Authors are grateful to C. Luzuriaga for the support in the porosimetry measurements. Authors also thanks to Ministerio de Econom&#237;a y Competitividad of Spain for financial support (Project MAT2014-59116-C2-2-R) and PIUNA (ref. 2018-15). The authors also would like to acknowledge Dr. Amir Maldonado from Departamento de F&#237;sica-UNISON for support and helpful comments and Dr. SE Burruel-Ibarra from DIPM-UNISON for SEM images and Rubio Pharma y Asociados S. A. de C. V. for financial support. ME Mart&#237;nez-Barbosa would like to thank CONACyT (M&#233;xico) projects No. 104931 and No. 256753, besides the financial support from Red Tem&#225;tica de Nanociencias y Nanotecnolog&#237;a del programa de Redes Tem&#225;ticas del CONACyT. And, also project USO316001081. MD Figueroa-Pizano would like to acknowledge CONACyT for financial support (scholarship 373321).

1. Preparation of chitosan-PVA hydrogels
<ol>
	<li>Prepare 2% (w/w) chitosan and 10% (w/w) PVA solutions. Dissolve 0.2 g of chitosan in 10 mL of 0.1 M CH3COOH solution (previously filtered) at room temperature and maintain continuous mechanical stirring overnight. Dissolve 1 g of PVA in 10 mL of distilled water and stir at 80 &#176;C for 1 h.</li>
	<li>Mix both solutions 1:1 using a magnetic stirrer until they are homogeneous at room temperature, and pour the mixtures on Petri dishes. Leave the samples for 2 h at atmospheric pressure to degas.</li>
	<li>Freeze the hydrogels at -4 &#176;C, -20 &#176;C or -80 &#176;C for 20 h and 4 cycles (samples CP4-4, CP4-20 and CP4-80, respectively). Freeze another hydrogel at -80 &#176;C for 20 h using 5 or 6 freezing cycles (samples CP5-80 and CP6-80). After the third freezing cycle, wash the hydrogels with deionized water. At the end, freeze-dry the hydrogels at -46 &#176;C for 48 h and store for further characterization (methodology adapted from2).</li>
</ol>
2. FT-IR characterization
<ol>
	<li>Place a little piece (1 mm x 2 mm) of hydrogel in the FT-IR spectrometer in ATR mode. Take the FT-IR spectra from 4000 to 600 cm-1 (2 cm-1 of resolution and average of 32 scans).</li>
</ol>
3. Swelling assays
<ol>
	<li>Cut out discs (13 mm in diameter and 10 mm in height) from the hydrogel and weigh them. Incubate the discs in 50 mL of deionized water with shaking at 25 &#176;C. Repeat three times.</li>
	<li>Every 30 min remove the sample from the medium, blotter to eliminate the excess of water, and weigh. Calculate the swelling degree using the equation 1 and calculate the equilibrium state of swelling, <img alt="Equation 1" src="/files/ftp_upload/59636/59636eq1.jpg" />, at 24 h using the equation 2. 
	<img alt="Equation 2" src="/files/ftp_upload/59636/59636eq2.jpg" />) 
	Where <img alt="Equation 3" src="/files/ftp_upload/59636/59636eq3.jpg" /> is the weight of the dry hydrogel and <img alt="Equation 4" src="/files/ftp_upload/59636/59636eq4.jpg" /> is the weight of the wet hydrogel. 
	<img alt="Equation 5" src="/files/ftp_upload/59636/59636eq5.jpg" /></li>
</ol>
4. Electronic Microscopy
<ol>
	<li>Cover a little piece of hydrogel with a thin gold layer (30 s and 10 mA) in a sputter coater.</li>
	<li>Put the sample in a scanning electron microscope (SEM). Analyze the samples under vacuum at 20 kV and take the images with a 500x and 1500x magnification.</li>
</ol>
5. Porosimetry
<ol>
	<li>Place discs 15 mm in diameter weighing around 0.26 g into the penetrometer (a solid penetrometer, having a bulk volume of 0.3660 mL and 5.7831 mL of stem volume). Analyze the porosity and pore size by Mercury Intrusion Porosimetry (MIP).</li>
	<li>Conduct the experiment in the hysteresis mode (intrusion-extrusion). Measure the total intrusion volume (mL/g), total pore area (m2/g), pore diameter (&#181;m), porosity (%), permeability (mDarcy) and tortuosity. Repeat twice.</li>
</ol>
6. Drug loading and release
<ol>
	<li>Before loading, prepare 4 L of 15 mg/L diflunisal solution and stir overnight. Confirm the concentration of the solution by UV-Vis spectroscopy (initial concentration). Indeed, swell 400 mg of freeze-dried samples of hydrogel in 6 mL of distilled water for 24 h.</li>
	<li>For loading, fill a flask with 50 mL of diflunisal solution and maintain at 25 &#176;C with constant stirring. Submerge each swelled hydrogel in the flask.
	<ol>
		<li>Take aliquots of remaining diflunisal solution (2 mL) at different times in order to determine the plateau region of the curve, for example: 3, 6, 24, 27, 30 and 48 h. After 24 h replace the solution with a fresh one.</li>
	</ol>
	</li>
	<li>Measure the absorbance at 252 nm of each aliquot, and determine the concentration of diflunisal present in the solution, using a calibration curve of diflunisal. Calculate the amount of diflunisal retained in the hydrogel at 24 and 48 h, as the difference of initial and final concentrations, taking into account the total volume (56 mL).
	<ol>
		<li>Determine the encapsulation efficiency (EE) using the equation 3. 
		<img alt="Equation 6" src="/files/ftp_upload/59636/59636eq6.jpg" /></li>
		<li>Freeze the loaded hydrogels at -80 &#176;C and lyophilize them at -50 &#176;C.</li>
	</ol>
	</li>
	<li>For drug release, submerge 300 mg of freeze-dried diflunisal loaded hydrogels in 50 mL of phosphate buffer (pH 7.4) at 25 &#176;C. Maintain constant stirring. Withdraw aliquots of 2 mL at different times and replace with fresh medium to keep a constant volume.
	<ol>
		<li>Determinate the diflunisal released spectrophotometrically at 252 nm, according to a calibration curve.</li>
	</ol>
	</li>
	<li>Deduce the predominant drug release mechanism in the hydrogels adjusting the drug release data corresponding to the first 60%, to the Korsmeyer-Peppas model (Equation 4), to obtain the kinetic (k) and the diffusion (n) constants. The n values indicate the mechanism of drug release24,25. Then, n values close to 0.5 are related to Fickian diffusion, meanwhile values of 0.5-1.0 for anomalous transport, where are involved diffusion and relaxation chains, and finally, values of 1.0 are related to case II transport. 
	<img alt="Equation 7" src="/files/ftp_upload/59636/59636eq7.jpg" />
	<ol>
		<li>To confirm the results, use the Higuchi, First order, and Zero order mathematical models (Equations 5 to 7) and select the better fit.</li>
		<li><img alt="Equation 8" src="/files/ftp_upload/59636/59636eq8.jpg" /> 
		<img alt="Equation 9" src="/files/ftp_upload/59636/59636eq9.jpg" /> 
		<img alt="Equation 10" src="/files/ftp_upload/59636/59636eq10.jpg" /> 
		where t represents the release time, Mt the amount of drug delivered at a given time, and M&#8734; the total amount of drug delivered at the end of the process.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

The freeze-thawing method is a suitable process to prepare biocompatible hydrogels focused in biomedical, pharmaceutical or cosmetical applications34,35,36. The most important advantage of this method, compared with other well-known methods to prepare hydrogels, is that crosslinking agent use is avoided, which could cause an inflammatory response or adverse effects in the human body34. This is a versatile method because it offers the possibility to prepare hydrogels from PVA or their mixtures with different polymers11,37 in such way that new characteristics from the other polymers can be obtained in the new material (e.g., major capacity to absorb water, antimicrobial or antioxidant properties2,18,35). However, it is important to consider that the incorporation of other polymers could decrease the strength of the hydrogels19,37.
The principal parameters to consider in the freeze-thawing method are the temperature of freezing, the time and the number of freezing cycles, and also, the polymer ratio (in case of polymer mixtures)2,19,20. A wide range of swelling, morphological and mechanical properties can be obtained with this method when the freeze conditions are controlled. These parameters affect directly the three-dimensional network configuration in chitosan-PVA hydrogels because the freezing conditions promotes the arrangements in PVA chains, which are joined by physical interactions, called crystalline zones12,38. These crystalline zones are concentrated regions of hydrogen bonds that act as crosslinking points in the hydrogels, which maintain and form the three-dimensional network and it is a retractor force when hydrogels are in the swelling state2,39,40.
In this study, we evaluated the effect of a new range of freeze-thawing temperatures (-4 &#176;C, -20 &#176;C and -80 &#176;C) combined with a different number of freezing cycles (4, 5 and 6) but the same time of freezing (20 h), to prepare 1:1 chitosan-PVA hydrogels. The lowest swelling capacity was observed at the lowest temperature (-80 &#176;C). Indeed, hydrogels at this lowest temperature obtained the smaller pores and the more porous networks. These differences in chitosan-PVA hydrogels are useful for different applications such as drug delivery systems or scaffolds. In general, chitosan-PVA hydrogels present high rates of swelling, due to chitosan hydrophilic groups (-NH2)41,42, and they are soft, flexible, easy to handle and resist the manipulation because of PVA characteristics. In that sense, the freeze-thawing method is easy, cheap and fast to produce chitosan-PVA hydrogels with different properties, avoiding toxic crosslinking.
Although the freeze-thawing is an easy and friendly method, it has some drawbacks. A complete homogenization of chitosan, in this case, and of the polymer mixtures is very important. Hydrogels can present more fragile zones and an irregular porous structure. Also, it is necessary to make a correct dissolution of PVA in water by heating at 70-80 &#176;C42,43 for 1 h under magnetic stirring. The cooling of this PVA solution must be slow with constant stirring to prevent the formation of a solid layer of PVA.
A limitation of this method, for cell culture assays, is the formation of whitish or semitransparent hydrogels. In this case, the application of glycerol or DMSO (toxic compound at room temperature) could be used to improve the appearance of hydrogel23,44. The freeze-drying step of the freeze-thawing method to prepare CS-PVA hydrogels is a critical step, because the hydrogels could present a constriction in the middle zone, which complicate the work and the characterization. To avoid this, the sample must be kept completely frozen before lyophilization. Concerning the drug loading and release studies, it is very important to ensure that there is no interference with the signals from the hydrogel components and the drug to be quantified.

Recently, hydrogels have attracted great interest in the biomedical field because they are three-dimensional networks with high water content and are soft and flexible, so they can mimic natural tissues easily1. Also, they do not dissolve in aqueous medium at physiological temperature and pH but present a large swelling2. Hydrogels can act as tissue engineering scaffolds, hygiene products, contact lenses, and wound dressings; because they can trap and release active compounds and drugs, they are used as drug delivery systems3. Depending on their application, hydrogels can be made from natural or synthetic polymers, or a combination of both, in order to obtain the best characteristics4.
The properties of hydrogels are a consequence of many physical and chemical factors. At the physical level, their structure and morphology depend on their porosity, pore size and pore distribution5. At the chemical and molecular level, the polymer type, the hydrophilic group content in the polymer chain, the crosslinking point type, and the cross-linking density are the factors that determine the swelling capacity and the mechanical properties6,7.
According to the type of crosslinking agent used to form the network, the hydrogels are classified as chemical hydrogels or physical hydrogels. Chemical hydrogels are joined by covalent interactions between their chains, which are formed through UV and gamma irradiation or using a crosslinking agent7,8. Chemical hydrogels usually are strong and resistant but, generally, the crosslinking agent is toxic to the cells and its removal is difficult, so its application is limited. On the other hand, physical hydrogels form by the connection of the polymer chains through non-covalent interactions, avoiding the use of crosslinking agents4,9. The main non-covalent interactions in the network are hydrophobic interactions, electrostatic forces, complementary and hydrogen bounds7.
Poly(vinyl alcohol) (PVA, Figure 1a) is a synthetic and water-soluble polymer with excellent mechanical performance and biocompatibility that can from crosslink agent-free hydrogels through the freeze-thawing method10,11. This polymer has the capacity to form concentrated zones of hydrogen bonds between -OH groups of their chains (crystalline zones) when they are freezing12. These crystalline zones act as crosslinking points in the network, and they are promoted by two events: the approaching of the polymer chains when the crystal water expands and the PVA conformational changes from isotactic to syndiotactic PVA during freeze13. Because of the freeze-drying, the water crystals are sublimated, leaving void spaces that are the pores in the hydrogel14. To obtain hydrogels with better properties, PVA can been easily combined with other polymers.
In that sense, chitosan constitutes an option as it is the only biopolymer from natural sources with positive charges. It is obtained by the deacetylation of chitin and it is composed of random combinations of &#946;-1,4 linked D-glucosamine (deacetylated unit) and N-acetyl-D-glucosamine (acetylated unit)15,16 (Figure 1b). Chitosan is biodegradable by human enzymes and it is biocompatible. Also, by its cationic nature, it can interact with the negative charge of the cell surface, and this property has been associated with its antimicrobial activity17. This polymer is easy to process; however, their mechanical properties are not sufficient and some materials have been added to form complexes with better characteristics.
Considering specific characteristics of chitosan and PVA, the successful manufacture of hydrogels has been reached by the freeze-thawing method2,18 to avoid the use of toxic crosslinking agents. In chitosan-PVA hydrogels, the crystalline zones of PVA are also formed, and chitosan chains are interpenetrated and form simple hydrogen bonds with -NH2 groups and -OH groups in PVA. The final chitosan-PVA hydrogel is mechanically stable, with high rates of swelling and low toxicity, and with antibacterial effect18. However, depending on the freezing conditions used in the preparation (temperature, time and number of cycles), the final characteristics may change. Some studies report that increasing the number of freezing cycles decreases the swelling degree and increases the tensile strength19,20. In order to strengthen the network, other agents such as gamma and UV radiation and chemical crosslinkers have been used additionally after the freeze-thawed preparation21,22,23. Hydrogels with a higher chitosan proportion have a more porous network and high swelling capacity but less strength and thermal stability. In this context, it is important to consider the preparation conditions to obtain suitable hydrogels for their target application.
The purpose of this work is to present in detail how the freezing conditions (temperature of freezing and number of cycles) affect the final characteristics of CS-PVA hydrogels. FT-IR spectra, morphological and porosity characteristics and swelling capacity were evaluated, as well as drug loading and release capacity. In the release studies, diflunisal (Figure 1c) was used as model drug, due to its size suitable to the hydrogel structure.

<table><tbody><tr><td>&lt;strong&gt;Materials:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Chitosan medium molecular weight</td><td>Sigma-Aldrich</td><td>448877</td><td>Mw determined by capillary viscometry (637,000 Da) and deacetylation degree of 70%</td></tr><tr><td>Diflunisal (2&#039;-4&#039;-difluoro-4-hydroxy-3-biphenyl-carboxylicacid)</td><td>Merck</td><td></td><td></td></tr><tr><td>Glacial acetic acid</td><td>Sigma-Aldrich</td><td>1005706</td><td></td></tr><tr><td>Poly(vinyl alcohol)</td><td>Sigma-Aldrich</td><td>341584</td><td>Mw 89,000-98,000, 99+% hydrolyzed</td></tr><tr><td>&lt;strong&gt;Equipment:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Cressington Sputter Coater 108 auto</td><td>TED PELLA INC</td><td></td><td></td></tr><tr><td>Cryodos Lyophilizator</td><td>Telstar</td><td></td><td></td></tr><tr><td>Falcon tubes</td><td>Thermo Fisher Company</td><td></td><td></td></tr><tr><td>FT-IR spectroscopy</td><td>Nicolet iS50</td><td>in ATR mode</td><td></td></tr><tr><td>Lyophilizator</td><td>LABCONCO</td><td></td><td></td></tr><tr><td>Micromeritics Autopore IV 9500</td><td>Micromeritics</td><td></td><td></td></tr><tr><td>Scanning electron microscope</td><td>Pemtron SS-300LV</td><td></td><td></td></tr><tr><td>UV-visible spectrophotometer</td><td>Agilent 8453</td><td></td><td></td></tr></tbody></table>

a freeze-thawing method to prepare chitosan-poly(vinyl alcohol) hydrogels without crosslinking agents and diflunisal release studies

Chitosan-poly(vinyl alcohol) hydrogels can be produced by the freeze-thawing method without using toxic crosslinking agents. The applications of these systems are limited by their characteristics (e.g., porosity, flexibility, swelling capacity, drug loading and drug release capacity), which depend on the freezing conditions and the kind and ratio of polymers. This protocol describes how to prepare hydrogels from chitosan and poly(vinyl alcohol) at 50/50 w/w % of polymer composition and varying the freezing temperature (-4 &#176;C, -20 &#176;C, -80 &#176;C) and freeze-thawing cycles (4, 5, 6 freezing cycles). FT-IR spectra, SEM micrograph and porosimetry data of hydrogels were obtained. Also, the swelling capacity and drug loading and release of diflunisal were assessed. Results from SEM micrographs and porosimetry show that the pore size decreases, while the porosity increases at lower temperatures. The swelling percentage was higher at the minor freezing temperature. The release of diflunisal from the hydrogels has been studied. All the networks maintain the drug release for 30 h and it has been observed that a simple diffusion mechanism regulates the diflunisal release according to Korsmeyer-Peppas and Higuchi models.

Hydrogels preparation 
Chitosan-PVA hydrogels were obtained at -4 &#176;C, -20 &#176;C and -80 &#176;C with 4 freezing cycles and at -80 &#176;C with 5 and 6 freezing cycles by the previously reported freeze-thawing method2. All hydrogels were homogeneous, semi-transparent, flexible and resistant against manipulation.
FT-IR characterization 
The FT-IR spectra are shown in Figure 2. Seven characteristics signals of chitosan and PVA polymers were detected: at 3286 cm-1 the stretching vibration mode of PVA hydroxyl group (-OH) and at 2918 cm-1 the stretching vibration mode of -CH group26,27. The signals of amide groups, representative of chitosan structure, were found at 1652 cm-1 to the stretching vibration mode of C=O (amide I), at 1560 cm-1 to the flection vibration mode of N-H (amide II) and 1325 cm-1 to the vibration of amide III28,29,30. Other signals, at 1418 cm-1 to the flection vibration mode of C-H and at 1086 cm-1 to the stretching vibration mode of C-O groups, both of PVA, were detected27,31,32.
Electronic Microscopy 
All CS-PVA hydrogels showed a highly porous surface (Figure 3, from left to right) and distinctive changes were observed according to the preparation conditions. Hydrogels prepared at -4 &#176;C (CP4-4) presented larger pores than the hydrogels prepared at -80 &#176;C (CP4-80). Moreover, the latter appears to have a more porous network. This effect may be due to the fact that, at lower temperature, the water crystal formation was faster and many small crystals emerged and were sublimated during the freeze-drying process, leaving void pores14,33. Meanwhile, the effect of the number of freezing cycles seems to promote more defined and circular pores in hydrogels CP6-80 (Figure 3, from top to bottom).
Porosimetry 
Samples CP4-4, CP4-80 and CP6-80 presented more pronounced changes; in order to complement the information about morphology, they were analyzed by MIP (Table 1). The comparison between hydrogels CP4-4 and CP4-80 (Figure 3a) showed that, at a lower temperature of freezing, hydrogels developed a more porous network, which presented a large total intrusion volume and higher total pore area. However, hydrogels CP6-80 showed less permeability than CP4-80 (Figure 3b), probably due to their high tortuosity, which was also reflected in a lower total intrusion volume. Figure 3 presents the different pore sizes of these hydrogels. Two pore sizes were distinguished, one between 0.3-5.0 &#956;m and other between 5.0-30 &#956;m. In hydrogels CP4-80 and CP6-80, the porous network had a greater number of small pores than large ones, compared with CP4-4 hydrogel. These results were similar to those observed by SEM micrographs and suggested that, at lower temperature greater interactions between the PVA chains were favored and more crystalline zones were formed. In such a way, the formation of crystalline zones by PVA chains, was stimulated at low temperature.
Swelling assays 
The swelling behavior of CS-PVA hydrogels can be seen in the Figure 4. They quickly absorbed large amounts of water; for the first 5 hours they retained 10x their weight, and after 20 hours they retain up to 15x their weight (equilibrium point). However, in relation to hydrogels prepared at the same number of freezing cycles, the hydrogel CP4-80 showed less swelling capacity in the first 5 hours as a consequence of the temperature that was used for its preparation (-80 &#176;C). In the case of hydrogels prepared at different number of freezing cycles (CP4-80, CP5-80 and CP6-80) no differences were found at any time. Probably, the decreased swelling capacity observed in hydrogels prepared at -80 &#176;C was caused by the small pore size of the hydrogel network.
Drug loading and release 
To evaluate the capacity of CS-PVA hydrogels as drug delivery systems, the anti-inflammatory drug diflunisal was loaded in the network and subsequently released. The encapsulation efficiency (EE) in all these systems was around 70%; however, the CP4-80 hydrogel presented more slightly EE at 73% (Table 2). Meanwhile, the releasing kinetics of diflunisal from the CS-PVA hydrogels were maintained for about 30 h in all cases. The CP4-80 hydrogel released the highest amount of diflunisal (Figure 5). This may be due to the fact this hydrogel showed a more porous structure in comparison with the other two types of hydrogel. This feature allowed the small molecule of drug to easily enter in the hydrogel network and, then, to be released. Between CP4-80 and CP6-80 hydrogels not differences were observed during release times (Figure 6). No burst effect was observed in any of the CS-PVA hydrogels, which is promising for pharmaceutical applications. Mathematical models were used to determine the main release mechanism in CS-PVA hydrogels. The results were adjusted to different mathematical models (Table 3) and according to the n values, it was found that the Fick diffusion dominates the drug release process.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59636/59636fig1v2.jpg" /> 
Figure 1: Chemical structure of PVA (a), chitosan (b) and diflunisal (c). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59636/59636fig1largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59636/59636fig2.jpg" /> 
Figure 2: FT-IR spectra of pure chitosan and PVA and, chitosan-PVA hydrogels prepared at different conditions of freezing. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59636/59636fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59636/59636fig3.jpg" /> 
Figure 3: SEM micrographs of chitosan-PVA hydrogels at 1500x magnification. Pore size distributions of chitosan-PVA hydrogels: a) hydrogels prepared with 4 cycles of freezing and at -4 &#176;C and -80 &#176;C. b) Hydrogels prepared at -80 &#176;C and, 4 and 6 cycles. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59636/59636fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59636/59636fig4.jpg" /> 
Figure 4: Swelling kinetics of chitosan-PVA hydrogels: a) hydrogels with 4 cycles of freezing and b) hydrogels prepared at -80 &#176;C. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59636/59636fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59636/59636fig5.jpg" /> 
Figure 5: Diflunisal release profiles in mg (a) and Mt/<img alt="Equation 11" src="/files/ftp_upload/59636/59636eq11.jpg" /> (b) for hydrogels CP4-4 and CP4-80. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59636/59636fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59636/59636fig6.jpg" /> 
Figure 6: Diflunisal release profiles in mg (a) and Mt/<img alt="Equation 11" src="/files/ftp_upload/59636/59636eq11.jpg" /> (b) for hydrogels CP4-80 and CP6-80. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59636/59636fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Hydrogel</td>
			<td>Total intrusion volume (mL/g)</td>
			<td>Total pore area (m2/g)</td>
			<td>Porosity (%)</td>
			<td>Permeability (mdarcy)</td>
			<td>Tortuosity</td>
		</tr>
		<tr>
			<td>CP4-4</td>
			<td>5.16</td>
			<td>10.19</td>
			<td>67.13</td>
			<td>132.43</td>
			<td>10.46</td>
		</tr>
		<tr>
			<td>CP4-80</td>
			<td>7.36</td>
			<td>15.14</td>
			<td>85.95</td>
			<td>151.16</td>
			<td>5.83</td>
		</tr>
		<tr>
			<td>CP6-80</td>
			<td>6.69</td>
			<td>12.86</td>
			<td>84.82</td>
			<td>129.28</td>
			<td>12.2</td>
		</tr>
	</tbody>
</table>
Table 1: Porosimetry parameters of the porous structure of chitosan-PVA hydrogels.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample</td>
			<td colspan="2">Diflunisal loaded</td>
			<td>Diflunisal released</td>
		</tr>
		<tr>
			<td></td>
			<td>mg/g hydrogel</td>
			<td>Encapsulation efficiency (%)</td>
			<td>% released respect to loaded</td>
		</tr>
		<tr>
			<td>CP4-4</td>
			<td>3.05&#177; 0.09</td>
			<td>71</td>
			<td>79 &#177; 3.33</td>
		</tr>
		<tr>
			<td>CP4-80</td>
			<td>3.22 &#177; 0.47</td>
			<td>73</td>
			<td>86 &#177; 0.4</td>
		</tr>
		<tr>
			<td>CP6-80</td>
			<td>3.19 &#177; 0.05</td>
			<td>68</td>
			<td>80 &#177; 3.9</td>
		</tr>
	</tbody>
</table>
Table 2: Encapsulation and release efficiencies for chitosan-PVA hydrogels.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample</td>
			<td colspan="3">Korsmeyer-Peppas</td>
			<td colspan="2">Higuchi</td>
			<td colspan="2">First Order</td>
			<td colspan="2">Zero Order</td>
		</tr>
		<tr>
			<td rowspan="2"></td>
			<td>kKP x 102</td>
			<td rowspan="2">n</td>
			<td rowspan="2">R2</td>
			<td>kH x 102</td>
			<td rowspan="2">R2</td>
			<td>k1 x 102</td>
			<td rowspan="2">R2</td>
			<td>k0 x 102</td>
			<td rowspan="2">R2</td>
		</tr>
		<tr>
			<td>(min-n)</td>
			<td>(min-0.5)</td>
			<td>(min-1)</td>
			<td>(min-1)</td>
		</tr>
		<tr>
			<td>CP4-4</td>
			<td>4.3 &#177; 0.39</td>
			<td>0.44 &#177; 0.02</td>
			<td>0.99</td>
			<td>3.1 &#177; 0.1</td>
			<td>0.98</td>
			<td>0.29 &#177; 0.03</td>
			<td>0.803</td>
			<td>0.18 &#177; 0.02</td>
			<td>0.54</td>
		</tr>
		<tr>
			<td>CP4-80</td>
			<td>3.6 &#177; 0.33</td>
			<td>0.50 &#177; 0.02</td>
			<td>0.99</td>
			<td>3.7 &#177; 0.1</td>
			<td>0.99</td>
			<td>0.42 &#177; 0.03</td>
			<td>0.894</td>
			<td>0.27 &#177; 0.02</td>
			<td>0.7</td>
		</tr>
		<tr>
			<td>CP6-80</td>
			<td>2.3 &#177; 0.24</td>
			<td>0.54 &#177; 0.02</td>
			<td>0.99</td>
			<td>2.9 &#177; 0.1</td>
			<td>0.99</td>
			<td>0.27 &#177; 0.02</td>
			<td>0.925</td>
			<td>0.17 &#177; 0.01</td>
			<td>0.77</td>
		</tr>
		<tr>
			<td colspan="4">k= kinetic constant; n= diffusion constant.</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 3: Kinetic parameters of diflunisal release from chitosan-PVA hydrogels.

Watch this Scientific Journal Video about A Freeze-Thawing Method to Prepare Chitosan-Polyvinyl alcohol Hydrogels Without Crosslinking Agents and Diflunisal Release Studies at JoVE.com

A Freeze-Thawing Method to Prepare Chitosan-Polyvinyl alcohol Hydrogels Without Crosslinking Agents and Diflunisal Release Studies

The freezing-thawing method is used to produce chitosan-poly(vinyl alcohol) hydrogels without crosslinking agents. For this method, it is important to consider the freezing conditions (temperature, number of cycles) and polymer ratio, which can affect the properties and applications of the obtained hydrogels.

A Freeze-Thawing Method to Prepare Chitosan-Poly(vinyl alcohol) Hydrogels Without Crosslinking Agents and Diflunisal Release Studies

Chitosan-poly(vinyl alcohol) hydrogels can be produced by the freeze-thawing method without using toxic crosslinking agents. The applications of these ...

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Nanyang Technological University

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Bioengineering

17.2K Views.  University of Sonora. The freezing-thawing method is used to produce chitosan-poly(vinyl alcohol) hydrogels without crosslinking agents. For this method, it is important to consider the freezing conditions (temperature, number of cycles) and polymer ratio, which can affect the properties and applications of the obtained hydrogels.

Video: A Freeze-Thawing Method to Prepare Chitosan-Polyvinyl alcohol Hydrogels Without Crosslinking Agents and Diflunisal Release Studies

Preparation and Characterization of SDF-1&#945;-Chitosan-Dextran Sulfate Nanoparticles

Chitosan (CS) and dextran sulfate (DS) are charged polysaccharides (glycans), which form polyelectrolyte complex-based nanoparticles when mixed under appropriate conditions. The glycan nanoparticles are useful carriers for protein factors, which facilitate the in vivo delivery of the proteins and sustain their retention in the targeted tissue. The glycan polyelectrolyte complexes are also ideal for protein delivery, as the incorporation is carried out in aqueous solution, which reduces the likelihood of inactivation of the proteins. Proteins with a heparin-binding site adhere to dextran sulfate readily, and are, in turn, stabilized by the binding. These particles are also less inflammatory and toxic when delivered in vivo. In the protocol described below, SDF-1&#945; (Stromal cell-derived factor-1&#945;), a stem cell homing factor, is first mixed and incubated with dextran sulfate. Chitosan is added to the mixture to form polyelectrolyte complexes, followed by zinc sulfate to stabilize the complexes with zinc bridges. The resultant SDF-1&#945;-DS-CS particles are measured for size (diameter) and surface charge (zeta potential). The amount of the incorporated SDF-1&#945; is determined, followed by measurements of its in vitro release rate and its chemotactic activity in a particle-bound form.

The objective of this protocol is to incorporate SDF-1&#945;, a stem cell homing factor, into dextran sulfate-chitosan nanoparticles. The resultant particles are measured for their size and zeta potential, as well as the content, activity, and in vitro release rate of SDF-1&#945; from the nanoparticles.

Chitosan (CS) and dextran sulfate (DS) are charged polysaccharides (glycans), which form polyelectrolyte complex-based nanoparticles when mixed under ...

Chemistry

Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field of 3D templates is rising due to the potential resemblance of those materials to the natural extracellular matrix. Protein-based hydrogels are particularly promising because they can easily be functionalized and can achieve defined structures with adjustable physicochemical properties. However, the production of macroporous 3D templates for cell culture applications using natural materials is often limited by their weaker mechanical properties compared to those of synthetic materials. Here, different methods were evaluated to produce macroporous bovine serum albumin (BSA)-based hydrogel systems, with adjustable pore sizes in the range of 10 to 70 &#181;m in radius. Furthermore, a method to generate channels in this protein-based material that are several hundred microns long was established. The different methods to produce pores, as well as the influence of pore size on material properties such as swelling ratio, pH, temperature stability, and enzymatic degradation behavior, were analyzed. Pore sizes were investigated in the native, swollen state of the hydrogels using confocal laser scanning microscopy. The feasibility for cell culture applications was evaluated using a cell-adhesive RGD peptide modification of the protein system and two model cell lines: human breast cancer cells (A549) and adenocarcinomic human alveolar basal epithelial cells (MCF7).

Different methods to manipulate three-dimensional architecture in protein-based hydrogels are evaluated here with respect to material properties. The macroporous networks are functionalized with a cell-adhesive peptide, and their feasibility in cell culture is evaluated using two different model cell lines.

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field ...

Biochemistry

Capillary Electrophoresis to Monitor Peptide Grafting onto Chitosan Films in Real Time

Free-solution capillary electrophoresis (CE) separates analytes, generally charged compounds in solution through the application of an electric field. Compared to other analytical separation techniques, such as chromatography, CE is cheap, robust and effectively requires no sample preparation (for a number of complex natural matrices or polymeric samples). CE is fast and can be used to follow the evolution of mixtures in real time (e.g., chemical reaction kinetics), as the signals observed for the separated compounds are directly proportional to their quantity in solution.
Here, the efficiency of CE is demonstrated for monitoring the covalent grafting of peptides onto chitosan films for subsequent biomedical applications. Chitosan&#39;s antimicrobial and biocompatible properties make it an attractive material for biomedical applications such as cell growth substrates. Covalently grafting the peptide RGDS (arginine - glycine - aspartic acid - serine) onto the surface of chitosan films aims at improving cell attachment. Historically, chromatography and amino acid analysis have been used to provide a direct measurement of the amount of grafted peptide. However, the fast separation and absence of sample preparation provided by CE enables equally accurate yet real-time monitoring of the peptide grafting process. CE is able to separate and quantify the different components of the reaction mixture: the (non-grafted) peptide and the chemical coupling agents. In this way the use of CE results in improved films for downstream applications.
The chitosan films were characterized through solid-state NMR (nuclear magnetic resonance) spectroscopy. This technique is more time-consuming and cannot be applied in real time, but yields a direct measurement of the peptide and thus validates the CE technique.

Free solution capillary electrophoresis is a fast, cheap and robust analytical method that enables the quantitative monitoring of chemical reactions in real time. Its utility for rapid, convenient and precise analysis is demonstrated here through analysis of covalent peptide grafting onto chitosan films for improved cell adhesion.

Free-solution capillary electrophoresis (CE) separates analytes, generally charged compounds in solution through the application of an electric field. ...

Preparation of Chitosan-based Injectable Hydrogels and Its Application in 3D Cell Culture

The protocol presents a facile, efficient, and versatile method to prepare chitosan-based hydrogels using dynamic imine chemistry. The hydrogel is prepared by mixing solutions of glycol chitosan with a synthesized benzaldehyde terminated polymer gelator, and hydrogels are efficiently obtained in several minutes at room temperature. By varying ratios between glycol chitosan, polymer gelator, and water contents, versatile hydrogels with different gelation times and stiffness are obtained. When damaged, the hydrogel can recover its appearances and modulus, due to the reversibility of the dynamic imine bonds as crosslinkages. This self-healable property enables the hydrogel to be injectable since it can be self-healed from squeezed pieces to an integral bulk hydrogel after the injection process. The hydrogel is also multi-responsive to many bio-active stimuli due to different equilibration statuses of the dynamic imine bonds. This hydrogel was confirmed as bio-compatible, and L929 mouse fibroblast cells were embedded following standard procedures and the cell proliferation was easily assessed by a 3D cell cultivation process. The hydrogel can offer an adjustable platform for different research where a physiological mimic of a 3D environment for cells is profited. Along with its multi-responsive, self-healable, and injectable properties, the hydrogels can potentially be applied as multiple carriers for drugs and cells in future bio-medical applications.

Here we describe a facile preparation of chitosan-based injectable hydrogels using dynamic imine chemistry. Methods to adjust the hydrogel&#8217;s mechanical strength and its application in 3D cell culture are presented.

The protocol presents a facile, efficient, and versatile method to prepare chitosan-based hydrogels using dynamic imine chemistry. The hydrogel is ...

Composite Scaffolds of Interfacial Polyelectrolyte Fibers for Temporally Controlled Release of Biomolecules

Various scaffolds used in tissue engineering require a controlled biochemical environment to mimic the physiological cell niche. Interfacial polyelectrolyte complexation (IPC) fibers can be used for controlled delivery of various biological agents such as small molecule drugs, cells, proteins and growth factors. The simplicity of the methodology in making IPC fibers gives flexibility in its application for controlled biomolecule delivery. Here, we describe a method of incorporating IPC fibers into two different polymeric scaffolds, hydrophilic polysaccharide and hydrophobic polycaprolactone, to create a multi-component composite scaffold. We showed that IPC fibers can be easily embedded into these polymeric structures, enhancing the capability for sustained release and improved preservation of biomolecules. We also created a composite polymeric scaffold with topographical cues and sustained biochemical release that can have synergistic effects on cell behavior. Composite polymeric scaffolds with IPC fibers represent a novel and simple method of recreating the cellular niche.

Scaffolds for tissue engineering need to recapitulate the complex biochemical and biophysical microenvironment of the cellular niche. Here, we show the use of interfacial polyelectrolyte complexation fibers as a platform to create composite, multi-component polymeric scaffolds with sustained biochemical release.

Various scaffolds used in tissue engineering require a controlled biochemical environment to mimic the physiological cell niche. Interfacial ...

Synthesis of Thermogelling Poly(N-isopropylacrylamide)-graft-chondroitin Sulfate Composites with Alginate Microparticles for Tissue Engineering

Injectable biomaterials are defined as implantable materials that can be introduced into the body as a liquid and solidify in situ. Such materials offer the clinical advantages of being implanted minimally invasively and easily forming space-filling solids in irregularly shaped defects. Injectable biomaterials have been widely investigated as scaffolds for tissue engineering. However, for the repair of certain load-bearing areas in the body, such as the intervertebral disc, scaffolds should possess adhesive properties. This will minimize the risk of dislocation during motion and ensure intimate contact with the surrounding tissue, providing adequate transmission of forces. Here, we describe the preparation and characterization of a scaffold composed of thermally sensitive poly(N-isopropylacrylamide)-graft-chondroitin sulfate (PNIPAAM-g-CS) and alginate microparticles. The PNIPAAm-g-CS copolymer forms a viscous solution in water at RT, into which alginate particles are suspended to enhance adhesion. Above the lower critical solution temperature (LCST), around 30 &#176;C, the copolymer forms a solid gel around the microparticles. We have adapted standard biomaterials characterization procedures to take into account the reversible phase transition of PNIPAAm-g-CS. Results indicate that the incorporation of 50 or 75 mg/ml&#160;alginate particles into 5% (w/v) PNIPAAm-g-CS solutions quadruple the adhesive tensile strength of PNIPAAm-gCS alone (p&#60;0.05). The incorporation of alginate microparticles also significantly increases swelling capacity of PNIPAAm-g-CS (p&#60;0.05), helping to maintain a space-filling gel within tissue defects. Finally, results of the in vitro toxicology assay kit, 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide (XTT) and Live/Dead viability assay indicate that the adhesive is capable of supporting the survival and proliferation of encapsulated Human Embryonic Kidney (HEK) 293 cells over 5 days.

Synthesis of Thermogelling PolyN-isopropylacrylamide-graft-chondroitin Sulfate Composites with Alginate Microparticles for Tissue Engineering

An injectable tissue engineering scaffold composed of poly(N-isopropylacrylamide)-graft-chondroitin sulfate (PNIPAAm-g-CS)-containing alginate microparticles was prepared. The adhesive strength, swelling properties and in vitro biocompatibility are analyzed in this study. The characterization techniques developed here may be applicable to other thermogelling systems.

Injectable biomaterials are defined as implantable materials that can be introduced into the body as a liquid and solidify in situ. Such materials offer ...

Preparation of Expanded Chitin Foams and their Use in the Removal of Aqueous Copper

Chitin is an underexploited, naturally abundant, mechanically robust, and chemically resistant biopolymer. These qualities are desirable in an adsorbent, but chitin lacks the necessary specific surface area, and its modification involves specialized techniques and equipment. Herein is described a novel chemical procedure for expanding chitin flakes, derived from shrimp shell waste, into foams with higher surface area. The process relies on the evolution of H2 gas from the reaction of water with NaH trapped in a chitin gel. The preparation method requires no specialized equipment. Powder X-ray diffraction and N2-physisorption indicate that the crystallite size decreases from 6.6 nm to 4.4 nm and the specific surface area increases from 12.6 &#177; 2.1 m2/g to 73.9 &#177; 0.2 m2/g. However, infrared spectroscopy and thermogravimetric analysis indicate that the process does not change the chemical identity of the chitin. The specific Cu adsorption capacity of the expanded chitin increases in proportion to specific surface area from 13.8 &#177; 2.9 mg/g to 73.1 &#177; 2.0 mg/g. However, the Cu adsorption capacity as a surface density remains relatively constant at an average of 10.1 &#177; 0.8 atom/nm2, which again suggests no change in the chemical identity of the chitin. This method offers the means to transform chitin into a higher surface area material without sacrificing its desirable properties. Although the chitin foam is described here as an adsorbent, it can be envisioned as a catalyst support, thermal insulator, and structural material.

This study describes a method to expand chitin into a foam by chemical techniques that require no specialized equipment.

Chitin is an underexploited, naturally abundant, mechanically robust, and chemically resistant biopolymer. These qualities are desirable in an adsorbent, ...

Magnetic and Thermal-sensitive Poly(N-isopropylacrylamide)-based Microgels for Magnetically Triggered Controlled Release

Magnetically and thermally sensitive poly(N-isopropylacrylamide) (PNIPAAm)/Fe3O4-NH2 microgels with the encapsulated anti-cancer drug curcumin (Cur) were designed and fabricated for magnetically triggered release. PNIPAAm-based magnetic microgels with a spherical structure were produced via a temperature-induced emulsion followed with physical-crosslinking by mixing PNIPAAm, polyethylenimine (PEI), and Fe3O4-NH2 magnetic nanoparticles. Because of their dispersity, the Fe3O4-NH2 nanoparticles were embedded inside the polymer matrix. The amine groups exposed on the Fe3O4-NH2 and PEI surface supported the spherical structure by physically crosslinking with the amide groups of the PNIPAAm. The hydrophobic anti-cancer drug curcumin can be dispersed in water after encapsulation into the microgels. The microgels were characterized by transmission electron microscopy (TEM), Fourier transform infrared spectroscopy (FT-IR), and UV-Vis spectral analysis. Furthermore, magnetically triggered release was studied under an external high frequency magnetic field (HFMF). A significant &#34;burst release&#34; of curcumin was observed after applying the HFMF to the microgels due to the magnetic inductive heating (hyperthermia) effect. This manuscript describes the magnetically triggered controlled release of Cur-PNIPAAm/Fe3O4-NH2 encapsulated curcumin, which can be potentially applied for tumor therapy.

Magnetic and Thermal-sensitive PolyN-isopropylacrylamide-based Microgels for Magnetically Triggered Controlled Release

This manuscript describes the preparation of magnetic and thermal-sensitive microgels via a temperature-induced emulsion without chemical reaction. These sensitive microgels were synthesized by mixing poly(N-isopropylacrylamide) (PNIPAAm), polyethylenimine (PEI) and Fe3O4-NH2 nanoparticles for the potential use in magnetically and thermally triggered drug-release.

Magnetically and thermally sensitive poly(N-isopropylacrylamide) (PNIPAAm)/Fe3O4-NH2 microgels with the encapsulated anti-cancer drug curcumin (Cur) were ...

Fusidic Acid-Loaded Hydrogel Films with Aloe vera for Enhanced Drug Delivery

Development and Characterization of Fusidic Acid-Loaded Alginate-Aloe vera Based Hydrogel Film

A new fusidic acid-loaded hydrogel film was prepared via the solvent casting technique using alginate and Aloe vera. The hydrogel films were optimized using different ratios of sodium alginate, Aloe vera, and glycerin. The films containing 10% glycerin (w/w of alginate) exhibited the best appearance. Incorporating Aloe vera influenced the thickness, swelling behavior, water vapor permeability, and drug release profile of the hydrogel films. Higher Aloe vera content resulted in thicker films (up to a certain ratio), increased swelling, reduced water vapor permeability, and a prolonged drug release of up to 93% over 12 h. Fourier-transform infrared spectroscopy (FTIR) analysis confirmed the presence of key functional groups and the interaction between the hydrogel components. The study suggests that the combination of sodium alginate, Aloe vera, and glycerin can improve the mechanical properties and drug release profiles of hydrogel films, making them a promising option for enhanced topical drug delivery and wound healing applications.

Development and Characterization of Fusidic Acid-Loaded Alginate-Aloe vera Based Hydrogel FilmWound Healing

A formulation of fusidic acid-loaded hydrogel film was developed with different Aloe vera ratios and characterized in this study.

A new fusidic acid-loaded hydrogel film was prepared via the solvent casting technique using alginate and Aloe vera. The hydrogel films were optimized ...

Formulations for Freeze-drying of Bacteria and Their Influence on Cell Survival

Cellular water can be removed to reversibly inactivate microorganisms to facilitate storage. One such method of removal is freeze-drying, which is considered a gentle dehydration method. To facilitate cell survival during drying, the cells are often formulated beforehand. The formulation forms a matrix that embeds the cells and protects them from various harmful stresses imposed on the cells during freezing and drying. We present here a general method to evaluate the survival rate of cells after freeze-drying and we illustrate it by comparing the results obtained with four different formulations: the disaccharide sucrose, the sucrose derived polymer Ficoll PM400, and the respective polysaccharides hydroxyethyl cellulose (HEC) and hydroxypropyl methyl cellulose (HPMC), on two strains of bacteria, P. putida KT2440 and A. chlorophenolicus A6. In this work we illustrate how to prepare formulations for freeze-drying and how to investigate the mechanisms of cell survival after rehydration by characterizing the formulation using of differential scanning calorimetry (DSC), surface tension measurements, X-ray analysis, and electron microscopy and relating those data to survival rates. The polymers were chosen to get a monomeric structure of the respective polysaccharide resembling sucrose to a varying degrees. Using this method setup we showed that polymers can support cell survival as effectively as disaccharides if certain physical properties of the formulation are controlled1.

Freeze-drying is often an easy and convenient way to obtain dry products of viable bacterial cells. An issue of the process is cell survival. We detail here a procedure to investigate how cell survival during freeze-drying is influenced by the properties of the formulation used.