The effect of freeze-drying on mucoadhesion and transport of acrylated chitosan nanoparticles

Journal Pre-proofs The Effect of Freeze-Drying on Mucoadhesion and Transport of Acrylated Chitosan Nanoparticles Shaked Eliyahu, Andreia Almeida, Maria Helena Macedo, José das Neves, Bruno Sarmento, Havazelet Bianco-Peled PII: DOI: Reference:

S0378-5173(19)30784-7 https://doi.org/10.1016/j.ijpharm.2019.118739 IJP 118739

To appear in:

International Journal of Pharmaceutics

Received Date: Revised Date: Accepted Date:

30 July 2019 22 September 2019 25 September 2019

Please cite this article as: S. Eliyahu, A. Almeida, M. Helena Macedo, J. das Neves, B. Sarmento, H. BiancoPeled, The Effect of Freeze-Drying on Mucoadhesion and Transport of Acrylated Chitosan Nanoparticles, International Journal of Pharmaceutics (2019), doi: https://doi.org/10.1016/j.ijpharm.2019.118739

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The Effect of Freeze-Drying on Mucoadhesion and Transport of Acrylated Chitosan Nanoparticles Shaked Eliyahu1, Andreia Almeida2,3,4, Maria Helena Macedo2,3,4, José das Neves2,3, Bruno Sarmento2,3,5, Havazelet Bianco-Peled1,6 * *Correspondence: [email protected]; Tel.: +972-4-829-3588 1 - The Russell Berrie Nanotechnology Institute, Technion-Israel Institute of Technology, Haifa, Israel 2 - i3S - Instituto de Investigação e Inovação em Saúde, Universidade do Porto, Porto, Portugal 3 - INEB – Instituto de Engenharia Biomédica, Universidade do Porto, Porto, Portugal 4 - ICBAS – Instituto Ciências Biomédicas Abel Salazar, Universidade do Porto, Porto, Portugal 5 - CESPU, Instituto de Investigação e Formação Avançada em Ciências e Tecnologias da Saúde, Gandra, Portugal 6 - Department of Chemical Engineering, Technion-Israel Institute of Technology, Haifa, Israel

Abstract Nanoparticle-based mucosal drug delivery is a promising method to increase the residence time of a drug in the mucosa. It is known that the stability of polysaccharidebased nanoparticles in aqueous solutions is limited, due to hydrolysis; hence the longterm stability of a formulation is usually improved by freeze-drying. The aim of this study was to investigate the effect of cryoprotection and freeze-drying on the physical and chemical properties of mucoadhesive acrylated chitosan (ACS) nanoparticles including the potential of these carriers to deliver drugs. The results showed that the most effective cryoprotection was achieved using sucrose. The incorporation of a hydrophilic macromolecular drug, dextran sulfate, increased the nanoparticle size and decreased the zeta potential for both fresh and freeze-dried nanoparticle formulations. In addition, the

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freeze-dried nanoparticles presented penetration across a mucus gel layer and the flow through technique revealed that short term mucoadhesive properties were not impaired. ACS nanoparticles were able to deliver a model drug across a mucin gel layer but could not improve drug penetration through the triple co-culture cell model that was used in order to mimic the small intestine epithelium. Keywords Acrylated chitosan; nanoparticles; mucoadhesive polymers; triple co-culture cell model; freeze-drying; cryoprotection.

1. Introduction Interactions with the mucosa are of prime importance in determining the fate and performance of nanosystems for mucosal drug delivery. Mucoadhesive nanocarriers may present desired features, such as increased residence time and intimate contact with the epithelium, which can ultimately contribute to enhanced drug bioavailability at both local and systemic levels (Sosnik and Sarmento, 2014). In an effort to increase the interactions and specificity towards the mucosa, second-generation mucoadhesive polymers were created (Andrews et al., 2009). This class of polymers possess side groups that can bind covalently to mucin glycoproteins covering the mucosal epithelium (Brannigan and Khutoryanskiy, 2019). The first sub-class of second-generation polymers that was described in the literature is thiolated polymers, termed thiomers (Bernkop-schnu, 2005). Thiolation was established as a method of improving the mucoadhesion based on disulfide bond formation with mucin glycoproteins (Albrecht et al., 2006; Dünnhaupt et al., 2015; Schattling et al., 2017). Another example for this type of polymers is polymers carrying acrylate end groups, that react with thiol residues of mucin glycoproteins via the Michael type addition reaction (Brannigan and Khutoryanskiy, 2019; Davidovich-Pinhas and Bianco-Peled, 2010; Schattling et al., 2017).

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Recently, we investigated a new nanoparticulate system based on acrylated chitosan (ACS) (Eliyahu et al., 2018), which was synthesized by grafting poly(ethylene glycol) diacrylate (PEGDA) chains on chitosan (CS) backbone (Shitrit and Bianco-Peled, 2017). Using different methods, we demonstrated that ACS nanoparticles crosslinked with tripolyphosphate present increased mucoadhesion compared to non-modified CS nanoparticles (Eliyahu et al., 2018). The improved mucoadhesion was attributed to the free acrylate end groups that were present in the nanoparticles surface, as well as entanglements and hydrogen bonds between PEGDA and mucin glycoproteins. It was suggested that this new type of nanoparticles can be useful for prolonging the residence time of associated drugs in the gastrointestinal tract, due to their mucoadhesiveness (Eliyahu et al., 2018). Aqueous suspensions of chitosan-based nanoparticulate systems display limited longterm stability due to hydrolysis (Almalik et al., 2017), which might hamper future clinical applications. Moreover, the ionic gelation process tends to produce nanoparticles with the ability to aggregate and swell, which is usually an undesirable phenomenon in the field of drug delivery. The addition of monovalent salts such as sodium chloride can help to produce narrow sized distributions (Jonassen et al., 2012b), however, the longterm physical stability is still insufficient. To reduce this chemical and physical instability, the freeze-drying technique can be performed (Abdelwahed et al., 2006). Freeze-drying is an industrial process which allows the removal of water by sublimation of frozen samples but can also generates stress causing aggregation and irreversible fusion. A cryoprotectant must be used in order to prevent this aggregation, as well as to increase the stability during storage. Sugarbased cryoprotectants such as sucrose, glucose and mannitol are commonly used to prevent nanoparticle aggregation and to protect them against the mechanical stress of ice crystal

formation

during freeze-drying

(Fonte

et

al.,

2016).

Polymer-based

3

cryoprotectants, including poly(ethylene glycol) (PEG), poly(vinyl pyrrolidone) and poly(vinyl alcohol) can also be useful (Abdelwahed et al., 2006). The overall goal of this study was to advance the understanding of ACS nanoparticles beyond fabrication methods. Our first specific aim was to investigate the effect of freezedrying on the physical and chemical properties of ACS nanoparticles. This was done using the flow through technique, which was designed to assess the mucoadhesion of liquid formulations (Khutoryanskiy, 2011) and Franz cell method, that examines the diffusion across mucus-like barriers (Colombo et al., 2013). To our knowledge, the effect of freeze-drying on the mucoadhesiveness of nanoparticulate systems was not systematically studied before. Our second specific aim was to examine the potential of these carriers as drug delivery vehicles in the gastrointestinal tract. For this purpose we used an in vitro cell model that was previously established to mimic the epithelium of the small intestine (Antunes et al., 2013). This triple co-culture cell model is composed of Caco-2 and HT29-MTX cell lines, which function as intestinal enterocytes and mucus producing cells, respectively. The ability of an epithelial cell to act as M-like cell and translocate nanoparticles is induced by a third cell line, Raji B cells, which is added to the basolateral side of the supporting membranes at the fourteenth day of co-culture, providing high resemblance to the small intestinal epithelium (Kerne et al., 1997). In this work, dextran sulfate was used as a model macromolecule drug and was incorporated into the nanoparticles. Low molecular weight dextran sulfates were shown to inhibit the activation of both the coagulation and the complement systems and were suggested to treat an instant blood-mediated inflammatory reaction in clinical islet transplantation (Johansson et al., n.d.). This macromolecule can also act as a model for other macromolecule drugs, such as different polysaccharides, peptides and proteins that have poor absorption in the gastrointestinal tract.

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2. Materials and Methods 2.1 Materials Low molecular weight chitosan (Mw 207 kDa, deacetylation degree 77.6%), D-glucose, fluorescein isocyanate (FITC), mucin type II from porcine stomach, L-αphosphatidylcholine, alcian blue and fluorescamine were purchased from Sigma Aldrich (Rehovot, Israel). Sodium tripolyphosphate (TPP) was obtained from Alfa Aesar (Lancashire, UK). Sodium taurocholate was obtained from Carbosynth (Compton, Berkshire, UK) and maleic acid from Arcos Organics (Geel, Belgium). Sodium chloride (NaCl), dichloromethane and sodium hydroxide were purchased from Bio-Lab Ltd. (Jerusalem, Israel). Acetic acid glacial and dimethyl sulfoxide (DMSO) were purchased from Merck (Darmstadt, Germany). Sucrose was purchased from J. T. Baker-Avantor (Allentown, PA, USA) and PEG 10 kDa from Merck (Kenilworth, NJ, USA). PEGDA with Mw of 10 kDa was obtained from the laboratory of biomaterials and regenerative medicine at the Department of Biomedical Engineering, Technion, Israel (Eliyahu et al., 2018). Dextran sulfate-rhodamine (DSR) 3.5 kDa was purchased from Creative PEGworks (Durham, NC, USA). Fresh porcine small intestine was obtained from the Pre-Clinical Research Authority, Technion, Israel. The porcine intestine was sliced and stored at -20 °C until further use. 2.2 Synthesis of Acrylated Chitosan ACS was synthesized as described before (Eliyahu et al., 2018). Briefly, one gram of CS was dissolved in 100 mL of 2% (v/v) acetic acid and stirred overnight at room temperature (RT). Next, one gram of PEGDA was added and the mixture was incubated at 60 °C for 3 h under shaking. The reaction mixture was dialyzed in the dark against five litters of double deionized water (DDW) for 72 h. The mixture was filtered with a Buchner funnel and freeze-dried. The product was stored at -20 °C until further use. ACS

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was evaluated for its acrylation degree using the fluorescamine assay, a method that was established in our previous work (Eliyahu et al., 2018). This value was found to be 54%. 2.3 Nanoparticle Fabrication Nanoparticle formulations were prepared according to a previously described protocol (Eliyahu et al., 2018). Briefly, 0.1% (w/v) of polymer (CS or ACS) was dissolved in 0.1% (v/v) acetic acid with 0.1 M NaCl, stirred for 20 h and filtered using 0.45 µm syringe filter. A volume of 1.25 mL of TPP (0.25% (w/v)) in NaCl (0.1 M) aqueous solution was added dropwise to 2.5 mL of polymer solution under magnetic stirring. The solutions were stirred for 15 minutes at RT. FITC-labeled nanoparticles were fabricated according to the same procedure using labeled polymers (Eliyahu et al., 2018). 2.4 Fabrication of Nanoparticle Loaded with Dextran Sulfate Loaded nanoparticles were prepared as described in section 2.3 with the addition of DSR. TPP solutions were mixed with DSR to a final concentration of 0.25 mg/mL DSR. Then, a volume of 1.25 mL of TPP/DSR solution was added dropwise to 2.5 mL of polymer solution under magnetic stirring. The solutions were stirred for 15 minutes at RT. The final concentration of DSR in the nanoparticle dispersion was 0.085 mg/mL. 2.5 Drug Loading Efficiency The amount of DSR entrapped in the nanoparticles was determined using precipitation and quantification of unloaded DSR in the supernatant. Loaded nanoparticle solutions were transferred to a Vivaspin® 6 mL centrifugal filter tube with 10 kDa polyethersulfone membrane (Merck Millipore) and centrifuged using a Megafuge 1.0 centrifuge (Heraeus, Hanau, Germany) at 3300 x g, 25 °C for 30 min to separate the nanoparticles from the free DSR solution. During the centrifugation, the nanoparticles precipitated on the membrane and the upper liquid was left with free molecules of DSR. Samples of 200 µL were placed in black 96-well plate and analyzed for their DSR content using a Synergy HT® microplate reader (Biotek Instruments Inc., Winooski, VT, 6

USA) at an excitation and emission wavelengths of 540 ± 25 nm and 590 ± 20 nm, respectively, for rhodamine. The entrapment efficiency (EE) was calculated as follows:

ሺͳሻ‫ܧܧ‬ሺΨሻ ൌ 

ܶ‫ ܴܵܦ݈ܽݐ݋‬െ ‫ܴܵܦ݁݁ݎܨ‬ ή ͳͲͲ ܶ‫ܴܵܦ݈ܽݐ݋‬

where Total DSR is the total mass of dextran sulfate rhodamine in the nanoparticle solutions, and Free DSR is the mass of dextran rhodamine sulfate in the supernatant. 2.6 Size and Zeta Potential Evaluation Dynamic light scattering (DLS) was used to determine the average size and size distribution, while laser Doppler anemometry was used to assess the zeta potential of the nanoparticles. Samples were diluted with 0.1% (v/v) acetic acid to half of the initial concentration at the time of preparation and the size and zeta potential were measured in triplicate using a Zetasizer Nano ZS (Malvern Instruments Ltd., Malvern, UK). 2.7 Freeze-Drying Freshly prepared nanoparticles were mixed with cryoprotectants (glucose, sucrose or PEG) at increased concentrations of 15%, 20% and 30% (w/v). The solutions were stirred for 5 minutes at RT until the cryoprotectant was fully dissolved, and then freezedried under vacuum at -30 °C. The freeze-dried nanoparticles were dispersed in DDW to the same concentration before the freeze-drying process and were used for further experiments as suspensions. 2.8 Franz Diffusion Cell Experiments The diffusion of FITC-labeled nanoparticles and the model drug DSR was studied using a homebuilt apparatus (Otmazgin, 2017), similar to others described elsewhere (Colombo et al., 2013). This apparatus consists of four donor and acceptor cells and a heating magnetic stirrer with a thermocouple to monitor the temperature of a water bath with four blocks in which each of the cells were placed. A volume of 9 mL of phosphate buffer saline (PBS 0.1 M, pH 6.8 which simulates the conditions in the small intestine, 7

prepared according to (Shtenberg et al., 2017)) was filled into each receptor compartment. The donor compartment was filled with 1.5 mL of nanoparticles or DSR, and their diffusion was examined through a mucus gel that was prepared by dissolving mucin type II from porcine stomach powder (200 mg/mL in PBS). A layer of 200 µm of mucin gel was spread on a filter paper (Nuclepore Whatman 25 mm, with pore size of 0.8 µm) and another filter paper was placed on top of the mucin layer. The filter papers assembly was placed between the donor and receptor compartments with a mean exposed area of 1.13 cm2. Both cells and the filter papers assembly were clamped together. During the experiment, the apparatus was kept at a constant temperature of 37 °C. Samples of 200 µL were withdrawn from the receptor compartment at predetermined time intervals and the same volume of fresh PBS solution was added to the receptor compartment immediately after sample withdrawal. The concentration of FITC-labeled nanoparticles or DSR in the samples was determined by measuring the fluorescence with a Synergy HT® microplate reader (Biotek Instruments Inc., Winooski, VT, USA) at an excitation and emission wavelengths of 485 ± 20 and 525 ± 20 respectively for FITC and 540 ± 25 nm and 590 ± 20 nm for rhodamine. The permeability percentage was calculated as follows:

ሺʹሻܲ݁‫ݕݐ݈ܾ݅݅ܽ݁݉ݎ‬ሺΨሻ ൌ 

‫ܯ‬௣௘௥௠௘௔௧௘ௗ  ή ͳͲͲ ଴

where M permeated is the mass of nanoparticles or drug that permeated across the barrier, and M0 is the mass of nanoparticles or drug that was placed in the donor compartment at time zero. 2.9 Flow Through Experiments Retention studies using porcine intestinal mucosa were performed according to a previously described method (Eliyahu et al., 2018). Frozen porcine intestine tissue was sliced into 1.5x3 cm portions and thawed for 5 min in 100% humidity and 37 °C. FITClabeled nanoparticles (50 µL of fresh or freeze-dried and re-dispersed) were placed on 8

the mucosal side of the tissue and were immediately washed or incubated at 37 °C and 100% humidity for 10 minutes. Using a syringe pump, a flow of simulating intestinal fluid (FaSSIF-V2 pH 6.8, prepared according to (Eshel-Green et al., 2016)) washed the formulation off the tissue. Aliquots of 200 µL were collected continuously and analyzed fluorescently using a Synergy HT® microplate reader (Biotek Instruments Inc.) at an excitation wavelength of 485 ± 20 and emission wavelengths of 525 ± 20. All measurements were performed in triplicate. The aliquots were analyzed for their nanoparticle concentration using a calibration curve of the labeled nanoparticles in FaSSIF-V2 buffer. 2.10 Cultured Cell Lines The Caco-2 cell line originated from human colon was obtained from the American Type Culture Collection (ATCC, USA). Mucus producing colon cell line HT29-MTX was kindly provided by Dr. T. Lesuffleur (INSERM U178, Villejuif, France) and Raji B cell line originated from the human lymph was kindly provided by Dr. Alexandre Carmo (Cellular and Molecular Biology Institute – IBMC, Porto, Portugal). 2.11 Cell Viability Assays Cell viability was assessed by the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) (MTT) reduction assay. The cytotoxicity of sucrose 30%, acetic acid 0.1% and NaCl 0.1 M, fresh and freeze-dried polymeric nanoparticles, free DSR and DSR-loaded nanoparticles was studied in Caco-2 and HT29-MTX cell lines. Cells were seeded at a density of 10,000 cells/well in 96-well plates using Dulbecco’s Modified Eagle’s Medium (DMEM, Lonza, Verviers, Belgium) supplemented with 10% (v/v) fetal bovine serum (FBS, Biochrom, Cambridge, UK) and 100 U/mL of penicillin and 100 mg/mL of streptomycin (Biowest, Nuaillé, France). After 48 h incubation at 37 ºC in a humidified atmosphere containing 5% CO2, the medium was removed, and cells were incubated in the presence of the treatment mentioned above in three volume fractions of 10%, 20% or

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30% (v/v) in cell culture medium, corresponding to 0.00833, 0.01667 or 0.2500 mg/mL of DSR, respectively. Cells with no treatment were used as positive control (100% cell viability), and incubation with Triton® X-100 (1% (v/v) in DMEM) was used as negative control (0% cell viability). Cells were incubated under the conditions described above for 3 h and 24 h. Then, the wells were washed twice with PBS (pH 7.4 at 37 ºC) and 200 µL of the MTT reagent (0.5 mg/mL in DMEM) were added to each well. Plates were incubated for an additional 4 h at 37 ºC. Subsequently, the content of the wells was carefully removed and 200 µL of DMSO was added to each well in order to solubilize the formazan crystals metabolized by viable cells. Plates were kept for 15 min in continuous shaking at 120 rpm. Next, the absorbance at 570 nm (test wavelength) and 630 nm (background wavelength) was measured using a Synergy 2 ® microplate reader (Biotek Instruments Inc.). Tests were performed in sextuplicate. 2.12 Permeability Assays Permeability experiments were performed on a triple co-culture cell model (Araújo and Sarmento, 2013). Co-cultures of Caco-2 and HT29-MTX at a 90:10 ratio were seeded in Millicell® inserts of 12 mm diameter with a translucent permeable membrane of polyethylene terephthalate (PET), pore size of 1 µm, and a growth area of 1.1 cm2. Cells were seeded at a density of 105 cells/cm2, in a final volume of 0.5 mL in the apical compartment of the insert, placed in 12-well plates, and 1.5 mL of supplemented DMEM was added to the basolateral compartment. The plates were maintained at 37 ºC in humidified atmosphere containing 5% CO2. The medium of the apical and basolateral compartments was changed every other day. The cell monolayers were monitored for their Transepithelial Electrical Resistance (TEER) values using an EVOM2 ® epithelial voltohmmeter with chopstick electrodes (World Precision Instruments, Sarasota, FL, USA). After 14 days, 105 Raji-B cells were added to the basolateral compartment. Once these cells were added, the medium in this compartment was not renewed until the permeability tests on day 21, while the medium in the apical side was changed every day. 10

To perform the in vitro permeability experiments, cell culture medium was carefully removed from the apical and the basolateral compartments; the inserts and wells were gently washed twice with PBS (pH 7.4) at 37 ºC to remove all the supplemented DMEM, and then filled with 0.5 and 1.5 mL of Hank’s balanced salt solution (HBSS) in the apical and basolateral compartments, respectively. The monolayers were maintained for 30 min at 37 ºC. Next, the media from the apical compartment was removed and 0.5 mL of free DSR at 0.085 mg/mL in HBSS was added. Nanoparticles with the same concentration of DSR were diluted in HBSS buffer and were placed in the apical compartment in the same manner. Plates were placed inside an orbital shaking incubator (IKA® KS 4000 IC, IKA, Staufen, Germany) at 100 rpm and 37 ºC. Aliquots of 200 µL were withdrawn from the basolateral compartment at predetermined time points (0, 15, 30, 45, 60, 90, 120 and 180 min) and immediately replaced with the same volume of pre-heated HBSS. At the end of the experiment, an aliquot from the apical compartment was collected. Tests were performed in triplicate and an insert without the addition of sample was used as a control. Before, during and at the end of the permeability experiments, the TEER was measured in order to monitor the integrity of cell monolayers. Experiments were performed in triplicate. The concentration of DSR in the samples was determined by measuring fluorescence with a Synergy 2® microplate reader at an excitation wavelength of 553 nm and emission wavelength of 627 nm. The drug apparent permeability coefficient (Papp) was calculated from the following equation (Artursson, 1991): ሺ͵ሻܲ௔௣௣ ൌ  ൤

݀‫ܥ‬ ͳ ή ൨ ή ൤ ൨ ݀‫ݐ‬ ‫ ܣ‬ή ‫ܥ‬଴

where Papp is the apparent permeability (cm/s); dC/dt (mg/mL∙s) is the rate of transport across the monolayer obtained from the slope of the curve of the amount of drug transported versus time; V (cm3) is the acceptor compartment volume, which in this case corresponds to 1.5 cm3; A (cm2) is the insert membrane growth area; and C0 (mg/mL) is the initial concentration in the apical compartment.

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2.13 Alcian Blue Staining The mucus production from the triple model was characterized by staining the glycoproteins present in the mucus secretions with alcian blue to allow the mucus visualization on the surface of the cells. After cultivating for 21 days (section 2.11), the monolayer was fixed using 2% (v/v) paraformaldehyde (PFA) for 30 min and then washed three times with PBS. Then, the membranes were removed from the Millicell ® supports and cut along the longitudinal axis using a Microm HM 550 cryostat (Thermo Scientific, Waltham, MA, USA) and put on glass slides. The slides were washed twice with DDW and once with 3% (v/v) acetic acid. Next, the slides were rinsed in 0.25% (w/v) alcian blue solution in 3% (v/v) acetic acid (pH 2.5) for 15 min and washed using 3% (v/v) acetic acid. The slides were rinsed in tap water for 10 min and left to dry overnight. Finally, cells were mounted onto glass slides with mounting medium (Cryomatrix, Thermo Scientific) and sealed with coverslips. Samples were refrigerated at 4 °C prior to analysis. Images of each sample were taken using an Olympus CX31 microscope (Olympus, Tokyo, Japan). 2.14 Statistical Analysis Data from independent experiments were analyzed for each variable using Microsoft ® Excel software. Standard errors of the mean (SEM) were calculated and presented for each treatment group. Comparison between multiple treatments was made with analysis of variance (ANOVA), and ad-hoc comparison between two treatments was performed using a one-tailed Student’s t-test. A p-value less than 0.05 was considered to be statistically significant.

3. Results and Discussion 3.1 Effect of Freeze-drying on Particle Size The poor stability of colloidal nanoparticles in aqueous solutions is an obstacle which considerably limits their usage in clinical applications. In particular, CS nanoparticles are 12

not stable for long periods of time, because they can undergo hydrolysis in aqueous solutions (Jonassen et al., 2012a). Moreover, CS-TPP nanoparticles tend to aggregate in solution, since they are based on ionic gelation between the positively charged amine group of CS and the negatively charged TPP (Jonassen et al., 2012b). Freeze-drying is an industrial process that converts suspensions and solutions into solids by sublimation of the water content under vacuum. This technique is considered a suitable solution that enables preservation of nanoparticles in the solid state for a long time (Abdelwahed et al., 2006). However, nanoparticles tend to agglomerate after freezedrying and their size is not preserved after re-dispersion. Cryoprotectants, typically sugars or polymers, can help prevent aggregation and irreversible fusion, as well as increase the stability during storage (Fonte et al., 2016). In the current study, we used sucrose, glucose and PEG as cryoprotectants for CS and ACS nanoparticles and examined their size after freeze-drying and re-dispersion. PEG was not efficient as a cryoprotectant, as supported by the considerable increase in particle size after freeze-drying and re-dispersion (data not shown). Figure 1 presents the nanoparticle Z-average and polydispersity index (PDI) obtained following drying in the presence of glucose or sucrose. The Z-average is an intensity-based harmonic mean, which indicates the average size based on a specific fit to the raw correlation function data, while the PDI value is an indication for the polydispersity (Arzenšek et al., 2010). In the absence of cryoprotectants, the size of the nanoparticles increased significantly, from 180.6 ± 2.1 nm to 1499.3 ± 137.1 nm for CS nanoparticles and from 121.2 ± 0.9 nm to 2979.7 ± 128.1 nm for ACS nanoparticles after freeze-drying and re-dispersion (p < 0.001 and p < 0.05 respectively, Figure 1A, B). Poor quality of protection was observed when 150 mg/mL cryoprotectant was added, for both particle types and both cryoprotectants, leading to aggregates larger than 1000 nm and PDI value close to 1. For ACS nanoparticles, an increase in glucose concentration to 200 mg/mL decreased the particle size to 409.2 ± 30.1 nm (p < 0.005), but a further increase did not change 13

Figure 1. The average size of ACS (white) and CS (grey) nanoparticles after freeze-drying and re-dispersion in DDW with the addition of increased concentrations of cryoprotectants: (A) glucose and (B) sucrose. The PDI values of the nanoparticles after freeze-drying and re-dispersion in DDW with the addition of increased concentrations of cryoprotectants: (C) glucose and (D) sucrose. (*) refers to statistically significant difference (p < 0.05), (***) refers to statistically significant difference (p < 0.001) and (****) refers to statistically significant difference (p < 0.0001). The bars represent standard error of the mean, n = 3.

significantly the Z average value (Figure 1A). Glucose concentration of 200 mg/mL decreased the size of CS nanoparticles to 699.6 ± 134.2 nm compared to glucose concentration of 150 mg/mL (p < 0.05), and a further decrease was observed using 300 mg/mL (p < 0.05), resulting in a particle size of 488.2 ± 46.3 nm (Figure 1A). Nevertheless, the PDI values of the nanoparticle solutions using glucose as cryoprotectant were high, which indicate that the distribution is polydisperse (Figure 1C). The use of sucrose as a cryoprotectant did not change significantly the post freeze-drying size compared to glucose in the case of CS nanoparticles, but decreased the size of ACS nanoparticles (p < 0.05, Figure 1B, D). PDI values of CS nanoparticles were found to be

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significantly dependent in the sucrose concentration (ANOVA, p < 0.0001). The best size preservation for both formulations was observed when 300 mg/mL sucrose was added, as the Z-average size was the lowest for ACS nanoparticles and PDI values were similar to those measured immediately after fabrication for CS nanoparticles. The post-freezedrying size of ACS nanoparticles with 300 mg/mL added sucrose was 267.9 ± 3.4 nm, while CS nanoparticles aggregated to a final size of 470.8 ± 9.3 nm (Figure 1B). The process of freeze-drying involves extraction of water molecules that form hydrogen bonds with the particles and contribute to the stability of the particle in solution. In the dry state, intramolecular hydrogen interactions replace the interactions with water (Crowe et al., 1993). When the particles are re-hydrated, intramolecular bonds are only partially broken, thus increasing the average particle size in the suspension (Allison et al., 1996). The addition of sugar molecules prevents particle aggregation because the interactions with sugar molecules replace the intramolecular hydrogen interactions. Upon re-hydration, the sugar-particle linkages are broken by water molecules and the particle regains its shape and size (Fonte et al., 2016). The results presented above are in agreement with previous studies on CS-TPP nanoparticles. In a study performed by Almalik et al., the best size preservation was obtained using sucrose (Almalik et al., 2017). It was claimed that this sugar efficiently forms hydrogen bonds with the CS-TPP nanoparticles because it does not have internal hydrogen bonding. ACS nanoparticles are composed of hydrophilic PEG chains that could also absorb water molecules (Umerska et al., 2018) and would probably contribute to the effective cryoprotection (Figure 1B, D). Following the results presented in this section, sucrose concentration of 300 mg/mL was chosen for further experiments.

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3.2 Effect of Freeze-Drying on Drug-Loaded Nanoparticles Since mucoadhesive nanoparticles are intended for use as drug carriers, it is essential to study the effect of drug loading and to characterize the effect of freeze-drying on drugloaded nanoparticles. For this purpose, we used dextran sulfate, which is known to inhibit the activation of both the coagulation and the complement systems (Johansson et al., n.d.). In addition, it can be used as a model for other macromolecule drugs such as polysaccharides, peptides and proteins. Nanoparticles were fabricated in the presence of DSR. The encapsulation efficiency was calculated from eq. 1 and found to be 87.5% ± 2.7% for ACS nanoparticles and 90.0% ± 4.1% for CS nanoparticles. The resulting drug-loaded nanoparticles were characterized in terms of size and zeta potential (Figure 2). Two parameters were examined: the influence

Figure 2. (A) Mean hydrodynamic diameter and (B) zeta potential of fresh and freeze-dried empty ACS and CS nanoparticles (white) and DSR loaded nanoparticles (grey). (*) refers to statistically significant difference (p < 0.05), (**) refers to statistically significant difference (p < 0.005) and (***) refers to statistically significant difference (p < 0.001). The bars represent standard error of the mean, n = 3.

of incorporating the model drug DSR on fresh nanoparticles, and the effect of sucrose as a cryoprotectant when the particles were freeze-dried and re-dispersed. The addition of DSR during nanoparticle fabrication caused a significant increase in the particle size, of about 80% for ACS nanoparticles and 35% for CS nanoparticles for fresh nanoparticles (p < 0.05, Figure 2A). After freeze-drying and re-dispersion in DDW, the size of the

16

loaded particle was increased in about 65% for ACS nanoparticles and 55% for CS nanoparticles, in comparison to the fresh loaded nanoparticles (p < 0.05, Figure 2A). Overall, similar to empty nanoparticles, addition of sucrose did not prevent completely the aggregation of the loaded particles during freeze-drying.

The chitosan-based nanoparticles used in this study were prepared using an ionic gelation process, where the positively charged CS backbone is crosslinked by the negatively charged TPP. When the negatively charged DSR is included in the formulation, it can be either encapsulated inside the nanoparticles, or adsorbed on their positively charged surface. The bulky DSR imposes less compact structure of the nanoparticles, causing their size to increase. Other studies on CS-TPP nanoparticles reported the same trend upon loading with negatively charged drugs. Trapani et al., used low molecular weight heparin and showed that the size of the nanoparticles has increased compared to empty nanoparticles (Trapani et al., 2013). Another study demonstrated that the size of CSpectin microparticles increased when loaded with insulin (Maciel et al., 2017). One More important characteristic of nanoparticles is the zeta potential value, that can be used to describe the electric double-layer potential of colloidal dispersions (Lu and Gao, 2010). The zeta potential values of CS nanoparticles (fresh or freeze-dried) were found to be significantly greater than those of ACS nanoparticles (p < 0.005, Figure 2B). This result is in line with our previous report (Eliyahu et al., 2018). The grafting of PEGDA chains on the CS backbone occupies some of the positively charged amine groups, leading to a decrease in zeta potential. The freeze-drying process and the sucrose presence did not affect the zeta potential of empty or loaded nanoparticles (Figure 2B), with similar values for fresh and freeze-dried nanoparticles. These results are in agreement with previous studies that used sucrose for cryoprotection [6, 26] and demonstrated that the increased size of CS-TPP nanoparticles

17

was not associated with a change in the zeta potential. The incorporation of DSR inside fresh and freeze-dried ACS nanoparticles resulted in a significant decrease in zeta potential (p < 0.005 and p < 0.001 respectively, Figure 2B). During the nanoparticle fabrication, CS or ACS were added in mass excess of about 8:1 relative to DSR. As previously suggested by Thierry Delair, when CS is in excess, neutral segments are formed by charge neutralization and segregating to form the core of the nanoparticle, such that an outer shell is formed from the excess CS (Delair, 2011). Thus, the final positive value of the zeta potential reflects the CS or ACS excess and implies that the stability is relatively high. As was mentioned before, CS nanoparticles have initially more positively charged amine groups compared to ACS, allowing more drug molecules to occupy these groups without affecting the zeta potential. Moreover, the high zeta potential of loaded CS nanoparticles could enhance the repulsion between the particles and explain the relatively low increase in size after freeze-drying, in comparison to ACS nanoparticles (Gala et al., 2015). It is commonly accepted that stable particles have a zeta potential value of between +30 mV and –30 mV (Riddick, 1968) and, since ACS nanoparticles display values outside this range, one might suggest that they are not stable. However, it is known that the addition of PEG chains contributes to the stability of the particles and oppose aggregation (Soo et al., 2016). The pendent PEG chains seem to contribute to the stability of ACS nanoparticles, despite the fact that their zeta potential is close to zero. The incorporation of the drug caused an additional significant decrease in the zeta potential value of ACS nanoparticles compared to both empty fresh and empty freezedried nanoparticles (p < 0.005, Figure 2B). The zeta potential of CS nanoparticles was not affected by the addition of the drug.

18

3.3 In Vitro Diffusion Studies Drug encapsulated within mucoadhesive nanoparticles may pass through the mucus and reach the underlying cell layer by two main routes: either remain entrapped in the nanoparticle and progress due to nanoparticle diffusion or be released from the nanoparticle and diffuse as a free drug. Obviously, these two mechanisms can co-exist, and their relative importance depends on the rates of drug release, nanoparticle diffusion and drug diffusion. In order to better understand the transport mechanisms in the system under study, we characterized the rate of nanoparticle and drug transport through a mucus layer surrogate using a Franz diffusion cell (Figure 3A). Following a previous research by Otmazgin et al., that used a layer of mucus gel to mimic the barrier and study the permeability of blue dextran (Otmazgin, 2017), we adopted the same concept to study the penetration of FITC-labeled nanoparticles and rhodamine-labeled drug. The properties of the mucin layer were adjusted in order to best mimic the properties in the gastrointestinal tract. The thickness was selected to be 200 µm, as this is the average mucin thickness in the small intestine [30, 31]. The experiment duration was determined considering the mucin turnover of 4 hours in the small intestine (Pepić et al., 2013). Control experiments were performed to verify the permeability of the nanoparticles through the supporting membrane (Figure 1S).

19

Figure 3. (A) A schematic illustration of the Franz diffusion cell was taken with permission from Otmazgin et al., [21]; (B) Nanoparticle permeability across the mucus gel as a function of time; (C) The rate of penetration calculated as P app. (**) refers to statistically significant difference (p < 0.005). The bars represent standard error of the mean, n = 3.

20

Generally speaking, the penetration or adhesion of a particle to the mucus is controlled by characteristics such as particle geometry, surface properties and size (Schattling et al., 2017). In the case of mucoadhesion, the mucus layer and a mucoadhesive material come into physical contact and then the particle interpenetrates to the mucin matrix leading to entanglement and formation of physical or chemical bonds between entangled chains. Consequently, the transport of mucoadhesive particles will be slow. Mucoadhesion is usually achieved with hydrophilic polymers bearing charged chemical groups that establish electrostatic interactions with mucin; thiol groups can also be added to create disulfide bridging with mucin (Khutoryanskiy, 2011). Contrary, mucus-penetrating particles do not form many interactions with the mucus layer and therefore diffuse faster. These systems aspire to minimize the interactions with mucus (Netsomboon and Bernkop-schnürch, 2016). Figure 3B shows the nanoparticle permeability (calculated from eq. 2) vs. time. Fresh CS nanoparticles exhibited low permeability percentage of only 0.453% ± 0.089%. This finding can be attributed to their high mucoadhesive behavior causing them to adhere to mucin. Similarly, fresh ACS nanoparticles presented low penetration of 2.97% ± 0.98% arising from their enhanced mucoadhesion. After freeze-drying with 30% sucrose, the permeability of CS nanoparticles increased, but remained low at a value of 1.36% ± 0.41%. Unpredictably, freeze-dried ACS nanoparticles presented significantly higher permeability of 18.29% ± 0.62%, compared to fresh ACS nanoparticles and fresh and freeze-dried CS nanoparticles (p < 0.0001, Figure 3B). The permeability coefficients, representing the rate of penetration, were calculated using eq. 3 and followed the same trend (Figure 3C). CS nanoparticles penetrated through the mucus layer at a rate 2.7-fold faster after freeze-drying, while for ACS nanoparticles the rate of permeability was increased by 8.6-fold (p < 0.005, Figure 3C). The sucrose included in the formulation could affect the results in two ways. First, it could increase the osmotic pressure in the upper cell and encourage water transport from 21

the mucin layer into this cell. As a result, the effective concentration of the glycoproteins in the tested layer would increase, which in turn should decrease the nanoparticle transport. However, the results suggest otherwise. A second feasible mechanism involves water transport from the lower cell, through the mucin layer, and into the upper cell. This scenario would lead to dilution of the nanoparticle solution that will effectively decreased diffusion rates. Since the diffusion rates are in fact higher after freeze drying (Figure 3C), dilution of nanoparticles can be overruled as well. Furthermore, the experimental results indicate that the sucrose did not affect the transport of CS nanoparticles in the same manner as ACS nanoparticles, as could be expected if its only effect were inducing dilution or osmotic pressure changes. Thus, a probable reason for the enhancement in the mobility of freeze-dried ACS nanoparticles is loss of mucoadhesion due to the drying process. It is believed that mucoadhesive systems are not able to penetrate across the mucus layer because of their strong adhesiveness (Lai et al., 2009). Thus, in order to further investigate the phenomenon of increased permeability after freeze-drying and understand whether it is related to changes in mucoadhesion, we performed experiments using the flow through technique. This method is a quantitative method used to assess mucoadhesion after different incubation times and investigate the kinetics of the nanoparticle binding (Eliyahu et al., 2018). For these experiments we used portions of porcine small intestine. Fluorescently labeled nanoparticles were placed on a porcine mucosal membrane and were either incubated for 10 minutes or immediately washed using heated buffer simulating the fluids in the intestine. Figure 4A demonstrates that the retention profiles of fresh and freeze-dried ACS nanoparticles without an incubation stage are similar, resulting in the same mass retention at the end of the experiment (Figure 4C). Surprisingly, after incubation for 10 minutes, freeze-dried ACS nanoparticles did not present improvement in adherence, although the retention of fresh ACS nanoparticles increased (Figure 4B). The curves were used to determine the steady

22

state retention, previously defined as 95% of the final retention at the end of the experiment (Eliyahu et al., 2018). After 10 minutes of incubation, a steady state of 71% retention was observed for fresh ACS nanoparticles, while freeze-dried ACS nanoparticles reached a steady state of 58% retention. Figure 4C summarizes the retention percentages at the end of the experiment, wash volume of about 4 mL. A significant improvement in mucoadhesion with the increase in the incubation time was observed in case of fresh ACS nanoparticles (t-test, p < 0.05, n = 3). However, the retention percentage of freeze-dried ACS nanoparticles was not enhanced following an increase in incubation time (Figure 4C). Enhanced mucoadhesion after incubation was described for fresh CS and ACS nanoparticles under similar conditions (Eliyahu et al., 2018). These phenomena can be partially explained using the diffusion theory, according to which the interpenetration of a liquid formulation depends directly on the contact time with the mucosal surface (Khutoryanskiy, 2011). Furthermore, incubation allows more time for secondary interactions that are formed upon contact with the mucosa, such as hydrogen, Van der Waals or electrostatic interactions. Coating the surface of particles with long PEG chains was previously shown to increase the mucoadhesiveness of nanoparticles [31, 32] as a result of hydrogen bonds between glycosylated residues of mucin and the ether oxygen of the EO segment [34, 35]. In the case of ACS nanoparticles, covalent bonding via Michael-type addition reaction may also occur between free acrylate end groups on the surface of the nanoparticles and thiol groups present in mucin glycoproteins (Shitrit and Bianco-Peled, 2017). Therefore, for fresh ACS nanoparticles, the increase in mucoadhesion with incubation time can be attributed to both additional entanglements and chemical interactions of long PEG chains and acrylate end groups, respectively. Freeze-dried ACS nanoparticles adhere to the examined mucosal surface, however an increase in mucoadhesion after longer incubation was not observed. These results imply that freeze-drying partially impairs mucoadhesion on a time-dependent manner. The only 23

difference between the ingredients included in the fresh and freeze-dried formulations is the sugar. As a cryoprotectant, sucrose is added in large excess and might occupy the ether sites on PEG chains forming hydrogen bonds. Hence, even when the freeze-dried ACS nanoparticles are given more time to interact with the mucosa, these sites are not available to bind to the glycosylated residues of mucin. Furthermore, the freeze-drying stage caused the particles to aggregate (Figure 1). As a result, some of the grafted PEG chains could be “buried” inside the nanoparticles and were unable to interact with mucin. Therefore, it could be that at long incubation periods, the strengthening of the interactions by entanglements and Michael-type addition reaction is damaged because less acrylate groups and PEG chains are present on the surface. Using sucrose is a common technique for cryoprotection, yet it has drawbacks in terms of damaging the mucoadhesion.

24

Figure 4. Retention profile of: (A) Fresh ACS (triangles) and freeze-dried ACS nanoparticles (squares) without incubation; (B) Fresh ACS (triangle) and freeze-dried ACS nanoparticles (squares) with 10 minutes of incubation; (C) Final retention percentages of fresh and freeze-dried ACS nanoparticles obtained after different incubation time intervals (t test, p < 0.05, n = 3). (*) refers to statistically significant

25 difference (p < 0.05). The bars represent standard error of the mean, n = 3.

Next, we performed Franz cell experiments to examine the drug transport through the mucin layer. Fresh and freeze-dried DSR-loaded nanoparticles were placed in the donor compartment and the medium in the receptor compartment was monitored for the presence of the drug. The results demonstrate that the permeability of the drug increased with time for all examined systems (Figure 5A, B). Figure 5C depicts the permeability coefficient (Papp) of DSR diffusion across the mucin gel. The highest percentage of mass permeability was observed for DSR in its free state. There were no significant differences between the permeability percentage of fresh and freeze-dried nanoparticles, however both types of ACS nanoparticles displayed higher permeability compared to fresh and freeze-dried CS nanoparticles (fresh ACS and CS nanoparticles p < 0.05, freeze-dried ACS and CS nanoparticles p < 0.01). Clearly, since CS and fresh ACS nanoparticles are immobilized in the mucus layer (Fig. 3B), the drug is transported only after its release from the particles. The fact that drug permeability and permeability rate is similar for fresh and freeze-dried ACS nanoparticles suggests that the main transport mechanism for those nanoparticles is diffusion of free drug as well (Fig. 5). It stands for reason that the higher permeability in the case of ACS nanoparticles may reflect higher release rates (Fig. 5C), arising from the more hydrophilic nature of these nanoparticles as compared to the CS nanoparticles. The higher release rate from the ACS nanoparticles may be considered as beneficial since mucus turnover is expected to remove the particles at times longer than about four hours.

26

Figure 5. (A) Fresh nanoparticles mediated DSR permeability across the mucus gel as a function of time; (B) Freeze-dried nanoparticles mediated DSR permeability across the mucus gel as a function of time; (C) The rate of penetration calculated as P app. (*) refers to statistically significant difference (p < 0.05). The bars represent standard error of the mean, n = 3.

27

3.4 Cell Viability Assays The in vitro cytotoxicity of the nanoparticles was studied using the MTT reduction assay in order to explore any potential safety issues. Different formulations were tested after 3 h and 24 h of incubation to assess the immediate and prolonged effect on the cell viability. The experiments were performed on two cell lines of the triple cell co-culture model (Caco-2 and HT29-MTX) to determine a safe concentration for the permeability assay. Values of 70% cell viability or higher were considered as potentially safe [36, 37]. Different constituents of the nanoparticle formulations were examined including acetic acid 0.1% (v/v) with NaCl 0.1 M, sucrose 30% (w/v) and free drug (Figure 2S). No cytotoxic effects were found to be triggered by these materials. Positive and negative controls were assessed using cells without any treatment and 1% Triton X-100, respectively. Figure 6 displays the viability of cells incubated with 30% (v/v) DSRloaded ACS and CS nanoparticles freshly prepared and freeze-dried and re-dispersed. It was observed that after 3 h of incubation with the cells, CS nanoparticles did not cause any cytotoxic effect on both cell types (Figure 6). The cell viability after 3 h of incubation with ACS nanoparticles was generally lower than for CS nanoparticles, but the viability percentage was still above the established threshold. Incubation of freezedried loaded ACS nanoparticles with Caco-2 cell line for 3 h and 24 h resulted in significantly greater viability percentage as compared to fresh nanoparticles (p < 0.05, Figure 6A). The same effect was not observed with CS nanoparticles. Mild cytotoxicity was observed for HT-29 MTX cells that were incubated for 24 h with fresh loaded ACS nanoparticles, as suggested by the viability value of 60.8% ± 3.0% (Figure 6B). An increase in viability percentage was observed when freeze-dried loaded ACS nanoparticles were incubated with HT-29 MTX cell line for 3 h and 24 h, compared to fresh nanoparticles (p < 0.05, p < 0.005 respectively, Figure 6B). Incubating fresh loaded CS nanoparticles for 3 h with HT-29 MTX cell line decreased the viability compared to

28

freeze-dried loaded CS nanoparticles (p < 0.05), while incubation for 24 h increased the viability of the cells (p < 0.005, Figure 6B). It is well established that CS is biocompatible and presents low toxicity potential (Andersen et al., 2017), making it a widely used polymer in the food and pharmaceutical industries [40, 41]. Moreover, TPP is approved by the U.S. Food and Drug Administration for human use (Jonassen et al., 2012b), while DS was previously shown to present low cytotoxicity potential (Choksakulnimitr et al., 1995). Hence, it is not surprising that DSR-loaded CS nanoparticles did not cause cytotoxic effects. However, compared to CS, ACS nanoparticles slightly decreased the cell viability (Figure 6). The addition of PEG chains on the surface of nanocarriers is widely used in drug delivery (Schattling et al., 2017). The chemical structure of PEG makes it inert in most cases, allowing the improvement of water solubility and the half-life of medications in the blood (Liu et al., 2017). However, recent studies showed that the PEG chain end groups and the molecular weight influence the cytotoxicity of PEG derivatives. In a study by Liu et al., PEG-based monomers with PEG methyl ether acrylate, showed noticeable cytotoxicity as compared to PEG oligomers (Liu et al., 2017). Acrylate end group is recognized as a reactive group, thus may lead to cell toxicity. Nonetheless, materials carrying this acrylate end group, including PEGDA polymers, are used extensively in bioengineering applications (Yoshii, 1996). In the results showed here, ACS nanoparticles were more cytotoxic than CS nanoparticles. In contrast to cell culture, the mucosal membrane in the human body is thick and covered with a layer of mucin gel that protects the cells from viruses and toxic bacteria (Johansson et al., 2013). Furthermore, this mucin layer in the small intestine is replaced every 4-6 h (Pepić et al., 2013). The acrylate end group is designed to attach to the mucin gel layer and allow improved drug absorption without damaging the tissue, as it is being washed after the time noted. In this viability assay, the cells were more exposed to cytotoxic effects because they were separated and did not form a continuous monolayer.

29

According to the viability assays, a working concentration of 30% (v/v) of particle suspension in buffer was chosen for the permeability studies, which corresponds to a polymer concentration of 0.2 mg/mL and a drug concentration of 0.025 mg/mL.

Figure 6. (A) Viability percentage of: (A) Caco-2 and (B) HT-29 MTX cell lines after 3 (white) and 24 h (grey) of incubation with different nanoparticle formulations (mean ± SD). (*) refers to statistically significant difference (p < 0.05), (**) refers to statistically significant difference (p < 0.005). The bars represent standard error of the mean, n = 6.

3.5 Drug Permeability using the Triple Cell Model In vitro cell models are useful tools to examine the permeability of drugs after oral administration and allow screening and predicting the fate of new carriers that deliver drugs in the body. Here we tested the drug permeability as mediated by nanoparticles using an intestinal triple co-culture cell model. This in vitro model has been previously established and found to be a valid model to mimic the epithelial barrier in the small intestine and evaluate permeability of different drugs and nanoparticulate systems [12, 16]. The model is composed of an epithelial layer of Caco-2 cells and mucus producing HT29-MTX cells to form a continuous cell monolayer with low electrical resistance, similar to the intestine (Li et al., 2013). The incorporation of Raji B cells allows inducing differentiation of a fraction of Caco-2 epithelial cells into M-cells, which play an important role in the intestinal translocation of nanoparticles [12, 16]. Figure 7A depicts a schematic representation of the triple co-culture cell model. DSR-loaded nanoparticles were placed in the apical chamber and the basolateral chamber was examined for DSR 30

presence in order to estimate its intestinal permeability. Since the triple co-culture cell model is characterized by the production of mucins, the thickness and continuousness of the secreted glycoprotein layer was studied by cutting the supporting membrane vertically and staining it with alcian blue. The results in Fig. 7B reveal positive staining for glycoproteins on the cell surface, which are, presumably, mucins [12, 16]. In general, the thickness of the mucin layer on the triple co-culture cell model varied between 7-15 µm (Figure 7B). However, the estimated thickness can suffer from measurement inaccuracies because of the preparation technique. Permeability profiles of DSR as mediated by ACS nanoparticles and CS nanoparticles, either fresh or re-dispersed in DDW after freeze-drying, were compared (Figure 7C). The amount of drug that was transported across the cell monolayer model increased with time for all the examined systems. Student’s t-test analysis revealed that CS nanoparticles increased the permeability of DSR as compared to its permeability in the free form (p < 0.005, Figure 7C). However, the cryoprotection caused a decrease in the final percentage of mass permeated using CS nanoparticles as carriers for DSR (p < 0.001, Figure 7C). Fresh and freeze-dried ACS nanoparticles did not improve significantly the final permeability of DSR (Figure 7C). The permeability profiles were fitted to a linear curve in order to obtain the slope, which was used to calculate Papp values using eq. 3 (Figure 7D). The Papp value of fresh CS nanoparticles was higher compared to fresh ACS nanoparticles (p < 0.05, Figure 7D). However after freeze-drying and re-dispersion, CS nanoparticles presented a lower Papp value as compared to the fresh state (p < 0.005, Figure 7D) that was similar to coefficients calculated for fresh and freeze-dried ACS nanoparticles.

31

Overall, only fresh CS nanoparticles improved the permeability rate compared to free DSR (p < 0.01, Figure 7D). For all studied samples, the permeability values were considerably lower as compare to those obtained with the in vitro Franz cell system (section 3.3), despite the fact that the mucin layer used in the Franz cell experiments is much thicker and imposes high resistance to mass transfer. This finding suggests that the resistance to mass transfer arises largely from the cell monolayer, and not from the mucin layer.

Figure 7. (A) Schematic illustration of the permeability assay; (B) Alcian blue mucus staining of vertical cuts of triple model membranes, scale bar of 50 µm; (C) Mass percentage of DSR permeated across the co-culture model monolayer over time; (D) Papp coefficient of the transport of DSR across the cell monolayer. (**) refers to statistically significant difference (p < 0.005). The bars represent standard error of the mean, n = 3.

32

TEER values of the monolayer were also monitored during the permeability assay and are presented in Figure 8. These values are used as indicators for the integrity of the cellular barriers before and during transport experiments of drugs or chemicals (Srinivasan et al., 2015). The results showed a decrease in TEER for the case of fresh CS nanoparticles (ANOVA, p < 0.0001), contrasting with no change in TEER of cell monolayers that were incubated with fresh ACS nanoparticles or free drug (Figure 8). ACS and CS nanoparticles that were freeze-dried with 30% (w/v) sucrose and redissolved in DDW caused an increase in the TEER values of the monolayer at the beginning of the experiment followed by a continuous decline (p < 0.001 and p < 0.05 respectively, Figure 8). This initial increase in TEER may be related with the electrolytic disturbance originated by the higher levels of sucrose introduced in the apical compartment, that eventually equilibrated along the experiment. Fresh ACS nanoparticles did not affect the TEER. However, freeze-dried ACS nanoparticles caused a decrease of 45% in TEER values after 3 h (p < 0.05, Figure 8A). Fresh CS nanoparticles caused a decrease of about 65% from the initial TEER value at the beginning of the experiment, while freeze-dried CS nanoparticles decreased TEER value in 53% resulting in a decrease in the drug permeability, as compared to fresh nanoparticles (p < 0.05, Figure 8B). TEER values are intrinsically related with the integrity of the cell monolayer, which is mainly maintained by tight junction formation between adjacent epithelial cells (Chen et al., 2015). CS is known to open tight junctions, allowing the penetration of several molecules and drugs (Khutoryanskiy, 2011). The modulating effects of epithelial tight junctions by CS were previously shown to be mediated by its cationic amine groups (Vllasaliu et al., 2010a). Furthermore, in a study performed by Driton Vllasaliu et al., CS-TPP nanoparticles showed a decrease in tight junction opening compared to CS solution, as a result of amine crosslinking forming the nanoparticulate system (Vllasaliu et al., 2010b). In this study, the influence of two parameters on tight junction modulation

33

was investigated, namely the acrylate modification of CS and the effect of freeze-drying. The results presented in Figure 8A suggest that fresh ACS nanoparticles did not affect the tight junctions, probably because of the acrylate modification and the drug incorporation, which reduced the amine groups available for interacting with the cells. Moreover, the bulky PEG chains could affect the interactions between the remaining amine groups and the cells. The results of nearly neutral zeta potential for fresh ACS nanoparticles (Figure 5B) indeed seem to support these claims. Conversely, CS nanoparticles presented markedly positive zeta potential, which backs up the availability of amine groups to interact with the cells, resulting in tight junction opening. Freezedried CS nanoparticles had a moderate effect compared to fresh particles, probably due to aggregation post freeze-drying and the presence of sucrose that can form hydrogen bonds with amine groups of the nanoparticles. Consequently, these amine groups are hindered and cannot interact with cells for tight junction opening. Previous studies have shown that mannitol induces osmotic stress, resulting in a reversible decrease in TEER of brain endothelial monolayers (Brown et al., 2004). This decrease correlates with an increase in permeability of different paracellular markers (Deli, 2009). The addition of freeze-dried ACS nanoparticles with sucrose to the apical side caused changes in the osmolarity of the monolayer, and perhaps the same mechanism of inducing osmotic stress applies in this case. It should be noted that in healthy gastrointestinal tract, the mucin layer varies along the intestine. Atuma et al. found that the average thickness of the total mucin layer in rat's small intestine is close to 250 µm (Atuma et al., 2000). Moreover, the thickness can change due to different conditions that affect the mucin secretion from goblet cells such as infections and chronic diseases (Kim and Ho, 2010). The mucin layer produced by the triple co-culture model (Figure 7C) is clearly much thinner. As demonstrated by the in vitro diffusion assays, CS nanoparticles are not able to diffuse through a thick mucus

34

layer. Thus, it is likely that CS nanoparticles are capable of opening the tight junctions in the model but will not be able to do so in vivo. High DSR permeability is correlated with low TEER values (Figure 7 and 8), likely because the opening of the cell tight junctions allows molecules to permeate the cell monolayer. The TEER values remained constant in the case of incubation with free drug and fresh ACS nanoparticles. However, the drug transport continued throughout the experiments. Similar phenomenon was reported in a study by Sakai et al., where fluorescein isocyanate dextran with similar molecular weight permeated through a monolayer of Caco-2 cells, but did not affect the TEER (Sakai et al., 2003). Diffusion processes are involved in the permeation of the drug across the barrier of the monolayer, hence there is a basal concentration that passes through even without tight junction opening.

35

Figure 8. TEER measured as a function of time for a triple co-culture model incubated with: (A) free drug and drug loaded ACS nanoparticles and (B) free drug and drug loaded CS nanoparticles. The bars represent standard error of the mean, n = 3. Connection lines between the symbols are added to guide the eyes.

It is noted that in vivo experiments must be conducted in order to determine the feasibility of this nanoparticulate system in delivering drugs to the small intestine. For clinical applications, we envision incorporating the dry NPs inside a protective coating of Eudragit® that will dissolve upon arrival to the small intestine. We expect that the nanoparticles and sucrose will be highly diluted upon their release in the small intestine, which will significantly decrease any toxic effects.

36

4. Conclusions In this work, we studied the effect of freeze-drying and cryoprotection on the physical and chemical properties of ACS and CS nanoparticles. Cryoprotection of both type of nanoparticles was achieved using sucrose and revealed that ACS nanoparticles are less sensitive to freeze-drying in terms of size. Freeze-dried ACS nanoparticles were found to partially bypass the mucin gel simulating a mucus layer. Moreover, the mucoadhesiveness of these nanoparticles in short contact times with porcine intestinal mucosa was not harmed by freeze-drying. The model drug DSR was successfully incorporated into CS and ACS nanoparticles. The effect of freeze-drying on the nanoparticles was assessed using the ability to modulate permeability of the drug across mucin gel and in a triple co-culture cell model mimicking the intestinal barrier. The diffusion of the drug through the mucin gel was greater when mediated by ACS nanoparticles compared to CS nanoparticles. However, cell monolayer permeability results showed that fresh CS nanoparticles could increase the translocation of DSR compared to the free drug, likely due to enhanced opening of tight junctions at the cell monolayer. Fresh ACS nanoparticles did not seem to extensively affect permeability due to tight junctions opening. A possible way to increase the drug transfer through the cell barrier is to use a multi-particle suspension composed of the two polymers ACS and CS that will be administered orally. Acknowledgements The authors wish to acknowledge the European Bioadhesion Community for the financial support for this work, COST Action CA 15216 ENBA (European Network of Bioadhesion Expertise). The article is also a result of the project NORTE-01-0145FEDER-000012, supported by Norte Portugal Regional Operational Programme (NORTE 2020), under the PORTUGAL 2020 Partnership Agreement, through the European Regional Development Fund (ERDF). This work was also financed by FEDER - Fundo Europeu de Desenvolvimento Regional funds through the COMPETE 2020 37

Operacional Programme for Competitiveness and Internationalisation (POCI), Portugal 2020, and by Portuguese funds through FCT - Fundação para a Ciência e a Tecnologia/ Ministério da Ciência, Tecnologia e Ensino Superior in the framework of the project "Institute for Research and Innovation in Health Sciences" (POCI-01-0145-FEDER007274). Andreia Almeida (grant SFRH/BD/118721/2016) and Maria Helena Macedo (SFRH/BD/131587/2017) would like to thank Fundação para a Ciência e a Tecnologia (FCT), Portugal for financial support.

5.

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Declaration of interests

‫ ܈‬The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

☐The authors declare the following financial interests/personal relationships which may be considered as potential competing interests:

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