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Chapter 3
Hybrid Nucleic Acid Nanocapsules for Targeted, Enzyme-Specific Drug Delivery and Intracellular Gene Regulation Joshua J. Santiana, Saketh Gudipati, Alyssa K. Hartmann, and Jessica L. Rouge* Department of Chemistry, University of Connecticut, Storrs, Connecticut 06033, United States *E-mail:
[email protected].
A combination of surfactant self-assembly and enzymatic ligation is used to build a nanoscale platform for therapeutic oligonucleotide delivery that is highly modular and biochemically responsive to cellular location. Each individual component of the nanocapsule is designed to maximize control over the timing, location, and degree of cargo release. The end product consists of a nucleic acid functionalized nanocapsule built from a micellular core that can controllably degrade or maintain its structure, dictated by the chemical nature of the crosslinking agent that holds the structure intact (i.e., diol linker, ester linker, or peptide, etc.). The versatility of the delivery system enables its potential for broad application. In particular, the nanocapsule has the capacity to deliver both small molecule cargo in combination with therapeutic oligonucleotides, including siRNA, microRNA, DNAzymes, and aptamers. This chapter presents specific applications of this delivery vehicle. To date, the delivery system has been shown to be non-toxic in cell culture, and it can effectively deliver dyes and drugs, among other small molecules, into cells. The release of these cargo can be tuned to the presence of specific enzyme expression levels in cells. Notably, the complete degradation of the particle results in the release of oligonucleotides modified with a terminal hydrocarbon chain, which aid its disruption and release from endosomal
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compartments. Targeted gene knockdown has been achieved using DNAzyme-functionalized nanocapsules, resulting in up to 60% reduction in gene expression without the use of transfection agents, making it an ideal co-delivery vehicle for drugs and oligonucleotides.
Introduction The past 20 years have seen significant advancements in delivering exogenous nucleic acids into cells, enabled by chemical modifications (1), viral expression (2), or encapsulation in nanomedicine formulations (3–5). The diversity of therapeuetic nucleic acids that are of interest to deliver into cells has also expanded over the last decade, including antisense oligonucleotides (ASOs), short interfering RNA (siRNA), short hairpin RNA (shRNA), and catalytic oligonucleotides such as DNAzyme and ribozymes, useful for a range of gene silencing and gene therapy applications (6, 7). Generally speaking, the mechanism that many therapeutic oligonucleotides use is targeted association with endogenous mRNA, catalyzing its cleavage or decreasing association with translational machinery, thereby resulting in altered gene expression. In recent years, nanomedicine formulations have markedly improved researchers’ ability to deliver nucleic acids into cells, doing so by shielding or encapsulating the therapeutic oligonucleotides as a way to protect them from rapid chemical and nuclease-mediated degradation. Many of the most promising formulations that have been developed are in the form of liposomes, which can effectively deliver nucleic acids into cells by fusing with a cell’s lipid bilyaer. To date, liposomes have also been highly effective commercially for delivering small-molecule drugs, such as the well-known Doxil® formulation. However, for nucleic acid delivery, their clinical application has been slower to progress (8). In a recent and significant step forward, Alnylam Pharamceuticals was granted FDA approval in 2018 for the first nucleic acid therapy involving a liposome, accomplished by incorporating chemistries that enable better biodegradability. This was an exciting step for liposomal siRNA formulations. Despite this promising approval, there remains to be a broad application of liposomal formulations for nucleic acid delivery in the clinic due to often toxic concentrations required to achieve responses in cells. It was not until 2006 that a major paradigm shift in the way nucleic acids can be delivered into cells occurred that opened up a new approach to nucleic acid delivery, and with it, a new avenue for achieving therapeutic effects in cells using oligonucleotides (9). This new route can be traced back to 1996, when researchers showed that thiolated DNA adhered to a gold nanoparticle could be controllably assembled at a nanoparticle surface for a variety of applications (10). This new structure was later used for the delivery of oligonucleotides into cells (9). For the first time, chemically unmodified nucleic acids could rapidly be delivered intact into cells, despite their outward surface display unshielded from cellular nucleases. This was surprising, considering the particles’ overall negative charge and that nucleic acids would seem vulnerable in this orientation. It was later found that 60
scavenger receptors on the outside of cells were responsible for their uptake, showing affinity for this structure resulting in rapid endocytosis (11). In addition, such structures provided a route to nucleic acid stability due to their spherical arrangement of oligonucletides. Such an arrangement could sterically shield neighboring molecules from nucleases due to their high loading density at the particles’ surface. These structures, now collectively known as spherical nucleic acids (SNAs) (12), have been shown to have numerous advantageous attributes for nucleic acid delivery, ranging from rapid and high levels of cellular entry, to low toxicity and extended half lives of their nucleic acid ligands. One of the most significant results of this new structure was that the density of the nucleic acid ligands at the surface of the particles were found to protect RNA and DNA ligands from degradation, and were the main reason why the nucleic acid ligands did not require any additional chemical modifications (13). Since this discovey, many different SNA-inspired particle types and compositions have been generated that take advantage of this formulation style, achieving rapid cellular uptake and knockdown of target genes (14, 15). Despite the SNAs’ structure success to date, there are two major areas in which the formulation could be further improved to help transition therapeutic nucleic acids into the clinic. First, the ability to target specific cell types has yet to be built into these structures, a limitation still facing many nucleic acid delivery platforms. Second, the fact that the core is not fully biodegradable means that there is a need to change the core composition so more of the structure can play an active role in the material’s overall therapeutic capability. Fortunately, the SNAs’ advantageous properties have been show to be universal, independent of the core’s nanoparticle composition. Some of these issues have therefore been partially addressed in the formation of the liposomal SNA (16), which combines the advantageous properties of SNAs with those of traditional liposomes, and through more direct methods of SNA-drug pairing, seen through conjugation of the SNA’s oligonucleotide ligands with drug molecules such as cisplatin (17). However, even with better degradability there needs to be greater control over which cells receive the therapeutic oligonucleotides—something that would program the structures to release the nucleic acid cargo in diseasesed cells rather than healthy cells.
Designing a Hybrid Nanocapsule for Targeted Nucleic Acid Delivery In order to address these outstanding challenges facing the therapeutic nucleic acid field, our group has recently developed a new nanocapsule specifically designed for targeted nucleic acid delivery. This new construct, which we refer to as a nucleic acid nanocapsule (NAN) (18), takes inspiration from the cellular uptake advantages of the SNA structure and combines it with a recent advancement in the assembly of nucleic acids at the nanoscale level using enzymes (Figure 1). In 2014, it was shown that T4 DNA ligase could be used to further functionalize DNA-coated nanostructures, including SNAs, in order to accommodate siRNA (19) and more highly folded thereapuetic nucleic acids such 61
as ribozymes (20). Once assembled, these hybride RNA-DNA SNA structures could enter cells and effectively result in gene knockdown, directed by the siRNA and ribozyme structures.
Figure 1. Assembly of a nucleic acid nanocapsule. Starting with the surfactant in water, a micelle structure can be rapidly formed. This is followed by copper catalyzed click chemistry using the diazido cross-linker (Figure 2, compound 1). Once cross-linked, the remaining alkyne sites on the surface of the crosslinked micelle are functionalized with thiolated DNA strands using a thiol-yne reaction, uv light, and a photoinitiator. This structure is then broken down by esterases upon endocytosis. Adapted with permission from ref (18). Copyright 2017 American Chemical Society.
The NAN formulation presented in this chapter describes a hybrid SNA-like structure that uses this enzyme-mediated approach to both assemble the nucleic acid ligands on the surface of a nanoscale material and utilize enzymes to controllably disassemble the oligonucleotide ligands, making it a fully biodegradable, nucleic acid delivery platform with the potential for targeted gene knockdown (18). This new hybrid structure has a micellular core built from a multifunctional surfactant molecule and consists of a densely coated nucleic acid functionalized surface, similar to an SNA (Figure 1). We specifically refer to the structure as a nucleic acid nanocapsule in order to indicate that unlike a traditional SNA structure, the core is able to serve a therapeutic function as well. In this case, it has two functions. The first is more traditional—due to its micellular core, it has the ability to encapsulate a cargo for delivery within its hydrophobic interior. The second and more unique role is with respect to how the nucleic acid is released during the particle’s degradation. As the nanocapsule degrades, it releases a hydrophobically modified oligonucleotide, a modification that 62
has been shown to be advantageous for crossing lipid bilayers (21, 22). The NAN’s hydrophobic core, unlike traditional nucleic acid delivery platforms of liposomes that encapsulate nucleic acids, is able to deliver small molecules with the oligonucleotide ligands situated at the surface. This is similar to the structural advantages of DNA-amphiphile assemblies, where nucleic acid ligands can be controllably released in response to external triggers such as light, pH, and redox conditions (23, 24). However, one significant difference in the NAN’s formulation is the incorporation of biochemically specific triggers, namely enzymes through the incorporation of ester and peptide linkages described later in this chapter. These properties make the NAN a unique and highly attractive nucleic acid delivery platform for its ability to co-deliver oligonucleotides and small molecules simultaneously, and builds upon the cellular uptake advantages of the SNA. Notably, the enzyme-mediated approach to assembling nucleic acid ligands at the surface of the NAN provides a route to multiplexing it with combinations of nucleic acid ligands, ranging from targeting oligonucleotides to therapeutic sequences. The structure and synthesis of the NAN and its applications are described in detail below.
Nucleic Acid Nanocapsule (NAN) Synthesis and Assembly The first step in the synthesis of a NAN involves self-assembly of its micelle core. The surfactant (Figure 1) that forms the nanocapsule’s core was inspried by the work of Zhao and coworkers (25), sharing a similar trialkylated head group. However the surfactant’s tail is slightly modified, specifically to make the tail a simplified (C12) chain (synthesized in-house through an SN2 reaction between dodecylamine and propargyl bromide) (18). This was done in order to generate an unstable micelle building block that only becomes stabilized if its head group is crosslinked post assembly. This is an important step in the nucleic acid delivery function of the surfactant and will be described in detail later in this chapter. NAN assembly begins in water, where the surfactant is stirred for 30 min to dissolve and form micelles at a concentration of 10 mM (the critical micelle concentration determined for this surfactant). At this step, a cargo can be loaded into the micelle core through hydrophobic association between the small molecule and the surfactant tail. Once assembled, the hydrophilic head group of each surfactant molecule can be reacted with azides or thiols in a variety of combinations, as the head group contains three alkyne moieties that it presents at the micelle’s surface. These alkyne moieties are utilized to undergo various “click” reactions in order to both cross-link and functionalize the surface of the NAN. Two common click reactions that are performed for the synthesis of the NAN include copper(I)-catalyzed alkyne-azide cycloaddition (CuAAC) and the thiol-yne reaction. CuAAC reactions utilize a copper(II) salt and a reducing agent such as sodium ascorbate in order to reduce copper(II) to copper(I) for catalyzing the cycloaddition between the alkyne and azide compounds. The thiol-yne reaction is driven by UV light and a water soluble photoinitiator, 2-hydroxy-4-(2-hydroxyethoxy)-2-methylpropiophenone (DHEMPP). The 63
next step after formation of a loaded or unloaded micelle is to stabilize it by cross-linking with diazido or thiol terminated cross-linkers (Figure 2). Here, it is very important to consider the stoichiometric relationship between the alkyne moieties and the cross-linker. A 1:1.2 ratio of terminal azide cross-linker to alkyne moiety is used for CuAAC reactions, while a 1:2 ratio of terminal thiol cross-linker to alkyne moiety is used for thiol-yne reactions. The ratio of the cross-linker is reduced for the thiol-yne reaction in order to avoid polymerization of micelles. In both cases, the stoichiometric ratios leave at least one alkyne moiety unreacted and available for further functionalization.
Figure 2. Cross-linkers utilized in the synthesis of nucleic acid nanocapsules. (1) A diazido ester cross-linker for cleavage by esterases, (2) a negative control cross-linker, containing a diol linkage (25), and (3) thiolated peptide cross-linker used in the synthesis of peptide nucleic acid nanocapsules.
After the selected cross-linking reaction is complete, the newly surface crosslinked micelles (SCMs) can be purified by size exclusion chromatography using G25 Sephadex columns. The final step in the NAN synthesis is the functionalization of the remaining alkyne moieties. This is done using a thiol-yne reaction between the alkyne moiety at the SCM surface and thiolated DNA. Here, the stoichiometric ratio of DNA to alkyne is approximately 2:1. This ratio is used because the thiolyne reaction can form covalent bonds with up to two thiolated DNA molecules via formation of carbon-sulfur bonds. This is preferred because increasing the amount of DNA reacting with the alkyne moieties increases the grafting density of the DNA on the NAN surface, leading to the benefits of SNA-like structures, such as high cellular uptake and steric protection of the nucleic acid ligands without chemical modification (18). Once functionalization with DNA is complete, the NANs can be purified again using size exclusion chromatography to remove any DNA that is not covalently attached to the NANs’ surface. 64
Figure 3. Characterization of NANs. A) DLS of crosslinked micelles and DNA functionalized NANs. B) Electron microscopy of NANs post-uranyl acetate staining—uniform micelle-like structures can be observed. C) Zeta potential measurements showing the shift in charge pre- and post-DNA functionalization. D) Agarose gel electrophoresis showing the changing surface charge of crosslinked micelle in lane 1 versus the DNA functionalized NAN in lane 2. Adapted with permission from ref (18). Copyright 2017 American Chemical Society.
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Nanoscale Characterization of NANs During the course of the NAN’s synthesis, it is important to monitor the nanocapsule’s properties at each step (SCMs and NANs) using various characterization methods. The methods most commonly used for characterization of NANs include dynamic light scattering (DLS), zeta potential measurements, transmission electron microscopy (TEM), and agarose gel electrophoresis. DLS is a useful technique for measuring the hydrodynamic size of nanoparticles based on the principle that changing material size will change the amount of light scattered from a solution. It is particularly advantageous for measuring the hydrodynamic size of SCMs and NANs because the DLS measurement will show a shift in size after functionalization with DNA. The change in size can be estimated and compared to the known size of the ligand that is being attached to the surface of the SCM. For example, SCMs have an average size of 20 nm. A single base pair in a DNA strand is approximately 3.4 Å. If a 30mer DNA strand is tethered to the surface of the SCM, the overall size of the nanocapsules should increase by 20 nm. An example of the shift in size from SCM to NAN can be observed in Figure 3A. TEM images can be paired with DLS measurements to characterize the size as well as degree of aggregation, if any. Zeta potential is then used to determine the degree of DNA loading on the surface of the NAN. If the structures of both the SCM and NAN are considered, there is a positively charged quaternary amine at the surface of the SCM and there is negatively charged DNA at the surface of the NAN. Taking zeta potential measurements at each of the two stages (SCM and NAN) shows a shift from positive to negative (+50 mV to –50 mV, respectively), indicating successful attachment of DNA. The significant change in surface charge of the nanocapsules can also be applied in agarose gel electrophoresis. Both SCMs and NANs are able to migrate through the gel; however, they travel in opposite directions. In order to track the nanocapsules through an agarose gel, a fluorescent dye, such as Rhodamine B, can be encapsulated inside of the construct during assembly. As seen in Figure 3D, the SCMs and NANs travel in opposite directions, toward the corresponding electrode. This is another indication of the successful attachment of DNA to the surface of the SCMs.
Incorporation of Enzyme Cleavable Cross-Linkers During the assembly process of a NAN, a variety of different cross-linkers can be incorporated as the trigger that will dictate when and where a NAN will degrade. It is therefore the cross-linker that will target the delivery of the oligonucleotide and/or small molecule drug. This modular aspect of the NAN is one of the structure’s major advantages. The only requirement is that the cross-linker must have either terminal azides or thiols. Ideally, the cross-linker that is selected should have the ability to degrade in the presence of a particular biological target in order to release its cargo. As a proof-of-concept experiment, the first cross-linker was a simple azide terminated ester compound (compound 1, Figure 2). The idea behind this cross-linker was that it can be cleaved by 66
the ubiquitous enzyme esterase. If esterase can cleave the ester linkages at the surface of the nanocapsules, the cargo could escape during degradation of the nanocapsules. As esterase is known to be concentrated inside cellular endosomes (26), it was hypothesized that this is where the nanocapsule’s degradation and cargo release would occur. In order to test this hypothesis, a fluorometric assay was developed. In this assay, a fluorescent dye (Rhodamine B) was encapsulated inside of an ester cross-linked NAN and exposed to porcine liver esterase. The fluorescence signal was monitored over time to see if the cargo release could be observed. As seen in Figure 4, the fluorescence signal of the ester cross-linked NANs increased after exposure to esterase over time. In order to ensure that the increase in fluorescence signal was due to the enzymatic degradation of the nanocapsules, a control cross-linker (compound 2, Figure 2) was synthesized and incorporated into an additional batch of NANs. This cross-linker is also azide terminated; however, there are no functional groups that can be degraded by an esterase. This nonester cross-linked NAN underwent the same assay and showed no release of cargo (Figure 4B). This lack of dye release shows the importance of the cross-linker and enzymatic environment of the NAN for controlling its degradation and subsequent release of cargo.
Figure 4. Profiles of NAN dye release using fluorescence assays. Adapted with permission from ref (18). Copyright 2017 American Chemical Society. 67
The next step was to show successful release of cargo in an in vitro cell assay. Two separate batches of NANs were synthesized and encapsulated with camptothecin, (a well-known cytotoxic cancer drug), then cross-linked with either the ester or non-ester cross-linker. HeLa cells were then incubated with the nanocapsules for 24 h with these ester cross-linked NANs (Figure 4C) and non-ester cross-linked NANs (Figure 4D). The results of this experiment reiterated the relationship between the NANs’ stability and its cross-linker and enzymatic environment. The ester cross-linked NANs were successfully degraded in the endosome, releasing the camptothecin and killing the cells in a dose-dependent manner (Figure 4C). The non-ester cross-linked NANs, however, stayed intact, keeping the cargo from being released into the endosome. While the development of this nanocapsule depended on the proof-of-concept that using an ester cross-linker would engage esterases, it was of interest to expand the identity of the cross-linkers to other more biochemically specific enzyme targets. The biggest disadvantage to the ester cross-linker is its lack of specificity. If the NAN is to be used as a therapeutic option for a disease such as cancer, it is vital that the specificity level of the NAN’s degradation be highly targeted to cancer cells. Since esterase is ubiquitous, a cytotoxic cargo can be released in both healthy and diseased cells, potentially causing undesirable side effects. However, for the delivery of non-toxic drugs such as anti-inflammatories, the use of ester-crosslinked NANs could be a viable option.
Modular Nature of the NAN’s Cross-Linker Provides a Route to Targeted Delivery The next phase in the development of the NANs was to introduce a new type of cross-linker that could increase the degree of specificity between the NAN and its intended biological target or environment. To do this, we took advantage of the fact that in certain diseases, specific enzymes tend to be overexpressed, particularly during periods of inflammation and/or during increased cell proliferation, as seen in many cancers. For example, it has been shown that cathepsins, a class of proteases, have high levels of expression in pancreatic and mammary carcinomas (27) and matrix metalloproteinases (MMPs, another class of proteases) are upregulated in numerous cancers (28). With this in mind, we focused on testing two specific enzymes, cathepsin B and MMP9. These enzymes were chosen as they both have well-studied peptide substrates (29, 30) and have been shown to have a high degree of specificity for their peptide substrate (31). These peptide substrates were modified with terminal cysteine residues (Figure 2, compound 3) in order to introduce terminal thiol residues to each substrate for incorporation into the NAN. Here, the thiol-yne reaction was utilized as a cross-linking method to form two batches of NANs that were cross-linked together using either the MMP9 peptide substrate GPLGLAGGERDG or the cathepsin B substrate GFLG (Figure 5). After successful characterization of the new peptide NANs in the same manner described previously, fluorescent assays were carried out in order to test the release of dye in the presence of the appropriate enzymes (32). 68
Figure 6A and 6B show release of cargo from both MMP9-NANs and CathB-NANs in the presence of their respective enzyme. However, it was important to demonstrate the resilience of the NANs at the approproiate pH as well as in the presence of other proteases. Shifting the focus to MMP9-NANs, the dependence on pH was tested first. This is important because MMP9 is an extracellular protease that functions optimally at pH 7. There was a lack of cargo release at pH 5 while in the presence of MMP9 (Figure 6C). In addition to pH dependence, it was important to show that there was a lack of cargo release in the presence of proteases within the same protease (MMP) family. Figure 6D shows the lack of cargo release when MMP9-NANs are exposed to MMP1 and MMP2. This was a particularly interesting result, as other reports have shown that MMP2 can in fact cleave MMP9 substrates (33). Our current hypothesis for why we see cleavage by MMP9 and not MMP2 for our MMP9 peptide cross-linked NANs is that the orientation of the peptide cross-linker is more favorable for MMP9 than MMP2. Another factor that makes a difference is the enzymes’ accessibility to the tethered peptide substrate. Ongoing kinetic analysis of the two enzymes on the NAN substrates will be compared to determine whether the MMP2 enzyme would eventually cleave the MMP9 peptide crosslinked NANs, if given sufficient time. For now, the observed difference in degradation rates is promising proof that specific peptide cross-linkers can enhance the overall specificity of the NAN’s implementation in biological systems using enzymes.
Figure 5. Peptide NAN assembly. Using a method similar to the original ester-cross-linked NAN assembly, surfactant molecules are first assembled in water, followed by cross-linking via thiol-yne chemistry between a cysteine-modified peptide and the alkynes of the surfactant. Thiol-yne reaction conditions are similar to those used for the functionalization of the NAN with DNA ligands. Adapted with permission from ref (32). Copyright 2017 American Chemical Society. 69
Figure 6. Peptide NAN cargo release. A) Fluorescence monitoring of MMP9 peptide NANs treated with MMP9. B) Dye release from Cathepsin B NANs treated with Cathepsin B enzyme. C) Comparison of MMP9 treatment of MMP9-NANs at pH 5 vs pH 7, and compared to treatment with Cathepsin B. D) MMP9-NANs treated with closely related MMP enzymes (MMP1 & MMP 2). Adapted with permission from ref (32). Copyright 2017 American Chemical Society. In addition to the fluorometric analyses, it was important to conduct in vitro cell experiments to ensure that the cargo was being released in the same, enzyme-specific fashion when internalized into cells. In order to test this, peptide cross-linked NANs, along with the original ester and diol cross-linked NANs were assembled around gold nanoparticle cores. This was done by using a gold nanoparticle that is functionalized with an alkanethiol, thus making it hydrophobic. This method was then used as the template for assembling the NAN and further cross-linked with either an MMP9 peptide, a cathepsin B peptide, an ester cross-linker, or a diol cross-linker. All four kinds of NANs were incubated separately with HeLa cells. After incubation, the cells were fixed with paraformaldehyde, embedded in resin, and sectioned for imaging using a transmission electron microscope. In all four cases, the presence of the NANs could be observed inside endosomal comparments of the cells (Figure 7). Notably, the diol cross-linked control shows that the NANs stay intact as seen in Figure 7A. This same effect is seen in the cathepsin and MMP9 Au NANs (Figure 7C and 7D). However, the ester crosslinked NANs appeared to lose their outer organic shells, indicated by the lack of gray halo in the TEM image as well as its apparent aggregation, typical of gold nanoparticles that have no passivating ligands at their surface (Figure 7B). 70
Figure 7. Various AuNP NANs observed in HeLa cells by electron microscopy. A) Diol-crosslinked AuNP NANs. B) Ester-crosslinked AuNP NANs. C) Cathepsin B AuNP peptide crosslinked NANs. D) MMP9 AuNP peptide crosslinked NANs. E) PMA treatment of HeLa cells with simultaneous treatment with MMP9 peptide crosslinked NANs. F) PMA treatment followed by treatment with MMP9 peptide crosslinked NANs 20 h later. Adapted with permission from ref (32). Copyright 2017 American Chemical Society.
Having confirmed the uptake of cargo via the peptide NANs using the traceable gold nanoparticle (Au NP) tag, it became of interest to test whether the presence of a specific enzyme in a cell could initiate the opening of the NAN to release a drug. In order to test this, phorbol 12-myristate 13-acetate (PMA) was used to induce the expression of MMP9 in cultured HeLa cells. Since MMP9 is an extracellular protein, timing was very important during the NAN treatment process. It was essential that PMA was incubated with HeLa cells before the MMP9-NANs (encapsulated with camptothecin) were introduced to assess drug release as a function of change in total cell viability. The translation of MMP9 takes longer than the endocytosis of the NANs. If both MMP9-NANs and PMA were incubated with HeLa cells at the same time, the MMP9-NANs would be endocytosed before a sufficient amount of MMP9 was present in the extra cellular space, meaning the degradation of the MMP9-NANs would not be possible (Figure 7E). In allowing additional time for the translation of MMP9, the viability of HeLa cells is affected (Figure 7F). The results indicate a dependence on the concentration of PMA incubated with the cells. Overall, we were able to show an increased level of specificity with the new pep-NAN system as it relates to the expression level of the trigger enzyme target. These experiments suggested that the NAN platform has signficant future potential due to its ability to program the cross-linker to a target enzyme in a specific biological location. 71
Nucleic Acid Nanocapule Ligands for Gene Regulation Using the nucleic acid nanocapsule platform, it is possible to deliver chemically unmodified, therapeutically active nucleic acid ligands without needing transfection agents to achieve gene knockdown. This was tested using a DNAzyme as a proof-of-principle oligonucleotide. The specific DNAzyme that was chosen for this study is the GATA-3 DNAzyme that was recently isolated via in vitro selection for the specific and targeted cleavage of the mRNA sequence that encodes GATA-3 protein (34, 35). GATA-3 is a transcription factor that is involved in the differentiation of Th2 cells in inflammation pathways. Because of this critical role, GATA-3 has shown promise as a potential target for the treatment of inflammatory diseases (36). DNAzymes are catalytically active nucleic acid molecules that specifically bind and cleave mRNA with high rates of turnover. However, DNAzymes are commonly chemically modified to increase their stability, and transfection agents are often used to aid their entry into cells, much like the modifications for siRNA and ASOs. In order to assess whether the NAN’s structure could afford a similar stability to oligonucleotides, as seen by other SNA-like structures, we functionalized the NAN with the GATA-3 DNAzyme and assessed its activity both by in vitro fluorescence assays and by cell culture experiments (37). DNAzyme-functionalized NANs (DNz-NANs) were assembled as previously described (15). The GATA-3 DNAzyme was synthesized in-house, and was chemically unmodified with the exception of a monophosphorylated 5’ end. This end is enzymatically ligated to the 3’ hydroxyl of the DNA presented on the NAN surface. The size of the NAN can be determined by DLS pre- and post-DNAzyme ligation. The DLS data suggest that the NANs and DNz-NANs are both monodisperse in size, and successful ligation is seen through the increase in size from roughly 20 nm to 60 nm. The DNz-NAN construct can also be characterized through agarose gel electrophoresis and zeta potential, which indicate that the DNz-NANs carry a -30 mV charge. The number of DNAzymes per surfactant molecule was also determined to compare this formulation with more traditional delivery methods for nucleic acids, such as Lipofectamine 2000. To assess this, the DNAzyme was modified with a 3’ TYE563 dye and ligated to the NAN surface (Figure 7). Post-ligation, free DNAzymes were removed by size exclusion chromatography. The fluorescence of the DNz-NAN construct was measured and compared to a standard curve of the TYE563 DNAzyme, resulting in the total concentration of DNAzymes. Comparing this to the known surfactant concentration, we were able to determine that there were 2.3 ± 0.2 DNAzymes per surfactant molecule (37). This result indicates the high efficiency of the NAN assembly and ligation steps. Due to the ester cross-linker, it was anticipated that the esterase-mediated cleavage of the nanocapsule would result in DNAzymes that are covalently modified with individual surfactant molecules (Figure 8). It was also anticipated that the hydrophobic chain modification on the DNAzyme will aid in endosomal escape.
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Figure 8. Dye-labeled DNAzyme-NANs treated with esterase.
Figure 9. Gel electrophoresis assay monitoring truncated GATA-3 mRNA cleavage. A) Denaturing 8% polyacrylamide gel showing the extent of cleavage by the DNAzyme NAN (Dnz-NAN) compared to the free DNAzyme. Mutated DNAzymes were used as negative controls. B) The relative extent of mRNA cleavage was also monitored using a fluorescence quencher assay, which resulted in fluorescence, as seen in the plots in B. C) Schematic depiction of the assay used in A. Adapted with permission from ref (18). Copyright 2017 American Chemical Society. 73
After functionalization and characterization of the DNz-NANs, it was of interest to determine if the DNAzyme retained its catalytic activity while immobilized on the nanocapsule surface. To achieve this, we performed an in-solution activity assay comparing cleavage of the Cy3-labeled mRNA target sequence between free DNAzyme and DNz-NANs under the selection conditions (100 mM NaCl, 10 mM MgCl2, 37 ºC). Free DNAzyme results in complete cleavage, and the DNz-NAN results in almost complete mRNA cleavage (Figure 9A). When four point mutations are introduced in the binding region of the DNAzyme, there is no longer cleavage of the mRNA target sequence. These results encouraged us to proceed with the DNz-NAN in cell culture to see if it could undergo cellular uptake and successful knockdown of GATA-3 mRNA. To investigate this, MCF-7 cells were treated with 100 nM GATA-3 siRNA, 300 nM free DNAzyme, and 250 nM DNz-NAN for 4 h. It is important to note that both the siRNA and free DNAzyme required Lipofectamine 2000 to undergo cellular uptake. Preliminary qPCR analysis of the mRNA expression showed that the DNz-NAN was able to achieve 60% knockdown of GATA-3 mRNA without the need for a transfection agent (Figure 10).
Figure 10. Preliminary gene knockdown results using the DNAzyme-NAN. Compared to untreated cells, the DNzyme-NAN can knock down the expression of GATA-3 by ~ 60% as quantified by qPCR. Adapted with permission from ref (37). Copyright 2018 Wiley-VCH.
These studies have shown that functionalization of the NAN core with a chemically unmodified DNAzyme results in cellular uptake and mRNA knockdown without the need for transfection agents. This is likely due to a similar engagement with scavenger receptors, as seen with SNA-like structures that undergo endocytosis. However, a major difference between the SNA and the NAN is that this form of nucleic acid delivery can be paired with small-molecule delivery. 74
Conclusions and Perspectives In summary, the NAN platform provides a number of exciting advancements for targeting nucleic acids delivery within cells. Although this chapter only discusses DNAzyme delivery as the proof-of-concept therapeutic nucleic acid ligand, we are actively working on gene knockdown using siRNA and shRNA functionalized NANs. We are also pursuing aptamer-functionalized NANs to bias the uptake of nanocapsules into specific cell types. Due to the versatility of the enzyme-mediated assembly of the nucleic acid ligands, one can use the ligation method to reliably mix and match a variety of unique sequences at desired stoichiometries at the nanocapsule’s surface. The next phase of NAN synthesis will focus on the potential synergistic therapeutic effects made possible by the co-delivery of a gene silencing oligonucleotide and a small-molecule drug, both of which can be controllably released under unique biological conditions dicated by the enzyme expression levels within cells. With this level of biochemical responsiveness designed into the NAN’s overall structure, it is easier to attain greater specificity over the resulting therapeutic response.
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