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Biomaterials 33 (2012) 2791e2800

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Biomaterials

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The effect of chitosan on transcellular and paracellular mechanismsin the intestinal epithelial barrier

Rita Rosenthal a,*,1, Dorothee Günzel a,1, Caroline Finger a, Susanne M. Krug a,Jan F. Richter a, Jörg-Dieter Schulzke b, Michael Fromma, Salah Amasheh a

a Institute of Clinical Physiology, Charité, Campus Benjamin Franklin, Freie Universität and Humboldt-Universität, 12200 Berlin, GermanybDepartment of Gastroenterology, Div. of Nutritional Medicine, Charité, Campus Benjamin Franklin, Freie Universität and Humboldt-Universität, 12200 Berlin, Germany

a r t i c l e i n f o

Article history:Received 17 October 2011Accepted 19 December 2011Available online 9 January 2012

Keywords:Chitin/chitosanDrug deliveryEpithelial cellElectrophysiology

* Corresponding author. Institute of Clinical PhysioBenjamin Franklin, 12200 Berlin, Germany. Tel.: þ498445 4239.

E-mail address: rita.rosenthal@charite.de (R. Rosen1 Shared first authorship.

0142-9612/$ e see front matter � 2011 Elsevier Ltd.doi:10.1016/j.biomaterials.2011.12.034

a b s t r a c t

Chitosan is employed as an absorption enhancer for drug delivery strategies. Aim of this study was toinvestigate the rapid effects on barrier properties of the intestinal epithelial cell model HT-29/B6. Chi-tosan (0.005%) induced a fast decrease in transepithelial resistance (Rt) which was completely reversibleafter wash-out. Two-path impedance spectroscopy revealed that chitosan affects both, the paracellular(Rpara) and the transcellular (Rtrans) resistance. pH-dependence and inhibition of both effects by nega-tively charged heparin indicated a chitosan action only in the protonated form. The decrease in Rtrans wasmediated by activation of a chloride-bicarbonate exchanger involved in intracellular pH regulation. Thisactivation was coupled to the decrease in Rpara which was associated with an increase in ion permeabilityand permeability for paracellular flux markers up to 10 kDa. No effects on expression and subcellulardistribution of tight junction (TJ) proteins or the actin cytoskeleton were observed. Accordingly, inhi-bition of actinemyosin interaction, Ca2þ-dependent intracellular signaling, PKC, PI3K/Akt, MAP kinasep38, and endocytosis pathways did not impair the chitosan effect. These results suggest that the rapidand reversible absorption-enhancing chitosan effect is due to changes in intracellular pH caused by theactivation of a chloride-bicarbonate exchanger resulting in the opening of the TJ.

� 2011 Elsevier Ltd. All rights reserved.

1. Introduction

Drug absorption across epithelial barriers can occur via thetranscellular and the paracellular route. Whereas lipophilic drugscan permeate across cell membranes and thereby achieve thedesired therapeutic responses, hydrophilic molecules, such aspeptide and protein drugs, can not penetrate the cell membranesand can only be absorbed via the paracellular route to a minorextent. Thus, the lowabsorption results in an inadequate availabilityof such drugs. A promising approach to enhance the absorption ofhydrophilic macromolecular drugs is to co-administrate absorp-tion-enhancing agents that reversibly open the paracellular barrier.

Chitosan, a non-cytotoxic polymer, is currently a prominentcandidate for the development of drug targeting and drugdelivery strategies, as it has been reported as an absorption

logy, Charité Berlin, Campus30 8445 2792; fax: þ49 30

thal).

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enhancer affecting the barrier of a broad variety of different targetsincluding dermal [1], nasal [1e3], ocular [4e6], pulmonary [7], andgastrointestinal epithelia [8,9]. Moreover, chitosan-based micro-emulsions were tested in mice as potential candidates for drugdelivery into the brain [10].

In Caco-2 cells, an intestinal epithelial cell model derived fromhuman colorectal carcinoma, a reversible opening of TJs by chitosanresulted in a decrease in transepithelial electrical resistance (TER)[11e15] and an increase in paracellular marker fluxes [2,16e18].Some authors found a chitosan- or chitosan derivative-inducedreorganization of the actin cytoskeleton in Caco-2 cells [13,19,20],whereas others did not find any morphological changes in the actincytoskeleton [15,21]. Furthermore, changes in the distribution ofthe TJ protein occludin and the TJ-associated protein ZO-1 wereobserved in these cells [13e15,18,20,22]. Inhibition of proteinkinase C (PKC) was found to prevent the chitosan-mediateddecrease in TER and changes in the cellular localization of ZO-1 inCaco-2 cells [23]. Thus, in Caco-2 cells chitosan appears to activatea PKC-dependent signaling pathway that affects the integrity of theTJ. In addition, an effect of chitosan on claudin-1 was observed.Dorkoosh et al. showed that opening of the TJ is associated with an

R. Rosenthal et al. / Biomaterials 33 (2012) 2791e28002792

alteration in the position of claudin-1 within Caco-2 cells and anincrease in its expression [14].

Several studies demonstrated that the effect of chitosan on theparacellular permeability is dependent on its positive charge[11,12]. Chitosan is positively charged at pH values lower than7.0 (pKa of chitosan: 5.5e7.0). To overcome chitosan’s limitedeffectiveness as absorption enhancer at neutral and alkalinepH values as present in the small intestine, chitosan derivativeshave been developed, such as trimethyl chitosan (TMC) or Mono-N-carboxymethyl chitosan (MCC), which had comparable effectson the TJs [8,24].

In addition to soluble chitosan and chitosan derivatives, chito-san nanoparticles and chitosan-coated microspheres were devel-oped for macromolecular drug delivery. While chitosan solutionsimprove drug absorption through TJ modulation and increasedparacellular drug transport, drug delivery by means of chitosannanoparticles and chitosan-coated microspheres has been reportedto take place partly via endocytosis and transcytosis [25]. Moreover,chitosan nanoparticles or chitosan-coated phospholipid vesiclesalso caused a decrease in TER and an increase in permeability ofFITC-dextrans [18,26], which means that they also affect the para-cellular transport. Until now, most of the in vitro studies wereperformed on Caco-2 cells and focused on the effects of chitosan onthe paracellular pathway for drug delivery across an epitheliallayer. In this study, we used the highly differentiated cell line HT-29/B6, a sub-clone of the human colon cancer cell line HT-29[27], a well characterized model epithelium suitable for studyingepithelial and/or intestinal properties. HT-29/B6 cells are mucin-producing cells with inducible chloride secretion and a broadexpression of claudins within the TJ, which also determine thein vivo barrier in human epithelia [28]. The use of HT-29/B6monolayer as intestinal model epithelium has previously beenvalidated by parallel studies in native rat and human colonepithelia, which were in accordancewith the cell model concerningresponses to proinflammatory cytokines, secondary plantcompounds, and general physiological barrier regulation [29e31].

The aim of this study was to analyze chitosan effects on thisintestinal cell model, regarding the effects on the trans- and para-cellular pathway and to get more insight into the underlyingmolecular mechanisms.

2. Materials and methods

2.1. Chitosan solutions

Chitosan with low molecular weight (50e190 kDa based on viscosity) and75e85% deacetylation was used for all experiments. A 1.0% w:v stock solution ofchitosan was prepared in bath solution with 2.5% acetic acid to obtain a pH value ofpH 6.5 which is necessary for solubility of chitosan. For cell treatment, chitosan wasadded to the apical bath solution in final concentrations of 0.001e0.01%. For controlexperiments the solvent was added to the perfusion solutions to ensure that thechanges following application of chitosanwere not due to changes in pH value. Thus,the control cells were treated at the same time with exactly the same volume ofsolvent.

2.2. Cell cultures

Monolayers of the human colon carcinoma cell line HT-29 clone B6 [27] weregrown in 25-cm2 culture flasks containing DMEM (Biochrom, Berlin, Germany)supplementedwith 10% (v/v) foetal bovine serum,100mg/ml streptomycin, and 100U/ml penicillin (Biochrom). Cells were cultured at 37 �C in a humidified 5% CO2

atmosphere. For all investigations, epithelial cell monolayers were grown on cultureplate inserts (pore size 3.0 mm, effective area 0.6 cm2, Millicell-PCF, Millipore, Bed-ford, MA). Confluent cell layers were used on days 6e8 after seeding.

2.3. Lactate dehydrogenase release assay

To examine chitosan-mediated toxic effects on cultured monolayers, a lactatedehydrogenase (LDH) release assay was performed according to Madara and Staf-ford [32]. LDH, a cytosolic enzyme, was released into the culture mediumwhen cells

are injured. For this, the LDH level in 500 ml of the apical medium and in 500 ml of thecell lysate was determined under control conditions, 30 min and 2 h after chitosanapplication and 24 h after wash out of the polycation. Cell lysis was performed with2% Triton X-100 in culture medium for 20 min.

2.4. Electrophysiology

2.4.1. Ussing chamber measurementsFor electrophysiological measurements, inserts were mounted in Ussing

chambers specially designed for insertion of Millicell filters [27], and water-jacketedgas lifts were filled with 10 ml circulating fluid on each side. The standard bathsolution (Ringer’s) contained: 137.6 mM NaCl, 2.4 mM Na2HPO4, 0.6 mM NaH2PO4,5.4 mM KCl, 1.2 mM CaCl2, 1.2 mM MgCl2, 10 mM HEPES, and 10 mM D(þ)-glucose.The solution was constantly bubbled with 95% O2 and 5% CO2. The temperature ofthe bath solutionwas kept at 37 �C. Ussing chamber measurements were performedin the open-circuit mode unless otherwise stated. The transepithelial voltage (V)was measured in reference to the basolateral side. Transepithelial resistance (TER,Rt) was calculated fromvoltage changes (DV) induced by short current pulses (50 mA,0.3 s). All experimental data were corrected for the values of the empty filter and thebath solution. The equivalent short circuit current (ISC) was calculated by Ohm’s lawfrom V/Rt.

2.4.2. Dilution potential measurementsDilution potential measurements were performed for investigation of changes

in cation and anion permeability following treatment with chitosan. Dilutionpotentials were measured in the Ussing chambers with modified bath solution onthe apical or basolateral side of the epithelial cell layer. In themodified bath solution,50% of NaCl was iso-osmotically replaced by mannitol for determination of Naþ andCl� permeability. Ion permeabilities were calculated by means of the Goldman-Hodgkin-Katz equation, as described by Günzel et al. [33].

2.4.3. Two-path impedance spectroscopyTwo-path impedance spectroscopy was performed as described by Krug et al.

[34]. Basis of the method is a model describing the epithelial resistance (Repi) asa parallel circuit consisting of the transcellular resistance (Rtrans) and the paracellularresistance (Rpara). The subepithelial resistance (Rsub) is in series toRepi and is causedbythefilter support in cell culture experiments. Bothmembranes, apical andbasolateral,of the confluent cell layerare representedby resistors and capacitors inparallel (Ra, Ca,and Rb, Cb, respectively). Rtrans is the sum of Ra and Rb. Application of alternatingcurrent (35 mA/cm2, frequency range 1.3 Hze65 kHz) resulted in changes of epithelialvoltage which was detected by phase-sensitive amplifiers (402 frequency responseanalyzer, Beran Instruments; 1286 electrochemical interface; Solartron Schlum-berger). Complex impedance (Zreal, Zimaginary) values were calculated and plotted ina Nyquist diagram. This plot yields a semicircular arc as long as the electrical timeconstants (s¼ R$C) of apical and basolateral membranes are similar (sa z sb), whichwas true for the experiments of this study. Rtrans and Rpara were determined fromexperiments in which impedance spectra and fluxes of fluorescein as a paracellularmarker were obtained before and after chelating extracellular Ca2þ with EGTA. Thiscaused TJ to partly open by internalization of TJ proteins and to increase paracellularmarker flux which was inversely proportional to Rpara changes [34].

2.4.4. Flux measurementsAll flux studies were performed in Ussing chambers under voltage-clamp

conditions (0 mV). Unidirectional tracer flux measurements were performed with25 Bq/ml [3H]-mannitol (American Radiolabeled Chemicals, St. Louis, MO) from theapical to the basolateral side in Ussing chambers as reported previously [35]. Theperfusion solution also contained nonlabeled mannitol (10 mM). Three flux periodsof 10 min were analyzed under each condition (control, chitosan treatment,recovery). Upon initiation and completion, a 100 ml sample was taken from thedonor (apical) side, and 900 ml Ringer’s and 4 ml of Ultima Gold high flashpointliquid scintillation cocktail (Packard Bioscience, Groningen, Netherlands) wereadded. Samples (1 ml) of the receiving (basolateral) side were replaced with freshRinger’s and mixed with 4 ml of the liquid scintillation cocktail. All 5 ml sampleswere subsequently analyzed with a Tri-Carb 2100TR Liquid Scintillation counter(Packard, Meriden, CT).

For fluorescein fluxes, Ussing chambers were filled with 10 ml perfusion solu-tion at each side and 10 ml of fluorescein (100 mM) was added to the apical side.Basolateral samples (300 ml) were taken every 10 min before and after application ofchitosan (four flux periodes, respectively) and replaced with fresh perfusion solu-tion. Fluorescein fluxes were calculated from the apical tracer concentrationmeasured with a fluorometer at 535 nm (Spectramax Gemini, Molecular devices,Sunnyvale, CA). For dextran flux analysis the perfusion solution contained 10 mMunlabeled dextran on each side. After application of 100 mM FITC-labeled dialyzeddextran (FD-4, FD-10, FD-20 with 4, 10, and 20 kDa, respectively) to the apical side,basolateral samples were collected every 30 min and replaced by fresh perfusionsolution. Three flux periods of 30 min were analyzed before and after application ofchitosan. Dextran fluxes were calculated from the amount of FITC-dextran in thebasolateral compartment which was measured with a fluorometer at 520 nm(Spectramax Gemini).

R. Rosenthal et al. / Biomaterials 33 (2012) 2791e2800 2793

2.5. Immunocytochemistry

Confluent cell monolayers grown on culture plate inserts were incubated for30 min with chitosan or solvent before processing for confocal microscopy. Cellswere rinsed with PBS, fixed with methanol, and permeabilized with PBS containing0.5% Triton X-100.

Primary antibodies employed were anti-occludin, anti-ZO-1, and antibodiesraised against different claudins (Zymed Laboratories, Invitrogen Immunodetection,South San Francisco, CA).

Concentrations of primary antibody were 10 mg/ml. Secondary antibodies AlexaFluor 488 goat anti-mouse, and Alexa Fluor 594 goat anti-rabbit (both used atconcentrations of 2 mg/ml) were purchased from Molecular Probes (Eugene, OR).DAPI (40 ,6-diamidino-2-phenylindole dihydrochlorid) was used to stain cell nuclei,and FITC-phalloidin to stain F-actin. Fluorescence images were obtained witha confocal microscope (LSM 510 meta, Carl Zeiss, Jena, Germany) using excitationwavelengths of 543, 488, and 405 nm, respectively.

Fig. 1. Effect of chitosan on cell viability of HT-29/B6 cells. Cell viability was tested withthe lactate dehydrogenase (LDH) release assay. As an indicator of cell disruption theLDH release from the cells was measured in apical medium and in whole cell lysates.The total LDH content of the cells was set as 100% and LDH released into the mediumwas calculated.

2.6. Freeze-fracture electron microscopy

Freeze-fracture analysis was carried out as described before [36]. Cells grown oncell-culture inserts were treated with chitosan or acidified control solution for 1 h,fixed with phosphate-buffered glutaraldehyde (2%), incubated in glycerol (10% and30% (v/v)), and finally frozen in liquid-nitrogen-cooled Freon 22. The preparationswere fractured at �100 �C and shadowed with platinum and carbon in a vacuumevaporator (Denton DV-502, Denton Vacuum, Cherry Hill, NJ). The replicas of thesamples were bleached with sodium hypochloride, mounted on special coppergrids, and analyzed using a Zeiss 902A electron microscope (Carl Zeiss NTS, Ober-kochen, Germany) with a digital camera iTEM Veleta (Olympus Soft Imaging Solu-tions, Münster, Germany).

Freeze-fracture electron micrographs (51,000� magnification) of all TJregions in which an apical and a contra-apical strand of the meshwork could beclearly distinguished were used for the morphometric analysis. The distancebetween the apical and the contra-apical strands was defined as the meshworkdepth. Vertical grids were drawn at 200 nm intervals perpendicular to the mostapical TJ strand. The number of horizontal strands within the main TJ meshworkwas counted at intersections with grid lines. Strand discontinuities of >20 nmwithin the horizontally oriented strands were defined as ‘breaks’. Dependent onthe linearity of strand formation ‘particle type’ and ‘continuous’ strands weredistinguished.

2.7. Chemicals

All chemicals, unless otherwise noted, were purchased from Sigma Aldrich.Antibodies raised against claudins and occludin were purchased from ZymedLaboratories (San Francisco, CA). Secondary antibodies, Alexa Fluor 488 goat anti-mouse and Alexa Fluor 594 goat anti-rabbit were purchased from MolecularProbes (MoBiTec, Göttingen, Germany).

2.8. Statistical analysis

Data are expressed as means � standard error of the mean. Statistical analysiswas performed by using Student’s t-test and the Bonferroni-Holm correction formultiple comparisons. p < 0.05 was considered significant. Significance levels aredenoted n.s.¼ not significantly different, *¼ p< 0.05, **¼ p< 0.01, ***¼ p< 0.001.The number (n) refers to the number of experiments.

3. Results

3.1. Analysis of cell viability

Since it is known that cationic polymers exert cytotoxic effects,the cell viability under control conditions and after 30 min and 2 hexposure to 0.005% and 0.001% chitosan was tested with the LDHrelease assay. The activity of LDH, a cytosolic enzyme released intothe medium when cells are injured, is evaluated under toxicconditions. As shown in Fig. 1, HT-29/B6 cells exposed to bothchitosan concentrations for 2 h had a strong, dose-dependentincrease in extracellular LDH activity. After 30 min only the expo-sure to 0.01% chitosan increased LDH activity, whereas 0.005% wasnot different from control values. A complete recovery, even afterexposure to 0.01% chitosan, could be observed within 24 h incu-bation in medium (n ¼ 3e7).

3.2. Transepithelial resistance measurements

Treatment of HT-29/B6 monolayers with chitosan applied to theapical compartment of the Ussing chamber resulted in a fast anddose-dependent reduction of transepithelial resistance (Rt) to17.3 � 3.4% (n ¼ 14) with 0.005% and 4.7 � 0.3% (n ¼ 9) with 0.01%after 30 min in comparison to the values before application ofchitosan. Treatment with acidified control solution or 0.001% chi-tosan did not significantly affect Rt (99.1 � 0.7% (n ¼ 13) and98.1 � 1.0% (n ¼ 6), respectively) (Fig. 2A). Complete recovery of Rt

values after wash out of the polymer occurred within 6 h for bothconcentrations (Fig. 2B). Longer incubation (60 min) with chitosandid not enhance the effect on Rt (14.8 � 1.6% (n ¼ 24) with 0.005%chitosan and 3.0 � 0.1% (n ¼ 5) with 0.01% chitosan). Since thebreak-down of Rt after incubation with 0.01% chitosan could partlybe due to an effect on cell viability, all further investigations wereperformed with 0.005% chitosan.

Two-path impedance spectroscopy was performed to analyzewhether the effect on Rt was due to a decrease of the transcellularresistance (Rtrans), reflecting theapical andbasolateral cellmembrane,or the paracellular resistance (Rpara), reflecting the resistance of the TJ.AsshowninFig. 2C,preincubationwithchitosan resulted inadecreaseof both, Rtrans and Rpara (n ¼ 4) in comparison to control conditions(n¼ 5). Thus, chitosan affects the transcellular conductance aswell asthe TJ. The increase in paracellular conductance was caused by anincrease in ion permeability (control: PNa 4.3 � 0.4.10�6 cm/s, PCl3.0 � 0.3.10�6 cm/s, chitosan: PNa 12.5 � 1.3.10�6 cm/s, PCl10.4 � 1.3.10�6 cm/s, n ¼ 8, respectively) and a loss in cation prefer-ence. The permeability ratio PNa/PCl changed from 1.46 � 0.04 incontrol cells to 1.22� 0.02 in chitosan-treated cells.

3.3. Paracellular marker permeability

Fluxes of paracellular markers of molecular weights between182Da and20kDaweremeasured to analyze the effect of chitosan onthe barrier function of the TJ (Fig. 3). As seen for 3H mannitol, thepermeability increased after application of chitosan and decreased tocontrol values after wash out of the substance. A strong increase inpermeability after addition of chitosan could be observed for 4 kDadextran, and the increase in permeability for 10 kDa dextranwas stillmarkedly detectable. In contrast, chitosan had no effect on thepermeability for20kDadextran, andcarrier solutionalonecontaining

Fig. 2. Effect of chitosan on transepithelial resistance. A. Dose-dependent effect ofchitosan on transepithelial resistance and recovery (after 6 h) following wash-out ofthe polycation (n ¼ 6e14). B. Original recording of resistance over time under controlconditions (acidified control) and after application and wash-out of 0.005% chitosan. C.Two-path impedance spectroscopy. Chitosan decreases transepithelial resistance by aneffect on transcellular and paracellular resistance.

Fig. 3. Effect of chitosan on the permeabilities of paracellular flux markers. A.Permeability measurements of paracellular markers revealed an increase in thepassage of mannitol (180 Da) in the presence of chitosan which was completelyrecovered after wash-out of the polycation. B. Permeability of 4 and 10 kDa dextranwas also increased in the presence of chitosan, whereas the passage of 20 kDa dextranwas unchanged.

R. Rosenthal et al. / Biomaterials 33 (2012) 2791e28002794

acetic acid (0.0125%) did not increase permeability at all. These datademonstrate a size-selective increase in permeability for paracellularmarkers up to 10 kDa in HT-29/B6 cells treated with chitosan.

3.4. Effect on the actin cytoskeleton

To test whether the chitosan effect on the TJ is associated withan effect on the actin cytoskeleton, an immunostaining against F-

actinwas performedwith chitosan-treated cells at the maximum ofthe chitosan effect and control cells at the same time. As shown inFig. 4A, the actin cytoskeleton was not changed by treatment withchitosan. This result is consistent with the missing effect of inhib-itors of actinemyosin interaction like ML-7 (10�5 M, apical andbasolateral, n ¼ 9), a myosin light chain kinase inhibitor, and Y-27632 (10�5 M, apical and basolateral, n¼ 3), a Rho kinase inhibitor.Furthermore, preincubation of the cells with BAPTA-AM (2$10�5 M,apical and basolateral, n ¼ 8), a membrane-permeable Ca2þ

chelator, does not diminish the chitosan effect on the cells (Fig. 4B).

3.5. Effects on TJ proteins

For further analysis of the chitosan effect on the paracellularbarrier function, the expression and subcellular distribution of TJproteins was investigated in control cells and chitosan-treated cells.

No difference was detected in the membrane expression ofdifferent TJ proteins in control cells and cells treated with chitosan(Fig. 5A). Thereupon, the subcellular localization of these TJproteins was analyzed, but no changes could be observed for any ofthe proteins after chitosan treatment, here shown for claudin-4 andthe TJ-associated protein ZO-1 (Fig. 5B). Immunostainings of otherTJ proteins were also performed, and no changes could be detected.Also, no alteration in the localization of tricellulin, which

Fig. 4. Effect of chitosan on the actin cytoskeleton. A. Staining of F-actin revealed thatthe actin cytoskeleton was unaffected by chitosan. B. Inhibitors of the actinemyosininteraction like ML-7 and Y-27632 and the membrane-permeable Ca2þ chelatorBAPTA-AM do not attenuate the chitosan-mediated decrease in Rt.

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delimitates the macromolecular passage via the paracellular route,could be observed in chitosan-treated cells (Fig. 5C). Thus, thechitosan effect on the TJ barrier could not be due to a redistributionof TJ components.

This result is consistent with the fact that inhibitors of endo-cytotic pathways that could affect the subcellular distribution oftight junction proteins like chlorpromazine (50 mg/ml apical),methyl-b-cyclodextrin (50 mg/ml apical and basolateral), anddynasore (80 mM apical and basolateral) did not affect the chitosan-mediated decrease in Rt (Fig. 5D). Furthermore, inhibition ofintracellular signaling pathways which are known to be involved inTJ protein expression, distribution, degradation or function (e.g. viaphosphorylation) like PKC, PKA, PKG, PI3K/Akt kinase, and MAPkinase p38 pathways, did not attenuate the effect of chitosan on Rt

(Fig. S1).

3.6. Effect on transepithelial ion transport

Since the decrease in Rtrans is due to the activation of trans-epithelial ion transport, experiments with inhibitors were per-formed to determine the ions involved in this process. BaCl2 (5 mMapplied basolateral), a potassium channel blocker and in thisfunction an inhibitor of chloride secretion in HT-29/B6 cells, did notattenuate the chitosan effect on Rt and ISC. NFA (niflumic acid,0.5 mM apical and basolateral), a chloride channel blocker, reduced

the chitosan-activated ISC (Fig. 6A), whereas no effect on Rt could beobserved. In contrast DIDS (4,40-Diisothiocyano-2,20-stilbenedi-sulfonic acid, 100 mM apical), an anion exchange inhibitor,completely blocked the chitosan-induced activation of ISC and therapid chitosan effect on Rt (n ¼ 12, Fig. 6A,B). This fact suggested aneffect of DIDS on both, the transcellular as well as the paracellularpathway. To test this, fluxes of the paracellular marker fluoresceinin the presence of DIDS before and after application of chitosanwere investigated (Fig. 6C). Fluorescein fluxes increased, firstslowly, later faster, after addition of chitosan (n ¼ 5), this increasewas totally blocked in the presence of DIDS (n¼ 4). A comparison ofthe increase in fluorescein flux and conductance after chitosanapplication revealed a faster increase in conductance, indicatingthat the activation of ISC occurs before opening of the TJ. Themissing effect of chitosan during equilibration of the perfusionsolution with oxygen instead of carbogen (n ¼ 10) suggested theinvolvement of a chloride-bicarbonate exchanger (Fig. 6D). This isconsistent with the finding that no chitosan effect on ISC and Rt

could be observed when cooling the bath solution to 8 �C (n ¼ 6).

3.7. Charge dependence of the chitosan effect

Since it is known that the effect of chitosan on the apicalmembrane permeability could be reduced by heparin [20], it wastested whether heparin, a negatively charged polysaccharide, hasan impact on the chitosan effect on Rt and ISC. After preincubation ofHT-29/B6 monolayers with heparin (5000 IU) at the apical side ofthe cells, the chitosan-mediated decrease in Rt and activation of theanion exchanger was completely prevented (n ¼ 8, Fig. 7A,B)whereas the application of heparin at the basolateral side has noeffect. Moreover, apical application of heparin at the maximum ofthe chitosan effect strongly attenuated the chitosan-induced effect(Fig. S2A, B). These findings indicate that the effects of chitosan ontranscellular and paracellular parameters are mediated via itspositive charge. Another indication for a charge-dependent effect isthe fact that the efficiency of chitosan was dependent on the pH ofthe extracellular solution, an increase in the pH value from pH 6.60to 7.05 inhibited the chitosan effect almost completely (Fig. 7C,D).

3.8. Effect on TJ ultrastructure

Freeze fracture electron microscopy was employed for analysisof chitosan effects on TJ ultrastructure. However, no changes withinthe TJ network were detected (Table S1).

4. Discussion

4.1. Chitosan is not toxic and causes reversible effects on HT-29/B6epithelial cells

Chitosan, due to its antimicrobial properties, has a variety ofapplications such as wound healing, tissue engineering and tissuerepair, and furthermore is an ingredient of a variety of nutritionaladditives. The molecule derives from the chitin molecule, and itsatisfies all requests for an absorption enhancer as it is not toxic inthe effective concentrations and it rapidly acts in a reversible way,with a partial loss of paracellular barrier function only to molecules< 20 kDa and thus avoiding unwanted penetration of bacterialantigens and other harmful immunoreactive substances throughthe gut wall.

As a valuable epithelial in vitro model for analysis of intestinalbarrier function, HT-29/B6 cells have been established [27]. Advan-tages of this model are a high reproducibility in accordance withparallel studies in rodent and human intestinal tissues, and espe-cially an endogenous expression of predominant gastrointestinal TJ

Fig. 5. Effect of chitosan on TJ proteins. A. Western blot analysis performed with membrane lysates of control and chitosan-treated cells revealed no effect of chitosan on TJ proteins.B. Immunofluorescence staining was performed for all TJ proteins shown in A and indicated that chitosan does not affect the subcellular distribution of TJ proteins, here shown onlyfor Cldn-4 and ZO-1. C. The localization of tricellulin in the tricellular TJ is not changed after chitosan treatment. D. Inhibitors of endocytosis pathways like chlorpromazine (n ¼ 6),methyl-b-cyclodextrin (n ¼ 3), and dynasore (n ¼ 4) have no effect on the chitosan-mediated decrease in Rt.

R. Rosenthal et al. / Biomaterials 33 (2012) 2791e28002796

Fig. 6. Effect of ion channel inhibitors on the chitosan effect on ISC, Rt, and fluorescein flux. A. Summary of the chitosan effect on ISC (left) and Rt (right) after preincubation withBaCl2, NFA, and DIDS. B. DIDS completely inhibited the chitosan-induced activation of ISc (left) and decrease in Rt (right). C. the increase in the flux of the paracellular markerfluorescein after addition of chitosan compared with the chitosan-induced conductance increase. D. No chitosan effect on ISC (left) and Rt (right) occurred during gassing of theperfusion solution with oxygen. After changing to carbogen, the effect on ISC and Rt could be observed.

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Fig. 7. Charge dependence of the transcellular and paracellular chitosan effect. A, B. Inhibition of the chitosan effect by heparin. Preincubation of HT-29/B6 cells with negativelycharged heparin completely inhibited the chitosan-mediated decrease in Rt (A) and the activation of ISC (B). C, D. Influence of the pH value on the chitosan effect. Chitosan-mediateddecrease of Rt at pH values of the extracellular solution before chitosan application between 6.60 and 7.05. A fast and strong decrease occurred at pH 6.60, whereas nearly no effectoccurred at pH 7.05 (C). Chitosan-mediated activation of ISC with a strong effect at pH 6.60 and nearly no effect at pH 7.05 (D). The answer of the control cells is always independentof the pH value.

R. Rosenthal et al. / Biomaterials 33 (2012) 2791e28002798

proteins such as occludin, claudin-1, -2, -3, -4, -5, -7, and -8[30,31,37]. In our experiments, LDH and Rt measurements revealedthat a concentration of 0.005%was not toxic for the cells and causeda strong decrease in Rt which was completely reversible, whereaslower concentrations (0.001%) had no effect on Rt and higherconcentrations (0.01%) were cytotoxic for the cells.

4.2. Chitosan effect on the paracellular pathway

Two-path impedance spectroscopy experiments revealed thatthe chitosan-mediated decrease in Rt of HT-29/B6 cells was causedby a decrease of Rpara as well as Rtrans. Major molecular determinantfor Rpara is the TJ, which is primarily composed of tetraspanproteins, occludin [38], the claudin family [39], and tricellulin [40]which are associated with the cytoskeleton via scaffolding proteins,such as ZO-1. Western blot analysis of membrane lysates of controland chitosan-treated cells revealed no difference in the expressionof TJ proteins and ZO-1. This is confirmed by immunostainings,which showed no changes in the subcellular localization of TJproteins and ZO-1, and no effect on the actin cytoskeleton aftertreatment with chitosan. The lacking effect of chitosan on subcel-lular TJ protein localization corresponds with missing effects ofendocytotic pathway inhibitors on the chitosan-mediated decreasein Rt. In contrast to these results on HT-29/B6 cells, other authorsobserved an association between the decrease in Rt and changes in

TJ proteins in Caco-2 cells. In these cells, some authors found anaccumulation of occludin at the cell borders [14,22], whereas othersobserved a decrease in the peripheral staining for occludin aftertreatment with chitosan [13]. The ZO-1 staining revealed a loss inthe continuous staining along cellecell contact points after incu-bation with chitosan [13,15,22].

While the subcellular localization of occludin and ZO-1 wasintensively investigated, information on the role of the barrier-determining claudins is scarce. Dorkoosh et al. found a higherexpression of claudin-1 within the TJ after treatment with tri-methyl chitosan [14], and a recently published study demonstrateda redistribution of claudin-4 in Caco-2 cells as a result of chitosantreatment [41].

In addition, different effects of chitosan on the actin cytoskel-eton were observed in Caco-2 cells, an accumulation of actin at thecell borders [14,22] and, in contrast, also a reduction in the F-actinstaining [13,20]. In HT-29/B6 cells, the F-actin cytoskeleton isunchanged after 1 h of chitosan treatment at the maximal decreasein Rt. Furthermore, inhibitors of the actinemyosin interaction andchelation of intracellular calcium did not diminish the chitosaneffect on Rt.

Taken together, in HT-29/B6 cells, the rapid chitosan-mediateddecrease in Rpara and the increase in permeability of paracellularmarker molecules seems to be rather not associated with analtered expression and localization of TJ proteins and ZO-1 or

R. Rosenthal et al. / Biomaterials 33 (2012) 2791e2800 2799

morphological changes in the F-actin cytoskeleton but relies to another mechanism.

As a consequence of TJ opening indicated by the decrease in Rt,the permeability to paracellular markers such as mannitol, FD-4,and FD-10 increased, whereas no change could be observed forFD-20. This is a further difference to Caco-2 cells, in whicha minor increase in the paracellular transport of FD-20 wasobserved after chitosan treatment [20,42]. Thus, chitosan causesa rapid, reversible and size-selective opening of the TJ formacromolecules. These are essential requirements for the appli-cation as an absorption enhancer, restricting an unwantedpassage of macromolecules which otherwise would lead toadverse reactions such as immune responses. As an importantdeterminant for the paracellular passage of macromolecules, tri-cellulin has been reported [36], which is located in tricellular TJs,the meeting point of three cells. Thus, we also tested thehypothesis that a redistribution or endocytosis of tricellulin couldexplain the chitosan-mediated increase in permeability tomacromolecules. However, no effect on tricellulin expression andon the subcellular localization within the tricellular junctioncould be observed in HT-29/B6 cells.

4.3. Intracellular signaling pathways

Inhibitor studies revealed that PKC-, PKA-, PKG-, PI3K-, andMAPkinase p38-dependent pathways are not involved in the chitosan-induced decrease in Rt and activation of the ISc. Thus, thesekinases do not participate in the chitosan effect in HT-29/B6 cells,whereas in Caco-2 cells treatment with chitosan results in theactivation of PKC-dependent signal transduction pathways whichaffect TJ integrity [23]. MAP kinase was also not involved in Caco-2cells. Effects on the phosphorylation status of kinases, however,have to be takenwith caution in case of the rapid chitosan effect, asthis starts immediately after application and therefore might beexplained by a primarily direct interaction. As a response tomarkedeffects on barrier integrity, activation of signaling pathways mightfollow, but might even represent a compensatory reaction of thecells.

4.4. Chitosan effect on transcellular ion movement

Ussing chamber measurements in combination with inhibitorstudies revealed that chitosan induces the transient activation ofa chloride-bicarbonate exchanger contributing to the decrease inRtrans. Interestingly, inhibition of this transporter with DIDScompletely abolished the chitosan-mediated decrease in Rt indi-cating not only an effect on Rtrans but also on Rpara. This wasconfirmed by the inhibitory effect of DIDS on the chitosan-inducedincrease in fluorescein fluxes. Furthermore, the comparison of thetime course in conductance and fluorescein flux increase afterchitosan application suggested that the activation of the anionexchanger occurs first, and with a delay, the TJ is affected. Thus,activation of the anion exchanger seems to be the prerequisite foropening of the paracellular pathway. A further hint for theinvolvement of an energy-dependent transport process was thefact that chitosan had no effect at low temperature. The involve-ment of endocytosis could be excluded by the missing effect ofendocytosis inhibitors. Since the chloride-bicarbonate exchanger isinvolved in cytosolic pH regulation, it seems probable that changesin intracellular pH cause an effect on the TJ barrier, maybe by aneffect on scaffold proteins. Even though the paracellular effect iscrucial for the use as absorption enhancer, the transcellular effect ofchitosan has to be kept in mind since unwanted secondary actionscould develop during application.

4.5. Chitosan effect on trans- and paracellular parameters ismediated by the positive charge

A further hint for the connection between trans- and para-cellular chitosan actions was the charge dependence of both effects.The loss of the chitosan effect on Rt and ISC after preincubation ofthe monolayers with negatively charged heparin and the restora-tion of Rt and deactivation of the ISC by addition of heparin at themaximum of the chitosan effect indicate that the rapid andconsistent chitosan effect is primarily mediated by its positivecharge. Interestingly, addition of heparin after the onset of thechitosan effect caused a rapid decrease of ISC probably due tothe binding of heparin on membrane-bound chitosan resulting inthe deactivation of the chloride-anion exchanger. In contrast, theincrease in Rt occured much slower which could be attributed toa long-lasting recovery of the intracellular homeostasis which inturn restores the TJ barrier function. Furthermore, both effects aredependent on the pH value of the perfusion solution. Increasing thepH and by that way deprotonating the cation diminished the chi-tosan effect. Thus, interaction of positively charged chitosan withanionic components on the surface of the epithelial monolayerseems to be the initial step in the absorption-enhancing effect ofthe polycation. A similar mechanism might also be responsible forthe rapid chitosan effect in Caco-2 cells, as both, inhibition of theabsorption enhancing effects of chitosan by the addition of heparin[20], and pH-dependent effects of chitosan derivatives on Rt [43]were reported. Similar effects on transcellular and paracellularmechanisms were induced by the polycation poly(L)-lysine, whichinduced a dose-dependent increase in transepithelial conductancein tight (MDCK I) and leaky (MDCK II) MDCK (Mardin-Darby caninekidney) epithelial monolayers, an increased transepithelial flux ofparacellular markers, and the activation of short circuit current.Similar effects on monolayers of T84 and HCT-8 human intestinalcells were found indicating that polycation action may be generalfor a range of epithelial types [44].

4.6. Chitosan alters paracellular permeability independent of TJprotein changes

The chitosan-mediated decrease in Rt and increase in para-cellular permeability seems to be similar in Caco-2 and HT-29/B6cells even though both cells differ in a variety of properties.Major differences have been shown for example for their endoge-nous expression pattern of TJ proteins, with only marginal orcompletely lacking expression of e.g. claudin-2 and -5 in confluentmonolayers in contrast to HT-29/B6 [28,31,45].

Since the cell lines differ in the claudin composition whichdetermines barrier properties of the paracellular pathway, it seemsprobable that a similar mechanism is the first step leading to thebreak-down of Rpara. Such a mechanism could be the change in thecytosolic pH resulting from the activation of an anion exchangerwhich in turn might affect distinct TJ proteins. As a consequence ofchitosan binding, a redistribution or degradation of TJ proteins hasbeen reported to occur which may account for an additional para-cellular effect. In Caco-2 cells, chitosan treatment induced a redis-tribution of claudin-4 intracellularly and a subsequent degradationin lysosomes. The recovery was accompanied by an increase inclaudin-4 gene transcription [41]. Furthermore, a full recovery ofthe chitosan effect after wash out was prevented by simultaneousaddition of the protein synthesis inhibitor cycloheximide [22].

The assumption of a similar initial mechanism is based on theobservation that chitosan-induced a fast decrease in Rt in differentcell lines, such as different intestinal epithelial cell lines, and also inhigh- and low-resistance MDCK cells, and this effect was alwaysconnected with the activation of a transepithelial current (Fig. S3).

R. Rosenthal et al. / Biomaterials 33 (2012) 2791e28002800

This is in line with the assumption that the ubiquitous rapid andconsistent effect of chitosan on paracellular barrier propertiesmight not depend on changes of just one member of the TJ proteinsbut rather on the whole TJ complex.

5. Conclusions

Chitosan induces a fast and reversible decrease in Rt in HT-29/B6cells, which is mediated by a specific binding of the polycation tonegative charges on the surface of the membrane. The bindingcauses the activation of a chloride-bicarbonate exchanger, which isthe prerequisite for the opening of the TJ barrier and the increase inthe permeability for ions and paracellular flux markers up to10 kDa. The chitosan effect on TJ barrier properties might mediatedby changes in cytosolic pH.

Acknowledgements

We thank Detlef Sorgenfrei, In-Fah Lee, and Anja Fromm fortheir expert technical assistance. This work was supported bygrants of the Deutsche Forschungsgemeinschaft (DFG FOR 721/2)and the Sonnenfeld-Stiftung Berlin.

Appendix. Supplementary material

Supplementarymaterial associatedwith this article canbe found,in the online version, at doi:10.1016/j.biomaterials.2011.12.034.

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