Article pubs.acs.org/JACS
Cite This: J. Am. Chem. Soc. 2018, 140, 1632−1638
Oligonucleotide−Peptide Complexes: Phase Control by Hybridization Jeffrey R. Vieregg,†,# Michael Lueckheide,‡,# Amanda B. Marciel,† Lorraine Leon,§ Alex J. Bologna,† Josean Reyes Rivera,∥ and Matthew V. Tirrell*,†,⊥ †
Institute for Molecular Engineering, University of Chicago, Chicago, Illinois 60637, United States Department of Chemistry, University of Chicago, Chicago, Illinois 60637, United States § Department of Materials Science & Engineering, University of Central Florida, Orlando, Florida 32816, United States ∥ Departamento de Ciencias Biológicas, University of Puerto Rico at Rio Piedras, San Juan, Puerto Rico 00925, United States ⊥ Institute for Molecular Engineering, Argonne National Laboratory, Argonne, Illinois 60439, United States ‡
S Supporting Information *
ABSTRACT: When oppositely charged polymers are mixed, counterion release drives phase separation; understanding this process is a key unsolved problem in polymer science and biophysical chemistry, particularly for nucleic acids, polyanions whose biological functions are intimately related to their high charge density. In the cell, complexation by basic proteins condenses DNA into chromatin, and membraneless organelles formed by liquid−liquid phase separation of RNA and proteins perform vital functions and have been linked to disease. Electrostatic interactions are also the primary method used for assembly of nanoparticles to deliver therapeutic nucleic acids into cells. This work describes complexation experiments with oligonucleotides and cationic peptides spanning a wide range of polymer lengths, concentrations, and structures, including RNA and methylphosphonate backbones. We find that the phase of the complexes is controlled by the hybridization state of the nucleic acid, with double-stranded nucleic acids forming solid precipitates while single-stranded oligonucleotides form liquid coacervates, apparently due to their lower charge density. Adding salt “melts” precipitates into coacervates, and oligonucleotides in coacervates remain competent for sequence-specific hybridization and phase change, suggesting the possibility of environmentally responsive complexes and nanoparticles for therapeutic or sensing applications.
■
INTRODUCTION In addition to storing information in their sequence, nucleic acids are among the most highly charged polymers known (axial charge density of 6 e−/nm for double-stranded B-form DNA), and interact strongly with other charged molecules in the cell. A striking example of this is chromatin formation, in which genomic DNA is condensed several thousand-fold into μm-size chromosomes, a process mediated by basic histone proteins and polyamines.1,2 RNA plays a key role in many membraneless organelles (also known as “biomolecular condensates”), intracellular phase-separated droplets assembled at least partially by electrostatic interactions. These are implicated in numerous essential cellular processes, including regulation of gene expression, embryonic development, and metabolism, and have recently been linked to diseases of aggregation such as Alzheimer’s and ALS.3−6 Phase separation can also occur when individual nucleotides are mixed with simple polycations, and the resulting droplets readily encapsulate oligonucleotides, an observation of great interest to early life scenarios and the “RNA world” hypothesis.7 Understanding the interactions of long, double-stranded DNA © 2018 American Chemical Society
(dsDNA) with cationic small molecules and polymers has been one of the classic problems of biophysical chemistry for nearly 60 years, but, despite its importance, we know far less about the behavior of short or single-stranded nucleic acids, a deficit that motivates the present study. Electrostatic interactions are also the dominant tool used to assemble therapeutic nucleic acids for delivery into cells in vitro and, increasingly, in vivo. Synthetic polycations such as polyethylenimine and polylysine (pLys) readily form polyelectrolyte complexes with nucleic acids, and have been used for many years to deliver dsDNA to cells, as well as more recently to encapsulate large DNA nanostructures.8−10 If the polycation is conjugated to a neutral hydrophilic block such as polyethylene glycol (PEG), then phase separation occurs on the nanoscale, producing “polyelectrolyte complex core micelles” that show great promise for gene delivery in vivo.11,12 Oligonucleotides and RNA should also be amenable to polyelectrolyte delivery, but our knowledge of even basic Received: April 9, 2017 Published: January 9, 2018 1632
DOI: 10.1021/jacs.7b03567 J. Am. Chem. Soc. 2018, 140, 1632−1638
Article
Journal of the American Chemical Society
This manuscript describes results of an investigation of the effects of polymer length, structure, concentration, and salt concentration on the complexation of oligonucleotides by cationic peptides and polyamines. The use of defined sequence oligomers enables exploration of a large, yet well-defined parameter space in molecular charge, length, and structure. By working at relatively high concentration, we are able to visualize the complexes directly, enabling a clear determination of their properties. These differ in several interesting ways from those obtained with longer polymers, and should inform both basic questions of biopolymer complexation and delivery of therapeutic nucleic acids.
structure−property relationships is incomplete, with different research groups reporting widely divergent results when RNA oligonucleotides are complexed with cationic peptides.8,12−17 Improved knowledge of the complexation properties of oligonucleotides is therefore required to enable rational design of more effective delivery vehicles. In aqueous solution, long dsDNA forms phase-separated polyelectrolyte complexes when mixed with cations with charge ≥3 or with cationic polymers. The dominant driving force for complexation is entropy gain from release of low-valence counterions;18 at low DNA concentrations, Poisson−Boltzmann models accurately predict key quantities such as the fraction of DNA charge that must be neutralized by polycations for phase separation to occur.19 Upon complexation, dsDNA of length ∼700 base pairs (bp) or more forms toroidal solids with the DNA helices wrapped circumferentially, while shorter dsDNA forms disordered rod-like precipitates, presumably because of a prohibitive energetic cost to bend the stiff DNA (persistence length 50 nm ≈ 150 bp) into smaller structures.1,20 Complexation has also been shown to stabilize the DNA double helix against thermal denaturation and allow formation of normally unfavorable structures such as Z-form and triplex DNA, presumably by reducing electrostatic backbone repulsion.1,21 The complexation behavior of oligonucleotides (N < 100 nt), single-stranded nucleic acids, and RNA has not been studied to a comparable degree, and may differ substantially from long dsDNA. The electric field around a charged polymer decreases near its end, and thermodynamic measurements show that these “end effects” can extend for as much as 10 nt,22,23 suggesting that shorter oligonucleotides may behave much less like polyelectrolytes than longer ones. Single-stranded nucleic acids are more mechanically flexible and have a lower charge density than double-stranded DNA, and RNA duplexes (which adopt an A-form helical structure, vs B-form dsDNA) have been shown to be complexed much less effectively by the small cations cobalt hexamine and spermine than is DNA.24,25 No systematic understanding of these effects on complexation and phase separation currently exists, despite the importance of these molecules to both natural and synthetic systems. Polymer science provides some, albeit incomplete, guidance for understanding polyelectrolyte phase separation. In 1929, de Jong reported the separation of gum arabic (a carbohydrate polyanion) and gelatin (a polycation below pH 4.8) solutions into polymer-rich and -poor liquid phases upon mixing, a phenomenon they named “complex coacervation”.26 Subsequent investigations revealed this to be a general occurrence when oppositely charged polyelectrolytes of sufficient charge and length are mixed; as with dsDNA, entropy gain from counterion release drives complex formation and phase separation, resulting in either hydrated liquids (coacervates) or solid precipitates.27−29 Polyelectrolyte complex materials are widely used in industry,27,30 despite an incomplete understanding of the molecular basis of their properties. Several models have been developed to describe the complexation process, differing in the relative importance of ion pairing, hydration effects, polymer structure, and electrostatics.28,29,31,32 In many cases, detailed molecular features such as hydrogen bonding and chirality also play key roles in determining complex behavior.33 Despite significant advances in describing certain systems, basic properties of polyelectrolyte complexes (e.g., the phase: liquid coacervate vs solid precipitate) cannot currently be predicted from the structures of their components.
■
RESULTS Figure 1A shows typical micrographs of complexes formed by 22 nt oligonucleotides and 50 amino acid pLys peptides mixed at equal charge (amine and phosphate) concentration. A striking qualitative difference is immediately apparent: doublestranded oligonucleotides form irregular solid precipitates, while single-stranded oligonucleotides form spherical coacervate liquid droplets. Upon mixing, we observe very rapid
Figure 1. Oligonucleotides and poly(L)lysine (pLys) form phaseseparated complexes upon mixing. (A) 22 nt single-stranded DNA and 50 aa pLys form liquid droplets when mixed at 2.5 mM amine and phosphate concentration. 22 bp double-stranded DNA and 50 aa pLys form solid precipitates when mixed under the same conditions. Images taken 4 h after mixing. (B) Quantification of noncomplexed DNA shows that the complexes appear nearly neutral (black line) regardless of bulk charge ratio and polymer length: [N]/[P] ≡ [pLys amines]/ [DNA phosphates]. Total charge ([amine] + [phosphate]) is fixed at 5 mM. Solution DNA values are normalized to 1 at [pLys] = 0 and 0 at [DNA] = 0. (C) Phase separation is consistent across a wide range of polyanion: polycation concentration ratios ([N]/[P] = 1 shown in Panel A). 1633
DOI: 10.1021/jacs.7b03567 J. Am. Chem. Soc. 2018, 140, 1632−1638
Article
Journal of the American Chemical Society (seconds) formation of small droplets and precipitates, which then coalesce into larger aggregates through Brownian diffusion. Droplets containing single-stranded DNA complexes undergo rapid hydrodynamic relaxation upon coalescence (movie Coacervate coalescence.avi of the Supporting Information, SI), while the dsDNA precipitate complexes stick together without significant changes in component shape (movie Precipitate coalescence.avi). The phase of the complexes is insensitive to the order of polymer addition, although the size distribution varies somewhat with the rates of addition and mixing. Fluorescence recovery after photobleaching (FRAP) measurements show recovery for the coacervate droplets but not for the precipitates (SI Figure S1), consistent with liquid and solid phase formation, respectively. We observed the same phase behavior across all polymer length combinations and charge ratios measured (Figures 1A,C, S2, and S3), spanning nearly 2 orders of magnitude in both polymer length and charge ([amine]/[phosphate], N/P) ratios. Under our experimental conditions, the complexes’ (liquid and solid) size stops increasing after 30−60 min, presumably due to coalescence of all phase-separated material in the local area of each complex. Quantification of the images (SI Figure S4) shows that droplets appear largest at bulk neutrality (N/P = 1) but that the droplet size is largely independent of polymer length when mixed under identical conditions. The irregular shape of the dsDNA precipitates makes accurate quantification difficult, but the trends appear similar. Phase behavior was also identical at all concentrations we measured, from 0.1 to 10 mM total charge (SI Figure S5), although the complex size and rate of growth increased with concentration. Centrifuging the samples collects the mixtures into two macroscopic phases, as observed for other complex coacervates.27 Quantifying the amount of DNA remaining in the supernatant thus provides a measure of the fraction of DNA incorporated into phase-separated complexes and (when pLys is the limiting reagent) the complex stoichiometry. As shown in Figure 1B, we observe apparently stoichiometric incorporation of nucleic acid into phase-separated complexes regardless of the length of either polyelectrolyte or the bulk charge ratio. Fourier transform infrared spectroscopy (FTIR) measurements (SI Section 2.2 and Figures S6 and S7) indicate that doublestranded DNA maintains its B-form helical structure in the precipitate complexes, while strong lysine-phosphate interactions are observed for both types of complexes. No additional absorbance peaks are noted upon complexation of the singlestranded oligonucleotides, consistent with the idea that hybridization and complexation are largely independent at the molecular length scale.
■
Figure 2. Oligonucleotide−peptide polyelectrolyte complex behavior vs salt concentration and temperature. (A) Representative phasecontrast images of complexes (2.5 mM each in amines and phosphates) formed between single-stranded DNA and polylysine (scale bar 50 μm). At very low [NaCl], large numbers of micron-sized coacervate droplets form. At intermediate concentrations, droplets are larger, and then shrink again before dissolving (not shown) as [NaCl] increases further. (B) Complexes between double-stranded DNA and polylysine form precipitates at low and moderate [NaCl]. For longer polymers, the precipitates appear softer, with rounded edges, as a critical [NaCl] concentration is approached. Above this concentration, liquid droplets are observed; these eventually dissolve at concentrations similar to those required for single-stranded DNA of the same length. (C) Phase diagram for oligonucleotide−pLys complexation. Complexes with single-stranded (left bar of pair) and double-stranded DNA (right bar) show similar stabilities, which increase with polymer length. By contrast, the precipitate/coacervate transition concentration
EFFECT OF SALT AND TEMPERATURE Salt ions play a key role in polyelectrolyte behavior, as they screen electrostatic interactions. Figure 2A shows typical results (see also SI Figure S8) for the single-stranded complexes with increasing NaCl concentration: the coacervate droplets first swell, then shrink and ultimately dissolve for salt concentrations greater than a critical value. Double-stranded complexes display more complex phase behavior: as salt concentration is increased, the precipitates (Figures 2B and S9) become “softer”, with rounded edges. Continued increase in salt concentration produces a transition from solid precipitates to liquid coacervates, followed by eventual dissolution as for the single-strand complexes. This precipitate−coacervate transition is unlikely to reflect denaturation of the DNA duplex, as salt 1634
DOI: 10.1021/jacs.7b03567 J. Am. Chem. Soc. 2018, 140, 1632−1638
Article
Journal of the American Chemical Society
either mixing separately formed complexes or by adding additional DNA to existing complexes. Results with preformed complexes are shown in Figure 3: droplets containing non-
Figure 2. continued (split in right bar) is only weakly dependent on polymer length. (D) Fraction of DNA complexed vs [NaCl] and polymer length for singleand double-stranded DNA (66 nt/bp shown for clarity). Oligonucleotides are released to solution as observed complex size shrinks, but little loss is observed during the precipitate−coacervate transition: see solid orange (30 aa pLys; 500−600 mM NaCl), solid green (50 aa pLys; 500−700 mM NaCl) curves. Values are normalized to the average value at 1 M NaCl to aid visual comparison. (E) At 300 mM NaCl, complexes between 10 bp double-stranded DNA and pLys undergo a melting transition at ∼51 °C: solids with rounded edges are observed just below this temperature and spherical liquids are observed at higher temperatures. Scale bar = 25 μm.
stabilizes nucleic acid hybridization,34,35 and has been observed with synthetic polymers and peptides, albeit at lower salt concentrations.36,37 Quantifying the salt concentrations required for the precipitate−coacervate transition and for dissolution over a range of oligonucleotide and pLys lengths produces the phase diagrams shown in Figure 2C. We find that the stability of the complexes against dissolution is quite similar for the singlestranded and double-stranded complexes, despite their differences in microscopic structure and macroscopic phase. For both types, stability increases dramatically with the length of either polymer (from 200 mM NaCl for the 10 nt/10 aa complexes to 1.2 M for the 88 nt/100 aa complexes). The precipitate−coacervate transition, by contrast, occurs within a narrow range of salt concentrations (500−700 mM) that appears only weakly dependent on polymer length. Measuring the amount of DNA in the supernatant (Figure 2D) shows that the complexes’ shrinking and dissolution at high salt concentrations reflects loss of DNA to solution, but that the precipitate−coacervate transition is not accompanied by significant nucleic acid release. The images shown in Figure 2A−D are for complexes mixed at the stated conditions; we observe similar behavior, including rapid “melting” of precipitates into coacervates, when concentrated salt solutions are added to preformed complexes (SI Figure S10, movie NaCl melting move.avi). We also investigated the effect of temperature on the complexes by mixing them at temperatures up to 55 °C and by heating and cooling preformed complexes. The single-stranded complexes showed no visible change with temperature, however we observed a precipitate−coacervate transition at ∼50 °C for complexes containing the shortest (10 bp) doublestranded DNA at 300 mM NaCl (Figure 2E). The transition is reversible and rapid (within a minute upon temperature change). The observed transition temperature is very similar to the melting temperature of the 10 bp dsDNA duplex in the absence of polycations (SI Table S4), suggesting that the complexes may be responding dynamically to disruption of oligonucleotide base pairing rather than changing phase due to weakened electrostatic interactions. Consistent with this hypothesis, we observed no precipitate−coacervate transitions for complexes containing longer oligonucleotides, whose melting temperatures are higher than 55 °C, the highest temperature we could access.
Figure 3. Coalescence of coacervate droplets formed from noncomplementary and complementary DNA oligonucleotides. Coacervate complexes are formed separately with TAMRA (red) and fluorescein (FAM, green)-labeled 22 nt oligonucleotides and 50 aa pLys. At T = 0, FAM-labeled coacervates are added to microplate wells containing the TAMRA-labeled coacervate droplets. (A) Droplets with non-complementary DNA sequences merge into larger spheres with uniform color and remain liquid. (B) Droplets with complementary DNA sequences stick on contact, forming patchy, irregular solids in which individual domains can be followed for long times. Movies of both processes are available in the SI.
complementary DNA mix readily to form larger spherical liquid droplets (movie Panel A, Non-complementary fluor movie.avi). By contrast, when droplets containing complementary DNA touch, they stick together without hydrodynamic relaxation to form irregularly shaped, solid aggregates in which individual domains can be followed for long times (movie Panel B, Complementary fluor movie.avi). After agitation, these aggregates appear very similar to those formed by mixing pLys with prehybridized double-stranded oligonucleotides (SI Figure S11). Interestingly, both the liquid and solid complexes display noticeable FRET signal (SI Figure S12), implying close proximity of oligonucleotides even (for the coacervates) in the absence of hybridization. Adding free DNA to preformed coacervate droplets showed a similar result (SI Figure S13): the complexes undergo a liquid-to-solid phase transition only upon addition of complementary oligonucleotides.
■
DETERMINANTS OF COMPLEX PHASE To determine what controls the phase of the complexes, we carried out several lines of experiments in which one or the other polymer was modified structurally or chemically. Changing the chirality or charge density of the cationic peptide did not affect the phase of the complexes, and the same behavior was also observed with the polyamines spermidine (3+) and spermine (4+) (SI Figures S14−S16). Single- and double-stranded RNA oligonucleotides displayed the same phase behavior as DNA (Figure 4): single-stranded RNA formed coacervates and double-stranded RNA formed precipitates.
■
POSTCOMPLEXATION BEHAVIOR We investigated whether the single-stranded DNA in the coacervate complexes remained competent for hybridization by 1635
DOI: 10.1021/jacs.7b03567 J. Am. Chem. Soc. 2018, 140, 1632−1638
Article
Journal of the American Chemical Society
length or structure, and our results may help explain several puzzling results from the past several decades, as well as informing understanding of liquid phase separation in cells. In 1969, Shapiro et al. described phase separation of long dsDNA and polylysine into highly hydrated, spherical aggregates when mixed at monovalent salt concentrations of ∼1 M.38 In light of the solid−liquid transition with increased NaCl we observe for longer oligonucleotides, it seems reasonable to suspect that these aggregates were coacervate droplets “melted” by the high salt concentration. In 1979, Porschke reported a turbid suspension, ascribed to coacervation, when trilysine or triarginine was mixed with long (N > 750) polyribonucleotides,39 but the phase of the complexes was not determined experimentally. Aumiller and Keating observed coacervate formation when poly(U) or yeast tRNA was mixed with arginine-rich cationic peptides or polyamines.40,41 Recently, Yin et al. reported microdroplet formation upon mixing of polylysine and 21 nt ssDNA.42 Our results suggest that coacervation is likely a general phenomenon for single-stranded nucleic acids complexed with cationic polymers. Interestingly, the poly(U)−spermine coacervates were only observed above a critical temperature, likely connected to denaturation of single-stranded stacking interactions in the long (2000−3000 nt) polyribonucleotide. Temperature cycling produced reversible phase change, similar to our results with the 10 bp DNA complexes. Similarly, Jain et al. recently published results showing that partially complementary tripletrepeat RNA sequences associated with Huntington’s disease and ALS form coacervate droplets in the presence of Mg2+ cations, followed by gelation of the droplets ascribed to formation of intermolecular base pairs.6 These results are consistent with our observation of solidification when coacervate droplets containing complementary oligonucleotides are mixed, and suggest a unified picture in which nucleic acid base pairing interactions control the dynamics of phaseseparated bodies formed through electrostatic interactions. The ability of nucleic acid polyelectrolyte complexes to respond dynamically to changing environments and base pairing states may be important for their role in living cells, as well as providing opportunities for engineering responsive molecular devices and sensors. Single- and double-stranded nucleic acids differ in several ways that might explain the link between hybridization and phase: double-stranded DNA has a charge density ∼2.4 times larger than the single-stranded form, is less flexible (persistence length 50 nm vs ∼1 nm), more hydrophilic,43 and adopts a defined helical structure that could give rise to specific binding interactions with cations. Site-specific cation binding has not been observed with the cations studied here,1 and the diversity of polyelectrolytes studied also argues against this hypothesis. Backbone hydrophobicity is correlated with precipitate formation for synthetic sulfonate polyanions,44 but the opposite trend is observed here. The methylphosphonate (MP) substitutions were designed to decouple DNA persistence length and charge density. We are not aware of any direct measurements of persistence length for MP nucleic acids, but substantial evidence indicates that DNA flexibility is determined primarily by base stacking.45,46 This is largely unperturbed by MP substitution,47 implying minimal difference in flexibility between the MP and DNA helices. An upper bound can be estimated from EPR measurements,48 which show an increase of as much as 40% in P−O bond mobility for MP vs DNA. Assuming a similar decrease in bending rigidity (i.e., no
Figure 4. Effects of nucleic acid backbone chemistry on complex phase. (A) Sequence of 21 nt luciferase siRNA from Hayashi et al.27 (B) Methylphosphonate substitution (right) at 10 of 22 nt reduces the charge of the oligonucleotides by 45% compared to native DNA (same sequence as Figures 1−3). (C) When mixed with 50 aa pLys, ssRNA (left) forms coacervates while dsRNA (right) forms solid precipitates, just as for DNA. (D) When mixed with 50 aa pLys, mDNA−DNA* duplexes form precipitates like native DNA, while mDNA−mDNA* duplexes form coacervate complexes.
We next investigated partially hybridized DNA duplexes, either by annealing complementary oligonucleotides of different lengths or by designing hairpins with a constant total length of 44 nt and double-stranded stems of 5−20 bp. Both systems gave similar results (SI Figure S17): precipitates were formed when the DNA was 40% or more double-stranded and coacervates formed when lower fractions were hybridized. Finally, we prepared reduced-charge DNA analogs by substituting uncharged methylphosphonate (MP) linkages for phosphodiesters at 10 positions in a 22 nt oligonucleotide and its complement and preparing complexes with 50 aa pLys. Figure 4B, D shows the results: DNA−DNA and DNA−MP (77% of total DNA charge) duplexes produced precipitates, but the MP−MP duplex (55% charge) produced coacervates that were visually indistinguishable from those formed by singlestranded DNA. UV melting confirmed that duplexes were formed in both combinations (SI Figure S18), indicating that the phase difference is due to the presence of the MP substitution rather than lack of hybridization. We also designed a range of oligonucleotide sequences to determine whether the observed phase behavior is sequencedependent. While most sequences formed coacervates as single strands, a few (3/14, 21%) formed solid complexes without their complement being present. Further investigation (SI Section S3) suggests that this is due to either dimerization as a result of partial self-complementarity or to a high density of purine nucleotides, which is known to promote noncanonical folded structures, particularly in the presence of polycations.1,21
■
DISCUSSION Our measurements show that the phase of complexes formed between nucleic acids and cationic peptides is controlled by nucleic acid hybridization over nearly 2 orders of magnitude in polymer length ratio, charge ratio, and total concentration, as well as with mixed lysine-glycine peptides and small polyamines. To our knowledge, this is the first systematic investigation of this phenomenon, particularly in connection with hybridization. As discussed earlier, long double-stranded DNA is known to form solid complexes when mixed with polycations or polyamines, but we are aware of few reports characterizing coacervate formation by nucleic acids of any 1636
DOI: 10.1021/jacs.7b03567 J. Am. Chem. Soc. 2018, 140, 1632−1638
Article
Journal of the American Chemical Society
that oligonucleotide/oligopeptide systems can provide important insights into intracellular phase separation by exploring the roles of RNA structure and peptide sequence.
stiffening from base stacking) and neglecting that 55% of the MP helix remains native DNA, we can estimate a persistence length of at least ∼30 nm, which is still several times larger than the 7.5 nm axial length of the duplex−it should behave as a rigid rod. This suggests charge density as the primary determinant of complex phase, a result consistent with our and others’ observations49 of a precipitate−coacervate transition as increasing counterion concentration decreases the strength of electrostatic interactions. Our results provide insight into two aspects of polyelectrolyte complexation that are not well understood: the precipitate/coacervate division, and mixtures with uneven polymer lengths and charge stoichiometry. Few theoretical approaches address the former: mean-field theories of the Voorn-Overbeek type and random phase approximation-based models predict conditions for phase separation but do not differentiate between coacervates and solid complexes.31,32,50 Schlenoff and co-workers have developed an appealing model based on the fraction of polyion charges paired with lowvalence counterions that is qualitatively consistent with the observed solid−liquid transition with increasing salt, as well as the observed swelling of the coacervates, but requires empirical parameters for each polyelectrolyte/counterion combination.49 Most models also assume either equal length polyelectrolytes or that one is much larger than the other, as well as bulk neutrality. Our measurements show that complexes form readily over nearly 2 orders of magnitude in both polymer length and charge ratios, and the uncomplexed DNA measurements (Figure 1B) indicate that the complexes are very nearly neutral, in apparent conflict with models that predict overcharging by the more-abundant polyelectrolyte.51,52 The observation of qualitatively identical phase behavior from the shortest (N = 10) to longest polymers (N = 100) studied here, as well as the polyamines, is also interesting, as several experimental and theoretical studies suggest a qualitative difference between short and long polyions, with only the latter capable of true polyelectrolyte behavior.22,51,53 We are presently undertaking measurements with even shorter peptides to probe the breakdown of polyelectrolyte behavior further. As discussed above, polyelectrolyte complexes are attractive delivery vehicles for therapeutic RNA oligonucleotides, but the literature is conflicted on the complexation properties of RNA, particularly duplexes such as siRNA. In particular, two reports suggested that double-stranded RNA is incapable of forming polyelectrolyte complexes with pLys and other cationic peptides due to its larger bending rigidity compared to singlestranded RNA,15,16 while a third reported effective complexation.17 As shown in Figure 4, the Fluc dsRNA sequence used by Hayashi et al. readily forms complexes with 50 aa pLys in our hands, though the complexes are solids rather than liquids as for ssRNA. Their concentrations (up to 10 mg/mL, or ∼30 mM charge)15 are comparable to ours, which suggests that either solid-phase complexes are incompatible with micelle formation or that the PEG block inhibits complex formation. We have observed that polyelectrolyte complex micelles with solid cores can be formed from homochiral peptides, though the rate is much slower than for micelles with liquid cores.33 We are presently conducting experiments to test whether the PEG block inhibits complex formation in some way specific to nucleic acids; resolving this question is crucial for applying polyelectrolyte complexes to nucleic acid delivery. The ability to explore a diverse range of complex properties also suggests
■
MATERIALS AND METHODS
■
ASSOCIATED CONTENT
The DNA sequence complementary to human microRNA-21 (TCAACATCAGTCTGATAAGCTA) was used as a basis for constructing a family of oligonucleotides of various lengths, with extensions and other oligonucleotides designed using the NUPACK software tool54 to avoid unwanted secondary structures and selfdimerization. DNA and RNA oligonucleotides were purchased from Integrated DNA Technologies and the methylphosphonate oligonucleotides were purchased from TriLink BioTechnologies. Sequences are listed in SI Tables S1−S6. Poly(L)lysine peptides (pLys, 10−100 aa) were purchased from Alamanda Polymers as the chloride salt (bromide salt for P(D)K100). Prior to use, the peptides were neutralized with NaOH and resuspended in water at 10 mM charge (NH2) concentration. Synthesis and characterization of the (KG)15 and (KGG)10 peptides are described in SI Section S1, along with complete experimental details. Polyelectrolyte complexes were prepared at pH 7 and room temperature (RT). Double-stranded DNA was prepared by annealing complementary strands at 45 °C for 5 min followed by slow cooling to RT. 18.2 MΩ water and concentrated NaCl solutions were mixed, followed by addition of the nucleic acid and then the peptide or polyamine. Samples were mixed thoroughly after addition of each polyelectrolyte. For the charge ratio measurements (Figure 1), total charge concentration ([DNA phosphate] + [pLys amine]) was fixed at 5 mM. In all other experiments, the charge concentration of each polyelectrolyte was 2.5 mM unless otherwise specified. Phase and morphology of the complexes were observed by brightfield and phase contrast optical microscopy using a Leica DMI-6000B inverted microscope with white light illumination and 5−20× magnification. 100 μL aliquots of the complex suspensions were placed in ultralow attachment 96 well plates (Costar, Corning). Images were taken shortly after mixing and then again 4 h later, with the latter used unless noted to the contrary. Coacervate droplet sizes were determined using ImageJ 64; detailed analysis methodology can be found in SI Section S1.3. Fluorescence microscopy data were acquired using an Olympus DSU spinning disk microscope in wide field mode. Fluorescein- and TAMRA-labeled oligonucleotides (SI Section S1.6) were doped into unlabeled oligos at 10% (fluorescein) and 1% (TAMRA) ratios; complexes were otherwise prepared identically to the other experiments. To determine the amount of DNA remaining in solution after complex formation, 100 μL aliquots were centrifuged for 10 min at 20 000g to collect complexes at the bottom of 1.5 mL microcentrifuge tubes. 50 μL of supernatant was removed from the top of the solution and diluted 4× with 5 M NaCl solution. DNA was quantified by absorbance at 260 nm using nearest-neighbor extinction coefficients provided by IDT. Typical fractions of DNA recovered under high-salt conditions where complexes were not seen were 80−90%, with no apparent dependence on the length of either polymer. For the charge ratio results in Figure 1B, the fraction of soluble DNA was normalized by comparison to the DNA-only (defined as 100% free) and peptideonly (ADNA = 0) mixtures. The data in Figure 2D was normalized to the average fraction of DNA recovered from these samples at 1 M NaCl (78.7% of the nominal input; the 50 aa complexes may not be totally dissociated) to aid visual comparison. S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/jacs.7b03567. Additional meterials, methods, and results (Figures S1− S21 and Tables S1−S7), and additional references (PDF) Coacervate coalescence.mov (AVI) 1637
DOI: 10.1021/jacs.7b03567 J. Am. Chem. Soc. 2018, 140, 1632−1638
Article
Journal of the American Chemical Society
■
(23) Shkel, I. A.; Record, M. T. Soft Matter 2012, 8 (36), 9345−11. (24) Li, L.; Pabit, S. A.; Meisburger, S. P.; Pollack, L. Phys. Rev. Lett. 2011, 106 (10), 108101−108104. (25) Katz, A. M.; Tolokh, I. S.; Pabit, S. A.; Baker, N.; Onufriev, A. V.; Pollack, L. Biophys. J. 2017, 112 (1), 22−30. (26) de Jong, H. B.; Kruyt, H. R. Proc. Koninklijke Nederlandse Akademie Wetenschappen 1929, 32, 849−856. (27) de Kruif, C. G.; Weinbreck, F.; de Vries, R. Curr. Opin. Colloid Interface Sci. 2004, 9 (5), 340−349. (28) Priftis, D.; Laugel, N.; Tirrell, M. Langmuir 2012, 28 (45), 15947−15957. (29) Fu, J.; Schlenoff, J. B. J. Am. Chem. Soc. 2016, 138 (3), 980−990. (30) Gouin, S. Trends Food Sci. Technol. 2004, 15 (7−8), 330−347. (31) Kudlay, A.; Ermoshkin, A. V.; Olvera de la Cruz, M. Macromolecules 2004, 37 (24), 9231−9241. (32) Spruijt, E.; Westphal, A. H.; Borst, J. W.; Cohen Stuart, M. A.; van der Gucht, J. Macromolecules 2010, 43 (15), 6476−6484. (33) Perry, S. L.; Leon, L.; Hoffmann, K. Q.; Kade, M. J.; Priftis, D.; Black, K. A.; Wong, D.; Klein, R. A.; Pierce, C. F.; Margossian, K. O.; Whitmer, J. K.; Qin, J.; de Pablo, J. J.; Tirrell, M. Nat. Commun. 2015, 6, 1−8. (34) Santalucia, J.; Hicks, D. Annu. Rev. Biophys. Biomol. Struct. 2004, 33 (1), 415−440. (35) Vieregg, J.; Cheng, W.; Bustamante, C.; Tinoco, I. J. Am. Chem. Soc. 2007, 129 (48), 14966−14973. (36) Chollakup, R.; Smitthipong, W.; Eisenbach, C. D.; Tirrell, M. Macromolecules 2010, 43 (5), 2518−2528. (37) Priftis, D.; Tirrell, M. Soft Matter 2012, 8 (36), 9396−9405. (38) Shapiro, J. T.; Leng, M.; Felsenfeld, G. Biochemistry 1969, 8 (8), 3219−3232. (39) Porschke, D. Biophys. Chem. 1979, 10 (1), 1−16. (40) Aumiller, W. M.; Keating, C. D. Nat. Chem. 2016, 8 (2), 129− 137. (41) Aumiller, W. M.; Pir Cakmak, F.; Davis, B. W.; Keating, C. D. Langmuir 2016, 32 (39), 10042−10053. (42) Yin, Y.; Niu, L.; Zhu, X.; Zhao, M.; Zhang, Z.; Mann, S.; Liang, D. Nat. Commun. 2016, 7, 1−7. (43) Costa, D.; Miguel, M. G.; Lindman, B. J. Phys. Chem. B 2007, 111 (37), 10886−10896. (44) Fu, J.; Fares, H. M.; Schlenoff, J. B. Macromolecules 2017, 50 (3), 1066−1074. (45) Mills, J. B. Nucleic Acids Res. 2004, 32 (13), 4055−4059. (46) McIntosh, D. B.; Duggan, G.; Gouil, Q.; Saleh, O. A. Biophys. J. 2014, 106 (3), 659−666. (47) Kan, L. S.; Cheng, D. M.; Miller, P. S.; Yano, J.; Ts'o, P. O. P. Biochemistry 1980, 19 (10), 2122−2132. (48) Okonogi, T. M.; Alley, S. C.; Harwood, E. A.; Hopkins, P. B.; Robinson, B. H. Proc. Natl. Acad. Sci. U. S. A. 2002, 99 (7), 4156− 4160. (49) Wang, Q.; Schlenoff, J. B. Macromolecules 2014, 47 (9), 3108− 3116. (50) Raspaud, E.; Olvera de la Cruz, M.; Sikorav, J. L.; Livolant, F. Biophys. J. 1998, 74 (1), 381−393. (51) Zhang, R.; Shklovskii, B. I. Phys. A 2005, 352 (1), 216−238. (52) Oskolkov, N. N.; Potemkin, I. I. Macromolecules 2007, 40 (23), 8423−8429. (53) Toma, A. C.; de Frutos, M.; Livolant, F.; Raspaud, E. Biomacromolecules 2009, 10 (8), 2129−2134. (54) Zadeh, J. N.; Steenberg, C. D.; Bois, J. S.; Wolfe, B. R.; Pierce, M. B.; Khan, A. R.; Dirks, R. M.; Pierce, N. A. J. Comput. Chem. 2011, 32 (1), 170−173.
Precipitate Coalescence.mov (AVI) NaCl melting movie (AVI) Non-complementary fluor movie (AVI) Complementary fluor movie (AVI)
AUTHOR INFORMATION
Corresponding Author
*
[email protected] ORCID
Jeffrey R. Vieregg: 0000-0002-4786-5867 Michael Lueckheide: 0000-0001-8310-0557 Notes
The authors declare no competing financial interest. # These authors conttributed equally to this work.
■
ACKNOWLEDGMENTS This work was supported by the U.S. Department of Energy Office of Science, Program in Basic Energy Sciences, Materials Sciences and Engineering Division.
■
REFERENCES
(1) Bloomfield, V. A. Curr. Opin. Struct. Biol. 1996, 6 (3), 334−341. (2) Teif, V. B.; Bohinc, K. Prog. Biophys. Mol. Biol. 2011, 105 (3), 208−222. (3) Shin, Y.; Brangwynne, C. P. Science 2017, 357, 6357. (4) Banani, S. F.; Lee, H. O.; Hyman, A. A.; Rosen, M. K. Nat. Rev. Mol. Cell Biol. 2017, 18 (5), 285−298. (5) Hnisz, D.; Shrinivas, K.; Young, R. A.; Chakraborty, A. K.; Sharp, P. A. Cell 2017, 169 (1), 13−23. (6) Jain, A.; Vale, R. D. Nature 2017, 546 (7657), 243−247. (7) Vieregg, J. R.; Tang, T. Y. D. Curr. Opin. Colloid Interface Sci. 2016, 26, 50−57. (8) Lächelt, U.; Wagner, E. Chem. Rev. 2015, 115 (19), 11043− 11078. (9) Ponnuswamy, N.; Bastings, M. M. C.; Nathwani, B.; Ryu, J. H.; Chou, L. Y. T.; Vinther, M.; Li, W. A.; Anastassacos, F. M.; Mooney, D. J.; Shih, W. M. Nat. Commun. 2017, 8, 1−9. (10) Tian, T.; Zhang, T.; Zhou, T.; Lin, S.; Shi, S.; Lin, Y. Nanoscale 2017, 485, 623−11. (11) Voets, I. K.; de Keizer, A.; Cohen Stuart, M. A. Adv. Colloid Interface Sci. 2009, 147−148, 300−318. (12) Miyata, K.; Nishiyama, N.; Kataoka, K. Chem. Soc. Rev. 2012, 41 (7), 2562. (13) Kuo, C.-H.; Leon, L.; Chung, E. J.; Huang, R.-T.; Sontag, T. J.; Reardon, C. A.; Getz, G. S.; Tirrell, M.; Fang, Y. J. Mater. Chem. B 2014, 2, 8142−8153. (14) Scholz, C.; Wagner, E. J. Controlled Release 2012, 161 (2), 554− 565. (15) Hayashi, K.; Chaya, H.; Fukushima, S.; Watanabe, S.; Takemoto, H.; Osada, K.; Nishiyama, N.; Miyata, K.; Kataoka, K. Macromol. Rapid Commun. 2016, 37 (6), 486−493. (16) Kwok, A.; McCarthy, D.; Hart, S. L.; Tagalakis, A. D. Chem. Biol. Drug Des. 2016, 87 (5), 747−763. (17) Chou, S.-T.; Hom, K.; Zhang, D.; Leng, Q.; Tricoli, L. J.; Hustedt, J. M.; Lee, A.; Shapiro, M. J.; Seog, J.; Kahn, J. D.; Mixson, A. J. Biomaterials 2014, 35 (2), 846−855. (18) Record, M. T.; Anderson, C. F.; Lohman, T. M. Q. Rev. Biophys. 1978, 11 (2), 103−178. (19) Mascotti, D. P.; Lohman, T. M. Proc. Natl. Acad. Sci. U. S. A. 1990, 87 (8), 3142−3146. (20) Laemmli, U. K. Proc. Natl. Acad. Sci. U. S. A. 1975, 72 (11), 4288−4292. (21) Maruyama, A.; Katoh, M.; Ishihara, T.; Akaike, T. Bioconjugate Chem. 1997, 8 (1), 3−6. (22) Ballin, J. D.; Shkel, I. A.; Record, M. T. Nucleic Acids Res. 2004, 32 (11), 3271−3281. 1638
DOI: 10.1021/jacs.7b03567 J. Am. Chem. Soc. 2018, 140, 1632−1638
