Antibody-targeted chromatin enables effective intracellular delivery and functionality of CRISPR/Cas9 expression plasmids.

We report a novel system for efficient and specific targeted delivery of large nucleic acids to and into cells. Plasmid DNA and core histones were assembled to chromatin by salt gradient dialysis and subsequently connected to bispecific antibody derivatives (bsAbs) via a nucleic acid binding peptide bridge. The resulting reconstituted vehicles termed 'plasmid-chromatin' deliver packaged nucleic acids to and into cells expressing antigens that are recognized by the bsAb, enabling intracellular functionality without detectable cytotoxicity. High efficiency of intracellular nucleic acid delivery is revealed by intracellular expression of plasmid encoded genes in most (∼90%) target cells to which the vehicles were applied under normal growth/medium conditions in nanomolar concentrations. Specific targeting, uptake and transgene expression depends on antibody-mediated cell surface binding: plasmid chromatin of identical composition but with non-targeting bsAbs or without bsAbs is ineffective. Examples that demonstrate applicability, specificity and efficacy of antibody-targeted plasmid chromatin include reporter gene constructs as well as plasmids that enable CRISPR/Cas9 mediated genome editing of target cells.

Quantitative fluorescence imaging determines the absolute number of locked nucleic acid oligonucleotides needed for suppression of target gene expression.

Locked nucleic acid based antisense oligonucleotides (LNA-ASOs) can reach their intracellular RNA targets without delivery modules. Functional cellular uptake involves vesicular accumulation followed by translocation to the cytosol and nucleus. However, it is yet unknown how many LNA-ASO molecules need to be delivered to achieve target knock down. Here we show by quantitative fluorescence imaging combined with LNA-ASO microinjection into the cytosol or unassisted uptake that ∼105 molecules produce >50% knock down of their targets, indicating that a substantial amount of LNA-ASO escapes from endosomes. Microinjected LNA-ASOs redistributed within minutes from the cytosol to the nucleus and remained bound to nuclear components. Together with the fact that RNA levels for a given target are several orders of magnitude lower than the amounts of LNA-ASO, our data indicate that only a minor fraction is available for RNase H1 mediated reduction of target RNA. When non-specific binding sites were blocked by co-administration of non-related LNA-ASOs, the amount of target LNA-ASO required was reduced by an order of magnitude. Therefore, dynamic processes within the nucleus appear to influence the distribution and activity of LNA-ASOs and may represent important parameters for improving their efficacy and potency.

Transcytosis of payloads that are non-covalently complexed to bispecific antibodies across the hCMEC/D3 blood-brain barrier model.

A transcellular shuttle system was generated for the delivery of non-covalently linked payloads across blood-brain barrier (BBB) endothelial cells. Transcytosis-enabling shuttles are composed of bispecific antibodies (bsAbs) that simultaneously bind transferrin receptor (TfR) and haptens such as digoxigenin or biocytinamide. Haptenylated payloads are attached to these vehicles via non-covalent hapten-antibody complexation. This enables targeting to and internalization into human BBB-derived microvascular endothelial hCMEC/D3 cells. In contrast to other shuttles, this system does not require special affinities or formats of their TfR-binding moieties for transcytosis and subsequent release. Non-covalent payload complexation to bsAb is flexible and robust, works for a multitude of payloads and enables separation of payloads from shuttles during transcytosis. Released payloads can enter the brain without connected bsAb entities, minimizing potential interference with distribution or functionality. Intracellular separation of shuttle and payload and recycling to cell surfaces may also enable recharging of the cell-bound BBB shuttle with payload for subsequent (merry-go-round) transport cycles.

Real-Time Cellular Imaging of Protein Poly(ADP-ribos)ylation.

Poly(ADP-ribos)ylation (PARylation) is an important posttranslational protein modification, and is involved in major cellular processes such as gene regulation and DNA repair. Its dysregulation has been linked to several diseases, including cancer. Despite its importance, methods to observe PARylation dynamics within cells are rare. By following a chemical biology approach, we developed a fluorescent NAD(+) analogue that proved to be a competitive building block for protein PARylation in vitro and in cells. This allowed us to directly monitor the turnover of PAR in living cells at DNA damage sites after near-infrared (NIR) microirradiation. Additionally, covalent and noncovalent interactions of selected target proteins with PAR chains were visualized in cells by using FLIM-FRET microscopy. Our results open up new opportunities for the study of protein PARylation in real time and in live cells, and will thus contribute to a better understanding of its significance in a cellular context.

Bioorthogonally Functionalized NAD(+) Analogues for In-Cell Visualization of Poly(ADP-Ribose) Formation.

Poly(ADP-ribos)ylation (PARylation) is a major posttranslational modification and signaling event in most eukaryotes. Fundamental processes like DNA repair and transcription are coordinated by this transient polymer and its binding to proteins. ADP-ribosyltransferases (ARTs) build complex ADP-ribose chains from NAD(+) onto various acceptor proteins. Molecular studies of PARylation thus remain challenging. Herein, we present the development of bioorthogonally functionalized NAD(+) analogues for the imaging of PARylation in vitro and in cells. Our results show that 2-modified NAD(+) analogues perform remarkably well and can be applied to the in-cell visualization of PARylation simultaneously in two colors. This tool gives insight into the substrate scope of ARTs and will help to further elucidate the biological role of PARylation by offering fast optical, multichannel read-outs.

Endocytosis and Endosomal Trafficking of DNA After Gene Electrotransfer In Vitro.

DNA electrotransfer is a successful technique for gene delivery into cells and represents an attractive alternative to virus-based methods for clinical applications including gene therapy and DNA vaccination. However, little is currently known about the mechanisms governing DNA internalization and its fate inside cells. The objectives of this work were to investigate the role of endocytosis and to quantify the contribution of different routes of cellular trafficking during DNA electrotransfer. To pursue these objectives, we performed flow cytometry and single-particle fluorescence microscopy experiments using inhibitors of endocytosis and endosomal markers. Our results show that ~50% of DNA is internalized by caveolin/raft-mediated endocytosis, 25% by clathrin-mediated endocytosis, and 25% by macropinocytosis. During active transport, DNA is routed through multiple endosomal compartments with, in the hour following electrotransfer, 70% found in Rab5 structures, 50% in Rab11-containing organelles and 30% in Rab9 compartments. Later, 60% of DNA colocalizes with Lamp1 vesicles. Because these molecular markers can overlap while following organelles through several steps of trafficking, the percentages do not sum up to 100%. We conclude that electrotransferred DNA uses the classical endosomal trafficking pathways. Our results are important for a generalized understanding of gene electrotransfer, which is crucial for its safe use in clinics.

Visualization of Protein-Specific Glycosylation inside Living Cells.

Protein glycosylation is a ubiquitous post-translational modification that is involved in the regulation of many aspects of protein function. In order to uncover the biological roles of this modification, imaging the glycosylation state of specific proteins within living cells would be of fundamental importance. To date, however, this has not been achieved. Herein, we demonstrate protein-specific detection of the glycosylation of the intracellular proteins OGT, Foxo1, p53, and Akt1 in living cells. Our generally applicable approach relies on Diels-Alder chemistry to fluorescently label intracellular carbohydrates through metabolic engineering. The target proteins are tagged with enhanced green fluorescent protein (EGFP). Förster resonance energy transfer (FRET) between the EGFP and the glycan-anchored fluorophore is detected with high contrast even in presence of a large excess of acceptor fluorophores by fluorescence lifetime imaging microscopy (FLIM).

Direct Monitoring of Nucleotide Turnover in Human Cell Extracts and Cells by Fluorogenic ATP Analogs.

Nucleotides containing adenosine play pivotal roles in every living cell. Adenosine triphosphate (ATP), for example, is the universal energy currency, and ATP-consuming processes also contribute to posttranslational protein modifications. Nevertheless, detecting the turnover of adenosine nucleotides in the complex setting of a cell remains challenging. Here, we demonstrate the use of fluorogenic analogs of ATP and adenosine tetraphosphate to study nucleotide hydrolysis in lysates of human cell lines and in intact human cells. We found that the adenosine triphosphate analog is completely stable in lysates of human cell lines, whereas the adenosine tetraphosphate analog is rapidly turned over. The observed activity in human cell lysates can be assigned to a single enzyme, namely, the human diadenosine tetraphosphate hydrolase NudT2. Since NudT2 has been shown to be a prognostic factor for breast cancer, the adenosine tetraphosphate analog might contribute to a better understanding of its involvement in cancerogenesis and allow the straightforward screening for inhibitors. Studying hydrolysis of the analogs in intact cells, we found that electroporation is a suitable method to deliver nucleotide analogs into the cytoplasm and show that high FRET efficiencies can be detected directly after internalization. Time-dependent experiments reveal that adenosine triphosphate and tetraphosphate analogs are both processed in the cellular environment. This study demonstrates that these nucleotide analogs indeed bear the potential to be powerful tools for the exploration of nucleotide turnover in the context of whole cells.

Intracellular tracking of single-plasmid DNA particles after delivery by electroporation.

Electroporation is a physical method of transferring molecules into cells and tissues. It takes advantage of the transient permeabilization of the cell membrane induced by electric field pulses, which gives hydrophilic molecules access to the cytoplasm. This method offers high transfer efficiency for small molecules that freely diffuse through electrically permeabilized membranes. Larger molecules, such as plasmid DNA, face several barriers (plasma membrane, cytoplasmic crowding, and nuclear envelope), which reduce transfection efficiency and engender a complex mechanism of transfer. Our work provides insight into the way electrotransferred DNA crosses the cytoplasm to reach the nucleus. For this purpose, single-particle tracking experiments of fluorescently labeled DNA were performed. Investigations were focused on the involvement of the cytoskeleton using drugs disrupting or stabilizing actin and tubulin filaments as the two relevant cellular networks for particle transport. The analysis of 315 movies (~4,000 trajectories) reveals that DNA is actively transported through the cytoskeleton. The large number of events allows a statistical quantification of the DNA motion kinetics inside the cell. Disruption of both filament types reduces occurrence and velocities of active transport and displacements of DNA particles. Interestingly, stabilization of both networks does not enhance DNA transport.