Protocols are described for the fabrication of degradable thermoresponsive hydrogels based on hydrazone cross-linking of polymeric oligomers on the bulk scale, microscale, and nanoscale, the latter for preparation of both gel nanoparticles and nanofibers.
While various smart materials have been explored for a variety of biomedical applications (e.g., drug delivery, tissue engineering, bioimaging, etc.), their ultimate clinical use has been hampered by the lack of biologically-relevant degradation observed for most smart materials. This is particularly true for temperature-responsive hydrogels, which are almost uniformly based on polymers that are functionally non-degradable (e.g., poly(N-isopropylacrylamide) (PNIPAM) or poly(oligoethylene glycol methacrylate) (POEGMA)). As such, to effectively translate the potential of thermoresponsive hydrogels to the challenges of remote-controlled or metabolism-regulated drug delivery, cell scaffolds with tunable cell-material interactions, theranostic materials with the potential for both imaging and drug delivery, and other such applications, a method is required to render the hydrogels (if not fully degradable) at least capable of renal clearance following the required lifetime of the material. To that end, this protocol describes the preparation of hydrolytically-degradable hydrazone-crosslinked hydrogels on multiple length scales based on the reaction between hydrazide and aldehyde-functionalized PNIPAM or POEGMA oligomers with molecular weights below the renal filtration limit. Specifically, methods to fabricate degradable thermoresponsive bulk hydrogels (using a double barrel syringe technique), hydrogel particles (on both the microscale through the use of a microfluidics platform facilitating simultaneous mixing and emulsification of the precursor polymers and the nanoscale through the use of a thermally-driven self-assembly and cross-linking method), and hydrogel nanofibers (using a reactive electrospinning strategy) are described. In each case, hydrogels with temperature-responsive properties similar to those achieved via conventional free radical cross-linking processes can be achieved, but the hydrazone cross-linked network can be degraded over time to re-form the oligomeric precursor polymers and enable clearance. As such, we anticipate these methods (which may be generically applied to any synthetic water-soluble polymer, not just smart materials) will enable easier translation of synthetic smart materials to clinical applications.
Smart materials have attracted significant attention due to their potential for reversible "on-demand" responses to external and/or environmental signals. Temperature-responsive materials have attracted particular interest due to their lower critical solution temperature (LCST) behavior, resulting in temperature-driven precipitation at temperatures T>LCST1,2. In the context of thermoresponsive hydrogels, this lower critical solution temperature behavior is manifested by reversible swelling/de-swelling events that result in temperature-tunable bulk sizes (larger at T<LCST)3, pore sizes (larger at T<LCST)4, and interfacial properties (more hydrophilic at T<LCST)5. Such transitions have been widely applied in drug delivery (for external or environmentally-triggerable drug release4,6,7), tissue engineering and cell culture (for thermoreversible cell adhesion/delamination8,9,10), separations (for switchable membrane porosities and permeabilities or thermally-recyclable diagnostic supports11,12,13), microfluidic processes (for on-off valves regulating flow14,15), and rheological modifiers (for temperature-tunable viscosities16). The most commonly investigated thermoresponsive hydrogels are based on poly(N-isopropylacrylamide) (PNIPAM)17, although significant (and increasing) work has also been conducted on poly(oligoethylene glycol methacrylate) (POEGMA)2,18 and poly(vinylcaprolactam) (PVCL)19,20. POEGMA has attracted particular recent interest given its anticipated improved biocompatibility21,22and its facile-to-tune LCST behavior, in which linearly-predictable mixtures of monomers with different numbers of ethylene oxide repeat units in their side chains can alter the LCST from ~20 °C to >90 °C2,23. However, each of these polymers is prepared by free radical polymerization and thus contains a carbon-carbon backbone, significantly limiting the potential utility and translatability of such polymers in the context of biomedical applications in which degradation (or at least the capacity for clearance through renal filtration) is typically a requirement.
In response to this limitation, we have recently reported extensively on the application of hydrazone chemistry (i.e., the reaction between hydrazide and aldehyde-functionalized pre-polymers) to prepare degradable analogues of thermoresponsive hydrogels24,25,26,27,28,29. The rapid and reversible reaction between hydrazide and aldehyde groups upon mixing of the functionalized precursor polymers30 enables both in situ gelation (enabling facile injection of these materials without the need for surgical implantation or any type of external polymerization stimulus such as UV irradiation or chemical initiation) as well as hydrolytic degradation of the network at a rate controlled by the chemistry and density of the crosslinking sites. Furthermore, by maintaining the molecular weight of the pre-polymers used to prepare the hydrogels below the renal filtration limit, hydrogels made using this approach degrade back into the oligomeric precursor polymers that can be cleared from the body25,27,28. Coupled with the low cytotoxicity and low inflammatory tissue response induced by these materials25,26,27, this approach offers a potentially translatable method for the use of thermoresponsive smart hydrogels in medicine, particularly if well-controlled degradable analogues of such hydrogels on all length scales (bulk, micro, and nano) can be fabricated.
In this protocol, we describe methods for making synthetic thermoresponsive pre-polymers functionalized with controlled numbers of hydrazide and aldehyde groups as well as methods to apply these polymers to create hydrogels with well-defined dimensions on various length scales. In particular, this manuscript describes four distinct approaches we have developed to control the mixing of the reactive hydrazide and aldehyde-functionalized pre-polymers and thus create thermoresponsive hydrogel networks with well-defined geometries and morphologies:
To create degradable bulk hydrogels with defined sizes, a templating strategy is described by which the reactive pre-polymers are loaded into separate barrels of a double-barrel syringe equipped at its outlet with a static mixer and subsequently co-extruded into a silicone mold with the desired hydrogel shape and dimensions21,27 (Figure 1).
Figure 1: Schematic of bulk hydrogel formation. Hydrazide and aldehyde-functionalized polymer solutions (in water or aqueous buffer) are loaded into separate barrels of a double barrel syringe and then co-extruded through a static mixer into a cylindrical silicone mold. Rapid in situ gelation upon mixing forms a hydrazone crosslinked hydrogel, which is free standing (once the mold is removed) within seconds to minutes depending on concentration and functional group density of the precursor polymers. Please click here to view a larger version of this figure.
To create degradable gel particles on the micron-scale, a reactive microfluidics method is described in which precursor polymer solutions are simultaneously mixed and emulsified using a soft lithography-templated microfluidic chip design, enabling the formation of mixed reactive polymer droplets that subsequently gel in situ to form gel microparticles with sizes templated by the emulsion (Figure 2)31,32.
Figure 2: Schematic of gel microparticle formation via reactive microfluidics. (A,B) Hydrazide and aldehyde-functionalized polymer solutions (in water or aqueous buffer) are fed by syringe pump into separate reservoirs that are connected downstream across a zig-zag series of channels designed to create a pressure gradient preventing backflow. The polymers are then mixed just before being sheared by paraffin oil flowing from both sides (also driven by a syringe pump) and forced through a nozzle, resulting in flow-focusing production of aqueous (polymer solution) droplets in a continuous paraffin oil phase (see (B) for an illustration of the nozzle area and the droplet formation process). An additional two paraffin oil inlets are positioned after the nozzle to further separate the droplets in the collection channel to allow for complete gelation prior to particle removal from laminar flow, after which the resulting microparticulate gels are collected in a magnetically stirred beaker; (C) Picture of droplet generation process at the nozzle (note that hydrazide polymer is labeled as blue to illustrate mixing)
To create degradable gel particles on the nanoscale, a thermally-driven reactive self-assembly method is described in which a solution of one of the reactive precursor polymers (the "seed" polymer) is heated above its LCST to form a stable nanoaggregate that is subsequently crosslinked by the addition of the complementary reactive precursor polymer (the "crosslinking" polymer); the resulting hydrazone crosslinked nanogel has a size templated directly by the nanoaggregate (Figure 3)28.
Figure 3: Schematic of nanogel formation via thermally-driven reactive self-assembly. An aqueous solution containing the (thermoresponsive) hydrazide-functionalized polymer is heated above its lower critical solution temperature to create a stable uncrosslinked nanoaggregate. Following, an aldehyde-functionalized polymer is added to crosslink the nanoaggregate via hydrazone bond formation and thus stabilize the nanogel particle upon cooling below the LCST. Please click here to view a larger version of this figure.
To create degradable nanofibers, a reactive electrospinning technique is described in which a double barrel syringe equipped with a static mixer at its outlet (as used for making bulk hydrogels) is attached to a standard electrospinning platform (Figure 4)33.
Figure 4: Schematic of hydrogel nanofiber formation via reactive electrospinning. A double barrel syringe with a static mixer (loaded as described for bulk hydrogels but also including a fraction of high molecular weight poly(ethylene oxide) as an electrospinning aid) is mounted on a syringe pump, with the needle at the end of the syringe connected to a high voltage power supply. Hydrazone crosslinking occurs during the fiber spinning process so that when the stream hits the collector (either aluminum foil or a rotating aluminum disk) the nanofibrous morphology is maintained. Please click here to view a larger version of this figure.
The application of such methods for creating degradable smart hydrogel networks is demonstrated in this protocol using either PNIPAM or POEGMA as the polymer of interest; however, the basic approaches described can be translated to any water-soluble polymer, albeit with suitable adjustments for viscosity and (in the case of the self-assembly nanogel fabrication method) the stability of the pre-polymer in forming the seed nanoaggregate.
1. Synthesis of Hydrazide-functionalized Polymers
Note: The following specific recipe is provided for the PNIPAM-mimetic thermoresponsive POEGMA precursor polymer (PO10) with 30 mol% hydrazide functionalization. PNIPAM and POEGMA precursor polymers with different phase transition temperatures can be prepared using this same general method but modifying the type and ratio of the core monomers used (see section 1.2 for modifications for various POEGMA polymers)21,25,27.
2. Synthesis of Aldehyde-functionalized Polymers
3. Fabrication of Hydrazone Crosslinked Bulk Hydrogels
4. Fabrication of Hydrazone Crosslinked Gel Microparticles
5. Fabrication of Hydrazone Crosslinked Nanogels
6. Fabrication of Hydrazone Crosslinked Nanofibers
Bulk hydrogels extruded from a double barrel syringe into a silicone mold conform to the dimensions of the mold and become free-standing upon mold removal; gelation typically occurs seconds to minutes following co-extrusion depending on polymer precursors used. Typical characterization via swelling (measured gravimetrically using a cell culture insert to easily remove the hydrogel from the swelling solution), thermoresponsivity (measured using the same technique but cycling the incubation temperature above and below the phase transition temperature), degradation (measured using the same technique but over longer time periods), and shear or compressive modulus (measured using 2 mm thick and 7 mm diameter molded samples) demonstrates the tunability of the hydrogel responses depending on the chemistry of the precursor polymer (specifically, for POEGMA, the ratio of short to long chain OEGMA monomers used to prepare the hydrogel), the mole fraction of functional groups on the precursor polymers, and the concentration of those precursor polymers (Figure 5)27.
Microfluidics leads to the formation of well-defined gel microparticles on the size scale of 25-100 µm, with the size controllable based on the flow rates of the oil and/or the combined aqueous polymer phases (Figure 6A)31. Hot stage optical microscopy confirms that the gel microparticles maintain the thermoresponsive nature of the bulk hydrogels, showing reversible temperature-dependent swelling-deswelling with only a slight hysteresis on cycle 1 (attributable to irreversible hydrogen bond formation between neighboring amide groups in the collapsed state34) consistent with that observed in bulk PNIPAM hydrogels (Figure 6B)32. Furthermore, the gel microparticles degrade back to their oligomeric precursors over time, enabling renal clearance (Figure 6C)32.
Self-assembly driven by the nanoaggregation of a hydrazide-functionalized PNIPAM polymer in a heated solution followed by crosslinking with an aldehyde-functionalized PNIPAM polymer results in highly monodisperse nanogels (polydispersity <0.1) on the size range of 180-300 nm, depending on the process conditions used (Figure 7A)28. The nanogels retain the typical thermoresponsive behavior of conventional free-radical crosslinked PNIPAM nanogels, with lower degrees of thermal deswelling observed as more cross-linking polymer was added (Figure 7B). The nanogels can be lyophilized and redispersed without a change in particle size (Figure 7C) and degrade over time via hydrolysis to re-form the oligomeric precursor polymers used to formulate the nanogels (Figure 7D).
Reactive electrospinning creates a nanofibrous hydrogel structure (Figure 8A), with nanofiber diameters on the order of ~300 nm achievable without visible electrosprayed particles present33. Soaking the POEGMA-based nanofibers in water results in rapid hydration (roughly two orders of magnitude faster than achieved with a bulk gel of the same composition, Figure 8B) but maintains the nanofibrous morphology over 8-10 weeks prior to hydrolytic degradation at physiological conditions; faster degradation is observed in acid-catalyzed environments, as expected due to the potential for acid-catalyzed hydrazone bond degradation (Figure 8C). The nanofibrous structures are also mechanically robust in both the dry and swollen states over multiple cycles, enabling easy handling and repetitive straining (Figure 8D).
Figure 5: Properties of in situ-gelling bulk degradable thermoresponsive hydrogels. (A) Representative POEGMA gel network microstructures and bulk hydrogel images with corresponding gelation times as a function of the mole % incorporation of OEGMA475 in the precursor polymers; (B-C) Storage modulus of PO100 hydrogels by varying (B) precursor polymer concentration and (C) mole % functional group incorporation per precursor polymer; (D-F) Physiochemical properties of POEGMA hydrogels as a function of OEGMA475 mole % incorporation: (D) storage modulus (E) degradation profile in 1 M HCl, and (F) volume phase transition temperature in response to temperature change over the range 20-60 °C. All error bars represent the standard deviation of n=4 replicate measurements. Adapted from reference27 with permission from Elsevier. Please click here to view a larger version of this figure.
Figure 6: Properties of degradable gel microparticles from reactive microfluidics. (A) Effect of paraffin oil flow rate on (purified) gel microparticle size in water; (B) Thermoresponsivity of purified gel microparticles in water following a single thermal cycle above and below the volume phase transition temperature; (C) Visual assessment (photos) and gel permeation chromatography traces (graph) confirming degradation of gel microparticles back to their precursor polymer components (here, in 1 M HCl to facilitate accelerated degradation on the time scale of imaging); scale bar = 100 µm. Adapted from reference32. Please click here to view a larger version of this figure.
Figure 7: Properties of degradable nanogels from reactive self-assembly. (A) Particle size distributions of nanogels prepared with different aldehyde:hydrazide polymer mass ratios from dynamic light scattering (inset: transmission electron micrograph confirming the spherical nature of the nanogels); (B) Thermosensitivity of self-assembled particles as a function of the mass ratio between aldehyde:hydrazide polymer used to prepare the nanogels (from dynamic light scattering), with error bars representing the standard deviation of n=4 replicates; (C) Visual confirmation of the lack of nanogel aggregation both pre and post-lyophilization; (D) Visual confirmation of the acid-catalyzed degradation of nanogels (here in 1 M HCl for consistency with other studies above); (E) Gel permeation chromatograph traces of nanogel degradation products indicating their similarity to the hydrazide and aldehyde-functionalized precursor polymers. Adapted with permission from reference28. Copyright 2015, American Chemical Society. Please click here to view a larger version of this figure.
Figure 8: Properties of degradable nanofibers from reactive electrospinning. (A) Scanning electron microscopy images of nanofibers in the dry state (left), half dipped in water (middle, thin film), and fully soaked in water overnight (right, thick scaffold); (B) Swelling of nanofibrous hydrogel (red) relative to a bulk hydrogel (blue) of the same composition, with error bars representing the standard deviation of n=4 replicates; (C) Scanning electron microscopy and (inset) visual images tracking the acid-catalyzed degradation of nanofibers in 1M HCl; (D) Tensile cycling of dry (80 cycles, 20% elongation/cycle) and swollen (325 cycles, 10% elongation/cycle in 10 mM PBS) electrospun nanofibrous hydrogels. Figure modified from reference33 and reproduced with permission from the Royal Society of Chemistry. Please click here to view a larger version of this figure.
We have successfully applied all these fabrication techniques to multiple polymer systems using only slight variations of the methods described in detail above for PNIPAM and POEGMA; however, users of these protocols must be cognizant of the potential issues that can arise when other polymers are substituted into these processes. In particular, increasing the viscosity of the precursor polymers can negatively impact both the processibility (especially in the microfluidic method) as well as the efficiency of mixing of the two precursor polymers. In addition, the gelation time of the polymers must be controlled at a rate dependent on the morphology targeted in order to avoid premature gelation that serves to inhibit flow or prevent interdiffusion of the reactive pre-polymers, essential to form the desired homogeneous gel structures. The specific limitations of each strategy, as well as approaches we have used to adapt these approaches to address such limitations at each fabrication length scale, are described below.
Bulk hydrogels via double barrel syringe co-extrusion
Gelation time is the key variable to control to ensure the efficacy of the double barrel syringe technique for forming bulk hydrogels. Polymers that gel too fast upon contact (<1-2 s) can clog the static mixer, either stopping flow entirely or resulting in non-stoichiometric amounts of the two precursor polymers being extruded from the syringe. We have found that gelation times >5 s are preferable (although not required) for the use of this technique; this is particularly important if replicate hydrogels are being cast for physical or mechanical analysis to ensure that each hydrogel cast has the same composition. Gelation time can be easily altered by changing the density of reactive functional groups on one or both precursor polymers (lower functional group density leading to slower gelation) or changing the concentration of the precursor polymers used to form the gel (lower concentrations leading to slower gelation)21. Alternately, replacing the (more reactive) aldehyde group with the (less reactive) ketone group as the electrophile in the gelling pair significantly reduces the gelation time without significantly changing the composition of the resulting hydrogel35; polymers prepared with mixtures of aldehyde and ketone monomeric precursors can be used to tune the gelation time as desired without changing the concentration of precursor polymers used (and thus the mass percentage of solids in the resulting gel formed).
We would also note that the first hydrogel cast does not always have the same properties as subsequent hydrogels cast, an observation attributed to slight differences in the rate at which the contents of the two barrels actually reach the static mixer. As a result, we typically prime the double barrel syringe by extruding a small (<0.3 mL) fraction of gel prior to initiating the casting process to minimize such variability. Finally, while not typically problematic when using oligomeric synthetic pre-polymers, the viscosity of one or more precursor polymer solutions can pose a challenge in the context of this technique, both in terms of facilitating flow using simple thumb depression as well as promoting effective mixing within the static mixer. However, somewhat surprisingly, even precursor polymer solutions with sharply different viscosities still form relatively homogeneous hydrogels using the static mixer attachments described in the parts list (e.g., PNIPAM with a high molecular weight carbohydrate26), suggesting that concerns about inefficient mixing as a result of mis-matched viscosities may not be significant at least on the bulk scale. If required, the use of a syringe pump (instead of the thumb) to drive flow and/or the use of a larger gauge needle at the outlet can help overcome issues associated with extrudability in these systems.
Microscale hydrogels via reactive microfluidics
The key step associated with the microfluidics approach for gel microparticle fabrication is the priming of the microfluidics chip with the two reactive polymers. If the polymers are delivered with different pressures or at different rates into the chip, the differential pressure can drive the backflow of one precursor polymer solution into the reservoir (or at least toward the reservoir) of the other precursor polymer. This results in gelation upstream from particle formation, effectively blocking flow and thus requiring chip disposal. The torturous path imprinted between each reservoir and the mixing point creates a significant resistance to backflow; however, even a trained operator will occasionally gel a chip before a stable flow regime is achieved. Based on our experience, between 1-2 min is typically required to stabilize the flows following the initiation of droplet formation (over which time relatively polydisperse gel microparticles are produced); if no problems are observed within the first 5-10 minutes of operation, it is likely that several hours of continuous monodisperse particle production can be achieved. The use of precursor polymers with relatively well-matched viscosities as well as non-instantaneous gelation times (at least >15 s preferable) greatly assists in avoiding such problems and promoting the formation of stable flows.
Note that various flow rates ranging from 0.01-0.1 mL/h in the aqueous phase and 1.1-5.5 mL/h in the oil phase have been tested using this chip design, leading to the fabrication of particles on the size range of ~25-100 µm according to the shear applied at the flow-focusing junction; faster flow rates equate to higher shear and thus smaller particles formed31,32. Varying the oil flow rate while keeping the total aqueous flow rate low (~0.03 mL/h, as cited in the protocol) was found to be most efficient to control gel microparticle size without compromising either monodispersity or the lifetime of the device, both of which were observed to significantly decrease at the higher end of the cited total aqueous flow rates. Larger oil flow rates (>5.5 mL/h) to create smaller particles are possible, but increased the risk of chip delamination (a common limitation encountered with plasma-bonded PDMS microfluidic chips). Bonding the chips using another method may enable faster flow rates and thus smaller gel microparticle production, a strategy we are currently exploring. Decreasing the size of the nozzle may also help to reduce the size of the microparticles that could be produced, albeit at a heightened risk of premature gelation prior to particle formation. Slower flow rates tended to lead to flow instabilities and thus higher polydispersities and an increased risk of chip gelation; this limitation could be overcome by using a multichannel microfluidic flow control system that has higher stability and higher resolution than the standard syringe pumps used in this protocol.
The choice of oil was critical to the success of this protocol, as heavier oils (favorable in terms of preventing gel microparticle agglomeration after collection) led to much less consistent particle formation at the nozzle than the light silicone oil reported in the protocol. We hypothesize this reduced reproducibility is a result of lower consistency of syringe pumping of heavier oils, leading to more variable shear at the mixing point. Avoiding gel microparticle aggregation in the collection flask was also a challenge, particularly immediately at the exit from the microfluidic device at which point in situ gelation was not complete and large numbers of available reactive functional groups were available to form bridges between colliding particles in the collection bath. This challenge is addressed by: increasing the length of the exit channel on the microfluidic chip itself, maintaining the gel microparticles in laminar flow for a longer period of time to promote more complete gelation; adding the side channels after the nozzle to feed more oil into the chip and thus better separate the gel microparticles in this post-mixing channel without affecting the shear fields at the nozzle itself or the particle production rate; and adding a magnetic mixer to the collection flask to avoid gel microparticle sedimentation and maintain a larger average separation between adjacent particles. While very slow gelling polymers would likely improve the device stability and minimize issues with priming, such systems also were observed to significantly increase the risk of gel microparticle aggregation, as a larger number of reactive functional groups remains unreacted (and thus able to form inter-particle bridges) over a longer period of time. As such, gelation times on the order of 15-60 s appear to be optimal for this technique: slow enough to enable priming but fast enough to ensure most reactive functional groups are consumed prior to the gel microparticles exiting the laminar flow channel into the collection flask.
Finally, removal of the templating oil is essential to ensure that the resulting particles maintain the smart properties anticipated based on the composition of the pre-polymers added and enable use of these particles in a biomedical context. The pentane washing procedure described was highly effective in this regard for general gel microparticle production. However, the application of this technique in a direct biomedical context (e.g., on-chip cell encapsulation) would require re-evaluation of this protocol. We have also explored the use of olive oil, suggested to be a more inert oil in the context of contacting cells36, as the dispersant. While particle formation was possible, the gel microparticle populations were significantly more polydisperse than could be achieved with mineral oil, at least with the current chip design. Thus, while the chip appears to be adaptable to both synthetic polymer and natural polymer gel microparticle formation31, a modified design may be required to exploit this technique more broadly across all possible material combinations.
Nanoscale hydrogels via reactive self-assembly
Nanogels have been formed using a very wide range of processing conditions, including different concentrations of seed polymer (0.5-2 wt%), different ratios of crosslinking:seed polymer (0.05-0.2), different temperatures (40-80 °C), different mixing speeds (200-800 rpm), and different heating times following the addition of the crosslinker polymer (2-60 min)28. In terms of concentrations, the trends observed are generally as would be predicted, as higher concentrations of seed polymer lead to larger nanogels and higher ratios of crosslinker:seed polymer lead to nanogels with higher crosslink densities and thus lower thermoresponsivities. It should be emphasized that increasing the seed polymer concentration too high ultimately leads to bulk aggregation as opposed to nanoaggregation, consistent with what is observed in the conventional free radical precipitation process for forming thermoresponsive nanogels3. Shorter heating times were also found to be favorable for forming smaller and more monodisperse particles. We hypothesize that holding the nanoaggregate at longer times at a temperature above the LCST one or both of the precursor polymers increases the probability of aggregation upon nanogel collision, with the increased hydrophobicity of the hydrazone bond relative to either the precursor aldehyde or hydrazide functional groups making this aggregation more likely as the degree of crosslinking achieved is increased. Ultimately, shorter heating times are favorable from a process perspective, as a monodisperse nanogel population can be formed in as little as 2 min after crosslinker polymer addition; 10 min was found to be the longest time that could consistently produce monodisperse nanogels while also allowing for the production of more highly crosslinked nanogels. Interestingly, the method is remarkably insensitive to mixing, with nearly identical particle sizes and particle size distributions resulting from mixing at different speeds or even scaling the process to larger volumes. While initially surprised by this result, it likely speaks to the primary role of thermodynamics in regulating nanogel production.
To achieve low polydispersities, the colloidal stability and the degree of hydration of the nanoaggregate appear to be the key variables. For example, nanoaggregates prepared using the more hydrophilic hydrazide-functionalized polymers as the seed as opposed to the less hydrophilic aldehyde-functionalized polymers lead to nanogels with significantly lower polydispersities. The difference between the experimental assembly temperature and the LCST of the seed polymer is also critical. Operating at a temperature just above the seed polymer LCST ((T – LCST) < 5 °C) offers the highest probability of monodisperse nanogel formation; operating well above the LCST creates more hydrophobic and collapsed nanoaggregates that are more likely to aggregate and less likely to crosslink, while operating below the LCST results in a relatively non-compact seed polymer that cannot be effectively or reproducibly crosslinked. For the best prediction of particle monodispersity, we recommend first performing a UV/vis scan to measure the onset LCST of the seed polymer and subsequently performing the self-assembly process at a temperature 1-2 °C above that LCST.
Note that nanogels produced using this method could be lyophilized and redispersed without any change in colloidal stability, often not possible for self-assembled structures and in our view attributable to our crosslinking stabilization method. We also anticipate that only the seed polymer needs to be thermoresponsive for this method to work; use of cross-linking polymers that are either non-responsive or responsive to other stimuli may further broaden the ultimate applicability of this technique. Finally, since the mixing of the two reactive precursor polymers is in this case passive as opposed to active, gelation time is much less important in terms of process control relative to the other fabrication strategies described. However, even in this technique, keeping the total crosslinking time <30 min is desirable to minimize the risk of particle aggregation.
Nanofibrous hydrogels via reactive electrospinning
Controlling the gelation time of the reactive pre-polymers is again essential to the success of gel nanofiber production. In particular, approximately matching the residence time of the precursor polymers in the static mixer (controlled by changing the flow rate of solution from the double-barrel syringe as well as the length and tortuosity of the static mixer) with the bulk gelation time of the precursor polymers is essential both to preserve spinnability as well as ensure effective crosslinking of the spun fibers between the needle and the collector. Faster gelation leads to ineffective Taylor cone development and thus poor spinnability, while slower gelation results in an aqueous solution instead of a gel hitting the collector, resulting in spreading and the ultimate formation of a thin film gel instead of nanofibers. Working at residence times slightly below the bulk gelation time has also been found to be effective (and indeed preferable to reduce the risk of needle clogging) since water evaporation as the solution is spun effectively concentrates the precursor polymers in the stream and thus accelerates gelation kinetics during the spinning process. In this same vein, operating at higher needle-to-collector distances (>10 cm) is generally favorable in this process, as shorter distances reduce the time available for water evaporation and thus require more stringent control over the relationship between residence time and gelation time in order to preserve a nanofibrous product.
Note that the use of PEO (or another high molecular weight and easily electrospun polymer) is essential in this protocol to promote nanofiber formation, as the short and highly branched POEGMA oligomers cannot alone reach an adequate degree of entanglement to induce electrospinning; instead, electrospray results at all process conditions tested for POEGMA-only formulations (although this may also have applications for making degradable gel particles using this same chemistry). A minimum PEO concentration of 1 wt% (1 MDa molecular weight) is required to maintain a fully nanofibrous morphology. Note that the PEO can be removed from the fibers following a simple soaking procedure (deionized water, 24 h) without disrupting the integrity of the nanofibrous network; in this way, PEO acts more as a transient electrospinning aid than an essential component of the final nanofibrous product. Note also that various types of collectors, including simple aluminum foil (to create thin layer hydrogels that can delaminate from the collector upon soaking) as well as a rotating aluminum disk (to create thicker scaffolds) can be used in conjunction with this same technique, provided the other process variables controlling the rate of gelation, the rate of electrospinning, and the rate of water evaporation during electrospinning remain unaltered.
Interestingly, depending on the method used to prepare the different morphologies, significant differences have been observed in the degradation times of hydrogels prepared from the same hydrogel precursors. For example, POEGMA nanofibrous hydrogels degrade slower than bulk POEGMA hydrogels with the same composition despite their significantly higher surface area and thus access to water to hydrolyze the hydrazone bonds. We relate these differences to the inherent contrasts between the described protocols in terms of the geometry of mixing the precursor polymers, which may lead to internal gel homogeneities and/or morphologies that are significantly different and/or the in situ concentration of polymer precursors on the same time scale as gelation, particularly relevant in electrospinning due to the simultaneous water evaporation and crosslinking observed in this process. While this may somewhat complicate the choice of the precursor polymers if one polymer is targeted for use in each protocol, it may also offer a technical opportunity in terms of making hydrogels with one chemical composition but very different physical properties.
Overall, the methods described provide a strategy for fabricating degradable (or at least renally clearable) analogues of thermoresponsive polymers on multiple length scales (bulk, micro, and nano) and with multiple types of internal structures (particles or fibers). Such protocols address the key barriers to the successful translation of conventionally-prepared synthetic thermoresponsive materials to the biomedical field: injectability and degradability. We are continuing to explore the application of such materials in both drug delivery and tissue engineering applications ranging from the physical targeting of cancers, the transport of drugs across the blood-brain barrier, the therapeutic delivery of proteins at the back of the eye, the directional growth of tissues, and the thermoreversible adhesion and differentiation of cells, among other applications.
The authors have nothing to disclose.
Funding from the Natural Sciences and Engineering Research Council of Canada (NSERC), the NSERC CREATE-IDEM (Integrated Design of Extracellular Matrices) program, 20/20: NSERC Ophthalmic Biomaterials Research Network, and the Ontario Ministry of Research and Innovation Early Researcher Awards program is acknowledged.
Chemicals | |||
2,2 – azobisisobutryic acid dimethyl ester | Wako Chemicals | 101138 | |
Di(ethylene glycol) methyl ether methacrylate (M(EO)2MA) | Sigma Aldrich | 447927 | 188.2 g/mol, n=2 ethylene oxide repeat units |
Oligo (ethylene glycol) methyl ether methacrylate (OEGMA475) | Sigma Aldrich | 447943 | 475 g/mol, n=8-9 ethylene oxide repeat units |
Acrylic acid (AA), 99% | Sigma Aldrich | 147230 | |
Thioglycolic acid (TGA), 98% | Sigma Aldrich | T3758 | |
Dioxane, 99% | Caledon Labs | 360481 | |
Nitrogen, UHP grade | Air Liquide | Alphagaz1 765A-44 | |
Adipic acid dihydrazide (ADH), 98% | Alfa Aesar | A15119 | |
N'-ethyl-N-(3- dimethylaminopropyl)-carbodiimide (EDC, x%) | Carbosynth | FD05800 | |
Hydrochloric acid (HCl), 37% | Sigma Aldrich | 320331 | |
Sodium hydroxide (NaOH), 97% | Sigma Aldrich | 221465 | |
Aminoacetyl aldehyde dimethyl acetal, 99% | Sigma Aldrich | 121967 | |
4-Hydroxy-TEMPO, 97% | Sigma Aldrich | 176141 | |
Methacryloyl chloride,97x% | Sigma Aldrich | 523216 | |
Petroleum ether, 95% | Sigma Aldrich | 32047 | |
Magnesium sulfate, 99.5% | Sigma Aldrich | M7506 | |
tert-Butyl methyl ether, >99.0% | Sigma Aldrich | 443808 | |
Phosphate buffered saline | BioShop | PBS405.1 | 1x, pH 7.3-7.5 |
N-isopropylacrylamide, 99% | J&K Scientific | 258717 | Recrystallized from 60% hexanes/40% toluene |
Ethanol, anhydrous | Commerical Alchols | P016EAAN | |
Span 80 | Sigma Aldrich | S6760 | |
Heavy paraffin oil | Caledon Labs | 1326197 | |
Pentane, reagent grade | Caledon Labs | 1/10/7800 | |
Poly (ethylene oxide) average Mv 600,000 | Sigma Aldrich | 182028 | |
Supplies essential for synthesis and hydrogel fabrication | |||
Rotary evaporator | Heidolph | G3 | |
Dialysis tubing (3500 Da molecular weight cut-off) | Spectrum Labs | 28170-166 | Vol/length= 6.4mL/cm |
Double barrel syringe | Medmix | L series | L series, 2.5 mL, 1:1 volume ratio |
Static mixer | Medmix | L series | L series, 2.5 mL, 1:1 volume ratio, 1.5" length |
Silicone rubber sheet, 1/16" thickness | McMaster-Carr | 9010K12, 30A Durometer (Super Soft) | |
Syringe pump | KD Scientific | KDS Legato 200 | Infuse Only Dual Syringe Pump |
High voltage power supply | Spellman | 230-20R | 0 to 20 kV |
Microfluidic Chip Fabrication | |||
Silicon wafer | University Wafer | 2080 | D = 76.2 mm; 380 µm thickness; P-doped; <100> orientation |
SU-8 100 | MicroChem | Y131273 | |
SU-8 Developer | MicroChem | Y020100 | |
Custom 2.5" spincoater | Built in-house | N/A | |
Mask Aligner | KARL SUSS | MJB3 UV400 (with a 276 W lamp) | |
Masterflex L/S 13 Silicone Tubing | Cole Parmer | OF-96400-13 | Peroxide-cured |
Dow Corning Sygard 184 Silicone Elastomer Base | Ellsworth Adhesives | 4019862 | |
Dow Corning Sygard 184 Silicone Elastomer Curing Agent | Ellsworth Adhesives | 4019862 | |
High Power Plasma Cleaner | Harrick | PDC-002-HP | |
Characterization Instruments | |||
Mach 1 micromechanical tester | Biomomentum | LB007-EN | |
Cellstar tissue culture 12 well plate | Greiner Bio-one | 665 180 | |
Cell culture insert for 12 well plate | Corning | 08-771-12 | 8 µm pore size |
Optical microscope | Olympus BX51 optical microscope | BX51 | |
Temperature-controlled microscope stage | Linkam Scientific | THMS600 | |
Gel permeation chromatograph (GPC) | Waters | 590 HPLC Pump | Waters Styragel columns (HR2, HR3, HR4; 30 cm x 7.8 mm (ID); 5 mm particles), Waters 410 refractive index detector |
Dynamic light scattering (DLS) | Brookhaven | 90Plus Particle Size Analyzer | |
Transmission electron microscopy (TEM) | TEMSCAN | JEOL 1200EX | Accelerating voltage 100 kV |
Scanning electron microscopy (SEM) | Tescan | Vega II LSU | Accelerating voltage 10 kV |
Microsquisher | CellScale Biomaterials Testing | MS-50M-01 |