Nanocellulose hydrogels as bio-interface analogs for studying nanomaterial transport and accumulation

15 16 Nanomaterials have been proposed as drug delivery vehicles to enhance targeting and efficiency 17 of traditional and novel therapeutics and have subsequently been studied for potential ecotoxicity. 18 Previous studies have identified size, surface charge, and volume exclusion as factors that 19 influence nano material diffusion and retention. However, there is little accepted or successful 20 quantification of how these parameters influence nano material penetration relative to biological 21 adaptation and biological response . Part of the challenge is the response of living biological 22 interfaces to many of these nanomaterial delivery vehicles and nanosized drugs. This study aimed 23 to emulate key physicochemical barriers to diffusion found in living biomaterials by developing a 24 tunable, synthetic hydrogel. Through the controlled exposure of 150 kDa and 2 MDa nanodextrans 25 with neutral and negative surface charge , we evaluated the system’s ability to emulate three core 26 physicochemical features often implicated in biofilm -associated transport resistance: size 27 exclusion, charge interactions, and volume exclusion. We demonstrated a 30% statistically 28 significant decrease in partition coefficients for 2 MDa nanodextran from 150 kDa nanodextran, 29 confirming the ability of the nanocellulose-based microcaps to mimic the permeability of hydrated 30 biomaterial matrices. These findings reflect patterns observed in , for example, living biofilm 31 studies, where size-based diffusion hinderance is commonly reported, but charge-based interaction 32 and volume exclusion are more context-dependent. This controllable system can be coupled with 33 in silico modeling to understand interfacial transport phenomena for nanomaterial -biomaterial 34 interactions. 35 36

Biointerface materials, nanoparticles, antibiotic recalcitrance, transport phenomena 37 38 Graphical Abstract 39 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 2 40 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 3 1. Introduction 41 Biological interfaces, like microbial cells and biofilms, are more complex than 42 engineered porous membranes[1] and subsequently the transport relationships are more complex. 43 For example, the liquid-surface biointerface typically contains soft-layers comprised of fats, 44 sugars, and proteins of various charges [2]. The interior of the biointerface exhibits heterogenous 45 porosity and tortuosity [3], heterogeneous mechanical compliance, heterogenous hydrophilicity, 46 and heterogeneous charge distribution due to the presence of cells and extracellular components 47 [4]. These complex structures increase the difficulty of decoupling the impacts of charge, size, 48 geometry, and chemistry on the transport of novel drug and drug delivery vehicles through these 49 biological interfaces. In microbial biointerfaces, like bacterial biofilms [5-8], this complexity is 50 increased by the dynamic response that can occur at similar time scales to the time for total 51 penetration. The dynamic response from bacteria can include extracellular matrix production of 52 polymers like alginate, nanocellulose, proteins like flagellin [9, 10], and secretion of extracellular 53 DNA. These properties contribute to enhanced virulence [11] and treatment failure [12] in 54 diseases such as cystic fibrosis. Consequently, new therapeutic strategies, including the use of 55 nanomaterials as drug delivery vehicles, have been proposed to enhance penetration and efficacy 56 within biofilms[13-15]. 57 These new therapeutic strategies will come with their eventual environmental 58 dissemination. While engineered nanomaterials cannot be studied as a uniform class[16], 59 experimental and computational platforms that account for their fate and transport must be 60 created [17-21]. Periphyton biofilms are the primary sink for nanomaterials in estuary 61 environments [22]. The primary factors which effect ENM ecotoxicological parameters such as 62 toxicity and bioaccumulation for a given ENM-biofilm-environment system can be evaluated. 63 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 4 What emerges is a set of four pairwise factors of the biofilm-ENM system of interest which 64 shapes bioaccumulation: size (particle/pore) [23-29], charge (surface/ECM) [24, 25, 27, 30-32], 65 chemistry (surface/ECM) [25, 31, 33], and hydropathy (surface/ECM) [27, 30, 34]. 66 Previous studies on the impact of mechanical stress on biointerfaces have used alginate 67 since it is an essential component of some biofilms and has facile preparation [35-38]. However, 68 alginate has 1 -2 orders of magnitude higher storage and loss moduli than natural biofilms [37]. 69 Additionally, alginate has different chemical absorption properties, specifically it is less able to 70 absorb divalent metallic cations than natural biofilms [37]. We have previously shown the 71 nanocellulose preparation used here has mechanical and certain chemical properties in range 72 with natural biofilms [39]. Nanomaterial exclusion from biofilms has been proposed to be a 73 function of the various heterogeneous matrix components alginate, nanocellulose, proteins; the 74 matrix size; commensal phage; bacterial cells, particularly their charge, and the channels in the 75 biofilm [4, 30, 40]. As heterogeneity is difficult to achieve synthetically, this may require 76 systematic study. For example, the importance of alginate on antibody binding in P. aeruginosa 77 biofilms was discovered through adding alginate back alginate deficient mutants. Recent work 78 on diffusion in alginate showed the impact of matrix cross-linking on size-exclusion [41]. 79 We propose the biofilm matrix and cells with their respective charges can be studied with 80 synthetic systems. We synthesized microcaps from nanocellulose that are modified with divalent 81 calcium ions as the biofilm matrix. We added neutral and charged microspheres to represent cells 82 and their respective binding affinities. We tested the ability of charged and neutral nanodextran 83 of different molecular weights to accumulate into the defined nanocellulose matrix. 84 This study tested the ability of a microcap synthesis method to replicate the important 85 effects of a biofilm on species diffusion: (a) the size-effect, (b) the volume-exclusion effect, and 86 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 5 (c) the attachment effect. These important biofilm effects were mapped on to three hypotheses. 87 We hypothesize that if the size-effect is replicated that larger diffusing species would accumulate 88 at lower concentrations in the matrix. We hypothesize that if the volume-exclusion effect is 89 replicated, that diffusing species would accumulate at a lower concentration in a microcap with 90 impermeable particles compared to one without. We hypothesize that if the attachment effect is 91 replicated that diffusing species would accumulate at a higher concentration in the microcap 92 when attachment sites are present. Biofilms have distinct porosity profiles based on 93 environmental conditions [42, 43], distinct matrix components based on nutrient conditions and 94 external threat [36, 43], and distinct cell populations based on age and nutrient conditions. This 95 study and platform may aid in understanding how each component contributes to overall 96 accumulation. 97 2. Methods 98 The goal of this study is to test the ability of a synthetic biomaterial to replicate the 99 important effects of a biofilm on species diffusion. We leveraged our recently developed 100 nanocellulose hydrogel that has closer mechanical stiffness to natural biofilms. We embedded 101 the biofilm with charged and non-charged microspheres to test volume-exclusion and attachment 102 and charge based exclusion. We used two sizes of nanodextrans to test size exclusion. We used 103 standard optical tools for examining biofilms to measure the effects of the tested parameters, 104 Figure 1. 105 2.1 Microcap Design and Nanodextran Information 106 To generate the microcaps, hydrogels were used as the hydrated mesh domain to achieve 107 a size-effect. Hydrogels have been used in prior studies to simulate the extracellular matrix of the 108 biofilm [41]. The hydrogel platform chosen for our system was nanocellulose hydrogels. We 109 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 6 recently developed a nanocellulose that better matched the mechanical storage and loss moduli 110 [39], that is also easily chemically modifiable for which future studies could choose to utilize for 111 a method of generating attachment sites independent of microspheres. 112 Fluorescent polystyrene microspheres (Spherotech | Lake Forest, IL) ranging from 0.7-113 0.9 microns in diameter were used to simulate bacteria and achieve a volume-exclusion effect. 114 Three distinct microspheres, each with distinct microsphere surface chemistry, were used: no 115 surface modifications (Plain) (Prod# FP-0862-2), carboxylate-modified (Carboxyl) (Prod# FP-116 0862-2), and amine-modified (Amino) (Prod# FP-0862-2). The carboxylate and amine-modified 117 microspheres were coated with these functional groups, which are either negatively or positively 118 charged at neutral pH. These charged microsphere surfaces were designed to only act as 119 attachment sites for the diffusing species if the diffusing species was oppositely charged. The 120 microspheres were fluorescently labelled with a vendor proprietary fluorescent tag. 121 Nanodextrans conjugated to FITC (fluorescein isothiocyanate) were used as the diffusing 122 species for the system (TdB Labs, Uppsala, Sweden). Two nanodextran molecular weights used 123 were 150 kDa and 2,000 kDa, on the premise these molecules would have different sizes, to test 124 the size-exclusion effect. The fluorescently labelled nanodextrans were chemically modified to 125 have either carboxymethyl (CM) groups with negative charge (Prod# FITC-CM-dextran-150), 126 diethylaminoethyl (DEAE) groups with positive charge (Product # FITC-DEAE-dextran-150), or 127 no modification at all (Dx) (Prod# FITC-Dx-150 and Prod# FITC-Dx-2000). The use of charged 128 nanodextrans and charged microsphere surfaces was designed to produce the attachment effect. 129 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 7 b c Figure 1: A. Experimental protocol for testing diffusion into hydrogel. B. Experimental Apparatus Diagram C. Artistic representation of microcap. Hydrogels were made as described in [39]. After centrifugation, microspheres were added. Resultant , A flow cell (b) from Biosurface Technology was seeded with hydrogel inserts along the coupon recess/imaging window. Red circles represent microspheres. Microspheres with amine-coating represented. Nanodextran with no modification shown. 130 2.2 Nanocellulose Solution Synthesis 131 A nanocellulose synthetic biofilm was generated as described in [39]. Briefly, 4.00 ± 0.05 132 g of dry cellulose powder (CAS#9004-34-6 | Sigma-Aldrich | Burlington, MA) and slowly 133 mixing it with an ionic liquid solution composed of 7.00 ± 0.05 g sodium hydroxide (CAS#1310-134 73-2 | Sigma-Aldrich | Burlington, MA), 12.00 ± 0.05 g urea (CAS#57-13-6 | Sigma-Aldrich | 135 Burlington, MA), and 81.00 ± 0.05 g of reverse-osmosis filtered (RO) water and adding it to a 136 125 mL Erlenmeyer flask. The resulting suspension was mixed using an inert stir bar at 500 rpm 137 until it was homogenous and consistently cloudy. The suspension was then submerged in an ice-138 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 8 isopropyl alcohol bath with a temperature ranging between -10 and 0 OC while still being stirred. 139 After more than 1 hour of stirring and the dissolution of the cellulose, the liquid was separated 140 into two 50 mL centrifuge tubes. The tubes were centrifuged at 10,000 RPM (rcf=13,751) for 10 141 minutes while maintaining a temperature of 22 OC. After centrifugation, the supernatant was 142 decanted and saved as nanocellulose solution, with any settled solids discarded. 143 2.3 Nanocellulose-Microsphere Solutions 144 For each batch of nanocellulose solution made, four different nanocellulose-microsphere 145 solutions were made: one with Plain microspheres, one with Carboxyl microspheres, one with 146 Amino-microspheres, and one with no microspheres. These nanocellulose-microsphere solutions 147 were prepared by mixing 5.0±0.1 mL of nanocellulose solution with 50±0.1 µL of 1wt% 148 microsphere solution in the well of a 6-well culture plate. Once all four nanocellulose-149 microsphere solutions were prepared for each nanocellulose batch on the same plate, the plate 150 was covered, wrapped in aluminum foil, and mixed for 24 hours on a shaking plate at 250 RPM 151 at room temperature ranging from 22-26 ºC. After 24 hours of mixing, the resulting 152 nanocellulose-microsphere solutions were stored at 4 ºC. 153 2.4 Microcap and Flow Cell Preparation 154 With the nanocellulose-microsphere solutions prepared, a 5 mL syringe affixed with a 155 27G nozzle was used to fill the nozzle with nanocellulose-microsphere solutions. The 156 nanocellulose solution was then extruded through the nozzle using the syringe. Once a small 157 amount of nanocellulose solution was extruded, priming the nozzle, the nozzle would then “leak” 158 the solution semi continuously. The nozzle tip was then repeatedly (between 10-30 times) 159 pressed onto the surface of a polycarbonate coupon, with a microscopic amount of hydrogel 160 being left behind on the coupon as residue with each press. The coupon then sat at room 161 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 9 temperature in a plastic petri dish for greater than 48 hours to dry with no plastic cover but 162 aluminum foil cover, which prevented contamination and fluorescent overexposure, but allowed 163 for air flow. This let the hydrogel residues to dry onto the coupon and form microcaps. Before an 164 experiment, the microcap covered coupon was loaded into a flow cell (Prod# FC 275-AL | 165 BioSurface Technologies | Bozeman, MT, USA), where the flow cell chamber containing the 166 coupon was filled with reverse-osmosis water first to rehydrate the microcap. Almost 167 immediately after, the chamber was then filled with a 5 µM solution of Calcofluor White (CW) 168 stain dissolved in water (Cat# 29067 | Biotium | Fremont, CA, USA). The fluorophore would 169 then slowly diffuse into the microcaps, allowing for fluorescent imaging of the hydrogel domain 170 of the microcap. 171 2.5 Nanodextran Exposure and Imaging Procedure 172 Once the flow cell was loaded with coupons covered with CW-stained microcaps, the 173 flow cell was then connected to a microfluidic apparatus with two inlets: reverse-osmosis filtered 174 (RO) water and 30 mg/L fluorescently labelled-nanodextran dissolved in Dulbecco’s phosphate 175 buffer solution (PBS) (Figure 1). The flow cell was placed on a confocal laser scanning 176 microscope, which allowed for real-time 3-D fluorescent imaging of the microcaps. 177 Each experiment involved a two-phase procedure: characterization and accumulation. To 178 perform an experiment, the flow chamber was flushed with 1 mL/min RO water during the 179 characterization phase. While the flow chamber containing the microcap-covered coupon was 180 flushed with 1 mL/min RO water, the microcaps on the coupon were checked to determine if 181 there were three microcaps less than 250 microns in diameter (all tests had at least three). A z-182 stack, or a series of microscope images taken at close, regularly spaced focal lengths to 183 reconstruct a 3-D image of a sample, was taken of three microcaps on each coupon (with the 184 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 10 three smallest microcaps usually chosen). Field of view size was adjusted for each microcap to 185 take up the entire image. Nyquist sampling was used to select z-section thicknesses and image 186 resolution. These z-stacks were set up to image three features in each microcap: the hydrogel 187 mesh by imaging the CW stain (Laser: 405 nm/Detector Range: 410-546 nm), the impermeable 188 microspheres (Laser: 561 nm/Detector Range: 585-700 nm), and the presence of FITC (Laser: 189 488 nm/Detector Range: 410-546 nm). In addition to measuring the presence of FITC, this same 190 signal was also used to determine the location of the coupon within the image, as the coupon 191 reflected green light at its surface. This first image before any FITC was added to the system will 192 be referred to as the characterization image. 193 After the characterization image of each of the microcap of interest, the microcaps were 194 exposed to a continuous stream of nanodextran during the accumulation phase of the experiment. 195 This was done by flowing 1 mL/min of the 30 mg/L nanodextran dissolved in PBS into the flow 196 chamber continuously for 24 minutes. After this continuous exposure to a constant concentration 197 of nanodextran, the flow in the chamber was stopped, and the microcaps re-imaged using the 198 same parameters as those for the characterization image, producing an accumulation image for 199 each microcap. This was done to quantify two things: the concentration of nanodextran (via the 200 proxy measurement of FITC-signal) in the water immediately surrounding each microcap, and 201 the concentration of nanodextran within each microcap. 202 During accumulation phase of each experiment, time-lapse imaging of a single-z-plane 203 within one of the imaged microcaps was taken to monitor if the system reached equilibrium 204 during each phase (which all were confirmed to reach). 205 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 11 2.6 Experimental Design 206 To test the functional properties of these microcaps, three predicted experimental effects 207 were tested to determine the capacity of these microcaps to replicate the biofilm features of 208 interest, Figure 2. The size-exclusion effect was tested to determine if a hydrated mesh was 209 formed. The volume-exclusion effect was tested to determine if the embedded microspheres 210 were diffusing species impermeable. The attachment effect was tested to determine if the 211 diffusing species were immobilized via opposite charge interactions with the surface of the 212 microspheres, indicating diffusing species attachment sites. 213 a b c Figure 2: (a-c)Diagrams of hypothesized size-exclusion effect, charge based attachment effects, and volume exclusion effect. d-e Hypothesized outcomes of nanodextran concentration in hydrogels based on the given effect. .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 12 214 To test for the size-exclusion effect, the two different sized nanodextrans were used: 150 215 kDa and 2,000 kDa. The 2,000 kDa nanodextran was expected to have a larger particle size than 216 the 150 kDa case. This was tested and confirmed using Nanoparticle Tracking Analysis and 217 Dynamic Light-Scattering measurements, shown in Table 1. While the measured sizes were 218 similar sizes via NTA and DLS, we used them as it well known that while designed for specific 219 sizes, environmental nanomaterials are never pristine. These nanodextan molecular weights were 220 chosen because they have been used in biofilm penetration studies as both small and large sized 221 nanomaterials [23, 25, 44, 45]. Additional details on nanoparticle characterization are provided 222 in the Supplementary Information. 223 Table 1: Nanodextran Characterization. All nanodextrans dispersed in Dulbecco’s Phosphate Buffer Solution (PBS). NTA error reported as standard error from measured particle size distribution (n=5). *- Standard error not reported on instrument. FITC-Dx-2000 size measured twice on NTA, both sizes reported. DLS/ELS error reported as standard deviation of three triplicate measurements. Nanodextran Nanodextran Molecular Weight [kDa] Size via NTA [nm] Size via DLS [nm] Charge via ELS [mV] FITC-Dx-150 150 226 ± 14 115 ± 24 -1.2 ± 0.4 FITC-DEAE-150 150 178 ± 5 107 ± 16 -0.4 ± 0.4 FITC-CM-150 150 173* 771 ± 1000 -5.7 ± 3.2 FITC-Dx-2000 2000 297 ± 18 306 ± 24 81 ± 0.1 -2.4 ± 0.7 Since hydrated meshes such as a biofilm extracellular matrix and a hydrogel are 224 comprised of disorganized, overlapping biopolymer chains, they can allow particles to diffuse 225 through them, but only up to a certain size. As the particle reaches a characteristic “mesh size”, 226 less of the volume within the hydrated mesh network is available for it to occupy. Once the 227 particle reaches a critical size, no amount of particle can accumulate within the matrix. Thus, as 228 particle size increases, the equilibrium concentration it reaches within a hydrated matrix 229 decrease, as less volume is available for the particle to permeate into. Hence, a decrease in 230 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 13 diffusing species concentration with particle size within the matrix at equilibrium shows the 231 presence of a size-exclusion effect in the microcap, which in our system is likely indicative of 232 the formation of a hydrogel. Microcaps prepared using identical conditions were used in size-233 exclusion experiments. 234 To test for the volume-exclusion effect, hydrogels were prepared either with 235 microspheres within them or without microspheres. The microcaps without microspheres within 236 them would be expected to reach a higher concentration of nanodextran if the volume-exclusion 237 effect was observed. The reasoning for this is similar to the reasoning for the size-exclusion 238 effect: certain portions of the microcap volume, specifically the volume occupied by the 239 microspheres, would be unavailable for the nanodextran to occupy, leading to a lower 240 concentration overall in the microcap. Identical nanodextrans were used in volume-exclusion 241 experiments. 242 To test for the attachment effect, hydrogels embedded with three different microspheres 243 were prepared and tested against three different nanodextrans. The different microspheres tested 244 were no surface functionalization (Plain, neutrally charged surface), amine-functionalization 245 (Amino, positively charged surface), and carboxyl-functionalized (Carboxyl, negatively charged 246 surface). The different nanodextrans tested were no chemical modification (None, neutrally 247 charged), carboxymethyl-modified (CM, negatively charged) and diethylaminoethyl-modified 248 (DEAE, positively charged). Since we only expect oppositely charged combinations to lead to 249 attachment between microsphere surface and nanodextran, we expect to see significant increases 250 in nanodextran concentration within the microsphere only for the combination of CM-modified 251 nanodextran accumulating in microcaps with amine-modified microspheres, and DEAE-252 modified nanodextran accumulating in microcaps with carboxyl-modified microspheres. Two 253 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 14 different control cases are considered for each of these combinations: (a) charged nanodextran 254 and uncharged microspheres, and b) uncharged nanodextran and charged microspheres. 255 Decreases in nanodextran concentrations within the microcap would be expected in both cases 256 compared to the oppositely charged nanodextran/microspheres case. 257 2.7 Image Analysis 258 To determine nanodextran concentration for each z-stack image taken, each pixel was 259 assigned to be in one of four spatial domains in the image: (1) the water domain (liquid domain 260 Ω!), (2) the hydrogel domain (interstitial domain Ω") (3) the microsphere domain (Ω#), and (4) 261 the coupon domain (solid domain Ω$). 262 Figure 3: Labelled Microcap from Characterization Image. Red channel: fluorescent polystyrene microspheres (carboxyl-modified). Blue channel: CW-stained nanocellulose hydrogel. Green-channel: polycarbonate coupon-microcap interface. 263 This was done by segmenting the hydrogel using the CW signal, the microspheres using 264 the AF594 signal, and the coupon using the FITC-signal in each image, and assuming all 265 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 15 remaining pixels were within the water domain. The total microcap domain was the union of the 266 microcap and microsphere domains. 267 The microspheres were segmented using an Otsu threshold on the AF594 signal in 268 MATLAB. The hydrogel domain was segmented in ImageJ using a Otsu threshold, followed by 269 a region-filling algorithm (CW-stain only penetrated ~10 microns into hydrogel), followed by an 270 algorithm for discarding very small filled regions. The coupon was segmented using an edge-271 detection algorithm along the z-direction. An important note on this process was that coupon 272 segmentation was not perfect. The coupon segmentation algorithm was designed to favor a pixel 273 as coupon as opposed to water or hydrogel on purpose, since the accuracy of the water and 274 hydrogel signal was more important than the coupon signal. 275 With each pixel assigned to a domain, the average value of the nanodextran/FITC signal 276 in each domain in the microcap was quantified. Rather than use average FITC-signal in the 277 microcap, [𝐹𝐼𝑇𝐶]%&'()'*+ , directly, the ratio of the FITC-signal in the microcap to the FITC-278 signal in the water, [𝐹𝐼𝑇𝐶],*-.( , or the nanodextran-microcap partition coefficient, 𝐾/, was 279 calculated for each microcap’s accumulation image, Equation 1. This ensured any variation in 280 FITC concentration and signal within the flow cell was normalized between microcaps. 281 𝐾/ = [𝐹𝐼𝑇𝐶]%&'()'*+ [𝐹𝐼𝑇𝐶],*-.( Equation 1: Microcap Partition Coefficient The MATLB code for image analysis is included as Supplementary Information and is 282 available via Github. 283 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 16 2.8 Statistical Analysis 284 Figure 4: Design of Experiments and Replicates For each nanodextran-hydrogel combination tested, two types of replicates were used, all 285 on the hydrogel side: batch replicates and microcap replicates. Three batches of each hydrogel 286 tested (no microspheres, plain microspheres, carboxyl-microspheres and amine-microspheres) 287 were generated, leading to 12 different hydrogels with three batch replicates each. For each 288 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 17 batch, one coupon covered with microcaps was generated for each nanodextran tested, leading to 289 33 different coupons. On each coupon, three microcap replicates were tested on each. This led to 290 a total of 99 microcaps tested. 291 To compile the data for each experimental condition (nanodextran-microcap 292 combination), the nanodextran-microcap partition coefficient for each microcap tested was first 293 found (see Image Analysis for how this was calculated). This value was then averaged for the 294 three microcaps on each coupon tested, as these replicates were determined to be technical 295 replicates, which warrant averaging together for the purposes of statistical analysis. This left 296 three nanodextran-microcap partition coefficients for each experimental condition, each one 297 representing a different hydrogel batch. These three nanodextran-microcap partition coefficients 298 comprised the sample for each experimental condition. 299 For each set of experimental conditions compared for each hypothesis, the three 300 nanodextran-microcap partition coefficients for each batch were compared to each other using 301 Welch’s one-way t-tests using an alpha of 0.05. Welch’s test was used since variances were not 302 assumed to be equal. One-way tests were used since only an effect in one direction would 303 constitute evidence for each hypothesis. All samples passed the Shapiro-Wilks test for normality. 304 3. Results 305 The microcaps generated using this synthesis procedure were tested for their ability to 306 replicate the core features of a synthetic biofilm. These included size-exclusion effects, volume 307 exclusion effects and attachment effects. Different combinations of microcaps and diffusing 308 nanodextran were used to test for these effects through equilibrium nanodextran partition 309 coefficients in the microcaps. 310 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 18 Testing for Hydrated Mesh Property via the Size Effect 311 To test for the size-exclusion effect, nanodextran-microcap partition coefficients for 312 microcaps with plain microspheres were compared between a 150 kDa nanodextran and a 2,000 313 kDa nanodextran. The results, as shown in Figure 5, show a statistically significant size-314 exclusion effect in the hydrogel mesh was observed. 315 316 Figure 5: Size-Exclusion Effect Test for Verifying the Hydrogel Mesh Property of the Synthetic Biofilm System. Error bars represent 95% confidence intervals on the sample. P-value represents results from one-tailed Welch’s t-test. Experiments used 4 wt% hydrogels loaded with 0.01 wt% plain polystyrene microspheres with neutrally charged nanodextrans. Partition coefficient represents nanodextran-microcap partition coefficient. 317 Testing for Impermeable Sub-volume via the Volume-Exclusion Effect 318 To test for the volume-exclusion effect, nanodextran-microcap partition coefficient for 4 319 wt% hydrogel microcaps embedded with either no microspheres or 0.01 wt% plain microspheres 320 were compared using a neutrally charged, 150 kDa nanodextran. The results, as shown in Figure 321 6, show no volume-exclusion effect due to the presence of the microspheres was observed. 322 323 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 19 Figure 6: Volume-Exclusion Effect Test for Verifying the Impermeable Subdomain Property of the Synthetic Biofilm System. Error bars represent 95% confidence intervals on the sample. P-value represents results from one-tailed Welch’s t-test. Experiments used 4 wt% hydrogels loaded with variable wt% plain polystyrene microspheres and neutrally charged, 150 kDa nanodextran. Partition coefficient represents nanodextran-microcap partition coefficient. 324 Testing for Attachment Sites via the Attachment Effect 325 To test for the attachment effect, 4 wt% hydrogels embedded with 0.01 wt% neutrally, 326 negatively, and positively charged microspheres were tested against neutrally, negatively, and 327 positively charged 150 kDa nanodextrans, shown in Figure 7. Opposite charge combinations 328 were tested for increases in nanodextran-microcap partition coefficients compared to both the 329 same charged nanodextran and uncharged microspheres and the uncharged nanodextran and 330 same charged microspheres. No statistically significant attachment effect was observed across all 331 comparisons. A non-statistically significant attachment effect was observed for the microcaps 332 embedded with amine-modified microspheres with a carboxymethyl-modified nanodextran for 333 comparisons to both control cases. 334 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 20 Figure 7: Attachment Effect Test for Verifying Presence of Attachment Sites in the Synthetic Biofilm System. Error bars represent 95% confidence intervals on the sample. P-values represent results from one-tailed Welch’s t-test. Experiments used 4 wt% hydrogels loaded with variable 0.01 wt% plain-, carboxyl- and amine- modified polystyrene microspheres and neutrally, positively or negatively charged, 150 kDa nanodextrans. Partition coefficient represents nanodextran-microcap partition coefficient. 4. Discussion 335 This research sought to create a synthetic biofilm system for use studying species 336 transport in biofilms. To accomplish this, nanocellulose hydrogel microcaps loaded with 337 polystyrene microspheres were developed to proxy bacterial biofilms in the features of a 338 hydrated mesh, an impermeable subdomain, and presence of attachment sites. The replication of 339 these features in the generated microcaps were tested for by quantifying an important 340 experimental effect for each feature: the size-exclusion effect for verifying the hydrogel mesh, 341 the volume-exclusion effect for verifying the impermeable subdomain, the attachment effect for 342 verifying presence of attachment sites. 343 Experimental results indicated that the size-exclusion effect was verified, and therefore 344 the hydrogel mesh feature of the microcaps. However, the attachment effect and the volume 345 exclusion effect were not observed. This indicates that the microcaps as currently designed are 346 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 21 unable to replicate the key biofilm properties needed to perform transport studies of an 347 impermeable subdomain and presence of attachment sites. 348 Size-effect agrees with prior literature on chemical species diffusion in single bacterial 349 species biofilms [23, 46]. However, evidence of this effect is limited in multispecies biofilm 350 literature on nanoparticles [47]. This is most likely due to effects of agglomeration, dissolution 351 and changes to the nanoparticle biopolymer corona in multispecies biofilms studies having 352 strong effects on effective nanoparticle size during diffusion [48, 49]. This shows the need for 353 studies with more controls over nanoparticle stability in multispecies biofilm transport studies. 354 A few distinct hypotheses could explain why the presence of microspheres and the use of 355 chemically modified microsphere surfaces and nanodextrans did not elicit the effects of volume-356 exclusion and attachment. One is the use of a 0.01 wt% microsphere percentage within the 357 hydrogels was too low to observe the anticipated effects. If little volume is made impermeable 358 due to low concentration of microspheres, then the volume-exclusion effect would be anticipated 359 to be small and below any statistically significant threshold. In addition, too low of a 360 concentration of microspheres would mean there are very few sites for nanodextran attachment, 361 which would also predict a small attachment effect again below any statistically significant 362 threshold. Another possibility is that nanodextran can penetrate and diffuse into the polystyrene 363 microspheres. While there is evidence for adsorption onto the surface of polystyrene [50], there 364 is less evidence for penetration and diffusion into polystyrene microsphere. However, no 365 experimental evidence is offered in this research proving or disproving this hypothesis. 366 For the attachment effect, another possibility is that the charged microsphere surfaces do 367 not act as attachment and immobilization sites for its oppositely charged nanodextran. This could 368 be due to a variety of reasons including the charge-charge interactions not being strong enough 369 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 22 to immobilize the nanodextran, the microsphere or the nanodextran not being charged species as 370 anticipated, or even that the immobilization of the nanodextran does not lead to higher levels of 371 nanodextran to accumulate into the hydrogel to balance the chemical potential gradient. While it 372 is possible attachment does occur in these systems but too slowly to observe in this experiment, 373 direct charge-charge interactions are the strongest intermolecular forces in chemistry, which 374 would make for quick attachment, making this possibility seem unlikely. This lack of a clear 375 effect of charge is common in the scientific literature on diffusion of chemical species in 376 hydrogel matrices such as biofilms [41, 44, 45, 51, 52]. Future experiments on this system could 377 include testing the accumulation and desorption of ions instead of nanomaterials as these have 378 been shown to matter in the biofilm matrix [53]. 379 Assuming the hypothesis that the non-observed effects were primarily due to a low 380 concentration of microspheres within the microcaps, further experimental work could replicate 381 these experiments with higher concentrations of microspheres and look for the same effects on 382 attachment and volume-exclusion. Further evidence for the size-exclusion effect could be found 383 by comparing current results to higher weight percentage nanocellulose hydrogels, which 384 theoretically lead to smaller critical mesh size, which would be anticipated to cause lower 385 nanodextran concentrations in these higher weight percentage microcaps [23, 54]. 386 The data presented here does not lend itself to diffusivity calculations common in the 387 literature. However, Bryers and Frummond showed lumped parameter diffusivity calculations 388 are inaccurate for describing transport in biofilm [55]. The data can be compared to 389 bioconcentration factors (BCFs) which have been widely reported in ecotoxicological studies of 390 engineered nanoparticles effects on environmental biofilms. Since estimates for BCFs in 391 ecotoxicological studies are usually only order of magnitude estimates, logarithmic BCF values 392 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 23 (pBCF) will be used here to compare BCFs between different experiments [56-58]. pBCFs will 393 be calculated from dimensionless BCFs on a mass-by-mass basis (µg ENM/g biofilm divided by 394 µg ENM/g water) using the wet weight of the periphytic biofilms when possible. Since 395 periphytic biofilms are hydrated in the environment, wet weights are used to give a more 396 intuitive sense of the extent of the increase in concentration of ENM that would be seen in an 397 environmental system. If only dry-weight/AFDM BCFs are reported, wet-weight BCFs will be 398 estimated by assuming a 100:1 ratio of biofilm wet mass to biofilm dry mass. Finally, many 399 studies for the bioaccumulation of ENM quantify total metal uptake, not ENM specific uptake. 400 Unless controls for dissolution or specific quantification of ENM is stated, all BCFs reported 401 here will reflect this, limiting their representation of "true" ENM BCFs. Since most studies do 402 not report detailed kinetic analysis of ENM bioaccumulation, characteristic times, 𝑡0, of 403 bioaccumulation will be estimated when possible. Estimations of these are mostly qualitative, 404 representing approximately the time necessary to reach concentrations 50% of equilibrium 405 concentrations. 406 The data in this study are 2 – 10 times less than the pBCF reported in literature, Table 2. 407 While this hydrogel matches mechanical properties better than alginate, many biofilms contain 408 both alginate and nanocellulose polymers. Lastly, biofilms are much more than cells or beads 409 distributed in a hydrogel mesh, they also contain channels filled with pore water [55]. Equation 2 410 shows that if the amount of nanoparticles in the pore water is in equilibrium with the surrounding 411 water, the BCF in a porous biofilm will be less than an equivalent amount of nanomaterials being 412 added to a fixed amount of biofilm mass without. Equivalently, the pBCF of a biofilm with pores 413 will be greater than one without for the same amount of substance added to the biofilm, by a 414 factor of log2 for biofilm of equal weight to water. 415 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 24 𝑚!"#,% + 𝑚!"# 𝑚% + 𝑚& 𝑚!"#/𝑚& = 𝑚!"#,% + 𝑚!"# 𝑚% + 𝑚& 𝑚!"#/𝑚& = (𝑚!"#,% + 𝑚!"#)/𝑚% 1 + 𝑚&/𝑚% 𝑚!"#/𝑚& < 𝑚!"#,% + 𝑚!"# 𝑚% 𝑚!"#/𝑚& Equation 2 416 417 Table 2: Summary of literature on bioaccumulation and bioconcentration kinetic parameters of ENMs in periphytic biofilms. n.d. no data reported. *Some hydropathies are inferred. Titanium dioxide hydropathy depends on crystal phase which also modifies its toxicity [59]. Type Biofilm Size (nm) Hydropathy Charge tC pBCF Study Latex carboxylate Wetland 100 Hydrophilic (-) < 24 h ~2 [60] Gold Estuary 65 Hydrophobic (+) < 12 d 2.18 [61] Gold polystyrene sulfonate Estuary 50 Hydrophobic* -53 mV ~12 h 3.46 [22] Citrate capped silver Estuary 30, 115 Hydrophobic (-) ~12 h 1.4 – 1.5 [62] Citrate capped silver Marine 100 – 700 Hydrophobic -18 mV 10 min , < 4 d 4.6 [63] Titanium dioxide Algal 150 n.d.* -17 mV < 28 d 4 - 5.6 [64] Copper oxide Periphyton 100 – 800 Hydrophilic* -35 – +15 mV ~50 min 3.6 – 4.25 [65] Citrate capped silver Aquabacteri um citratiphilum 30, 70 Hydrophobic -60 mV < 20 h 0.8 [66] FITC-Dx-150 Hydrogel 92 – 240 Hydrophilic -1.2 ± 0.4 mV 24 min 0.153 FTIC-Dx-2000 Hydrogel + neutral spheres 81 – 330 Hydrophilic -2.4 ± 0.7 mV 24 min 0.319 FITC-Dx-150 Hydrogel + neutral spheres 92 – 240 Hydrophilic -1.2 ± 0.4 mV 24 min 0.164 FITC-DEAE-150 Hydrogel + neutral spheres 91 – 183 Hydrophilic -0.4 ± 0.4 mV 24 min 0.132 FITC-CM-150 Hydrogel + neutral spheres 173 – 1000 Hydrophilic -5.7 ± 3.2 mV 24 min 0.160 FITC-Dx-150 Hydrogel + carboxyl (-) spheres 92 – 240 Hydrophilic -1.2 ± 0.4 mV 24 min 0.145 FITC-DEAE-150 Hydrogel + carboxyl (-) spheres 91 – 183 Hydrophilic -0.4 ± 0.4 mV 24 min 0.179 FITC-CM-150 Hydrogel + carboxyl (-) spheres 173 – 1000 Hydrophilic -5.7 ± 3.2 mV 24 min 0.135 FITC-Dx-150 Hydrogel + amino (+) spheres 92 – 240 Hydrophilic -1.2 ± 0.4 mV 24 min 0.125 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 25 FITC-DEAE-150 Hydrogel + amino (+) spheres 91 – 183 Hydrophilic -0.4 ± 0.4 mV 24 min 0.158 FITC-CM-150 Hydrogel + amino (+) spheres 173 – 1000 Hydrophilic -5.7 ± 3.2 mV 24 min 0.068 418 In summary, a systematic analysis through a synthetic biofilm model adds to the toolkit 419 of biointerface studies. Size exclusion has been replicated in alginate [41] and now in 420 nanocellulose. While attachment and volume-exclusion were not replicated, future work using 421 similar matrices should also consider concentration of microspheres within the nanocellulose 422 matrix. For example, there may be a critical concentration of microspheres or bacterial cells 423 necessary for some of the absorption effects. While these can be studied with non-toxic 424 nanomaterials, these affects are often very difficult to uncouple in living systems [67]. 425 5. Conclusion 426 This study aimed to emulate key physicochemical barriers to diffusion found in natural 427 biofilms using tunable synthetic microcap biofilm matrix system. Through the controlled 428 exposure of nanodextrans with varying size and surface charge, we evaluated the system’s ability 429 to emulate three core physicochemical features often implicated in biofilm-associated transport 430 resistance: size exclusion, charge interactions, and volume exclusion. The results demonstrated a 431 statistically significant size-exclusion effect, confirming the ability of the nanocellulose-based 432 microcaps to mimic the selective permeability of hydrated biofilm matrices. However, the 433 designed system did not display statistically significant volume-exclusion or attachment effects, 434 suggesting that the current microsphere concentration and charge configurations were 435 insufficient to replicate these additional features. 436 These findings reflect patterns observed in natural biofilms studies, where size-based 437 diffusion hinderance is commonly reported, but charge-based interaction and volume exclusion 438 .CC-BY 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted February 4, 2026. ; https://doi.org/10.64898/2026.02.02.703274doi: bioRxiv preprint 26 are more context-dependent. The absence of strong attachment or volume exclusion effects may 439 be due to insufficient microsphere loading, incomplete charge immobilization, or the dynamic 440 behavior of the nanodextrans in the hydrated mesh. 441 Future studies should explore increased microsphere loading, use of covalently bound 442 attachment sites or incorporation of more biologically relevant surface chemistries to better 443 recapitulate these additional transport-limiting features. Ultimately, refining this synthetic 444 biofilm platform will enhance its utility in advancing the ecotoxicology of engineered 445 nanomaterials. 446 447

448 449 • DT was supported in part by the National Science Foundation under Grant No. DGE-450 2022040. 451 • Research reported in this publication was supported by the National Institute of General 452 Medical Sciences of the National Institutes of Health under Award Number NIH R35 453 GM142898 The content is solely the responsibility of the authors and does not 454 necessarily represent the official views of the National Institutes of Health. 455 456

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