Using Magnetic Resonance Imaging to Study Enzymatic

May 23, 2014 - Surface-Induced Hydrogelation for Fluorescence and Naked-Eye Detections of Enzyme Activity in Blood. Tengyan Xu , Chunhui Liang ...
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Using Magnetic Resonance Imaging to Study Enzymatic Hydrogelation Weijuan Wang,† Junchao Qian,‡ Anming Tang,† Linna An,† Kai Zhong,*,‡ and Gaolin Liang*,† †

CAS Key Laboratory of Soft Matter Chemistry, Department of Chemistry & Collaborative Innovation Center of Suzhou Nano Science and Technology, University of Science and Technology of China, Hefei, Anhui 230026, China ‡ High Magnetic Field Laboratory, Chinese Academy of Sciences, Hefei, Anhui 230031, China S Supporting Information *

ABSTRACT: Herein, we report, for the first time, the use of MRI methods to study enzymatic hydrogelation. Supramolecular hydrogels have been exploited as biomaterials for many applications. However, behaviors of the water molecules encapsulated in hydrogels have not been fully understood. In this work, we designed a precursor 1 which could self-assemble into nanofibers and form hydrogel I (gel I) upon the catalysis of phosphatase. The differences of mechanic property, pore size, water diffusion rate, and magnetic resonance relaxation times T1 and T2 of gel I containing different concentrations of 1 were systematically studied and analyzed. T1, T2, and diffusion-weighted 1H MR images from gel I phantoms were obtained at 9.4 T. Analyses of the MRI data uncovered how the density of the nanofiber networks affects the relaxation behaviors of the water protons encapsulated in such hydrogels. Rheological analyses and cryo-TEM observations showed increased gel elasticities with increased concentrations of 1 while the pore sizes of gel I decreased. This also resulted in an increase in the proton relaxation rate (i.e., shortened T1, T2, and apparent diffusion coefficient (ADC)) for the water encapsulated in the hydrogel. With MRI, our study provides a new in vitro method to potentially mimic and study in vivo diseases that involve fibrous aggregates.

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between the hydrogelators and the gelled water is crucial to this process. Interestingly, understanding the water motion plays a key role in magnetic resonance imaging (MRI), and study of the protons of water molecules within or surrounding the fibrous proteinic plaques in human brain has been successfully employed to investigate the progress of aging diseases (e.g., Alzheimer’s disease, Parkinson’s disease (PD), etc.) in clinic. During the past decade, the capabilities of using diffusion MRI method to study the anisotropical motion of water has been quite successful and have been widely adopted to depict the anatomy of the white matter fiber tracts in human brains.27−31 Therefore, in this work, we employed MRI method to study the behavioral differences of the water molecules before and after gelation in the enzymatic hydrogels.32−36 We hypothesize that, after hydrogelation, the water molecules that contribute the hydrogen bonds to stabilize the nanofibers would interact more strongly with the surfaces of nanofibers than the outer layer waters. This fixation of the conjugating waters will greatly shorten the longitudinal relaxation time T1, transverse relaxation time T2, and the ADC of their protons and those protons in outer layer waters, as illustrated in Figure 1. The denser the nanofibers in hydrogel, the more proportion of waters are fixed on the fiber surfaces, resulting in enhanced T1,

olecular self-assembly is ubiquitous but important in biological systems, underlying the formation of a wide variety of complex biological structures. For example, it is a central concern in structural biology to understand selfassembly and the associated noncovalent interactions (e.g., π−π stacking, hydrogen bonding) with which complementary interacting molecular surfaces are connected to form biological aggregates. On the basis of this understanding, people are able to design nonbiological mimics with new types of function.1−7 Supramolecular hydrogels, resulted from the self-assembly of small molecules to fixate large amount of water, have been extensively explored as biocompatible biomaterials in recent years and have shown promising applications in threedimensional cell culture, screening biomolecules, wound healing, drug delivery, tissue engineering, and microfabrication.8−16 Compared to other physically or chemically initiated hydrogelation (e.g., pH, temperature, ionic strength, ligand− receptor interactions, etc.), enzymatic regulations have drawn increased attention because they integrate the hydrogelation process with biological events and are more biocompatible.17−22 Up to date, many methods have been reported to study the processes of enzymatic hydrogelation, such as real time Fourier transform infrared (FT-IR) spectroscopy, circular dichroism (CD) spectra, rheology, high performance liquid chromatography (HPLC), fluorescence spectra, etc.23−26 However, to the best of our knowledge, there is no analytical method reported to study the water behavior accompanying the hydrogelation process, even though the hydrogen bonding © 2014 American Chemical Society

Received: March 17, 2014 Accepted: May 23, 2014 Published: May 23, 2014 5955

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G1322A pump and an in-line diode array UV detector using an Agilent Zorbax 300SB-C18 RP column, with CH3CN (0.1% of TFA) and ultrapure water (0.1% of TFA) as the eluent. The spectra of electrospray ionization-mass spectrometry (ESI-MS) were recorded on a LCQ Advantage MAX ion trap mass spectrometer (Thermo Fisher). 1H NMR and 13C NMR spectra were obtained on a Bruker AV 400. Rheological measurement was performed on an AR 2000ex (TA Instruments) system, 40 mm parallel plates was used during the experiment at the gap of 300 μm, or on a Haake RheoStress 6000 (Thermo Scientific), with cone-and-plate geometry (1 deg/20 mm) at the gap of 50 μm. Fluorescence spectra of 1 with alkali phosphatase in buffer as time elapse at pH 9.8 and 37 °C, and 2 in water at varying concentrations at pH 9 and room temperature were recorded on an F-2500 fluorescence spectrophotometer (Hitachi High-Techonologies Corporation, Japan) with excitation wavelength at 265 nm. Cryo transmission electron micrographs (Cryo-TEM) were obtained on a Tecnai F20 Transmission Electron Microscope from FEI company, operating at 200 kV. The cryo-samples were prepared as following: a special copper grid coated with carbon was put into Gatan SOLARUSTM plasma-cleaning system to remove hydrocarbon contamination on the sample holder, and then the sample was dropped on the copper grid in FEI Vitrobot sample plunger. The sample preparation was completed in the plunger. DSC Measurements. The thermoanalytical measurements were conducted on a TA DSC Q2000 equipped with a RCS 90 unit. The hydrogel samples were directly put into the DSC volatile sample pan. The following cycle was proposed: sample was cooled to −70 °C, maintained at −70 °C for 5 min, heated from −70 to 30 °C with a heating rate of 5 °C·min−1 and cooled from 30 to −70 °C, then heated from −70 to 30 °C with a heating rate of 5 °C·min−1. In Vitro Phantom MRI. The in vitro phantom MRI experiments were performed on a 9.4 T/400 mm (Agilent Technologies) wide bore scanner, using a 72 mm quadrature volume RF coil. The scanning procedure began with a localizer and then a series of inversion-prepared fast spin echo images were acquired for longitudinal relaxation time (T1) measurement (TR 6,000 ms, TE 5 ms, BW 25 kHz, slice thickness 2 mm, matrix 96 × 96, FOV 50 × 50 mm2, NEX 1). Twenty inversion times (TIs) were used varied from 10 to 2,500 ms. Signal intensity (SI) versus TI were fitted to the inversion− recovery model by nonlinear least-squares regression

Figure 1. Schematic illustration of how nanofibers in supramolecular hydrogel affect the relaxation of protons in waters conjugating to them and waters in outer layers.

T2, and ADC-weighted MRI of the hydrogels. On the basis of above hypothesis, we rationally designed enzymatic hydrogelator 1 which could self-assemble into nanofibers to form hydrogel upon the catalysis of alkaline phosphatase (ALP) in buffer (Figure 2). We chose phosphatase to initiate the self-

Figure 2. Schematic illustration of conversion of precursor 1 to hydrogelator 2 by alkaline phosphatase.

assembling of the hydrogelators into nanofibers and used this process to mimic the in vivo conditions due to the fact that many diseases (e.g., cancer, diabetes, Alzheimer’s disease, and multiple sclerosis) are associated with the abnormal activities of phosphatases and kinases.37−41 Thus, to study the enzymecontrolled hydrogelation with MRI would probably offer people with an alternative in vitro model to quickly study some aggregate-induced diseases in vivo which normally take a long progression time.

SI (TI) = A1 − A2·exp( −TI/T1) + σ

and the T1 relaxation time were determined for each sample. The transverse relaxation time (T2) were measured with a multiecho spin echo sequence, TR = 5500 ms and 12 echoes with echo times (TE) ranging from 25 to 300 ms, BW = 25 kHz, slice thickness 2 mm, FOV 50 × 50 mm2, matrix 96 × 96. T2 was then calculated with a monoexponential fitting model. Diffusion-weighted imaging was applied to determine the ADC maps using spin echo sequence (TR = 2300 ms, TE = 36.5 ms, NEX = 1, matrix = 64 × 64, FOV = 44 mm × 44 mm, slice thickness = 2 mm, BW = 52 kHz, b value 692 s·mm−2). δ (duration) and G (amplitude) of the pulsed gradients were 5 ms and 123.6 G·m−1, respectively. Δ (distance between the leading edges of the two pulsed gradients) was 25 ms. ADC values was calculated with a monoexponential model fitting. Syntheses and Characterizations. The preparations of 1 and 2 were described as below; Fmoc-Phe-Phe-Tyr-(PO-

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EXPERIMENTAL SECTION Materials. All the starting materials were obtained from Adamas or GL Biochem. Commercially available reagents were used without further purification, unless noted otherwise. All other chemicals were reagent grade or better. General Methods. HPLC analyses were performed on a Shimadzu UFLC system equipped with two LC-20AP pumps and an SPD-20A UV−vis detector using a Shimadzu PRC-ODS column, or on an Agilent 1200 HPLC system equipped with a 5956

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(OH)2)−OH (1) and Fmoc-Phe-Phe-Tyr(tBu)−OH (3) were synthesized following the literature method.42 Synthesis of Fmoc-Phe-Phe-Tyr-OH (2). The tBu protecting group of 3 was cleaved with 95% TFA in DCM for 3 h at room temperature to yield 2 after HPLC purification. (Supporting Information Scheme S1) Characterization of Fmoc-Phe-Phe-Tyr-(PO(OH)2)−OH (1). 1 H NMR of 1 (400 MHz, DMSO-d6) δ (ppm): 8.42−8.33 (d, 1 H), 8.15−8.07 (d, 1 H), 7.91−7.86 (d, 2 H), 7.66−7.59 (t, 2 H), 7.44−7.39 (t, 2 H), 7.29−7.23 (m, 10 H), 7.20−7.15 (t, 2 H), 7.13−7.09 (d, 2 H), 4.68−4.60 (m, 1 H), 4.52−4.43 (m, 1 H), 4.28−4.21 (m, 1 H), 4.21−4.17 (m, 1 H), 4.16−4.10 (m, 2 H), 3.13−3.07 (d, 1 H), 3.07−3.04 (t, 1 H), 3.00−2.93 (m, 1 H), 2.93−2.90 (m, 1 H), 2.88−2.79 (m, 1 H), 2.76−2.67 (m, 1 H) (Figure S1, Supporting Information). 13C NMR of 1 (100 MHz, DMSO-d6) δ (ppm): 172.58, 171.29, 171.05, 155.64, 150.22, 143.77, 143.67, 140.62, 140.60, 138.11, 137.57, 132.80, 130.14 (2C), 129.27 (2C), 129.16 (2C), 127.95 (4C), 127.58 (2C), 127.03 (2C), 126.21, 126.13, 125.31, 125.21, 120.03 (2C), 119.84, 119.79, 65.69, 56.04, 53.62, 53,59, 46.51, 37.59, 37.39, 35.96 (Figure S2, Supporting Information). MS: calculated for C43H56N9O16 [(M + H)+] 778.25, obsvd. ESIMS: m/z 778.10. Characterization of Fmoc-Phe-Phe-Tyr-OH (2). 1H NMR of 2 (400 MHz, DMSO-d6) δ (ppm): 8.42−8.25 (d, 1 H), 8.18−8.03 (d, 1 H), 7.97−7.83 (d, 2 H), 7.68−7.59 (t, 2 H), 7.46−7.39 (t, 2 H), 7.31−7.21 (m, 10 H), 7.08−7.04 (d, 2 H), 6.75−6.66 (d, 2 H), 4.69−4.59 (m, 1 H), 4.47−4.38 (m, 1 H), 4.30−4.22 (m, 1 H), 4.22−4.18 (t, 1 H), 4.18−4.11 (m, 2 H), 3.14−3.03 (m, 1 H), 3.03−2.98 (d, 1 H), 2.98−2.94 (t, 1 H), 2.93−2.88 (t, 1 H), 2.88−2.82 (m, 1 H), 2.77−2.67 (m, 1 H) (Figure S3, Supporting Information). 13C NMR of 2 (100 MHz, DMSO-d6) δ (ppm): 172.79, 171.23, 170.88, 156.02, 155.62, 143.75, 143.69, 140.63, 140.61, 138.12, 137.51, 130.05 (2C), 129.28 (2C), 129.16 (2C), 127.97 (4C), 127.60 (2C), 127.28, 127.04 (2C), 126.63, 126.17, 125.31, 125.22, 120.04 (2C), 115.03 (2C), 65.68, 56.07, 53.60, 53,56, 46.52, 37.58, 37.37, 35.97 (Figure S4, Supporting Information). MS: calculated for C43H56N9O16 [(M + H)+]: 698.29, obsvd. ESIMS: m/z 698.06.

Figure 3. Proposed molecular arrangement of 2 in gels I or II and morphological studies of gels I and II. (A) Hierarchical self-assembly of 2 into a five-membered supermacrocycle and result in nanofibers in gels I or II, top (left) and side (right) views (VMD). (B) Cryo-TEM image of gel I (inset shows 300 μL of 1 wt % 1 in water at pH 9.8 (left), and corresponding gel I obtained after the incubation of the solution with 200 U/mL of alkaline phosphatase at pH 9.8 and 37 °C for 30 min (right)). (C) Cryo-TEM image of gel II (inset shows 300 μL of water solution of 2 at 1 wt % (left), and corresponding gel II obtained after adjustment of the pH value of the solution to 9 with 10 M HCl (right)).

with 10 M HCl (right vial in the inset of Figure 3C). The transparent hydrogels formed with 1 or 2 suggested that there were no microcrystalline aggregates in the hydrogels to scatter visible light, in agreement with the cryo transmission electron microscopy (cryo-TEM) images of the hydrogels (Figure 3B and C). To study the enzymatic formation process of gel I, dynamic time sweep was chosen to examine the change of viscoelasticity of the solution containing 1 upon phosphatase addition. ALP (200 U/mL) was added to the solution of 1 wt % 1 in the buffer for the rheological measurement. As shown in Supporting Information Figure S5A, storage modulus (G′) and loss modulus (G″) of the solution are very small at the time when the enzyme was added, indicating that the solution of 1 indeed behaves as a low-viscosity liquid. Thereafter, both G′ and G″ gradually increase with time, and the values of G′ start to dominate over those of G″ around 60 s after the addition of the phosphatase, suggesting the approach of the gelling point. After 0.8 h, the G′ values are about 5-time larger than those of G″, indicating the extensive formation of a three-dimensional matrix in the hydrogel. After confirming that 1 is indeed an efficient precursor for enzymatic hydrogelation, we examined the transition of 1 to 2 at 60 s (i.e., gelling point at this condition) after the addition of ALP. HPLC analysis indicated that 23% of 1 was converted to 2 at this point (Figure S5B, Supporting Information). Morphological and Fluorescence Studies, Possible Molecular Arrangement. The microscopic structure of gel I under cryo-TEM exhibited regularly arranged, long fibers with an average width of 4.3 ± 0.6 nm (Figure 3B). Cryo-TEM

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RESULTS AND DISCUSSION Syntheses of 1 and 2. We began the study with the syntheses of the precursor 1 and hydrogelator 2.43,44 The syntheses of 1 and 2 are facile and straightforward as follows (Figure 2 and Supporting Information): tripeptides (FmocPhe-Phe-Tyr(PO(OH)2)−OH) for 1, or Fmoc-Phe-Phe-Tyr(tBu)−OH (3) for 2) were synthesized with solid phase peptide synthesis (SPPS). Deprotection of the tbutyl (tBu) group from 3 with 95% trifluoroacetic acid (TFA) in dichloromethane yields 2 after HPLC purification. Hydrogelation and Rheological Characterizations. After obtaining 1 and 2, we tested their gelation abilities. In brief, 3 mg of 1 or 2 was suspended in 300 μL of water (12.9 mM for 1, or 14.3 mM for 2). Adjustment of the pH values of suspension of 1 to 9.8 with solid Na2CO3 resulted in clear solutions (left vial in the inset of Figure 3B). After 200 U/mL of ALP (10 μL) was added into the solution and incubated for 30 min at 37 °C, transparent gel I was obtained (right vial in the inset of Figure 3B). Careful addition of solid Na2CO3 into the suspension of 2 in water resulted in clear solution (left vial in the inset of Figure 3C), and transparent gel II was afforded by carefully adjusting the pH value of the solution back to 9 5957

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Figure 4. Rheological analyses and cryo-TEM images of gel I containing different concentrations of 1. Upper panels indicate the dynamic frequency sweep of the storage modulus (G′) and the loss modulus (G″) of gel I at 1 wt % (left), 2 wt % (middle), and 4 wt % (right). Lower panels show cryo-TEM images of gel I at 1 wt % (left), 2 wt % (middle), and 4 wt % (right). All rheological measurements were carried out at 37 °C and strain of 1.0%.

image of gel II showed entangled long nanofibers with an average width of 4.7 ± 0.8 nm (Figure 3C). Participation of the fluorenyl moiety of the Fmoc group in the gelation was investigated by measuring the time-course fluorescence spectra of 1 upon the addition of the enzyme, or those of 2 at different concentrations in buffer (Figure S6, Supporting Information). At a diluted concentration of 0.94 mM without ALP, 1 was found to remain at the molecular state, suggested by its emission peak at 316.5 nm from the fluorenyl moiety excited at 265 nm. After the addition of ALP, its emission maximum redshifted to 327 nm over time. For 2, at a lower concentration of 0.94 μM, it has an emission peak at 306 nm, indicating that 2 is at the molecular state. When the concentration of 2 was increased to 0.94 mM, the emission peak of 2 red-shifted to 332.5 nm. Both of aforementioned redshifts of the emissions confirmed the participation of the self-assembly of the fluorenyl moieties in the hydrogelation processes of 1 or 2, with their aromatic rings overlapping in parallel orientation. According to the amphiphilic structure of the Fmoc-tripeptide hydrogelator 2, a possible molecular arrangement was proposed for the nanofibers in gel I obtained during the gelation process in water.45 As shown in Figure 3A, from top view, the nanofiber of gels I or II is likely a cylindrical micellar structure formed by the self-assembly of 2. In this model, molecules of 2 centripetally stack to form the hydrophobic inner tubular layer due to the strong π−π interactions of the fluorenyl moieties and the peptide segments of 2 stretching to the surrounding water to form the hydrophilic outer layer. Each molecule of 2 is placed under and between two molecules of 2 on the above layer with a layer distance of 0.4 nm. Five molecules of 2 constitute one repeating unit of the tubular

structure. From this molecular arrangement model, the nanofiber of gel I has a calculated diameter of 5.3 nm, in good agreement with the cryo-TEM measurement (i.e., 4.3 nm). Estimation of Pore Size of Gel I with DSC Measurement. Since the hydrogelation process of 1 at 1 wt % was confirmed with the catalysis of ALP, we further studied how the concentrations of 1 affects the pore sizes in gel I, as well as its elasticity. First, we used dynamic frequency sweep at the strain of 1.0% and 37 °C to study mechanic properties of gel I containing different concentrations of 1. The gels I were formed in Na2CO3−NaHCO3 buffer (pH 9, 0.2 M) treated with 64.3 nmol/U phosphatase at 37 °C for 10 h. As shown in Figure S7 in Supporting Information, the values of the storage modulus (G′) and the loss modulus (G″) of gel I exhibited a weak dependence from 0.5 to 3.0% of strain (with G′ dominating G″), indicating that the samples are hydrogels. Optical images showed the all the gels I at 1, 2, and 4 wt % are transparent (Figures S8, Supporting Information). In addition, for each sample, the G′ and G″ values of gel I slightly increased with the increase of frequency from 0.1 to 11 Hz. When concentrations of 1 were raised from 1 to 4 wt %, both G′ and G″ values of gel I increased (upper panels in Figure 4). At 1 Hz, The G′s for gels I at 1%, 2%, and 4% (wt) are 84.9, 396.2, and 1050.0 Pa, respectively, suggesting the contents of 1 affect the elasticity of the hydrogels dramatically. The microscopic images of gels I under cryo-TEM consistently showed that with the increase of 1, the fibrous networks in gels I became denser, as shown in the lower panels of Figure 4. Since the cryo-TEM images only show the two-dimensional fibrous networks in the hydrogels, the actual pore sizes formed by the nanofibers could 5958

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Figure 5. DSC heating curves for gel I at 1 wt % (A), 2 wt % (B), 4 wt % (C), and Na2CO3−NaHCO3 buffer (pH 9, 0.2 M) (D). Heating rate: 5 °C· min−1.

Table 1. T1, T2, and ADC of Gels I Formed by Different Concentrations of 1 (0, 1, 2, 4 wt %) in Na2CO3−NaHCO3 buffer (pH 9, 0.2 M) Measured with IRFSE, MEMS, and SEMS phantom

pore size R (nm)

G′ at 1 Hz (Pa)

buffer 1 wt % 2 wt % 4 wt %

NA 55.17 39.43 NA

NA 84.9 396.2 1050.0

T1 (ms) 2417 2164 2014 1839

± ± ± ±

68 39 58 33

T2 (ms) 220 198 180 141

± ± ± ±

16 7 6 3

ADC (×10−3 mm2·s−1) 1.89 1.85 1.81 1.79

± ± ± ±

0.12 0.08 0.13 0.06

the pore radius R of the porous materials can be calculated with equation

not be directly estimated. Therefore, differential scanning calorimetry (DSC) was applied to determine the pore radii for gels I formed with 1, 2, and 4 wt % of 1. The DSC method, originally developed by Ishikiriyama and co-workers, is based on the calorimetric analysis of a liquid−solid transition (e.g., water to ice) in liquid-filled porous materials.46−48 In brief, the pore size distribution of porous materials could be determined from the freezing or melting DSC curves of the freezable pore water. In our case, the hydrogels could be assumed as nanofiber-based, three-dimensional, porous materials. Water molecules sticking to the nanofibers via hydrogen bonding could be regarded as the fixed layer and those water molecules attaching to the fixed layer could be regarded as the adsorbed layer. The remaining water molecules can be regarded as bulk waters and could exchange with the waters in adsorbed layer. Different pore size regulates the melting point of the freezable pore water (i.e., water in fixed layer and adsorbed layer). Thus,

R = −64.67/ΔT + 0.57 (nm)

(1)

where ΔT = T − T0 is the decrease of freezing temperature of the water molecules fixed on the networks. In our case, T0 is the freezing temperature of water in the Na2CO3−NaHCO3 buffer (pH 9, 0.2 M), and T is the freezing temperatures of the water molecules fixed on the nanofibers in the hydrogels. The DSC heating curves for gels I at 1, 2, and 4 wt % are shown in Figure 5. Each DSC curve shows two endothermic peaks: the first peak at lower temperatures is attributed to the melting of water confined in the network pores (in our case, in the two layers conjugating to the nanofiber networks); the second peak, at higher temperatures, is assigned to the melting points of bulk water. Thus, the first peaks, associating with the pore sizes, are employed to calculate the pore radii with eq 1. For gels I at 1 5959

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and 2 wt %, T values are −5.884 and −6.364 °C, respectively. T0 of the Na2CO3−NaHCO3 buffer is −4.699 °C. Thus, the pore radius of gels I at 1 and 2 wt % are calculated to be 55.17 and 39.43 nm, respectively, which are well consistent with cryoTEM observations (Figure 4). From Figure 5, we also noticed that the endothermic peaks at lower temperature become weaker and broader and could not be well separated for gel I at 4 wt %. The G′ values at 1 Hz of gels I at 1, 2, and 4 wt %, as well as their pore sizes calculated from DSC data are summarized in Table 1. Using MRI to Study Hydrogelation. The changes of hydration properties in gels I with different concentrations of 1 lead us to hypothesize that within a defined volume, the denser the nanofibers, the more proportion of water molecules that will conjugate to their surfaces through hydrogen bonding. This will reduce the water diffusion rate around the nanofibers and shorten the water proton magnetic resonance (MR) relaxation time. Therefore, we applied MRI method to quantify the shortening of relaxation time T1, T2, and water diffusion rate of protons in the hydrogels, which in return could be utilized to characterize the hydrogelation process. For this purpose, ALPcontrolled gels I containing different concentrations of 1 (i.e., 1, 2, and 4 wt %) at pH 9 were investigated, with Na2CO3− NaHCO3 buffer at pH 9 and the same ionic strength (0.2 M) as control. T1-weighted, T2-weighted, and diffusion-weighted MR images of these hydrogel phantoms were acquired on a 9.4 T MR scanner (Figures S10−S12, Supporting Information). Analyses of signal intensity versus inversion time and echo time versus diffusion gradient gave the corresponding T1 and T2 relaxation times, as well as the ADC of each gel at given concentration, also shown in Table 1. T1-weighted MR images showed dramatic signal increase with increased concentration of 1, suggesting a significant decrease of the corresponding T1 values. This effect arises from the fact that as more and more bulk waters being loaded to the surface of nanofibers, their longitudinal relaxation is strongly reduced by the nanofibers. Exchange between the bulk water and this increased surface component therefore decreases the overall water T1 relaxation time, as observed in our measurement. The same process also affected the T2-weighted MR contrast, in which the signal intensity decreased dramatically with increased concentration of 1, due to the shortened transverse relaxation time. On the other hand, the water diffusion in the hydrogel was modified somewhat differently in comparison with T1 and T2 that are more due to the surface layers of the naofibers. Diffusionweighted MR images showed increased intensity with increased concentration of 1, and can be attributed to a slower water proton diffusion rate (Figure 6). The changes of the water diffusion rate are influenced by the nanofiber network (i.e., the pore size of the nanofibers that spatially restricted the volume that water protons could freely diffuse without much hindrance). With increased concentration of 1, the pore size of the network decreases to a value that would strongly affect the water diffusion property. Interestingly, both ADC measurements and T measurement from DSC stabilized at high concentration of 1, which saturated in its concentration of around 4 wt %. Therefore, MR measurements provide a quite sensitive approach to probe the microstructures of the nanofiber network, and could potentially be used to dynamically monitor the self-assembly of the nanofiber network initiated by enzymatic hydrogelation.

Figure 6. T1-weighted MR phantom images (upper row, inversion time 1714 ms), T2-weighted MR phantom images (middle row, echo time: 200 ms), and diffusion-weighted MR phantom images (third row, b = 687 s·mm−2, direction 2) of gels I formed by different concentrations of 1 (0, 1, 2, and 4 wt %) in Na2CO3−NaHCO3 buffer (pH 9, 0.2 M) treated with 64.3 nmol·U−1 phosphatase at 37 °C for 5 h and imaged with a 9.4 T MR scanner (TR 6000 ms, TE 5 ms).

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CONCLUSIONS In summary, by rational design of the precursor 1, we obtained the enzymatic hydrogel gels I constituted of 3D nanofiber networks and waters gelled. The nanofiber structures, formed from the phosphate-controlled self-assembly of the amphiphilic hydrogelator 2, could be controlled by the quantity of 1 in the hydrogels. The elasticity increases in Gels I, while the nanofiber pore sizes decrease with the increased concentrations of 1. Decrease of pore sizes was accompanied by more proportion of bulk water molecules being fixed to the nanofibers, which was validated by DSC measurements. Using T1, T2, and diffusionweighted MRI, we studied for the first time how the density of the nanofibers affects the proton relaxation in gels I. Our results indicated that with higher concentrations of 1 (i.e., denser of nanofibers and smaller pore sizes) in gels I, T1 and T2 are significantly shortened and water proton diffusion rates are greatly reduced in the hydrogels. Our study could provide a new method for the detection of enzyme activity in vitro or to mimic and study certain in vivo fibrous aggregate-induced diseases that normally would take a long progression time.

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ASSOCIATED CONTENT

S Supporting Information *

Synthetic route for 2, 1H NMR, and 13C NMR spectra of 1 and 2, HPLC conditions, Scheme S1, Figures S1−S12, and Tables S1−S3. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: kzhong@hmfl.ac.cn. *E-mail: [email protected]. Author Contributions

W.W., J.Q., and A.T. contributed equally. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors are grateful to Prof. Yi Cao for his assistance of rheology study. This work was supported by the National Natural Science Foundation of China (Grants 21175122, 91127036, 21375121, and U1232212), the Fundamental Research Funds for Central Universities (WK2060190018). 5960

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dx.doi.org/10.1021/ac500967x | Anal. Chem. 2014, 86, 5955−5961