extfieldsofstudy: Chemistry Biology

text:
Pseudomonas aeruginosa leucine aminopeptidase influences early biofilm composition and structure via vesicle-associated anti-biofilm activity

Pseudomonas aeruginosa, known as one of the leading causes of disease in cystic fibrosis (CF) patients, secretes a variety of proteases. These enzymes contribute significantly to P. aeruginosa pathogenesis and biofilm formation in the chronic colonization of CF patient lungs, as well as playing a role in infections of the cornea, burn wounds and chronic wounds. We previously characterized a secreted P. aeruginosa peptidase, PaAP, that is highly expressed in chronic CF isolates. This leucine aminopeptidase is highly expressed during infection and in biofilms, and it associates with bacterial outer membrane vesicles (OMVs), structures known to contribute to virulence mechanisms in a variety of Gram-negative species and one of the major components of the biofilm matrix. We hypothesized that PaAP may play a role in P. aeruginosa biofilm formation. Using a lung epithelial cell/bacterial biofilm coculture model, we show that PaAP deletion in a clinical P. aeruginosa background alters biofilm microcolony composition to increase cellular density, while decreasing matrix polysaccharide content, and that OMVs from PaAP expressing strains but not PaAP alone or in combination with PaAP deletion strain-derived OMVs could complement this phenotype. We additionally found that OMVs from PaAP expressing strains could cause protease-mediated biofilm detachment, leading to changes in matrix and colony composition. Finally, we showed that the OMVs could also mediate the detachment of biofilms formed by both non-self P. aeruginosa strains and Klebsiella pneumoniae, another respiratory pathogen. Our findings represent novel roles for OMVs and the aminopeptidase in the modulation of P. aeruginosa biofilm architecture. Importance Biofilm formation by the bacterial pathogen P. aeruginosa is known to contribute to drug- resistance in nosocomial infections and chronic lung infections of cystic fibrosis patients. In order to treat these infections more successfully, the mechanisms of bacterial biofilm development must be elucidated. While both bacterially-secreted aminopeptidase and outer membrane vesicles have been shown to be abundant in P. aeruginosa biofilm matrices, the contributions of each of these factors to the steps in biofilm generation have not been well studied. This work provides new insight as to how these bacterial components mediate the formation of a robust, drug-resistant extracellular matrix and implicates outer membrane vesicles as active components of biofilm architecture, expanding our overall understanding of P. aeruginosa biofilm biology.

Importance 23
Biofilm formation by the bacterial pathogen P. aeruginosa is known to contribute to drug-24 resistance in nosocomial infections and chronic lung infections of cystic fibrosis patients. In 25 order to treat these infections more successfully, the mechanisms of bacterial biofilm 26 2 development must be elucidated. While both bacterially-secreted aminopeptidase and outer 27 membrane vesicles have been shown to be abundant in P. aeruginosa biofilm matrices, the 28 contributions of each of these factors to the steps in biofilm generation have not been well 29 studied. This work provides new insight as to how these bacterial components mediate the 30 formation of a robust, drug-resistant extracellular matrix and implicates outer membrane 31 vesicles as active components of biofilm architecture, expanding our overall understanding of P. 32 aeruginosa biofilm biology. 33

Introduction 37
The Gram-negative bacterium Pseudomonas aeruginosa is a prominent opportunistic 38 pathogen capable of causing both acute and chronic disease in a variety of compromised hosts. 39 P. aeruginosa is most well known as a leading cause of morbidity and mortality in cystic fibrosis 40 (CF) patients 1 . In these individuals, the pathogen establishes chronic, biofilm-based infections 41 that may persist for decades in the unique, mucous-rich environment of the CF lung. In addition 42 to lung tissue, P. aeruginosa forms biofilms on a wide range of substrates relevant to human 43 infection, including corneal and skin tissue, as well as soil and water reservoirs and hospital 44 surfaces, which can contribute to infection initiation and spread 2 . 45 P. aeruginosa is considered a model organism for the study of biofilm formation, and 46 many of the cellular and matrix components contributing to this mode of growth have been 47 studied previously 3-5 . The process of forming biofilm communities has also been characterized, 48 revealing distinct growth phases. First the bacteria settle and attach onto a suitable host tissue 49 or abiotic surface, then they begin to form stationary microcolonies and secrete extrapolymeric 50 substance (EPS) to form a dense matrix. This matrix contains a variety of polysaccharides, with 51 Psl and Pel polysaccharides present in all strains, and alginate found in mucoid P. aeruginosa 52 isolates 6 . It also contains proteins, lipids, extracellular outer membrane vesicles (OMVs), and 53 extracellular DNA (eDNA) 4,5,7 ; however, the mechanisms by which many of these secreted 54 bacterial components specifically affect microcolony growth and development remain unclear. 55 As with other extracellular pathogens, P. aeruginosa secretes many of its virulence 56 determinants into the extracellular milieu to affect interactions with the host, including those in 57 the biofilm matrix. Interestingly, many of the virulence factors secreted specifically by P. 58 aeruginosa demonstrate proteolytic activities 8- 10 . These enzymes, including elastase, protease 59 IV, and alkaline protease, act on both bacterial and host proteins to directly impact host-60 pathogen interactions. For example, purified elastase has been found to cleave fibrin, laminin, 61 various immunoglobulins, and components of the complement system 11 . Their activity not only 62 8 164

qRT-PCR 165
Pellicle and coculture biofilms were grown for the indicated times as described. For coculture 166 biofilms, supernatants were removed and 1 mL of Trizol was added to each sample to lyse both 167 bacterial and A549 cells. For pellicle biofilms, 3 mL/well of cultures incubated in 6-well 168 polystyrene dishes (VWR) were used to obtain sufficient biofilm material for analysis over the 169 as outlined previously 28 ), LB+ 0.4% glucose, or tissue culture media (described above) with no 200 antibiotics. Cultures were grown to an OD 600 of 0.5 and diluted to OD 600 of 0.1. Aliquots of the 201 diluted culture (100 µL/well) were incubated in five 96-well flat bottom polystyrene dishes (VWR) 202 for 0, 2, 6, 10, or 24 h at 37°C with no shaking and evaluated using a static biofilm quantitation 203 assay 28 . Briefly, at the incubation endpoint, the plate was washed out using distilled water 204 (dH 2 0). After three washes, 150 µL of 1% aqueous Crystal Violet solution was added to each 205 well and the samples were incubated for 10 min at room temperature. The stained wells were 206 washed three times using dH 2 0, 200 µL of 30% aqueous acetic acid was added to solubilize the 207 dye, and the OD 495 was measured. 208 209

Pellicle biofilm imaging 210
Cultures were prepared as described above and diluted cultures were grown in 35 mm glass 211 bottom dishes for 5 h at 30°C. Media was removed and the pellicles were stained using 212 10 BacLight. The pellicles were imaged using a Zeiss 780 inverted confocal microscope and 213 biomass was calculated using Comstat 2.1. 214 215

Psl and matrix protein staining 216
To stain the biofilm extrapolymeric substance, 100 µg/mL HHA-FITC lectin (USA Biologicals) 217 was added to cocultures 2 h before the imaging timepoint, and the samples were incubated at 218 37°C with 5% CO 2 . Pellicle samples were treated with 100 µg/mL HHA-TRITC (USA Biologicals) 219 lectin for 2 h before the final imaging timepoint. For protein staining, pellicles were stained with 220 Ruby FilmTracer SYPRO Ruby Biofilm Matrix Stain (ThermoFisher Scientific) for 5 min before 221 imaging, as described in the reagent protocol. All samples were imaged using a Zeiss 780 222 inverted confocal microscope and staining was quantified using Comstat 2.1. 223

OMV isolation and supernatant fractionation 225
Bacterial cultures were grown overnight in LB at 37°C, diluted 1:100 into 1.5 L LB, and grown 226 (37°C, shaking) to OD 600 0.9-1.1. Cells were pelleted (10,000xg for 15 min 4°C), and 227 supernatants were collected and concentrated to 100 mL using tangential flow with a 10 kDa 228 samples were further concentrated using Millipore 10,000 MWCO centrifugal filters, and the 235 soluble proteins were separated from the OMVs using an iodixanol (Optiprep, Sigma) density 236 gradient. Concentrated proteins and OMVs were adjusted to 50% Optiprep/HEPES in 2 mL, and 237 applied to the bottom of the gradient tube, and subsequently layered with 2 mL of 40%, 2 mL of 238 35%, 4 mL of 30%, and 2 mL of 25% Optiprep/HEPES. The gradient was centrifuged overnight 239 at 40,000xg, 4°C in an ultracentrifuge (SW 40 Ti swing bucket rotor, Beckman Coulter). 240 Gradient fractions were removed from the top in 1 mL aliquots, and the lipid and protein content 241 for each fraction were identified using FM4-64 and Bradford assays as described previously 29 . 242 The OMV-containing fractions (typically 1-5) and soluble protein fractions (typically 7-12) were 243 pooled and diluted with HEPES, then centrifuged at 40,000xg for 2 h to pellet OMVs and 244 proteins and to remove Optiprep. Protein and OMV pellets were resuspended in HEPES, and 245 sterile-filtered (0.4 µm PVDF, VWR). 246 247 rPaAP purification 248 rPaAP expression, purification, and refolding were carried out as described previously 15 , and the 249 refolded protein was concentrated using 10,000 MWCO Amicon filter units (Millipore-Sigma). 250 The concentrated material was then dialyzed using the same method described above, applied 251 to a S200 Sephadex ion exchange column, eluted with a gradient of 0-2 M NaCl, pH 7.6, at a 252 flow rate of 0.02 mL/min, and 18-2 mL fractions were collected. Fractions were assayed for 253 leucine aminopeptidase activity as described 30 , and analyzed by SDS-PAGE and Ruby staining 254 as described above. Fractions containing pure rPaAP were pooled and concentrated using 255 Amicon 10,000 MWCO filter units to ~0.5 µg/mL. 256 257

Complementation and detachment experiments 258
For biofilm complementation assays, CFS, OMV fractions, soluble fractions, and rPaAP 259 samples were prepared as described above, total protein was measured by Bradford assay, and 260 aminopeptidase activity was measured as described above. To examine the hypothesis that PaAP impacts bacterial microcolony formation, we adapted a 291 previously published biofilm coculture assay for use with A549 lung epithelial cells 26  To first establish the coculture conditions for our biofilm assays in which we would use a 297 clinical P. aeruginosa isolate (S470 WT), the confluent A549 cells were inoculated with 298 fluorescently labeled S470 and incubated for up to 6 h, and the A549 cells were assessed for 299 damage and breaks in the cell layer. We established that this biotic substrate could be used to 300 investigate clinical P. aeruginosa strain biofilm development through the early stages (up to 5 h 301 post infection, hpi) of microcolony formation when LDH activity remained low (Fig S1) as 302 substantial breakdown of these cellular layers occurred after 6h. We noted that once these 303 cytotoxic bacteria infiltrated the host cell layer, the microcolony structure also broke down, as 304 the bacteria were able to colonize the underlying tissue culture dish and migrate to the air-liquid 305 interface. These results were consistent with the previously described toxicity of P. aeruginosa 306 clinical strains to host cells 31 and time courses for static coculture biofilm models 26 . In addition, 307 we confirmed that the bacterial colonies exhibited biofilm properties, as they displayed gene 308 expression patterns similar to those documented for biofilm-associated cells (Fig S2) as well as 309 increased antibiotic resistance when compared to planktonically grown bacteria (Fig S3). 310 To examine the effect of PaAP on P. aeruginosa biofilm microcolony growth, we 311 quantitatively compared cocultures of fluorescent S470 WT to S470 ∆PaAP, a previously 312 described isogenic knockout strain (S470 ∆PaAP) that contains a disruption of PA2939, the 313 gene encoding PaAP 24 . We observed significant differences between the two biotic biofilms at 5 314 hpi. Interestingly, the PaAP mutant strain formed biofilms with substantially greater cellular 315 biomass compared to the WT strain (Fig 1A, B). We also examined S470 WT and ∆PaAP 316 microcolony morphologies (Fig 1D, E). In addition to their greater quantitative biomass, S470 317 ∆PaAP colonies displayed greater cellular organization. Even at this early point in biofilm 318 development, characteristic mushroom-shaped colony structures were observed for the PaAP 319 mutant. The S470 WT microcolonies, by comparison, showed very little cellular organization 320 and were packed less densely. Based on these data, we can conclude that the lack of PaAP not 321 only increases cellular biomass, but also cellular organization at the microcolony level during 322 early coculture biofilm development. 323 To confirm this PaAP-dependent phenotype in a different strain background, we 324 compared the biofilm characteristics of PAO1, a laboratory P. aeruginosa strain commonly 325 studied in vitro, and the isogenic PAO1 ∆PaAP strain that harbors a transposon insertion in 326 PA2939. We note that our lab previously showed that OMVs isolated from PAO1 cultures show 327 very low aminopeptidase levels compared to S470 and other clinical P. aeruginosa isolates 13 . 328 Nevertheless, as with the clinical strain phenotype, PAO1 ∆PaAP formed significantly more 329 biomass as compared to the WT counterpart (Fig 1A, C). Our observations indicate that PaAP 330 is important to biofilm development, even in strains with low endogenous PaAP expression, and 331 that its role may be relevant to a broad range of P. aeruginosa strains. 332

Increased biofilm cellular density induced by PaAP deletion coincides with PaAP 334 expression and increased EPS production 335
To detail the time course of the PaAP-dependent phenotype, we imaged the cocultures 336 at hourly intervals post infection. Directly following the wash step at 1 hpi and up to 3 hpi, we 337 observed a similar level of biomass from the S470 WT and ∆PaAP strains. A significant 338 phenotype did not develop until 4 hpi (Fig 2A), suggesting that the aminopeptidase is critical in 339 limiting microcolony growth and development rather than during the initial steps of colonization 340 and attachment. Based on this timeline, it is expected that the observed phenotype would 341 develop in parallel with the level of aminopeptidase expression. Indeed, previous reports found 342 PaAP to be regulated by the las quorum sensing system 14,32 , and thus PaAP expression would 343 be expected to increase as bacterial density increases in our experimental system. Using qRT-344 PCR, we examined the expression of PaAP in S470 WT biofilms over time under the coculture 345 assay conditions. Consistent with quorum sensing density-dependent expression, our results 346 show expression of PaAP mRNA at 1 hpi, increasing steadily over the next several hours (Fig  347   S4) and PaAP protein in cocultures at the 5 hpi time point (Fig 2B). Based on these data, it 348 appears that the biofilm limiting phenotype may depend on either a threshold level of 349 aminopeptidase or a specific microcolony developmental phase. 350 Bacterial fluorescence images from the coculture assay in Fig 1 revealed significant  351 density and structural differences between the biofilms formed by WT and ∆PaAP P. aeruginosa 352 strains, but we were also interested in whether PaAP affected the composition of the biofilm 353 matrix. The P. aeruginosa matrix is composed of polysaccharides, proteins, DNA, and vesicles 354 secreted by cells in the microcolony. We imaged and quantitated the polysaccharide content of 355 the matrix using a lectin specific for Psl, a major component of non-mucoid matrices. As seen in 356 Fig 2C and 2D, the S470 microcolonies contained significantly more Psl per cellular biomass. 357 This also correlated with a modest, but significant increase in colistin resistance (Fig 2E), on a late-stage phenotype, they also observed that PaAP deletion led to increased biomass in 373 early biofilms. By growing the bacteria planktonically in Jensen's media to mid-log phase and 374 diluting and inoculating the culture containers, we were able to replicate this phenotype with the 375 S470 clinical strain and corresponding PaAP deletion mutant (Fig 3A, B, C, Untreated). In 376 Jensen's media, the ∆PaAP strain formed pellicles with significantly greater cellular biomass. 377 Interestingly, this phenotype was not found with cultures grown in other media used frequently 378 in biofilm studies (M63 minimal media and LB + glycerol) or the cell culture media used in the 379 coculture experiments (both fresh and conditioned F-12K) (Fig S5). These data lead us to 380 conclude that PaAP-dependent effects in early biofilms are growth-condition and nutrient 381 specific. 382 To examine whether PaAP affected the matrix composition of abiotic biofilms, pellicles 383 grown in Jensen's media were stained for Psl. As with the coculture biofilms, the S470 WT 384 pellicles consisted of significantly greater exopolysaccharide per cellular biomass as compared 385 to ∆PaAP pellicles (Fig 3D, Untreated). We were also able to stain for matrix protein content 386 using this model since the background protein levels were much lower as compared to the 387 coculture biofilms. Compared to the ∆PaAP strain, the WT pellicles exhibited a significantly 388 lower protein to Psl ratio (Fig 3E, Untreated), strengthening our conclusion that PaAP 389 expression can cause substantial changes in matrix composition of early biofilms under the 390 stated growth conditions. 391 392

PaAP + OMVs, but not soluble aminopeptidase, exhibit anti-biofilm activity 393
Complementation of the PaAP deletion phenotype was pursued next. Based on its 394 tightly-controlled, quorum sensing-regulated expression and secretion pathways, PA2939 395 complementation using plasmid-based expression has proven to be unfeasible. We therefore 396 performed biochemical complementation experiments to test the ability of secreted PaAP to 397 reconstitute the original biofilm composition in ∆PaAP biofilms. Complementation was tested 398 using cell-free supernatants (CFS) from mid-log phase cultures of S470 WT and S470 ∆PaAP 399 after confirming the respective presence and absence of PaAP activity in the S470 WT and 400 S470 ∆PaAP CFS by Western blot (Fig 4A) and aminopeptidase activity assays (Fig S6). 401 Equivalent amounts (by total protein) of CFS were added to washed S470 ∆PaAP coculture 402 biofilms 1 hpi, and the cocultures imaged 5 hpi. Addition of the PaAP-containing S470 WT CFS 403 complemented the phenotype of ∆PaAP biofilms to WT biomass levels, while the S470 ∆PaAP 404 CFS resulted in an insignificant change (Fig 4B). We concluded that P. aeruginosa CFS 405 harbors a PaAP-dependent biofilm modulating activity. 406 We wondered if secreted PaAP itself was responsible for the activity and recalled that 407 after secretion by the type II secretory system, PaAP is found both associated with OMVs and 408 as a soluble protein 24 . Therefore, to locate the source of the activity in the coculture 409 supernatants, soluble proteins and OMV-bound factors were separated using iodixanol density 410 gradients, and consistent with our previous reports 13 , aminopeptidase activity was detected in 411 both the light-and heavy-density fractions, representing the OMV-bound and soluble forms of 412 PaAP (sPaAP), respectively. Notably, whereas treatment with the OMV-containing S470 WT 413 fractions was able to significantly inhibit cellular biomass development by S470 ∆PaAP and 414 partially restore relative polysaccharide levels in these biofilms, the heavier, sPaAP-containing 415 fractions had no noticeable effect on biomass (Fig 4C, Fig S7). As expected, neither treatment 416 at 1 hpi of OMV-containing or heavy fractions of the CFS from S470 ∆PaAP cultures altered the 417 ∆PaAP coculture biomass at 5 hpi (Fig 4C). PaAP + OMVs, but not ∆PaAP OMVs, were also 418 able to complement the biofilm phenotype observed in Jensen's media pellicles and could 419 restore matrix composition in these samples (Fig 3A, C-E, Treated). These data suggested 420 OMVs were responsible for the biofilm modulating activity in a PaAP-dependent manner, and 421 that solubly secreted PaAP was not. 422 To confirm that aminopeptidase alone was not capable of restoring these phenotypes, 423 we purified and refolded recombinant PaAP (rPaAP) expressed in E. coli 15 . Consistent with the 424 results using sPaAP, purified rPaAP was also incapable of disrupting the formation of coculture 425 biofilms by the ∆PaAP strain (Fig 4D). Finally, we tested whether addition of rPaAP or sPaAP 426 could reconstitute the biofilm inhibitory activity of the OMVs isolated from ∆PaAP cultures 427 (∆PaAP OMVs). These rPaAP/∆PaAP OMV and sPaAP/∆PaAP OMV preparations also failed to 428 inhibit biotic biofilm development by the S470 ∆PaAP strain (Fig 4D). These data lead us to 429 conclude that OMVs derived from a PaAP + strain, mediate cell aggregation and matrix 430 composition at early stages of biofilm microcolony development. The fact that ∆PaAP OMVs 431 and PaAP alone or in combination with ∆PaAP OMVs were unable to limit biofilm development 432 additionally suggests that the aminopeptidase is not directly causing changes in biofilm 433 architecture, but rather modifying OMV activity and thereby indirectly changing the biofilm 434 microcolonies. 24  In order to characterize the biofilm-remodeling mechanism of the OMVs, we studied the 439 kinetics of the vesicle-mediated anti-biofilm activity. OMVs were added to cocultures either 440 before inoculation (pre-treatment), at 1 hpi, or at 4.5 hpi, (0.5 h before the final imaging time 441 point). Table 1  To test whether PaAP + vesicles could disrupt preformed microcolony structures, OMVs 448 were not added to the A549/S470 ∆PaAP cocultures until 4.5 hpi, and the biofilms imaged 30 449 minutes later. As seen in Fig 5A and B, this brief PaAP + OMV treatment resulted in a significant 450 and very substantial decrease in biomass of a preformed biofilm. To examine whether this 451 biofilm reducing activity was specific to S470 strain-derived vesicles, we purified and tested 452 OMVs derived from the PAO1 and PA14 P. aeruginosa WT strains and their corresponding 453 ∆PaAP mutants. WT OMVs from both strains disrupted the preformed S470 ∆PaAP coculture 454 biofilms whereas OMVs from the isogenic mutants did not (Fig 5A, B). OMVs also mediated 455 biofilm reduction for S470 pellicles grown in Jensen's media but not for pellicles that did not 456 show the original phenotype (e.g. those grown in M63 minimal media or tissue culture cell 457 media) (Fig S8). 458 During respiratory infections biofilms are often found as polymicrobial communities, and 459 components secreted by one species can impact the microcolonies of other strains or species in 460 nearby environments. To determine whether the anti-biofilm activity of the OMVs is species-461 specific, A549 cell layers were inoculated with K. pneumoniae, a bacterium which is often found 462 20 with P. aeruginosa in cases of ventilator-associated pneumonia 34 . S470 WT or ∆PaAP OMVs 463 were added at 4.5 hpi, and the cocultures were stained at 5 h with Congo Red to examine both 464 cellular and matrix biomass. As seen in (Fig 5C) susceptibility; however, cells undergoing a signaled dispersal response will lose this resistance 474 as they gain motility and release from the biofilm colonies. To characterize which of these 475 occurs during vesicle-mediated cellular release from coculture biofilms, the released cells were 476 collected and assayed for colistin resistance (Fig 6A). These cells displayed significantly higher 477 antibiotic resistance compared to planktonically grown cultures, suggesting that the mechanism 478 behind OMV-mediated biofilm disruption and remodeling likely involves the detachment of cells 479 or cellular aggregates, rather than a programmed dispersal response. Known biofilm-detachment factors include not only polysaccharide hydrolases, but also 490 DNase, Proteinase K and various endogenous P. aeruginosa proteases 36 . Based on the 491 importance of endogenous proteases to biofilm development, we tested PaAP + and ∆PaAP 492 OMVs for overall protease activity. Interestingly, we found that PaAP + OMVs display higher 493 protease activity, which cannot be directly accounted for by the presence of aminopeptidase in 494 these samples, as demonstrated by the inclusion of an rPaAP control matched to 495 aminopeptidase activity levels in the PaAP + vesicles (Fig 6C). By pre-incubating the vesicle 496 samples with a protease inhibitor (PI) cocktail, we were able to eliminate both protease and anti-497 biofilm activity in the PaAP + OMVs (Fig 6C, D) without impacting aminopeptidase activity in the 498 samples (Fig S9). While future work will be required to fully elucidate the proteases involved in 499 this phenotype, these results suggest that PaAP expression in P. aeruginosa increases 500 endogenous protease activity that becomes localized to secreted OMVs, which can in turn lead 501 to OMV-induced cell detachment and contribute to remodeling of overall biofilm architecture. 502 503

Discussion 504
We set out to investigate the role of the PaAP leucine aminopeptidase in biofilm 505 development, as PaAP is one of the most abundant proteins in the P. aeruginosa biofilm 506 extracellular matrix and one of the major protein components of OMVs from clinical P. 507 aeruginosa isolates 13,20 . When we compared coculture biofilms of wild-type P. aeruginosa and a 508 PaAP deletion mutant, we found the mutant developed early biofilms with a significantly higher 509 cellular biomass and a lower concentration of the matrix exopolysaccharide Psl, a phenotype 510 confirmed in specific pellicle biofilm growth conditions. The mutant phenotype included a less 511 robust biofilm architecture that rendered the cellular community more susceptible to the 512 antibiotic colistin. We further found that biofilm detachment activity resided in OMVs in 513 22 supernatant fractions from wildtype cultures, but not OMVs secreted by ∆PaAP cells. The effect 514 of PaAP expression was indirect, as neither recombinant nor purified soluble aminopeptidase 515 alone or in combination with ∆PaAP OMVs possessed detachment activity. Our data further 516 lead us to conclude that PaAP influences changes in biofilm architecture through proteolytic 517 activity, as WT OMVs exhibited higher proteolytic activity than ∆PaAP OMVs, and WT OMV-518 mediated detachment was sensitive to protease inhibition. OMVs have been implicated in the 519 process of biofilm formation 7,19 , but while their presence in these bacterial communities has 520 been documented, active roles for these particles and the bacterial products they carry have not on the involvement of PslG in biofilm disruption, which was also, intriguingly, PaAP-dependent 531 33 . However, in this case, biofilm disruption was found to take place in late log-stage biofilms 532 under nutrient-limiting conditions such that the lack of PaAP lead to cell lysis, releasing the 533 periplasmic PslG enzyme. By contrast, the biofilm phenotype we observed not only does not 534 depend on starvation, it is mediated by OMVs and is independent of PslG expression. 535 Additionally, AlgL and rhamnolipids, which have biosurfactant properties that can cause the 536 release of biofilm cells, were shown not to be involved in this effect. These results demonstrate 537 a distinct role for both PaAP and OMVs in the modulation of biofilm communities. 538

23
Other described mechanisms to influence bacterial detachment involve protease activity, 539 which is important in many aspects of P. aeruginosa biology 38 . Previous studies have implicated 540 endogenous proteases in biofilm detachment and the dissolution of cellular aggregates 39 , 541 specifically those with a protein rich matrix. Based on our data we can add new details to this 542 model: endogenous PaAP expression helps regulate protease activity in secreted vesicles 543 which, in turn, lead to the remodeling of biofilm architecture (Fig 7). The changes induced by 544 OMVs ultimately increase Psl/biomass ratios in early biofilm matrices and help to protect the 545 developing colony from disruption by antimicrobials. As for the specific protease(s) and 546 substrate(s) involved in his mechanism, protease expression and activation during P. 547 aeruginosa development is a highly complex, regulated process with many steps, including 548 expression, secretion, and processing. Additionally, many of these proteases are involved in the 549 activation and expression of others 15,38,40 . This complex system makes it challenging to specify 550 which protease(s) are affected by PaAP expression, whether this is the effect of one or several 551 endogenous proteases, and how PaAP increases vesicle-associated protease activity. The 552 protein substrate(s) for the OMV-activity are also unknown and also require further study, 553 however one interesting observation is that both early biofilms of P. aeruginosa and K. 554 pneumonia contain a substrate for the activity. 555 While some aminopeptidases have been implicated in pathogenesis phenotypes 41 , these 556 enzymes are generally thought to act during starvation responses, in which they can provide 557 necessary amino acids for cellular processes. This study, by contrast, presents a novel role for 558 a bacterial aminopeptidase, as PaAP impacts early biofilm formation processes early in 559 development and under nutrient-rich conditions. Additionally, because we have shown that this 560 phenotype is not directly caused by the vesicle-associated aminopeptidase, but rather other 561 vesicle-associated proteases, we have implicated PaAP in a more complex regulatory network 562 that influences the activity of bacterial molecules and structures important to the formation of 563 24 robust biofilm structures. Future work on this project will seek to elucidate the mechanism by 564 which PaAP influences OMV-mediated biofilm maturation.


extfieldsofstudy: Medicine Biology

s2fieldsofstudy: Biology Chemistry

text:
An Evolutionary View of Trypanosoma Cruzi Telomeres

Like in most eukaryotes, the linear chromosomes of Trypanosoma cruzi end in a nucleoprotein structure called the telomere, which is preceded by regions of variable length called subtelomeres. Together telomeres and subtelomeres are dynamic sites where DNA sequence rearrangements can occur without compromising essential interstitial genes or chromosomal synteny. Good examples of subtelomeres involvement are the expansion of human olfactory receptors genes, variant surface antigens in Trypanosoma brucei, and Saccharomyces cerevisiae mating types. T. cruzi telomeres are made of long stretches of the hexameric repeat 5′-TTAGGG-OH-3′, and its subtelomeres are enriched in genes and pseudogenes from the large gene families RHS, TS and DGF1, DEAD/H-RNA helicase and N-acetyltransferase, intermingled with sequences of retrotransposons elements. In particular, members of the Trans-sialidase type II family appear to have played a role in shaping the current T. cruzi telomere structure. Although the structure and function of T. cruzi telomeric and subtelomeric regions have been documented, recent experiments are providing new insights into T. cruzi's telomere-subtelomere dynamics. In this review, I discuss the co-evolution of telomere, subtelomeres and the TS gene family, and the role that these regions may have played in shaping T. cruzi's genome.

INTRODUCTION
Trypanosome cruzi causes Chagas disease a debilitating and often lethal malaise affecting millions of people in Latin American countries. T. cruzi populations are very variable and this variability is due in part to a clonal population structure, genome plasticity and abundance of repeated sequences. Nearly 50% of the total genome is made of repeated sequences, some of which code for protein families such as the Transialidases (TS), mucins (MUC), mucins associated proteins (MASP), Disperse gene family 1 (DGF-1), Retrotransposon Hot Spot proteins, and retrotransposons elements .
The Trans-sialidase family is divided into eight groups (Freitas et al., 2011), and the family name derives from enzymes included in the first group that can transfer host sialic acid onto the parasite's surface MUC. The rest of the TS groups have no enzymatic activity but code for surface glycoproteins involved in infectivity, adhesion, and evasion from the host immune response. MUC are not only receptors for sialic acid (Di Noia et al., 1995) but together with MASP proteins (Bartholomeu et al., 2009) provide a developmentally regulated shield that protects the parasite in hostile environments. DGF-1 are integrin-like proteins (Gonzalez et al., 2009;Kawashita et al., 2009;Lander et al., 2010) that are developmentally regulated, but whose role has not been deciphered. The Retrotransposon Hot Spot (RHS) family codes in T. brucei for nuclear and perinuclear proteins and their sequences are targets for the insertion of RIME/ingi non-LTR retrotransposons (Bringaud et al., 2002). In T. cruzi RHS genes are frequently interrupted by the non-LTR retrotransposon LTc1 (Bringaud et al., 2006).
Early experiments in Manning's lab (Peterson et al., 1989;Ruef et al., 1994) revealed that two members of the Transialidase family type II (TSII) located in the vicinity of the telomere were highly expressed, whereas another member, in a more internal chromosomal location, showed a lower expression level (Peterson et al., 1989). After this original report, Freitas-Junior et al. (1999) and Chiurillo et al. (1999) confirmed the presence of TSII gene members in the subtelomeres of several T. cruzi strains. The cloning of T. cruzi's telomeric and subtelomeric regions using a vector-adaptor complementing the last nine nucleotides of the telomere (Chiurillo et al., 1999(Chiurillo et al., , 2002Kim et al., 2005), allowed us to define the sequence of the telomeric repeat, and have a detailed view of a large segment of the subtelomere.
Subtelomeres are often described as regions of genomic instability (Baird, 2018), and this observation is supported by the abundance of retrotransposon fragments and pseudogenes in T. cruzi telomeric clones, but they also represent a buffer region to minimize chromosome damage when telomere attrition occurs. However, despite this unstable environment, complete gene copies of DGF-1, TS, and RHDEAD/H-RNA helicase and N-acetyltransferase in T. cruzi subtelomeres are expressed, indicating that a positive selection is preserving the integrity of these genes at these locations (Moraes Barros et al., 2012). At the end of subtelomeres, there is a 189 base pairs sequence with homology for the 3 ′ and 5 ′ UTRs sequences of a gp85 gene (TSII) that we dubbed the 189 bp junction (Kim et al., 2005). A recapitulation of the possible events associated with the creation of this junction was proposed by Kim et al. (2005), Chiurillo et al. (2017) in which tandem repeats of TSII genes suffered breakages, excisions and rejoining to generate the junction. After this junction, the chromosomes are capped by runs of hexameric repeats of variable length ending in a single strand terminus 5 ′ -TTAGGG-OH-3 ′ .

THE EVOLUTION OF T. cruzi TELOMERES AND THE ROLE OF MEMBERS OF THE TSII GENE FAMILY IN SHAPING ITS GENOME
The intimate association between telomeres and TSII (gp85) sequences raises many questions about the creation of T. cruzi telomeres, such as how and when TS II family members were incorporated into the 189 bp junction? Were they part of transposon elements that eventually evolved into a primitive telomere? Did they come along with the telomeric repeats?
When we examine the structure of eukaryotic telomeres we observe a tremendous diversity in telomeric repeats and sequences associated with telomeres, the extreme case of Diptera where transposons assumed the role of telomeres. Telomeres arose during eukaryogenesis as a need to protect the ends of linear chromosomes from exonucleases degradation, prevent chromosome rearrangements, and facilitate the replication of the DNA lagging strand. This process may have started with the endosymbiosis of an ancestral phagotrophic host cell (Cavalier-Smith, 2002) and an α-proteobacterium carrying Type II introns. Type II introns are Eubacterial mobile retroelements with ribozyme and reverse transcriptase activities. Eventually, Type II introns were transferred into the host cell genome and gave rise to the spliceosome, the non-LTR transposons and telomerase (Garavis et al., 2013;Podlevsky and Chen, 2016). According to this hypothesis, these non-LTR transposons were inserted at several genome locations, including the chromosome ends, originating the prototelomeres.
On the other hand, although still debatable, based on the sequence homogeneity of the transialidases within the animal kingdom, and their sequence homology with their very variable bacterial counterparts (Roggentin et al., 1993) proposed that sialidases originated in ancestors of the Echinodermata and Deuterostomate animals, and later these genes were horizontally transferred to bacteria via viruses. However, Schwerdtfeger and Melzig (2010) argued that the irregular appearance of sialidases in invertebrates does not support a common evolution of this gene family. In agreement with this view, in the Trypanosomatidae family, a taxon that includes pathogenic flagellates, we also observe this kind of patchy inheritance, since of all genera within this family the TS genes are only present in the genus Trypanosoma.
It is assumed that the first trypanosome to acquire sialidase genes was T. brucei or a common ancestor of both T. brucei and T. cruzi. The separation Trypanosoma brucei from Trypanosoma cruzi occurred approximately 100 millions of years during the Gondwanaland breakage (Briones et al., 1995;Stevens et al., 1999), and since then, the TS family suffered an expansion exclusively in the T. cruzi clade (Chiurillo et al., 2016a). During this long separation, the two species developed different survival strategies within their vectors and hosts. T. brucei uses the telomeric-subtelomeric compartment as a specialized expression site for variable antigenic determinants. On the contrary, T. cruzi acts like a stealth invader jamming the vertebrate host immune system through a simultaneous expression of multiple surface antigens (Millar et al., 1999), and among these, members of the TS family play an important role.
How the expansion of TSII genes occurred in T. cruzi is unknown, but likely, several events of gene mutation, duplication, recombination, transposition, and genetic drift generated the current picture of the TS family. Vestiges of these events are the salad of sequences derived from retrotransposon elements and Retrotransposons Hot Spot (RHS) genes scattered through the genome, and particularly in T. cruzi subtelomeres (Figure 1). The abundance of these elements at the subtelomere suggests the active participation of these regions in shaping the parasite genome (Kim et al., 2005). Regarding the telomeric repeat 5 ′ -TTAGGG-OH 3 ′ , it is found in Plantae, Chromoalveolata, Excavata (where Trypanosomatids are included) and Rhizaria supergroups, a reason why some authors (Dressen et al., 2007;Fulneckova et al., 2013) have suggested that it was the repeat of the ancestral Eukaryotic telomeres. Thus, a safe assumption is that the TS sequences in the subtelomeric transition (189 bp junction) appeared after the hexameric repeats were already fixed, and perhaps the fixation of the 189 bp junction has to do with its adaptation to the telomeric function, thus this sequence appears to be an integral part of T. cruzi telomere. Chiurillo et al. (2017) have proposed that the 189 bp junction may act as a seed to stabilize and/or create new telomeres via telomerase, but it is also possible that it was adopted as a recognition site for other proteins that contribute to the telomeric function.

GENE FAMILIES AND SUBTELOMERES
In T cruzi apart from the TS family other gene families suffered big expansions, namely RHS, DFG-1, MASP, and MUC and these genes tend to form clusters at multiple locations in the genome, in some cases covering whole chromosomes Weatherly et al., 2009;Berna et al., 2018;Callejas-Hernández et al., 2018). Except for MUC and MASP families, these expanded families are often found at chromosomal terminal locations Weatherly et al., 2009;Moraes Barros et al., 2012;Callejas-Hernández et al., 2018). Therefore it is not surprising to detect members of these families when T. cruzi telomeres and subtelomeres are cloned (Chiurillo et al., 1999;Freitas-Junior et al., 1999). Thus, the T. cruzi genome is organized in blocks of syntenic non-repeated gene sequences at more interstitial locations, and non-syntenic blocks consisting of repeated genes that are intermingled with retrotransposons and other highly repeated elements, some of which are located at subtelomeres . The presence of many repeated genes may propitiate recombination events that, although not desirable for housekeeping genes, favor the generation of variability for genes coding for surface proteins.
The uniqueness of terminal chromosomal locations (subtelomeres) as a ground for the generation of new capabilities has been well documented among other organisms in yeast (Haber, 1998), Plasmodium (Scherf et al., 2008), T. brucei (Horn, 2004), and the expansion of human odor receptors (Mefford et al., 2001). Based on these observations and in the composition of T. cruzi telomeres and subtelomeres, we proposed that subtelomeres were places where the variability of some of these multigene families was generated, and in later events, the gene variants were mobilized to different locations in the genome (Kim et al., 2005).

HYPOTHETICAL MECHANISMS TO GENERATE GENE VARIABILITY
What are the mechanisms that may generate gene variability in T. cruzi, and how has the mobilization of gene variants occurred? Is the variability generated by gene conversion or unequal crossing over? Are these processes still occurring in T. cruzi populations?
Our first attempt to address these questions was through in silico simulation studies using as inputs real gene sequences generated by the T. cruzi genome project (Azuaje et al., 2007). The simulations evaluated the generation of variability by introducing different mutagenic pressures in housekeeping genes, and members of gene families with presence at subtelomeres. The premise was that mutation rates would be a trade-off between the generation of variability to expand adaptive capabilities, and the need to keep important core functions. This study concluded that housekeeping genes were more robust against the introduction of random point mutations than genes coding for surface proteins and that the most effective mechanism to introduce variability was gene conversion. The energetic burden of keeping a large number of pseudogenes is an indication that they play an important role in the parasite, an observation that prompted us to include pseudogenes of RHS, TS, and DGF-1 in our simulations. The results confirmed the potential of pseudogenes to contribute to the generation of variability. This finding contradicts the idea that pseudogenes are merely relics of gene deterioration (Rogers et al., 2011) or pseudogenization.
An important piece of information came from experiments using T. cruzi artificial chromosomes (pTAC) carrying the 189 bp junction, hexameric repeats, and drug selection markers (Curto et al., 2014). When T. cruzi epimastigotes were transformed with these pTCAs, they were able to replicate with surprising stability for 150 generations in the presence of the selection drug, or 60 generations without it. In other words, the pTACs showed nondetectable sequence exchange with the host chromosomes and replicated and segregated without the presence of centromeres, or perhaps the telomere was fulfilling this role. In a more recent experiment (Chiurillo et al., 2016b), addressed the possibility of chromosomal sequence exchanges by introducing a cutting site for the rare meganuclease I-SceI within the RHS gene of one pTACs (pTAC-D6CISceI * ) harboring larger portions of T. cruzi subtelomeres. After transforming this pTAC into T. cruzi cells expressing the meganuclease I-SceI, and confirming that double-strand breaks (DBSs) were produced, the probing pTAC was examined to check whether the DSBs were repaired. Out of seven clones studied, six showed repairs, as evidenced by the disappearance of the I-SecI site, and the reversion to the original pTAC (pTAC-D6C * ). The most likely explanation is that the repair events used as template the host subtelomere homologous to the pTAC. The seventh clone was repaired losing the I-SecI site, but the sequences around the repair site shared homology with the subtelomere of another chromosome (ectopic recombination). From these experiments several conclusions can be derived: first chromosomal exchanges at the subtelomere can be promoted by the introduction DSBs; second the repair mechanism for these DSBs is homologous recombination (HR), or some version of it (Dressen et al., 2007); third, events involving exchanges with non-homologous chromosomes (ectopic recombination) can occur, with the possibility of generating gene variants. These experiments don't rule out that similar exchanges can occur at more interstitial locations. The absence of detectable recombination in earlier pTAC experiments was likely due to a lack of sufficient homology with any given chromosome subtelomere, and/or that DSBs are strictly necessary to induce recombination.
Contrary to most eukaryotes in T. brucei and T. cruzi the most important repair mechanism for DSBs repair is HR, with minor participation of microhomology repair (MHR). To detect MHR-DSB repair, it was necessary to abolish HR (Glover et al., 2008).
In T. cruzi out of the potential genes that participate in HR, RAD51 plays a central role in facilitating homologous strand invasion (Gomes Passos Silva et al., 2018). An interesting activity discovered after massive gamma radiation of T. cruzi cells was tyrosyl-DNA phosphodiesterase I (Tdr-I), an enzyme that plays an important role in Topoisomerase I mediate DNA DBS repair (Das et al., 2010). No genes for NHEJ have been found in T. cruzi (Gomes Passos Silva et al., 2018). MHR seems to occur in chromosomal rearrangements when single DSB is introduced by Cas9 endonuclease (Lander et al., 2015;Soares Medeiros et al., 2017).
An efficient HR repair mechanism in T. cruzi may explain the rapid karyotype and cell growth recovery after massive irradiation with 500 Gy of gamma radiation (Garcia et al., 2016), and the difficulty in generating mutations (indels) in CRISPR experiments involving a single DSB vs. gene replacement (Lander et al., 2015;Soares Medeiros et al., 2017). But also, the prevalence of a very stringent HR repair mechanism in genomes with a large number of repeated sequences, like T. cruzi, hampers (but does not eliminate) chromosomal-internal recombination events that can be detrimental to the organism.
Thus, we believe that an important source for the generation of variability in some T. cruzi surface proteins is subtelomeric recombination promoted by DSBs followed by a dispersion of these variants either by transpositions or ectopic recombination events. Further duplications events and genetic drift produced the current clusters that we see at several locations of the T. cruzi's genome.
Along this line, experiments with meganuclease I-Sce-I in T. brucei (Dressen et al., 2007) revealed that the introduction of a DSB in telomeric VSG gene promoted antigenic switching via gene conversion. The break is resolved by a replication mechanism induced by this break (Break Induced Replication, BIR). Similar detailed studies to determine how DBSs repair occurs in T. cruzi are missing due in part to the lack of an RNAi machinery and/or inducible promoters coupled to CRISPR-Cas9 vectors.
As mentioned before, despite this stringent HR mechanism interstitial chromosome rearrangement leading to karyotype changes can occur via ectopic recombination within multigene families through duplicated sites flanking these sequences (Figures 1A,B).

HOW DSBs CAN BE INTRODUCED IN T. cruzi SUBTELOMERES?
In T. cruzi retrotransposon elements occupy nearly 5% of its genome and among these elements, the L1Tc non-LTR retrotransposon has the necessary machinery for its mobilization (Macías et al., 2018). L1Tc genome codes for an AP endonuclease activity (NL1Tc) capable of introducing breaks for the insertion of the retrotransposon, but also plays a role in repairing DSBs produced by daunorubicin (Olivares et al., 2013). Other retrotransposon elements like SLAKS and CZAR code for sitespecific endonucleases (Macías et al., 2018). So, some DSBs may be introduced in the subtelomeres by retrotransposon nucleases, and HR repairs the break using as templates homologous chromatids, or on occasion non-homologous chromatids (ectopic recombination). In T. cruzi there are active L1Tc transposons and no RNAi machinery to counteract their activities. Several observations about the organization of multigene families reveal close associations with retrotransposon sequences i.e., most DGF-1 and TSII copies are flanked by RHS (Olivares et al., 2000;Kim et al., 2005) or L1Tc retrotransposons sequences, the duplications of RHS and L1Tc genes at both sides of these genes suggest the occurrence of ectopic recombination events (Figure 1; Olivares et al., 2000). Although L1Tc appears to be randomly distributed in T. cruzi's genome, 50% of its copies are associated with RHS genes flanked by the putative insertion site 5 ′ -TGCAGACAT-OH-3 ′ (Olivares et al., 2000; Figures 1B,C). Also, they are found inserted downstream the sequence GA (x) 2 AxGa (x) 5 txTATG↑A(x) 11 ↑ where arrows mark the single strand cleavage sites Bringaud et al., 2006). How frequent DBSs leading to genetic recombination and gene variability occurs is difficult to assess, since as shown in the experiments with meganuclease I-Sce-I (Chiurillo et al., 2016b), DBSs are mainly repaired by HR using as template homolog chromosomes.
The MASP superfamily is associated with the site-specific retrotransposon TcTREZO (Souza et al., 2007). These elements are frequently found flaking MASP genes, thus providing potential sites for HR. Since TcTREZO is species-specific, it must have appeared after the separation of T. cruzi and T. brucei, and it may have played an important role in the expansion of the MASP family. MASPs proteins present highly conserved N and C terminal sequences, and a variable middle region, also besides, their gene's 5 ′ and 3 ′ UTRs are highly conserved (Bartholomeu et al., 2009). So sequence evolution in this gene family is quite different from the rest of the repeated families. Interestingly other retrotransposons and members of the gp85, DFG-1, and RHS gene families frequently interrupt MASP clusters.
Duplication and mobilization of genes can also occur via piggybacking the retrotransposon reverse transcriptase machinery by the transduction of genes neighboring the retrotransposons insertion site ( Figure 1D). The termination signal for transposon transcriptases is usually weak, thus transcription can run through neighboring segments which can be duplicated and mobilized elsewhere (Xing et al., 2006). So far no experiments have been conducted to address this type of event in T. cruzi, although the size of the DNA segments that non-LTR retrotransposons can transduct is usually small (1 or 3 Kbp), making it an, unlike mechanism to mobilize large genes like DGF-1 (>10 Kbp).
In T. brucei, in the case of non-recombinational gene conversion for antigenic switching, alternatives for the generation of subtelomeres DSBs have been proposed, such as accelerated transcription, conflicts between replication and transcription machineries , and TERRA (Telomeric Repeat-containing RNA) transcription leading to the formation of R-loops (Nanavaty et al., 2017;Saha et al., 2019). Since antigenic variation is a vital phenomenon for the survival of T. brucei populations, it is not surprising that redundant mechanisms exist to make sure that antigenic switching occurs. In the case of T. cruzi, not such extensive studies have been done, and it is possible that R-loops formation or transcription-replication conflicts may also contribute to the generation of subtelomere DBSs. In this review, I favored the role of retrotransposons in the generation of DBSs, given the ubiquity of these elements in the T. cruzi genome, their close association with important surface antigens families, and the absence of RNAi machinery.

CONCLUSIONS
Once T. cruzi telomeres were fixed, members of the TS II family positioned at the subtelomeres co-evolved to be part of the transition to the telomeric repeat. Although the reason for the fixation of this junction is still unknown, it suggests a potential telomeric function for this region. The variability of some surface proteins and their localization at the subtelomeres together with retrotransposon elements suggests that these regions are grounds for the generation genetic variability. We propose that DBSs introduced in the subtelomeres by retrotransposon nucleases are repaired by homologous recombination, and when the repair includes non-homologous chromatids there is a possibility to generate gene variants. These variants are mobilized elsewhere either by transposition or ectopic recombination. Gene families increased their numbers by gene duplication to achieve higher expression levels. MUC and MASP superfamilies likely evolved in later events not related to the subtelomeres.

AUTHOR CONTRIBUTIONS
The author confirms being the sole contributor of this work and has approved it for publication.


extfieldsofstudy: Chemistry Medicine

s2fieldsofstudy: Biology Medicine

text:
Effect of Adhesion and Substrate Elasticity on Neutrophil Extracellular Trap Formation

Neutrophils are the most abundant type of white blood cells. Upon stimulation, they are able to decondense and release their chromatin as neutrophil extracellular traps (NETs). This process (NETosis) is part of immune defense mechanisms but also plays an important role in many chronic and inflammatory diseases such as atherosclerosis, rheumatoid arthritis, diabetes, and cancer. For this reason, much effort has been invested into understanding biochemical signaling pathways in NETosis. However, the impact of the mechanical micro-environment and adhesion on NETosis is not well-understood. Here, we studied how adhesion and especially substrate elasticity affect NETosis. We employed polyacrylamide (PAA) gels with distinctly defined elasticities (Young's modulus E) within the physiologically relevant range from 1 to 128 kPa and coated the gels with integrin ligands (collagen I, fibrinogen). Neutrophils were cultured on these substrates and stimulated with potent inducers of NETosis: phorbol 12-myristate 13-acetate (PMA) and lipopolysaccharide (LPS). Interestingly, PMA-induced NETosis was neither affected by substrate elasticity nor by different integrin ligands. In contrast, for LPS stimulation, NETosis rates increased with increasing substrate elasticity (E > 20 kPa). LPS-induced NETosis increased with increasing cell contact area, while PMA-induced NETosis did not require adhesion at all. Furthermore, inhibition of phosphatidylinositide 3 kinase (PI3K), which is involved in adhesion signaling, completely abolished LPS-induced NETosis but only slightly decreased PMA-induced NETosis. In summary, we show that LPS-induced NETosis depends on adhesion and substrate elasticity while PMA-induced NETosis is completely independent of adhesion.

INTRODUCTION
Neutrophilic granulocytes are the most abundant type of circulating white blood cells. In a process termed NETosis, they release neutrophil extracellular traps (NETs), web-like structures composed of decondensed chromatin decorated with antimicrobial proteins (1,2). During NETosis, the nuclear chromatin swells until both the nuclear envelope and the outer cell membrane rupture (3). NETosis is considered an important immune defense mechanism as neutrophils can bind and kill bacteria and other pathogens via NETs. Apart from physiological stimuli such as pathogens, chemokines (e.g., CXCL8), activated platelets or urea crystals there are several additional NET-inducers like phorbol 12-myristate 13-acetate (PMA) and lipopolysaccharides (LPS), which induce NETosis in vitro (4). Even though NETosis was initially described as part of the innate immune defense system, we know today that dysregulated NETosis is also involved in a variety of chronic inflammatory and autoimmune diseases such as atherosclerosis, systemic lupus erythematosus, preeclampsia, as well as malignant diseases (5)(6)(7)(8). Therefore, the question which environmental factors play a role in this process and may influence the course of diseases is highly important.
Mechanical properties of tissues are environmental signals that are able to modulate the functionality of surrounding cells. This has been demonstrated by a substantial amount of studies investigating the effect of physical factors on cellular functions (9)(10)(11)(12)(13). It has previously been shown that phenotype and functionality of immune cells such as macrophages and dendritic cells are affected by substrate elasticity/stiffness (14)(15)(16). It has also been reported that substrate elasticity affects neutrophil adhesion, migration, and chemotaxis (17)(18)(19). Transmigration of neutrophils through endothelium was also proven to be affected by sub-endothelial cell matrix stiffness (20). Tissue stiffness increases in multiple pathological processes including, most prominently, atherosclerotic plaques (21) but also fibrosis (22) and cancer (23). In general, cell adhesion is mediated through surface receptors interacting with specific ligands presented on surfaces (24)(25)(26). Integrin ligands have been previously shown to play an important role in leukocyte adhesion and migration (27)(28)(29). Additionally, the ligand density on the surface affects adhesion and migration of neutrophils (28,30). For example, neutrophils adhere via the integrin Mac-1 to the platelet receptor GPIbα and show the fastest adhesion maturation when ligands are distributed at a medium distance of approximately 100 nm (28).
In mice, it has been shown that blocking of the integrin LFA-1 prevented NETosis in a model of endotoxemia (31). Similarly, mice lacking the beta2 subunit of integrin receptors were largely protected from overwhelming NET production in a murine model of hantavirus infection (32).
In humans, there are several studies that have investigated the involvement of integrin signaling in NET formation, although the results are partially contradictory (33,34). Thus, even though the role of integrins on neutrophil adhesion has been addressed to a certain extent, there is still need to further characterize this complex interaction in a well-defined manner. Moreover, the impact of substrate elasticity in conjuncture with adhesionrelated processes on NETosis has not been investigated yet and thus remains entirely enigmatic.
In this paper, we explore the effect of substrate stiffness/elasticity (Young's modulus E) and general adhesion on NETosis in human neutrophils (Figure 1). We use collagen Iand fibrinogen-coated polyacrylamide (PAA) gels with stiffnesses within the physiologically relevant range (E = 1-128 kPa) to study the impact of elasticity and adhesion on NETosis induced by two different stimuli (LPS, PMA).

Polyacrylamide (PAA) Gel Preparation
Polyacrylamide (PAA) gels were prepared on round glass cover slides as previously reported (9,13). In brief, the cover glasses were plasma cleaned, coated with 3-aminopropyltriethoxysilane (Sigma, Munich, Germany, A3648) and afterwards incubated with glutaraldehyde solution (0.05%, Sigma, Munich, Germany, G7651). Appropriate mixtures of acrylamide (Bio-Rad, Munich, Germany, #161-0140) and bis-acrylamide (Bio-Rad, Munich, Germany #161-0140) diluted in Dulbecco's phosphate-buffered saline (PBS, Sigma-Aldrich, Munich, Germany) were freshly prepared, stored at +4 • C and used within 2 months (see Supplementary Table S1 for details). Polymerization was initiated by addition of 1/1,000 N,N,N,Ntetramethylethylenediamine (TEMED) and 1/100 ammonium persulfate (APS) solution. Thirty-five microliter of this solution was used per cover slip. A square hydrophobic cover glass was placed on top in order to equally distribute the solution on the bottom glass. Gels were polymerized for 60 min in a saturated water atmosphere to avoid evaporation and were ∼70 µm thick. Young's modulus E of PAA gels was quantitatively controlled by measurements on a bulk rheometer (MCR-501, Anton Paar, Austria). To prevent toxicity to cells, non-polymerized residues were thoroughly washed away using PBS. Substrates were treated with the photo-activatable cross-linker Sulfo-SANPAH (Thermo Scientific, Waltham, Massachusetts, 22,589; 0.4 mM in 50 mM HEPES buffer at pH 8), exposed to UV light (λ = 365 nm) for 10 min and then either coated with rat tail collagen I (0.02 mg/mL for data presented in all main figures and 0.2 mg/mL for Supplementary Figure S3) (Corning, New York, New York, #354236) or fibrinogen (0.02 mg/mL, from human plasma, Sigma, Munich, Germany # F-3879) overnight at 4 • C unless otherwise stated.

Human Neutrophil Isolation
All experiments with human neutrophils were approved by the Ethics Committee of the University Medical Center (UMG) Göttingen (protocol number: 29/1/17). Donors gave informed voluntary consent to the study. Neutrophils were isolated according to previously published standard protocols (3,35) from healthy donors' venous blood supplemented with EDTA.
In short, fresh blood was collected with S-Monovettes KE 7.5 ml (Sarstedt, Nümbrecht, Germany). Blood was gently layered in a 1:1 ratio on top of Histopaque 1119 (Sigma-Aldrich, Munich, Germany) and centrifuged at 1,100 × g for 21 min. Then, the transparent third and pink fourth layer containing the white blood cells were collected and mixed with HBSS (without Ca 2+ /Mg 2+ , Thermo Fisher Scientific, Waltham, Massachusetts). Cells were pelleted by centrifugation for 10 min at 400 × g. After discarding the supernatant, the pellet was resuspended in HBSS without Ca 2+ /Mg 2+ and layered on top of a phosphate buffered percoll (GE Healthcare, Chicago, Illinois) gradient with the concentrations 85, 80, 75, 70, and 65% and centrifuged at 1,100 × g for 22 min. The accumulated neutrophils were received by collecting half of the 70%, full 75% and half of the 80% layer and washed with HBSS. The remaining cell pellet was resuspended in 1 ml HBSS. Cells were counted using Trypan blue solution (Sigma, Munich, Germany) and cell viability was assessed simultaneously. Viability was always >98%. The desired cell number was resuspended in Roswell Park Memorial Institute (RPMI) 1640 medium (Lonza, Basel, Schweiz) containing 0.5% heat inactivated fetal calf serum (FCS; Biochrom GmbH, Berlin, Germany). Neutrophil purity was determined by a cytospin assay (Shanson, Cytospin 2 Centrifuge) followed by Diff Quick staining (Medion Diagnostics, Miami, USA). In all experiments neutrophil purity was >95%.

Quantification of Neutrophil Extracellular Trap Formation
Freshly isolated neutrophils were seeded (0.5 × 10 6 cells/well) on PAA gels and collagen I-or fibrinogen-coated glass and incubated for 30 min at 37 • C, 5% CO 2 in order to allow the cells to adhere. NETosis was either stimulated with 5 nM Phorbol-12-myristate-13-acetate (PMA, Sigma-Aldrich, Munich, Germany) or with 75 µg/mL lipopolysaccharide (LPS) from Pseudomonas aeruginosa (serotype 10.22, strain: ATCC 27316, Sigma-Aldrich, Munich, Germany) and incubated for 3 h (37 • C, 5% CO 2 ). As vehicle control, neutrophils on 4 and 128 kPa gels coated with collagen-I or fibrinogen were incubated with RPMI media for 3 h to control for spontaneous NETosis. NETosis was stopped with 2% paraformaldehyde (PFA, Roth, Karlsruhe, Germany) fixation overnight. The next day, the fixed neutrophil chromatin was stained using 1 µg/ml Hoechst 33342, trihydrochloride, trihydrate (Hoechst) (life technologies, Carlsbad, California). Cells were imaged with an Axiovert 200 microscope (16× magnification, Zeiss, Oberkochen, Germany) being set on the blue channel (Filter set49 DAPI shift free, 488049-9901-000, Zeiss, Oberkochen, Germany) connected to a CoolSNAP ES camera (Photometrics, Tucson, Arizona). Six images of different locations were taken per well. Phase contrast images for cell contact area measurements were taken for each setting and the cell contact areas were calculated by using ImageJ 4.46r for at least 50 cells per condition. NETosis was quantified in a standardized blinded fashion as percentage of condensed/lobulated nuclei vs. decondensed nuclei and strands of extracellularly localized chromatin, as described previously in the literature (36). For better clarity, we used "decondesed chromatin" when labeling axes.

PI3K Inhibition
Human neutrophils were treated with the PI3K inhibitor BAY 80-6946 (copanlisib) for 20 min on ice and then seeded on PAA gels (4, 128 kPa) and glass followed by a 30 min incubation (37 • C, 5% CO 2 ). Afterwards cells were stimulated, fixed, and imaged as described above.

Statistics and Data Analysis
Statistics were performed with GraphPadPrism (Version 6.0, GraphPadSoftware Inc., San Diego, California). Significance was tested using standard one-way-ANOVA with Bonferroni's multiple comparisons test (ns = not significant, * p < 0.05, * * p < 0.01, * * * p < 0.001, * * * * p < 0.0001), after testing for normal distribution, where applicable. n numbers refer to the number of single cells analyzed, N numbers refer to the number of independent donors. Mean results and standard error of the mean (SEM) were calculated on the basis of the results obtained from the independent donors. Fluorescence images were processed with ImageJ.46r (National Institutes of Health, Bethesda, Maryland) and all cell counts obtained using the Plugin "Cell Counter."

Substrate Elasticity Affects NETosis in a Stimulant-Dependent Manner
To investigate the effect of substrate elasticity/stiffness on NETosis, freshly isolated human neutrophils were seeded on PAA gels coated with either collagen I or fibrinogen which are both known to interact with integrins on neutrophils (30,37). The PAA substrate elasticity was varied within the physiological range (1,2,4,8,16,20,30,64, 128 kPa). The prepared gels cover a wide range of physiological elasticities for example that of brain tissue (<1 kPa), muscle (14-16 kPa), osteoids, pre-calcified bone (30 kPa) or cartilage (>100 kPa) (9). Importantly, PAA gels without coating do not provide a physiological surface and do not promote cell adhesion. This coating procedure leads to a uniform high density distribution of the proteins on the surface and was kept constant for different gels to ensure that possible differences in neutrophil behavior were not due to increased availability of surface interaction partners (13,38).
To understand how substrate elasticity and the presence of certain integrin ligands affect NETosis, neutrophils were seeded and incubated for 30 min and then stimulated with either PMA or LPS for 3 h (Figure 2; Supplementary Figures S1, S2). As vehicle control, neutrophils on 4 or 128 kPa PAA collagen-I-or fibrinogen-coated gels were incubated with RPMI media only and did not show significant NET formation (Supplementary Figures S1, S2). PMA is a well-known activator of protein kinase C (PKC) and frequently used to induce NETosis in vitro (39). LPS on the other hand, induces NETosis in a receptor-mediated fashion (40). PMA was applied at a very low concentrations (5 nM) to avoid the strong stimulation at typical concentrations [100 nM (3)] that could blur the impact of substrate elasticity.
Formation of NETs was confirmed (Figures 2B,C on glass and Supplementary Figure S5 on gels) by co-staining of decondensed intracellular or released extracellular chromatin by Hoechst and MPO, according to commonly used protocols (41). As histone citrullination in PMA-induced NETosis is highly controversial, staining of citrullinated histones as a means of verifying NET formation was not carried out (42,43).
NETosis was imaged and quantified on both surface coatings (collagen I and fibrinogen) for both ways of stimulation (PMA and LPS) (Figure 2). Representative images for 2, 16, 128 kPa PAA gels and a glass control are shown in Figure 2. Importantly, NETs can be distinguished from apoptotic cells by their characteristic, decondensed chromatin. Counting decondensed nuclei and extracellular chromatin is a standard procedure to quantify NET-formation rates (36).
Images of all other conditions are shown in the Supplementary Figures S1, S2.
PMA-stimulated NETosis was independent of stiffness. On collagen I-coated substrates, the NETosis rate was the same for stiffness values (Figure 3A). Similarly, on fibrinogen-coated substrates, PMA-induced NETosis ( Figure 3C) did not vary with stiffness. However, on glass surfaces, NETosis was higher compared to that on substrates with defined stiffnesses of 8, 16, 20, or 128 kPa, respectively. This observation might be explained by different effective concentrations of PMA due to diffusion into the gel or adsorption onto glass. We concluded that PMA-induced NETosis is not affected by substrate stiffness.
In contrast, NETosis was significantly affected by substrate elasticity under LPS stimulation on both collagen and fibrinogencoated surfaces (Figures 3B,D). On stiffer substrates (E > 20 kPa), NETosis was significantly increased. Interestingly, NET formation on collagen-I coated glass surfaces was comparably low after LPS stimulation, which correlated well with poor spreading of neutrophils on said glass surfaces (see below, Figure 4A).

Correlation Between Neutrophil Adhesion and NETosis
To further investigate the connection between NETosis, adhesion and substrate elasticity we assessed neutrophil adhesion on the Frontiers in Immunology | www.frontiersin.org FIGURE 2 | Substrate elasticity affects NETosis in a stimulant-dependent manner. (A) Human neutrophils were seeded on PAA gels coated with collagen-I or fibrinogen. They were stimulated with either PMA (5 nM) or LPS (75 µg/mL), as indicated, and incubated for 3 h. The fluorescence images show the nuclei or extracellular chromatin (Hoechst DNA/chromatin stain) of fixed cells. The images reveal that NETosis depends on all three parameters (substrate elasticity, stimulant, surface coating). See quantification in Figure 3 and images of all conditions in Supplementary Figures S1, S2. (B,C) Neutrophils were seeded on glass coverslips and stimulated with PMA (B) or LPS (C) as described above. After fixation with PFA, cells were stained with anti-MPO antibody (red) and Hoechst against chromatin (blue) and imaged by confocal microscopy to verify NET production. Representative images of neutrophils are shown. different substrates. Of note, fibrinogen is known to activate neutrophils and promote adhesion (37). However, as fibrinogen concentration was not varied, any changes in adhesion could be attributed to the changing stiffness of the substrate. Although it is known that upon stimulation with PMA or LPS neutrophils initially adhere strongly to the provided surfaces before starting NETosis (3), it remains controversial whether this initial adhesion is a necessary prerequisite and to which extent NETosis and adhesion are correlated with one another. During NETosis, the cells round up and the cytoskeleton is degraded. There have been contradictory studies on the role of integrin receptor engagement during NET formation (33,34). The substrate stiffness most likely influences adhesion. Indeed, neutrophils have been reported to better adhere to stiff surfaces compared to soft surfaces (17,18). Therefore, we next analyzed the spreading of human neutrophils on collagen I-and fibrinogencoated substrates and investigated the correlation between adhesion and NETosis. As a surrogate parameter for adhesion we determined the cell contact area (by phase contrast microscopy, representative images shown in Supplementary Figure S4). On both fibrinogen and collagen I-coated PAA gels the cell contact area/spreading area of neutrophils increased with increasing stiffnesses (Figures 4A,B), corresponding to the measured LPSinduced NETosis rates (Figures 3B,D). Next, a cross-correlation analysis for neutrophil spreading area vs. NETosis was carried out for LPS stimulation. LPS-induced NETosis correlated well with spreading area for both coatings (Figures 4C,D). Therefore, adhesion signaling appears to be highly relevant for LPS-induced NETosis. Differences in adhesion due to different stiffnesses directly translate into different NETosis rates.
In contrast, as expected from the results in Figure 3, PMA-induced NETosis did not correlate with the spreading area ( Figure 4E). Interestingly, when the gels were coated with a 10-fold higher (0.2 mg/ml) collagen I concentration, cells did not perform LPS-induced NETosis at all (Supplementary Figure S3). We hypothesized that there is an optimum density for NET formation on collagen I, in accordance with previously published results that show an optimum ligandreceptor ratio for different biological functions including cell adhesion or spreading (24,28,44,45). Similarly, the cells did not adhere well and did not spread above the area expected from a fully settled neutrophil (Supplementary Figure S3). These observations additionally corroborate our hypothesis that conditions that affect the adhesive phenotype change NETosis rates, which suggests that adhesion and NETosis are interconnected.

PI3K Inhibition Abrogates Stiffness-Dependent Variations in NETosis
PI3K activity is important for neutrophil mechanosensing and enables these cells to distinguish between substrates of different stiffnesses (18). To corroborate that the observed effects on LPSinduced NETosis could be attributed to variations in substrate stiffness, we treated neutrophils with a potent PI3K inhibitor, BAY 80-6946 (copanlisib), and analyzed adhesion and NETosis. Fibrinogen-coated PAA gels of 4 and 128 kPa, respectively, were chosen as representative conditions for adhesion and NETosis quantification as they had revealed significant differences in the experiments presented above. Adhesion was impaired when neutrophils on stiff surfaces (128 kPa and glass) were pre-treated with the inhibitor (Figures 5A,B). PI3K is a known signaling intermediate for PMA-induced NETosis, which is why moderate effects on PMA-induced NETosis were to be expected (40). To investigate the connection between adhesion and NETosis, we treated cells with PMA or LPS under PI3K inhibition. This treatment partially impaired PMA-induced (Figure 5C), and completely abrogated LPS-induced NETosis (Figure 5D).
To further prove the importance of adhesion for NETosis, we coated glass surfaces with poly-L-lysine (PLL) and poly-L-lysine-grafted-polyethylene glycol (PLL-g-PEG). PEG functionalization/passivation is well-established to prevent unspecific adsorption of proteins and adhesion of cells (28). This environment therefore does not provide any adhesive cues and can be used to test how adhesion affects NETosis. First, cells were seeded on glass, PLL-coated, and PLL-g-PEG-coated surfaces and imaged by using reflection interference contrast microscopy (RICM). Dark regions in RICM images indicate close proximity between the cell and the substrate (i.e., adhesion), while bright regions indicate non-adhesive sitting of the cell on the substrate. Cells adhered to both glass and PLL-coated surfaces, with images suggesting a slightly stronger adhesion on glass (Figures 6A,B). However, they did not adhere to PLL-g-PEG coated surfaces as expected ( Figure 6C). Cells were stimulated on these coatings and NETosis was found to be independent from adhesion under PMA stimulation as indicated previously (3) (Figure 6D). On the other hand, LPS-induced NETosis was completely inhibited on PLL-g-PEG, even with very high concentrations of LPS (150 µg/ml) (Figure 6E).

DISCUSSION
Neutrophils are the most abundant leukocytes in humans. Being the first responders to inflammation, neutrophils infiltrate different kinds of tissue and subsequently face different mechanical and chemical environments. Substrate elasticity has been shown to influence important cellular functions such as adhesion, differentiation and migration of many different types of cells including neutrophils (10,11,13,38,46). Therefore, we investigated for the first time how NETosis is affected by substrate elasticity in general and more specifically by substrate elasticity-modulated adhesion. Our results clearly indicate that LPS-mediated NETosis depends on the cells' adhesion and correlates with the adhesion/contact area, which itself correlates with stiffness.
NETosis is initiated and transmitted via diverse pathways, highly depending on the respective stimulus. PI3K is part of the intermediate signaling in PMA-induced and plateletinduced NETosis (40). PI3K also plays an important role in neutrophil mechanosensing and has been shown to be required for neutrophils to sense substrates of higher stiffness (18). In order to investigate the role of mechanosensing on neutrophil spreading as well as on NETosis, we tested a highly selective PI3K inhibitor (BAY 80-6946, copanlisib). Indeed, PI3K inhibition impaired spreading of neutrophils and subsequently LPS-induced NETosis. It also inhibited PMA-induced NETosis to a certain degree, which is understandable as PI3-kinase plays a role in PMA-mediated activation of neutrophils. However, it did not abrogate NET formation completely as it did after LPS-stimulation.
Another factor affecting cell adhesion is the available surface concentration of integrin ligands (25,44). By using nanotechnology approaches it is possible to control the exact distance and overall density of integrin ligands such as RGD and the Mac-1 ligand GPB1α or even link them to advanced near infrared fluorescent nanomaterials (28,47,48). For neutrophils, adhesion maturation and cellular functions such as spreading and migration depend on ligand density (28). Here, we showed that on collagen, LPS-induced NETosis not only depends on substrate elasticity but also on the amount of available surface cues. On a substrate that provided a very high density of surface cues (collagen I) cells did not adhere very well and consequently NETosis did not take place. For PMA, NETosis was once again independent of surface cues, in this case density of surface receptors. Indeed, PMA-induced NETosis does not appear to require adhesion at all (Figure 6). These differences between the stimulants can be explained, at least to some extent, by the different receptor and signaling pathways involved in PMA-and LPS-stimulated NETosis. PMA acts intracellularly and directly activates protein kinase C (PKC). It also triggers subsequent production of reactive oxygen species (ROS), which then interact with MEK, ERK, PI3K, mTOR, MPO, and NE (2,49,50). LPS also activates PKC in neutrophils, primarily through binding to toll-like receptor 4 (TLR4) (51). Our results show that there is an active connection between adhesion and stiffness signaling and LPS-triggered NETosis, putatively by a connection between integrin and TLR4 signaling, which remains to be explored in depth in the context of NETosis. This hypothesis is also corroborated by the complete abrogation of LPS-induced NETosis following phosphatidylinositol 3-kinase (PI3K) blocking. PMA-induced NETosis is also decreased after PI3K inhibition, thus indicating a role of PI3K in PMA-signaling. However, NETosis still took place, albeit at a lower level. It is important to stress that the medium conditions of NETosis assays affect absolute NETosis rates and might shift certain thresholds as previously shown (41).
Our results might bear considerable medical and pharmacological implications as neutrophils and other cells of the immune system are continuously confronted with surfaces of different stiffness/elasticity. One may speculate that reducing neutrophil responses on and in tissues with low substrate elasticity such as the brain or most importantly the blood itself may serve to keep aberrant immune responses in check. On the other hand, alterations of tissue stiffness as seen in arteriosclerotic vessels, tumor tissue, or organ fibrosis might lead to an increase of inflammatory NETosis-related processes (52,53). Indeed, NET-induced inflammation has been implicated in fibrotic organ changes and is likely to trigger tissue remodeling and lead to even more tissue stiffness, causing a pro-inflammatory, self-sustaining vicious circle (54). Our results may also be of importance to the explanation of how NETosis can be triggered by implant materials (55), which tend to have a higher stiffness than "natural" tissues within the body. An improved understanding of the environmental factors that affect NETosis could therefore lead to novel therapeutic approaches for diseases that coincide with alterations of tissue elasticity such as arteriosclerosis, lung, or liver fibrosis or cancer. This understanding will also be of great importance not only for unifying and understanding of NETosis experiments in general, but also for the design of future implant and exogenous materials, which are designed to remain in the body for a certain time, such as catheters.

CONCLUSION
In summary, we show how NETosis rates are affected by different levels of substrate elasticity and adhesion. Neutrophil adhesion increases with substrate stiffness, which leads to higher LPSinduced NETosis rates on stiff substrates. In contrast, PMAinduced NETosis does not require any adhesion at all.

DATA AVAILABILITY STATEMENT
The datasets generated during and/or analyzed during the current study are available from the corresponding author on reasonable request.

ETHICS STATEMENT
The studies involving human participants were reviewed and approved by Ethik-Kommission der Universitätsmedizin Göttingen. The patients/participants provided their written informed consent to participate in this study.

AUTHOR CONTRIBUTIONS
SK and LE designed the study. AG, GG, DM, EN, JB, and LE performed NETosis experiments. LE, AG, MS, and SK analyzed data. GK and FR prepared and characterized gels. LE, AG, GG, and SK wrote the manuscript with inputs from all authors.

FUNDING
This project was supported by the state of Lower Saxony (life@nano) and the German Research Foundation (DFG grant KR 4242/4-1 and ER 723/2-1). FR acknowledges funding through the DFG within the SFB 937 project A13.