Nondestructive Characterization of Biodegradable Polymer Erosion

Department of Mechanical and Aerospace Engineering, Case Western Reserve University, 2123 Martin Luther King Jr. Drive, Cleveland, Ohio 44106, United ...
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Nondestructive Characterization of Biodegradable Polymer Erosion in Vivo Using Ultrasound Elastography Imaging Haoyan Zhou,† Anna Gawlik,† Christopher Hernandez,† Monika Goss,† Joseph Mansour,‡ and Agata Exner*,§ †

Department of Biomedical Engineering and §Department of Radiology, Case Western Reserve University, 11100 Euclid Avenue, Cleveland, Ohio 44106, United States ‡ Department of Mechanical and Aerospace Engineering, Case Western Reserve University, 2123 Martin Luther King Jr. Drive, Cleveland, Ohio 44106, United States S Supporting Information *

ABSTRACT: Significant advancements in biodegradable polymeric materials have been made for numerous biomedical applications including tissue engineering, regenerative medicine, and drug delivery. The functions of these polymers within each application often rely on controllable polymer degradation and erosion, yet the process has proven difficult to measure in vivo. Traditional methods for investigating polymer erosion and degradation are destructive, hampering accurate longitudinal measurement of the samples in the same subject. To overcome this limitation, we have utilized ultrasound elastography imaging as a tool to nondestructively measure strain of poly(lactic-co-glycolic acid) (PLGA) phase sensitive in situ forming implants (ISFI), which changes with progressive loss of structural integrity resulting from polymer erosion. Using this tool, we investigated erosion kinetics of implants comprised of three different PLGA molecular weights (18, 34, and 52 kDa) in vitro and in vivo. The in vitro environment was created using a novel polyacrylamide based tissue mimicking phantom while the in vivo experiment was performed subcutaneously using a rat abdominal model. A strong linear relationship independent of polymer molecular weight was found between average strain values and erosion values in both the in vitro and in vivo environment. Results support the use of a mechanical stiffness-based predicative model for longitudinal monitoring of material erosion and highlight the use of ultrasound elastography as a nondestructive tool for measuring polymer erosion kinetics. KEYWORDS: ultrasound elastography, in situ forming implant, erosion, characterization, nondestructive

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INTRODUCTION

and erosion processes in vivo, which involve very specific and complex interactions with local tissue at the implantation site. Studies done in vitro have shown poor correlations with in vivo behavior, suggesting that in vitro models can rarely represent an in vivo system adequately. Therefore, new nondestructive technologies that can be used to study polymer degradation and erosion in situ are highly desirable. Traditionally, gel permeation chromatography (GPC) and gravimetric analysis have been used as tools for determination of molecular weight changes during polymer degradation and mass loss during the erosion process, respectively. However, these methods are destructive, making longitudinal studies of a

Continued rapid development of new biomaterials for applications in tissue engineering,1,2 regenerative medicine,3,4 gene therapy,5,6 and controlled drug delivery7−9 has stimulated the ongoing need for new tools facilitating high throughput in vitro characterization and streamlined evaluation of in vivo performance.10,11 One of the most important parameters in many of these applications is the rate of in vivo retention12 resulting from polymer degradation and subsequent erosion. Significant advances in the fields using polymeric biomaterials have led to increasingly sophisticated technology with new challenges including quantification of local and systemic pharmacokinetics associated with a polymeric implant drug delivery system13−15 or the process of cells migrating into a degrading tissue scaffold.16,17 Many of these sought-after parameters are, in turn, highly dependent on the degradation © XXXX American Chemical Society

Received: March 10, 2016 Accepted: April 20, 2016

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DOI: 10.1021/acsbiomaterials.6b00128 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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Figure 1. Void creation and ISFI implant injection process.

(PDMS). The detection range was found to be between 47 kPa and 4 MPa, with a detectable difference as low as 157 kPa based on the optimized scan setup during experimentation.30 For this study, USE technique was applied to a PLGA based in situ forming implant (ISFI) drug delivery system for characterization of implant erosion in vitro and in vivo. Implant solutions prepared with three different molecular weights of PLGA were examined. For the in vitro study, we injected implants into a novel polyacrylamide based tissue mimicking phantom, and then performed USE sequentially at designated time points. For the in vivo study, we implanted the system subcutaneously on the abdominal side of Sprague−Dawley rats, and then performed USE. In vitro and in vivo erosion of the implants was measured simultaneously using gravimetric analysis. Finally, the relationship between strain and erosion was analyzed by comparing both data sets. The goal of this study was to demonstrate the feasibility of establishing a mechanical stiffness-based predictive model using ultrasound elastography for noninvasive and nondestructive characterization of implant erosion in an example of ISFI drug delivery system.

single sample over time impossible, leading to an increased number of experimental animals required for a particular study. This nonsequential analysis of materials also leads to inaccurate conclusions, as there can be broad batch-to-batch and animalto-animal variations. Also, the necessity to surgically remove individual samples increases the likelihood of specimen damage prior to analysis. In an attempt to circumvent the issues associated with these invasive and destructive techniques, predictive degradation and erosion models have been developed. Models for degradation have used random theory to predict the random scission process of polymer chains,18 which is assumed to be a first- or second-order kinetic process.19 Erosion models have been especially complex because of the multitude of involved parameters such as swelling, dissolution of oligomers or monomers, and morphological changes. Even though very comprehensive models have been developed including the Monte Carlo model and the diffusion theory-based model,16,20 they have proven to be only somewhat predictive. Recently, research efforts have been shifting to the use of biomedical imaging to noninvasively monitor the degradation and erosion process of implants in vivo.12,21−25 These imaging techniques allow for high throughput, serial (i.e., longitudinal) studies of biodegradable materials in their intended implanted state. Mader et al. have already demonstrated the feasibility of using MR imaging to monitor polymer tablet behaviors such as size, water content and erosion in vivo.21 Most recently, Artzi et al. demonstrated noninvasive assessment of implanted hydrogel erosion using in vivo fluorescence imaging.12 Although these techniques have shown to be successful in preclinical animals, the high cost of MRI and the translatability of fluorescence imaging to human subjects have hindered these techniques from achieving widespread use. Ultrasound elastography (USE) is a dynamic technique that uses ultrasound to assess the mechanical stiffness of materials noninvasively and nondestructively by measuring material distortion or strain in response to external compression. The distortion is measured based on speckle tracking of two ultrasound B-mode frames before and after compression. In order to estimate the speckle displacement, either an A-line cross-correlation method or a tissue Doppler method is used. A-line cross-correlation is more widely applied due to its high sensitivity. The strain is calculated cumulatively over frames throughout the entire compression process and plotted as a color coded map. USE has already been widely used as a “computerized palpation” tool to characterize tissue mechanical properties for diagnostic purposes such as early detection of breast and prostate cancer.26,27 Previous studies done by Kim et al. have shown the feasibility of using ultrasound elastography for monitoring tissue scaffold degradation28 and tissue ingrowth.29 In our previous study, we characterized USE for its application in imaging stiffer materials by comparing it to the gold standard mechanical test using polydimethylsiloxane

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MATERIALS AND METHODS

Materials. All materials were used as received with no further purification. Poly(DL-lactide-co-glycolide) (PLGA 50:50: MW 18 kDa, inherent viscosity 0.19 dL/g; MW 34 kDa, inherent viscosity 0.29 dL/ g; MW 52 kDa, inherent viscosity 0.41 dL/g) was purchased from Evonik, Birmingham, AL. N-methylpyrrolidinone (NMP) was purchased from Fisher Scientific, Waltham, MA, and sodium fluorescein (MW 376.28) was purchased from Sigma-Aldrich, St. Louis, MO. Acrylamide, bis-acrylamide, ammonium persulfate (APS), N,N,N′,N′-tetramethylethylenediamine (TEMED), titanium dioxide, and phosphate buffered saline were purchased from Fisher Scientific, Waltham, MA. In Situ Forming Implant (ISFI) Solution Preparation. ISFI solutions of PLGA in NMP were prepared with 18 kDa, 34 kDa, or 52 kDa polymers. Sodium fluorescein was used as a mock drug because it has a similar chemical structure to the clinically used chemotherapeutic, doxorubicin. The polymer and mock drug were added to NMP in glass scintillation vials and allowed to mix overnight in an incubator shaker at 37 °C until the polymer had completely dissolved. The final ISFI solution had a 60:39:1 mass ratio of solvent:polymer:drug. The solution was used immediately after incubation. Tissue Mimicking Phantom Fabrication and ISFI Injection. Breast tissue mimicking phantoms (50 kPa) were made from 10% (w/ v) solutions of acrylamide and bis-acrylamide (37.5:1 ratio, respectively) in 1X PBS solution. Titanium dioxide was added as a scattering agent to more clearly define the boundaries between the phantoms and implant samples when performing ultrasound elastography. The acrylamide solutions were cross-linked by free radical polymerization using 1.7% and 0.1% (v/v) of TEMED and APS, respectively. The phantoms were polymerized in plastic molds with two polyacrylamide gel layers. One hundred milliliters of the above mixture was stirred and poured into the plastic phantom mold to form the first layer. After the first layer gelled, custom-made hemispheres of sucrose, 7 mm in diameter, were placed on the gelled B

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Figure 2. Animal study design: 100 μL of each polymer solution (18, 34, and 52 kDa) was injected subcutaneously at one of three locations on the abdominal side of the rat. The injection location of each polymer type was alternated between animals to account for variability between tissue locations. USE scan was performed every other day to assess the mechanical stiffness of the implants. To measure the erosion, implants were removed at day 1, 4, 8, 12, 16, and 22 for gravimetrical analysis. In Vitro Mechanical Testing. A sheet-shaped implant was created by injecting polymer solution into a sheet-shaped void within the tissue mimicking phantom. A disc-shaped (3 mm in diameter) PLGA implant was then obtained from the PLGA sheet using a biopsy punch prior to mechanical testing. The Young’s modulus of the implants was determined using unconfined compressive mechanical testing with a rheometer (Rheometrics RSAII, NJ). A strain rate of 0.01/s was used. Testing time was optimized for each sample to ensure at least 30% strain. After the stress−strain curve was generated, a threshold of 0.1 N was used to ensure consistent contact between the compressing probe and PLGA sample. Thus, measurements with associated force less than 0.1 N were discarded. A first order polynomial fit was used to determine the linear region of the stress−strain curve for modulus calculation. The Young’s modulus (E) was determined by finding the slope of the linear region with strain no greater than 20% on the stress−strain curve. In Vitro Erosion Study. At designated time points (days 1, 3, 6, 9, 12, 16, and 21), samples were removed from the phantoms. The procedure was carefully performed to avoid any damage to the samples to ensure 100% sample recovery. A gravimetrical analysis was performed after freeze-drying the harvested samples to obtain the implant dry mass. To ensure complete removal of water from the implants, the freeze-drying process was allowed to run for 72 h. Erosion was then calculated using the eq 1.

polyacrylamide. The second 100 mL acrylamide mixture was then poured on top of the first layer, embedding the sucrose hemispheres. Phantoms were allowed to swell in PBS for 6 days until the phantom volume reached plateau and the embedded sucrose hemispheres had fully dissolved. This resulted in voids in the phantom in the same shape as the sucrose hemispheres. These voids were flushed 3 times using PBS before implant injection. Then 150 μL of polymer solution was injected into the voids to form solid implants, which had the same dimensions and shape as the voids. PLGA implants embedded in tissue mimicking phantoms were kept in 300 mL of PBS at 37 °C using an incubator shaker (80 rpm, 37 °C) for 25 days. PBS was replaced daily. Three phantoms were made for polymer injection at each time points, and injections were performed in triplicate. The void creation and ISFI implant injection process is summarized in Figure 1. In Vitro USE Scan and Image Analysis. Implants were scanned daily using a clinical ultrasound system (Toshiba Aplio500) with a linear array transducer centered at 12 MHz to evaluate their mechanical stiffness change over time. A laboratory-designed stage and a linear actuator were used to induce uniform compression for all samples. The USE scans were carried out as described previously.30 Briefly, the ultrasound transducer was fixed to the laboratory designed stage, and the programmable linear actuator was used to provide uniform displacement. Since the ultrasound transducer was fixed, samples between the transducer and linear actuator were compressed by pushing up from below. An optimized 10% prestrain was applied to ensure contact between the transducer and polyacrylamide phantom. During the scan, a compression cycle composed of three compressions and three dilations was induced. The displacement of each compression and dilation equaled 1 mm. A time frame tissue doppler tracking (TDT) algorithm was used to calculate axial strain.31 Color doppler imaging (CDI) frequency was set at 8.8, whereas the velocity scale was 0.3 cm/s to accommodate the maximum value of the velocity profile. Velocity smoothing was used to optimize the strain calculation. After the scan, a color-coded strain map was generated and superimposed on top of the B-mode image. The regions of interest (ROI) were manually selected based solely on the B-mode images to remove potential operator dependent bias. The mean strain value of each implant was obtained by spatially averaging over the entire implant ROI.

erosion =

polymer dry mass initial polymer mass

(1)

In eq 1, polymer dry mass was obtained after freeze-drying and initial polymer mass was known from the initial formulation. Animal Preparation and Implant Injection. The animal study was performed using 6−10 week old male Sprague−Dawley rats (body weight 200−300g, Charles River Laboratories Inc., Wilmington, MA). The protocol followed the National Institutes of Health (NIH) guidelines for animal care and was approved by the Case Western Reserve University Institutional Animal Care and Use Committee. The anesthesia was induced using 3% isoflurane inhalation and maintained at 2% isoflurane inhalation with an oxygen flow rate of 1 l/min (EZ150 Isoflurane Vaporizer, EZ Anesthesia). The rat was placed in the supine position on a 37 °C heating pad, and once completely anesthetized, C

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ACS Biomaterials Science & Engineering the hair on the abdomen region was removed. Implant solutions were prepared as described earlier. 100 μL of each polymer solution was injected subcutaneously at one of three locations on the abdominal side of the rat using a 21-gauge hypodermic needle. The injection location of each polymer type was alternated between animals to account for variability between tissue locations (Figure 2). Three animals were used for each time point. In Vivo USE Scan. In vivo USE scans were performed using the same experiment set up as described earlier. The rats were anesthetized on a custom-made platform that was attached to the linear actuator. The ultrasound transducer was fixed to a customized stage, hence the animal was compressed by pushing up from below using the linear actuator. This set up was used to induce uniform strain on each animal. A standard B-mode image was used to determine the starting compression point, where the rat’s abdominal skin was just barely in contact with the ultrasound transducer. During the scan, unlike in the in vitro study, a displacement of 2 mm was used in the compression and dilation cycle to produce greater strain for detection. Other parameters were the same as those used in the in vitro experiment. The USE scan was performed every other day from day 1 to day 22 (Figure 2). In Vivo Erosion Study. At designated time points (days 1, 4, 8, 12, 16, and 22), the animals were sacrificed (Figure 2). The implants were excised and freeze-dried. The dry sample mass was obtained using gravimetrical analysis. The erosion was calculated using the equation described in the in vitro section.

from day 9 to day 23. It is worth noting that the sample size was n = 3 for most days but n = 2 for days 8, 11, 12, 16, 19, 21, 22, and 23. This is due to sample damage during the harvesting and preparation process at the later time points. The accumulated strain in reference to the original geometry of each sample was color coded and superimposed onto the Bmode image (Figure 4). The ultrasound transducer was located at the top of the image, while the linear actuator induced the compression cycle from the bottom. In this figure, the color scale bar on the right indicates the accumulated strain, deep blue to dark red indicating hard to soft. Reduction in strain was observed over time for all PLGA molecular weights: 18, 34, and 52 kDa, except for the period from day 0 to day 1. The implants were found to stiffen in the first day (Figure 4) due to the phase inversion process. Then, the mean strain value for each time point was calculated over 3 samples and plotted in Figure 5 A. In this plot, the same implant stiffening from day 0 to day 1 was observed. In addition, strain was found to change at different rates for implants with different molecular weights. 18 kDa implants had the highest rate of increasing strain, followed by 34 kDa then 52 kDa. These implants also reached their plateau stage at different time points: 18 kDa PLGA at 14 days, 34 kDa PLGA at 17 days and 52 kDa PLGA at 21 days. In Figure 5B, the rate of implant erosion was also found to be different depending upon the molecular weight of the implant. 18 kDa implants had the highest rate of erosion followed by 34 kDa then 52 kDa. Implant Erosion in Vivo. The color-coded strain map and B-mode ultrasound images for 18 kDa, 34 kDa and 52 kDa implant at day 1, day 10 and day 20 are summarized in Figure 6. For each implant type, the top row represents the color coded strain map and the bottom row represents B-mode images. The abdominal skin is located at the top of the image, with the ultrasound transducer fixed above and linear actuator pushing up from the bottom. The region of interest (white dashed circle) was selected based on B-mode images. As animal experiments progressed from day 1 to day 20, the implants became softer and the regions of interest became smaller (Figure 6). The mean strain values of each implant type were calculated for all 5 animals and plotted in Figure 7A. Again, the strain was found to change at different rates based on the molecular weight of the implants. Eighteen kDa implants had the highest rate of increasing strain, followed by 34 kDa then 52 kDa. In addition, the erosion of each implant type is plotted in Figure 7B. The erosion rate is also molecular-weight-dependent. Eighteen kDa PLGA eroded the fastest, followed by 34 kDa then 52 kDa.

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RESULTS Implant Erosion in Vitro. The mechanical test was performed on 34 kDa PLGA implants. As shown in Figure 3,

Figure 3. Young’s modulus of 34 kDa PLGA implants.

the Young’s modulus of the PLGA implants was found to decrease over time after injection into the tissue mimicking phantom. The highest modulus was found to be 878 kPa at day 1, whereas the lowest modulus was 7 kPa found at day 22. The decrease in modulus continued until day 8, then no clear trend was observed, and modulus values ranged from 7 to 99 kPa

Figure 4. In vitro: Color-coded strain map of implant over time. D

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Figure 5. In vitro: (A) strain of implants obtained from ultrasound elastography scan, where the dashed line indicates the water strain value under the same scan condition as explained in the discussion; (B) implant erosion in phantoms.

Figure 6. In vivo color-coded strain map of implant over time.

Figure 7. In vivo: (A) strain of implants obtained from ultrasound elastography scan, (B) implant erosion.

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Figure 8. (A) In vitro erosion and strain correlation, (B) in vivo erosion and strain correlation.

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DISCUSSION As stated earlier, some of the 34 kDa samples became fragile and broke, leading to difficulty in mechanical testing. Because of the challenges faced with this first set of PLGA implants, mechanical testing was not performed on the remaining two molecular weight implants. From mechanical testing of 34 kDa PLGA, a decreasing modulus trend was observed within the first 8 days, which indicates the loss of structural integrity during the degradation and erosion process. We anticipated the same decreasing modulus trend would be seen in the 18 kDa and 52 kDa PLGA implants when preforming mechanical tests on them. After 8 days, an inconsistent fluctuation of implant modulus was observed. We attribute this to the fact that some of the samples became very fragile and easy to break at this stage, which made the implant harvesting and preparation extremely difficult and thus led to inconsistent testing results. The challenges we faced in the mechanical testing also reflect the drawback of traditional methods as they are destructive, which may lead to inaccurate testing results. In the in vitro study section, the erosion was found to be greater than 1 in Figures 5B and 7B, this is because of the residue NMP that was plasticized with PLGA that cannot be removed using Lyophilization. The morphology and strain distribution of PLGA ISFI implants at every designated time point can be clearly observed in Figure 4. On the basis of the ultrasound B-mode images, the size of the implant did not change over the course of the entire experiment. We attribute this to the empty voids, which support the shape of the implants. This was confirmed when we removed the later time point samples. We observed that the PLGA implants became attached to the inside of the phantom voids, causing them to maintain their shape and size while becoming hollow in the center. On the basis of the USE images, the stiffness of the implants was found to increase from day 0 to day 1 and decrease afterward. The early stiffening process was due to the liquid to solid precipitation process of the implant. After day 1, the stiffness change of the implant was governed by the erosion of the implant and therefore became softer overtime. The same trend was also observed in quantification (Figure 5A). The rate of strain increase was found to be dependent on the molecular weight of the implants which corresponds very well with the rate of implant erosion (Figure 5B). This indicates that the softening process is a result of erosion of the implants. Due to the different hydrophilicities associated with different molecular weights of polymer used, the implants erode at different rates through hydrolysis. After increasing in strain, all implant types

seemed to reach a plateau phase, at different rates. For example, 18 kDa implants reached plateau at about 14 days while 34 kDa and 52 kDa implants reached this phase at about 17 days and 21 days, respectively (Figure 5A). This plateau is believed to be where the implants have eroded to the point at which their mechanical stiffness has decreased below the sensitivity of our technique and thus changes in strain could no longer be detected. We confirmed this by scanning water under the same condition. The strain value of water was about 7% (the dashed line in Figure 5A), which equaled the plateau strain of all implant types. The correlation between strain and implant erosion was also analyzed in the linear region of strain and erosion data and plotted in Figure 8A. A strong linear strain− erosion relationship was observed, with R2 = 0.9148. This relationship was also found to be independent of polymer molecular weight. On the basis of this finding, change in implant integrity, and thus mechanical stiffness, appears to reflect the polymer erosion with a simple linear relationship. In the in vivo study section, the observations were very similar to what was observed in vitro. Implants of all types became softer overtime after injection. The rate of implant softening was again based on the different molecular weight of polymer used. 18 kDa had the highest softening rate followed by 34 kDa then 52 kDa (Figure 7A). This corresponds well with the implant erosion rate (Figure 7B). Unlike in the in vitro studies, the implants we injected subcutaneously into the rats’ abdominal regions became smaller in size overtime (Figure 6), and this change seemed to also be dependent upon the molecular weight. For example, the 18 kDa implants were almost gone whereas 34 kDa and 52 kDa implants were still about half of their original size at 20 days after injection (Figure 6). We hypothesize that the surrounding tissue was compressing the implant while the tissue mimicking phantom was not because the implant filling space was precreated. This injection site compressive force has also been reported to alter the drug release profile of the ISFIs.7 In addition, the strain values of the implants in vivo were smaller than in vitro. Furthermore, the plataeu phase oberved in the in vitro studies was not found in the in vivo studies. This was anticipated because the stiffness of the surrounding tissue environment mainly consists of muscle and internal organs,32 and is not as high as the tissue mimiking phantom (50 kPa) that we used. Thus, most of the strain we applied was absorbed by the compliant surrounding tissue, resulting in a lower strain from implants when compared to the in vitro study. The correlation between strain and erosion was also analyzed in the linear region of strain and erosion data and plotted in Figure 8. The F

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findings were very similar to what we observed in vitro; a strong linear relationship between strain and erosion was found, suggesting the possibility of using the mechanical stiffness of implants to predict implant erosion independent of molecular weight. Combining findings in both in vitro and in vivo experiments, a predictive model based on strain-erosion relationship was built. In order to test this model, an in vitro single blind prediction study was performed. See details in the Supporting Information. By comparing the erosion estimation using the predictive model and the actual measured erosion for 9 samples up to 15 days, an average percent error of 17.3% was found. The percent error for 5 days, 10 days and 15 days estimation was found to be 9.23, 16.59, and 26.14%, respectively. See details in Table 1. Biodegradable polymeric materials have been widely used for various biomedical applications. A nondestructive and high throughput imaging technique for the longitudinal assessment of biomaterial erosion will help researchers to refine their designs in biomedical devices with erosive properties. USE has shown high potential for clinical use, but limited applications in preclinic. By examining the relationship between polymer erosion and strain in this presented study, we were able to build a predicative model for longitudinal nondestructive characterization of polymer erosion. Its accuracy for erosion estimation was also tested in a single blind study, showing only 17.3% difference from direct measurements (see details in the Supporting Information). Although this model is material and condition specific, the simple mechanical stiffness and erosion relationship can be adapted to build other models for prediction in various situations. When combining this model with the use of USE integrated diagnostic ultrasound, it is now possible to track and even predict the fate of erosive polymeric materials in a nondestructive and noninvasive manner.

ACKNOWLEDGMENTS This work was supported by the National Institute of Biomedical Imaging and Bioengineering of the National Institutes of Health under award numbers R01EB016960 and T32-EB007509. Views and opinions of and endorsements by the author(s) do not reflect those of the National Institutes of Health.

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CONCLUSION The mechanical stiffness of in situ forming implants was monitored using ultrasound elastography imaging. The softening process of implants over time was found to be dependent on the PLGA molecular weight used, which corresponds well with the implant erosion rate. By analyzing the correlation between implant mechanical stiffness and erosion, we observed a strong linear relationship, which is independent of PLGA molecular weight. This finding was observed in both in vitro and in vivo studies, demonstrating the possibility of building a mechanical stiffness-based predictive model using USE for longitudinal assessment of implant erosion. ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsbiomaterials.6b00128. Single blinded erosion prediction study using polyacrylamide tissue mimicking phantom (PDF)

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REFERENCES

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AUTHOR INFORMATION

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*E-mail: [email protected]. Notes

The authors declare no competing financial interest. G

DOI: 10.1021/acsbiomaterials.6b00128 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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DOI: 10.1021/acsbiomaterials.6b00128 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX