJ, Lawson ND, Wolfe SA. Nat Biotechnol 2008;26:695-701.

M. Knockout rats via embryo microinjection of zinc-finger nucleases. Geurts AM, Cost GJ, Freyvert Y, et al. Science 2009;325:433.

ZFN design rules

N. Toward controlling gene expression at will: selection and design of zinc finger domains recognizing each of the 5′-GNN-3′ DNA target sequences. Segal DJ, Dreier B, Beerli RR, Barbas CF, 3rd. Proc Natl Acad Sci U S A 1999;96:2758-63.

O. Insights into the molecular recognition of the 5′-GNN-3′ family of DNA sequences by zinc finger domains. Dreier B, Segal DJ, Barbas CF, 3rd. J Mol Biol 2000;303:489-502.

P. Validated zinc finger protein designs for all 16 GNN DNA triplet targets. Liu Q, Xia Z, Zhong X, Case CC. J Biol Chem 2002;277:3850-6.

Q. Development of zinc finger domains for recognition of the 5′-ANN-3′ family of DNA sequences and their use in the construction of artificial transcription factors. Dreier B, Beerli RR, Segal DJ, Flippin JD, Barbas CF, 3rd. J Biol Chem 2001;276:29466-78.

R. Development of zinc finger domains for recognition of the 5′-CNN-3′ family DNA sequences and their use in the construction of artificial transcription factors. Dreier B, Fuller RP, Segal DJ, et al. J Biol Chem 2005;280:35588-97.

TALENS

Breaking the code of DNA binding specificity of TAL-type III effectors. S. Boch, J., Scholze, H., Schornack, S., Landgraf, A., Hahn, S., Kay, S., Lahaye, T., Nickstadt, A. and Bonas, U. Science 2009;326,1509-12.

T.  The crystal structure of TAL effector PthXo1 bound to its DNA target. Mak, A.N., Bradley, P., Cernadas, R.A., Bogdanove, A.J. and Stoddard, B.L. Science 2012;335, 716-9.

U. A simple cipher governs DNA recognition by TAL effectors. Moscou, M.J. and Bogdanove, A.J. Science 2009;326,1501.

Golden gate

V. Efficient design and assembly of custom TALEN and other TAL effector-based constructs for DNA targeting. Cermak, T., Doyle, E.L., Christian, M., Wang, L., Zhang, Y., Schmidt, C., Baller, J.A., Somia, N.V., Bogdanove, A.J., Voytas, D.F., Geissler et al. Nucleic Acids Res 2011;39,e82.

W. Modularly assembled designer TAL effector nucleases for targeted gene knockout and gene replacement in eukaryotes. Li, T., Huang, S., Zhao, X., Wright, D.A., Carpenter, S., Spalding, M.H., Weeks, D.P. and Yang, B., Morbitzer et al. Nucleic Acids Res 2011;39, 6315-25.

X. A modular cloning system for standardized assembly of multigene constructs. Weber, E., Engler, C., Gruetzner, R., Werner, S. and Marillonnet, S. PLoS One 2011;6, e16765.

Y. Efficient construction of sequence-specific TAL effectors for modulating mammalian transcription. Zhang, F., Cong, L., Lodato, S., Kosuri, S., Church, G.M. and Arlotta, P. Nat Biotechnol 2011;29,149-53.

CRISPR

Z. A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Jinek, M; Chylinski K, Fonfara I, Hauer M, Doudna JA, Charpentier E. Science 2012;PMID 22745249.

AA. Multiplex genome engineering using CRISPR/Cas systems. Cong, Le; Ran FA, Cox D, Lin S, Barretto R, Habib N, Hsu PD, Wu X, Jiang W, Marraffini LA, Zhang F. Science 2013;PMID 23287718.

BB. RNA-guided human genome engineering via Cas9. Mali, P; Yang L, Esvelt KM, Aach J, Guell M, DiCarlo JE, Norville JE, Church GM. Science 2013;PMID 23287722.

CC. One-Step Generation of Mice Carrying Mutations in Multiple Genes by CRISPR/Cas-Mediated Genome Engineering. Wang, H; Yang H, Shivalila CS, Dawlaty MM, Cheng AW, Zhang F, Jaenisch R. Cell 2013;PMID 23643243.

Modified mRNA

DD. Incorporation of pseudouridine into mRNA yields superior nonimmunogenic vector with increased translational capacity and biological stability. Karikó K, Muramatsu H, Welsh FA, Ludwig J, Kato H, Akira S, Weissman D. Mol Ther. 2008;Nov;16(11):1833-40.