Method Article

DNA Nanotubes as a Versatile Tool to Study Semiflexible Polymers

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DOI:

10.3791/56056

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October 25th, 2017

 ,  ,  ,  ,  ,  ,  ,  , 
1Fraunhofer Institute for Cell Therapy and Immunology, 2Institute of Experimental Physics I, Universität Leipzig

* These authors contributed equally

In This Article

Summary

Semiflexible polymers display unique mechanical properties that are extensively applied by living systems. However, systematic studies on biopolymers are limited since properties such as polymer rigidity are inaccessible. This manuscript describes how this limitation is circumvented by programmable DNA nanotubes, enabling experimental studies on the impact of filament rigidity.

Abstract

Mechanical properties of complex, polymer-based soft matter, such as cells or biopolymer networks, can be understood in neither the classical frame of flexible polymers nor of rigid rods. Underlying filaments remain outstretched due to their non-vanishing backbone stiffness, which is quantified via the persistence length (lp), but they are also subject to strong thermal fluctuations. Their finite bending stiffness leads to unique, non-trivial collective mechanics of bulk networks, enabling the formation of stable scaffolds at low volume fractions while providing large mesh sizes. This underlying principle is prevalent in nature (e.g., in cells or tissues), minimizing the high molecular content and thereby facilitating diffusive or active transport. Due to their biological implications and potential technological applications in biocompatible hydrogels, semiflexible polymers have been subject to considerable study. However, comprehensible investigations remained challenging since they relied on natural polymers, such as actin filaments, which are not freely tunable. Despite these limitations and due to the lack of synthetic, mechanically tunable, and semiflexible polymers, actin filaments were established as the common model system. A major limitation is that the central quantity lp cannot be freely tuned to study its impact on macroscopic bulk structures. This limitation was resolved by employing structurally programmable DNA nanotubes, enabling controlled alteration of the filament stiffness. They are formed through tile-based designs, where a discrete set of partially complementary strands hybridize in a ring structure with a discrete circumference. These rings feature sticky ends, enabling the effective polymerization into filaments several microns in length, and display similar polymerization kinetics as natural biopolymers. Due to their programmable mechanics, these tubes are versatile, novel tools to study the impact of lp on the single-molecule as well as the bulk scale. In contrast to actin filaments, they remain stable over weeks, without notable degeneration, and their handling is comparably straightforward.

Introduction

Due to the complex behaviors enabled by their unique mechanical properties, semiflexible polymers are fundamental building blocks of living matter. In contrast to flexible polymers, semiflexible polymers adopt an outstretched configuration due to their non-vanishing backbone stiffness while still remaining subject to strong thermal fluctuations1. Thus, purely stochastic models cannot be applied to their behaviors, as with the extremes of fully flexible or rigid polymers. The so-called worm-like chain model2,3,4 was developed to quantify this stiffness via the lp, which is the decay constant of the tangent-tangent correlation along the filament4. If lp is comparable to the contour length (lc) of the filament, the polymer is considered semiflexible1. Analogous to the poles of a tent, their arrangements in networks or bundles stabilizes the entire collective system at low volume fractions, leading to unusual viscoelastic properties5,6,7,8,9. These structures provide high elasticities at large mesh sizes10, maintaining mechanical integrity while still facilitating diffusive and active transport processes. This property is especially suitable for biological systems such as the cytoskeleton or the extracellular matrix, but it is also widely used in food engineering1,11,12.

Going beyond their significance to living matter, it is crucial to comprehensively examine the physical properties of these structures in order to have the tools to develop biomimetic materials or novel hydrogels. In terms of semiflexible polymers, this implies the systematic determination of the collective properties of networks resulting from single-filament properties such as lp and the development of a descriptive theoretical framework. In pioneering studies, the cellular biopolymer actin was established as a model system for semiflexible polymers and is still widely considered the gold standard5,13,14,15,16,17. However, exhaustive studies are limited with this system since they are bound to the inherent properties of this protein. Various theoretical approaches have aimed at building a description of the non-trivial mechanical behaviors at the single-filament level and have led to notably different scaling predictions for the dependence of the linear elastic plateau shear modulus, G0 (i.e., the "elasticity" of the network), with respect to concentration (c) and lp6,7,13,14,15,18,19,20,21,22,23. While the concentration scaling is readily accessible in experiments with actin-based or other model systems and while theoretical predictions have been rigorously verified13,16,24,25, the scaling with respect to lp has remained experimentally inaccessible. This, however, is a major limitation since lp is also an independent variable that is the defining quantity of semiflexible polymers.

This central, natural limitation imposed by the fixed lp of actin or other biologically-derived polymers such as collagen has recently been resolved by employing tile-based DNA tubes, which are tunable in their mechanical properties9,26,27,28. Slight variations in the architectures of the tubes (e.g., different numbers of constituent DNA strands within the unit ring) yield distinct values for lp, which can be evaluated via fluorescence microscopy, either by analyzing one fluctuating tube or by evaluating the curved configurations of several adhered tubes, as described previously9,28. These analyses revealed that lp values of the different tube populations vary over more than one order of magnitude and that different evaluation techniques yield consistent results9,28.

Surprisingly, the overall scaling of the linear elastic plateau shear modulus G0 with respect to the concentration and lp has been reported to be inconsistent with all previous theoretical approaches9, in particular demonstrating a much stronger than predicted dependence upon lp. These findings emphasize the value of a new model system to study the central properties of semiflexible polymers. Employing n-helix DNA tubes dramatically broadens the scope of these investigations. Not only can lp be freely varied without changing the basic material, but the inherent programmable nature of DNA can enable the systematic examination of additional elements, such as crosslinks or kinetic switching processes. Additionally, these tubes are soluble in water and, in contrast to most proteins, stable in adequate pH and ionic conditions for several weeks, without detectable degradation9.

To assemble these tubes, a discrete set of DNA oligonucleotides is used, each of which contains two domains that share complementary base sequences to two neighboring strands (due to the specific sequences, a single strand cannot form structures such as hair pins). The complementary sequences hybridize in a cyclic manner, forming enclosed, half-overlapping rings of n interconnected double-helical segments (Figure 1A and B). These rings form at a discrete diameter (Figure 1C), and their half-overlapping configuration exposes axial sticky ends complementary to the sticky ends of another ring. This selective addition of matching oligonucleotides triggers a stacking of the rings, leading to the effective polymerization of filamentous DNA helix tubes of size n (nHT). Their contour lengths typically measure several microns in length, and their length distribution is comparable to that of actin filaments9,26,27,28. It has been shown for similar DNA nanotubes that they indeed exhibit polymerization kinetics similar to those of actin filaments and microtubules29. Depending on the number n of individual DNA strands making up the basic ring structure, the nHT architecture, as well as its circumference and diameter, can be controllably varied. Using more DNA strands increases the circumference of the rings/tubes, and the corresponding architectural change shifts the mechanical properties to higher lp values (Figure 1C), corresponding to a higher rigidity. On the mesoscopic scale, these larger lp values translate into less bent conformations due to the higher stiffness (Figure 1D and E).

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Protocol

1. Preparation of n HTs

NOTE: Here, n denotes the number of different single DNA strands involved in the formation of the helix tubes of a certain size. For n=8, eight different single DNA strands make up a unit ring.

  1. Purchase DNA sequences (HPLC purity grade or higher) from a suitable DNA synthesis service or perform high-quality synthesis and purification (exemplary sequences given in Table 1).
  2. Resuspend lyophilized oligonucleotides in purified water and follow the further resuspension steps given in the corresponding manual from the company. Adjust the amount of water to obtain a final concentration of 200 µM.
  3. Determine concentrations of the single DNA strands to verify the values given by the distributor (e.g., via UV-Vis spectroscopy at 260 nm) since the exact stoichiometry is crucial for the assembly process. Calculate the concentration of the oligonucleotides, taking into account the specific molar weight and the extinction coefficient for each single-stranded DNA oligonucleotide.
    NOTE: The extinction coefficient values inherently vary for the different sequences and are stated in the documentation of commercially purchased DNA. They can also be calculated according to the nearest-neighbor model using a number of freely available online tools. To comply with the linear range of the spectrometer, consider diluting the sample used for concentration determination.
  4. Using a micropipette, mix the DNA strands-each at the same concentration-in a container suitable for the thermocycler (e.g., a 200 µL PCR tube) to form the desired nHTs (final buffer conditions: 1xTE (10 mM Tris and 1 mM EDTA, pH 8) and 12.5 mM MgCl2).
    NOTE: Final DNA nHT samples consist of n- 1 strands, U1− Un−1, and one Tn strand. The concentration per strand can vary from sub-micromolar to more than 10 µM, depending on the needs of the user (e.g., <1 µm per strand for the subsequent dilution to observe single filaments or higher concentrations for the examination of dense network mechanics).
  5. Hybridize the nHTs in a thermocycler (denaturation for 10 min at 90 °C, with a subsequent quick drop to 65 °C; slow temperature decrease from 65 °C to 55 °C, with complementary base pairing in 20 temperature steps of -0.5 K for 30 min each; and a quick drop from 55 °C to 20 °C); see Figure 2A.
    NOTE: The slow decrease steps from 65 °C can be lengthened to increase the average tube length; however, the subsequent quick drop to 20 °C is crucial to avoid the aggregation of polymerized tubes.
  6. Store the hybridized nHTs for up to 3 weeks at 4 °C without detectable degradation.
    NOTE: For fluorescence microscopy, labeled nHTs are hybridized by (partly) substituting U1 with the modified U1-Cy3 (Table 1). For longer observation times, the addition of established anti-bleaching agents is advisable.
  7. Carefully dilute the sample to the desired final concentration (e.g., 4 µM for networks or 20 nM for single-filament observations) using the sample buffer, if necessary. To minimize possible error sources, assemble samples at the desired final concentration.

2. Shear Rheology

  1. Choose an appropriate geometry (plate-plate or cone-plate) for the dynamic shear rheometer. For small volumes (i.e., below 200 µL), use the cone-plate geometry.
  2. Load the sample on the dynamic shear rheometer.
  3. Passivate the air-water interface of the sample and environment using the following steps to prevent potential evaporation effects.
    1. Surround the sample with 2.5 mL of the sample buffer bath.
    2. Add a surfactant just below the micelle concentration.
    3. Use a Hamilton syringe to surround the sample with a lipid to avoid direct contact of the sample with air.
  4. Seal the sample chamber with a cap equipped with wet sponges and, if possible, with an additional water bath (not in contact with the sample) to further suppress evaporation.
  5. Start the measurement using the rheometer-specific software at room temperature; alternatively, perform the measurement at different temperatures, such as 37 °C (if physiological conditions are needed).
    NOTE: Cooling the sample is often accompanied by the condensation of water vapor from the surrounding air, while heating leads to enhanced evaporation. Both effects can uncontrollably change the concentration of the sample, so all measurements in a series should be performed at a constant temperature.
  6. Allow the sample to equilibrate for 2 h at room temperature. The time evolution may be monitored with a time sweep (one data point per minute; strain γ = 5%; frequency f = 1 Hz).
  7. Measure the mechanical properties with the desired test protocols. Start with a series of frequency sweeps (f-sweeps) because they leave the sample intact and allow for more measurements.
    NOTE: Additionally, short f-sweeps should be carried out before and after longer measurements (such as long f-sweeps with a much higher resolution) to verify that the sample remained unchanged throughout the full measurement protocol.
    1. Perform a short f-sweep (e.g., γ = 5%; f = 0.01 Hz to 30 Hz; 5 data points per decade).
    2. Perform a long f-sweep (e.g., γ = 5%; f = 0.001 Hz to 30 Hz; 21 data points per decade); the low-frequency regime can be omitted if not necessary to reduce measurement time.
    3. Perform a short f-sweep.
    4. Perform a γ-sweep (e.g., f = 1 Hz; γ = 0.0125% to 100%; 20 data points per decade).
      NOTE: Use the short f-sweep first, as it is usually inconsistent with previous f-sweeps because networks usually rupture during γ-sweep or detach from the plates. Alternatively, use γ-ramp (e.g., = 0.025 s-1, t = 60 s); however, the ramps usually require changing the motor mode, so subsequent short f-sweeps would be falsified.
  8. Bin and/or smooth the raw data with a Gaussian kernel.

3. Fluorescence-assisted Analysis of DNA Annealing

  1. Prepare 8 aliquots of 12 µL for each sample using 1-4 µM DNA per strand, 12.5 mM MgCl2 in 1x TE buffer, and 1x nucleic acid gel stain from a 10,000x DMSO stock. Make a negative control with no DNA, but include nucleic acid gel stain.
  2. Transfer the aliquots to a real-time PCR plate and seal with adhesive film (e.g., a 96-well reaction plate).
    NOTE: The high temperatures used in the thermal annealing protocols will evaporate samples. Thus, make sure that the wells are tightly sealed with adhesive film to prevent water evaporation and concentration increase, particularly during long-term experiments.
  3. Centrifuge the plate at 100 x g for 1 min at room temperature to remove the gas bubbles; if the gas bubbles expand, the wells cannot be analyzed.
  4. Load the sealed plate into a real-time quantitative PCR system.
  5. Run an annealing program. Denature the DNA at 90 °C for 10 min. Drop the temperature quickly to 72 °C. Then, decrease the temperature slowly by 0.5 °C every 30 min. After the signal has decayed, accelerate the temperature ramp.
    NOTE: 72 °C is sufficiently above the real hybridization temperature; thus, the signal is recorded over a broad temperature range suitable for determining the necessary temperature range.
  6. Make sure that the lid of the cycler heats to at least 100 °C to prevent the condensation of evaporated sample on the adhesive film.
  7. During assembly, excite the samples with an argon-ion laser at 488 nm and detect the fluorescence at 550 nm when using a nucleic acid gel stain (or spectrally similar). For other dyes, choose an appropriate excitation/emission combination. Measure the fluorescence signal as often as possible to increase the overall signal quality using the built-in camera.
    NOTE: Here, measurements were taken every 9 s.
  8. If preset melting and annealing programs are applied, use the provided software to analyze the data. If not, subtract the previous mean value from the mean value for each temperature step to obtain the relative change of the fluorescent signal.

4. AFM Imaging

  1. Cleave mica (e.g., mica "V1") by attaching commercially available adhesive tape and ripping it off; the uppermost layer of the mica should be removed.
  2. Using a micropipette, deposit pre-formed nHT in Mg2+ containing folding buffer on the freshly cleaved mica and let it settle for 10 min.
  3. Spin the samples in short 3-s intervals at 2,000 x g on a small table centrifuge to remove the remaining liquid. Several nHTs should still be attached to the mica surface.
  4. Use a cantilever tip with a high stiffness (e.g., 54 Nm-1) and determine its resonance frequency with the internal AFM software.
  5. Record the height profile with the AFM in air in tapping mode at low scan rates (e.g., 1 line/s) to avoid damaging or displacing the nHTs.

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Results

The assembly of DNA nanotubes via a temperature ramp (Figure 2) is a very reliable method to form these artificial semiflexible polymers. These polymers have comparable characteristics to their naturally occurring counterparts, such as actin filaments, but provide a much broader experimental framework since their mechanical properties can be controllably altered9,27. Like biopolymers, they can be arra...

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Discussion

To obtain properly formed networks, assembling the DNA nanotubes is a crucial step. Errors during the synthesis process negatively impact the tube quality; therefore, it is recommended that HPLC or a more stringent process be used to purify the oligonucleotides. Since the formation of discrete rather than aggregated DNA nanotubes, as well as their length distribution, depends upon the equimolar stoichiometry of the n constituent oligonucleotides within the set, it is necessary to remeasure the concentrations of ...

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Disclosures

The authors have nothing to disclose.

Acknowledgements

We acknowledge funding by DFG (1116/17-1) and the Leipzig School of Natural Sciences "BuildMoNa" (GSC 185). This work has been supported through the Fraunhofer Attract project 601 683. T. H. acknowledges funding from the European Social Fund (ESF-100077106).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
AFM cantilever ACTAAppNano
AFM - NanoWizard 3JPK Instruments
CCD cameraAndoriXon DV887
DMSOSigma-AldrichD2650
DNA oligonucleotidesBiomers.netFor sequences see Table 1
DNA Cy3-labeled oligonucleotidesBiomers.netFor sequence see Table 1
EDTASigma-AldrichE-9884
Epi-fluorescence micro-scopeLeicaDM-IRB
MgCl2Sigma-AldrichM-8266
Mica "V1", 12 mm roundPlano GmbH50-12
MicroAmp® Fast Optical 96-Well Reaction PlateThermo Fisher Scientific Inc.4346907
MicroAmp® Optical Adhesive FilmThermo Fisher Scientific Inc.4306311
NanoDrop 1000 SpectrophotometerThermo Fisher Scientific Inc.
100x objectiveLeica5506168
Purified waterMerk Millipore - Milli-Q & Elix
Sapphire PCR tubesGreiner Bio-One683271
TProfessional Standard PCR ThermocyclerCore Life Sciences Inc.070- Standard
7900HT Fast Real-Time PCR SystemApplied Biosystems4351405
RheometerTA InstrumentsARES
SYBR® Green I nucleic acid gel stainThermo Fisher Scientific Inc.S7567
TrisSigma-AldrichT4661
Triton X-100Sigma-Aldrich Co.X-100Suppresses evaporation of sample at air-water interface

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Persistence LengthRheological MeasurementDynamic Shear RheometerFluorescence DetectionThermocycler HybridizationReal time PCRPolymerization KineticsHydrogel Application

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