T cell priming is enhanced by maturation-dependent stiffening of the dendritic cell cortex

T cell activation by dendritic cells (DCs) involves forces exerted by the T cell actin cytoskeleton, which are opposed by the cortical cytoskeleton of the interacting APC. During an immune response, DCs undergo a maturation process that optimizes their ability to efficiently prime naïve T cells. Using atomic force microscopy, we find that during maturation, DC cortical stiffness increases via process that involves actin polymerization. Using stimulatory hydrogels and DCs expressing mutant cytoskeletal proteins, we find that increasing stiffness lowers the agonist dose needed for T cell activation. CD4+ T cells exhibit much more profound stiffness-dependency than CD8+ T cells. Finally, stiffness responses are most robust when T cells are stimulated with pMHC rather than anti-CD3ε, consistent with a mechanosensing mechanism involving receptor deformation. Taken together, our data reveal that maturation-associated cytoskeletal changes alter the biophysical properties of DCs, providing mechanical cues that costimulate T cell activation.


INTRODUCTION 14
The initiation of an adaptive immune response requires priming of naïve T cells by professional 15 antigen presenting cells (APCs). This process involves multiple receptor-ligand interactions, 16 which occur in concert at a specialized cell-cell contact site called the immunological synapse 17 (Dustin, 2014). Through these interactions, APCs transmit a highly orchestrated series of signals 18 that induce T cell activation and direct differentiation of T cell populations (Friedl, den Boer, & 19 Gunzer, 2005). While the biochemical aspects of this process have been the subject of many 20 studies, the contribution of mechanical cues is only now being uncovered. Thus, it appears that force application is mechanically coupled to the T cell's ability to 33 sense stiffness (mechanosensing). In other cell types, substrate stiffness has been shown to 34 affect a variety of cell functions including differentiation, migration, growth and survival (Byfield,35 Reen In this work, we aimed to better understand the relationship between DC cortical stiffness and T 63 cell activation. We show that during maturation, DCs undergo a 2-3 fold increase in cortical 64 stiffness, and that T cell activation is sensitive to stiffness over the same range. Moreover, we 65 find that stiffness sensitivity is a general trait exhibited by most T cell populations. Since 66 mechanosensing occurs lowers the threshold signal required for T cell activation, we conclude 67 that stiffness serves as a novel biophysical costimulatory mechanism that functions in concert 68 with canonical signaling cues to facilitate T cell priming. 69 We next sought to identify the molecular mechanisms controlling DC cortical stiffness. Several 119 actin regulatory mechanisms are known to change during DC maturation. In particular, mature 120 DCs upregulate the actin bundling protein fascin (Yamashiro, 2012) Surprisingly, the stiffness of BMDCs from fascin -/mice was indistinguishable from that of WT 130 BMDCs both before and after LPS-induced maturation ( Figure 2A). Next, we tested the 131 contribution of myosin contractility, which is known to control stiffness and membrane tension in 132 other cell types (Salbreux, Charras, & Paluch, 2012). As shown in Figure 2B, treating mature 133 BMDCs with the myosin II inhibitor blebbistatin reduced stiffness by a small, albeit statistically 134 significant amount. Similar results were obtained with the Rho-kinase (ROCK) inhibitor Y27623, 135 which indirectly inhibits myosin function. 136 We next considered the possibility that cortical stiffness is modulated by actin polymerization. 137 Broadly speaking, actin polymerization is induced by two sets of proteins: formins generate linear 138 actin filaments, while activators of the Arp2/3 complex produce branched actin structures. 139 Treatment of DCs with the pan-formin inhibitor SMIFH2 significantly reduced the cortical stiffness 140 of mature DCs ( Figure 2B). A similar reduction was observed after inhibition of Arp2/3-mediated 141 branched actin polymerization by CK666. DCs express multiple activators of Arp2/3 complex, of 142 which two have been implicated in maturation-associated changes in actin architecture: 143 Hematopoietic Lineage Cell-Specific Protein 1 (HS1), the hematopoietic homologue of cortactin 144 (Huang et al., 2011), and WASp, the protein defective in Wiskott-Aldrich syndrome (Bouma, Burns, 145 & Thrasher, 2007;Bouma et al., 2011;Calle, Chou, Thrasher, & Jones, 2004). To individually assess 146 the role of these two proteins, we used BMDCs cultured from HS1 and WASp knockout mice. As 147 shown in Figure 2A, loss of HS1 had no impact on cortical stiffness of either immature or mature 148 BMDCs. In contrast, mature WASp knockout BMDCs were significantly less stiff than WT controls. 149 This difference mirrors that seen after inhibition of Arp2/3 complex by CK666, suggesting that 150 WASp is the primary activator of Arp2/3 complex-dependent changes in cortical stiffness. The 151 defect in WASp knockout DCs was observed only after maturation; immature WASp knockout DCs 152 did not differ in stiffness from WT controls. This is consistent with our finding that the stiffness of 153 immature DCs is unaffected by actin depolymerizing agents. Taken together, these results show 154 that activation of formin and WASp-dependent actin polymerization pathways, and to a lesser 155 extent increased myosin contractility, all contribute to the increased cortical stiffness of mature 156

Stiffness lowers the threshold for activation of CD4 + T cells, but not CD8 + T cells. 158
Our findings raise the possibility that changes in DC cortical stiffness, like other maturation-159 induced changes, enhance the ability of these cells to prime a T cell response. Previous studies 160 have shown that T cells are sensitive to the stiffness of stimulatory surfaces (Judokusumo et al., 161 2012;O'Connor et al., 2012), and that that the TCR serves as the mechanosensory (Judokusumo 162 et al., 2012). However, the results are conflicting, and these earlier studies were not performed 163 within the physiological stiffness range that we have defined for DCs. Thus, we tested T cell 164 responses on hydrogels with a stiffness range spanning that of immature and mature DCs (2 -25 165 kPa). We verified the compliance of the hydrogel surfaces by measuring the elastic modulus of 166 the surfaces directly by AFM. Hydrogel stiffnesses were found to be similar to those reported by 167 the manufacturer (Figure 3 -Figure Supplement 1). Surfaces were coated with varying doses of 168 peptide-loaded major histocompatibility complex (pMHC) molecules, together with a constant 169 dose of anti-CD28. Surfaces were coated with H-2K b class I MHC loaded with the N4 (SIINFEKL) 170 peptide (pMHC-I), or I-A b class II MHC loaded with the OVA329-337 (AAHAEINEA) peptide (pMHC-171 II), to stimulate OT-I CD8 + or OT-II CD4 + T cells, respectively. Plastic surfaces, which are commonly 172 used for stimulation with surface-bound ligands, were included as a familiar reference point. To 173 test the effects of substrate stiffness on early T cell activation, we measured surface expression 174 of the activation markers CD25 and CD69, as well as the production of IL-2, all at 24 hours post 175 stimulation. As shown in Figure 3A-C, CD4 + T cells showed a profound stiffness-dependent 176 response at 24 hours across all measures. This response was most clearly seen for upregulation 177 of CD25 and CD69, where increasing substrate stiffness enhanced the CD4+ T cell response in a 178 graded manner. As expected, for any given substrate stiffness T cell activation increased with 179 increasing peptide dose. However, comparison between stimulatory surfaces revealed that 180 increasing substrate stiffness lowered the pMHC-II dose required to obtain the same level of 181 activation. Over the stiffness range associated with DC maturation (2-8kPa), the dose of TCR signal 182 needed to induce surface marker upregulation was shifted by 1-2 logs. Analysis of IL-2 production 183 revealed a similar effect, although the stiffness sensitivity was more bi-modal. IL-2 production 184 increased almost 3-fold when CD4+ T cells were stimulated on surfaces of 8kPa, as opposed to 185 2kPa, for the same antigen dose. 186 Strikingly, the robust stiffness response we observed in CD4 + T cells was not recapitulated for CD8 + 187 T cells, especially when T cell activation was assessed based on surface marker upregulation 188 ( Figure 3D,E). Analysis of IL-2 production did reveal stiffness-enhanced activation of naïve CD8 + T 189 cells, but this was largely restricted to substrates outside the stiffness range defined for DCs 190 (12kPa, 25 kPa and plastic). 191 To determine if the stiffness dependency of CD4 + T cells seen at early times after TCR engagement 192 is maintained at later times, we measured T cell proliferation based on CFSE dilution at 72 hours 193 post stimulation. Similar to what was observed for early activation markers, increasing substrate 194 stiffness produced graded increases in CD4 + T cell proliferation, and the threshold dose required 195 to induce robust proliferation shifted as a function of substrate stiffness ( Figure 4A,B). This effect 196 was particularly evident at low doses of pMHC-II (0.1-1ug/ml). Interestingly, although soft 197 hydrogels (2-4 kPa) elicited only very low levels of CD4 + T cell proliferation, these substrates did 198 induce upregulation of CD25 in a high percentage of CD4 + T cells, even in the undivided 199 populations ( Figure 4C,D). This indicates that an activating signal was received, but was 200 insufficient to drive proliferation. 201 Since the threshold stimuli (stiffness and dose of pMHC-II) required to induce significant IL-2 202 production and proliferation were very similar ( Figures 3C and 4B), we reasoned that the 203 threshold for proliferation might be driven by IL-2 availability. To test this, OT-II CD4 + T cells were 204 stimulated on hydrogels with or without addition of 25 U/ml exogenous IL-2. Interestingly, 205 addition of IL-2 did not rescue the proliferation of T cells stimulated on soft surfaces ( Figure 4E). 206 Thus, we conclude that in addition to influencing the signaling threshold for IL-2 production, 207 substrate stiffness also affects other IL-2 independent events needed for efficient T cell 208 proliferation. 209 Although early activation events in CD8 + T cells did not exhibit stiffness sensitivity, we reasoned 210 that proliferative responses might behave differently. As shown in ( Figure 5A,B), CD8 + T cells 211 showed a mild stiffness-dependent proliferation response. The concentration of stimulatory 212 ligand needed to induce at least one round of division was similar across the entire stiffness range 213 Over successive rounds, increased stiffness did enhance the extent of proliferation, but the 214 differences were relatively small ( Figure 5B). Interestingly, analysis of CFSE dilution as a function 215 of CD25 expression reveals evidence that CD8 + T cells exhibit a binary stiffness response ( Figure 5   Taken together, our findings point to a mechanism in which stiffer substrates have a sensitizing 221 effect on CD4 + T cells, similar to that of classical co-stimulatory molecules such as CD28 (Harding,222 McArthur, Gross, Raulet, & Allison, 1992). When considered in this way, the relative lack of 223 stiffness responses in CD8 + T cells fits with the fact that CD8 + T cells are much less dependent on 224 costimulatory signals (McAdam, Schweitzer, & Sharpe, 1998). 225

Degranulation of cytotoxic T cells shows mild stiffness sensitivity. 226
Whereas naïve T cells are activated by DCs, effector T cells interact with many cell types. In 227 particular, cytotoxic CD8 + T cells (CTLs) must respond to a variety of possible target cells, which 228 may differ widely with respect to stiffness. We therefore reasoned that CTL effectors might be 229 stiffness independent. To test this, ex vivo OTI CD8 + T cells were activated on plastic surfaces and 230 grown in the presence of exogenous IL-2 to produce mature CTLs. To induce and detect release 231 of cytolytic granules, CTLs were re-stimulated on hydrogels coated with a range of pMHC-I 232  Figure 6D-F); on soft surfaces, pMHC yielded the strongest response, and anti-TCR was more 261 effective than anti-CD3ε. Taken together, these results indicate that T cells sense substrate 262 stiffness best through direct engagement of TCRαβ. 263 The increased stiffness of mature DCs enhances their ability to prime T cells 264 Our hydrogel assays show that T cell activation is enhanced by changes in stiffness over the range 265 observed for DC maturation, consistent with the idea that modulation of cortical stiffness is a 266 biophysical mechanism by which DCs control T cell activation. To test this directly, we sought 267 conditions under which we could manipulate the stiffness of mature DCs. We took advantage of 268 our finding that mature WASp knockout BMDCs are approximately 20% softer than WT controls 269 ( Figure 2A; data are presented as absolute values in Figure 7A). WT and WASp -/-BMDCs were 270 pulsed with increasing concentrations of OVA323-339 peptide and co-cultured with OT-II CD4 + T cells. 271 T cell proliferation was measured by CFSE dilution. As shown in Figure 7B approximately 20% relative to WT cells ( Figure 7A), but these BMDCs failed to prime T cells more 280 efficiently ( Figure 7C). Expression of CA-WASp only enhances BMDC stiffness to approximately 281 5kPa, and based on our hydrogel studies, this increase is unlikely to be sufficient to enhance T cell 282 activation. It seems likely that conditions that stiffen DCs to 10kPa or more would further enhance 283 T cell responses, but we were unable to test this directly, and it is not clear whether this happens 284 in vivo. Nonetheless, the studies using WASp-/-DCs show that changes in DC stiffness within the 285 range observed during maturation has a significant impact on their ability to efficiently prime a T 286 cell response. 287

DISCUSSION 288
Recent work from several labs clearly shows that T cell activation involves mechanical cues. We 289 have previously shown that the DC cytoskeleton constrains the mobility of stimulatory ligands 290 on the DC surface, enhancing T cell activation by opposing the forces exerted by the T cell on the 291 corresponding receptors (Comrie et al., 2015). In the current study, we elucidate a second 292 mechanism whereby the DC cytoskeleton enhances T cell activation. We show that actin 293 remodeling during DC maturation increases the cortical stiffness of DCs by 2-3 fold, and that T 294 cell activation is enhanced by increases in stiffness over the same range. Importantly, increased 295 stiffness lowers the threshold dose of TCR ligand needed for T cell activation, as expected if 296 substrate stiffness serves as a costimulatory signal. In keeping with this concept, CD4 + T cells 297 showed more profound stiffness-sensitivity than CD8 + T cells, especially at early times in the 298 activation process. Together, these results indicate that stiffening of the DC cortex during 299 maturation provides biophysical cues that work together with canonical costimulatory cues to 300 enhance T cell priming. 301 Modulation of actin architecture has long been appreciated as an essential feature of DC 302 maturation. Changes in the DC actin cytoskeleton facilitate the transition from highly endocytic 303 tissue-resident cells to migratory cells specialized for antigen presentation ( The observed increase in stiffness depends on changes in actin architecture; whereas 316 depolymerization of actin filaments has no effect on the stiffness of immature DCs, the increase 317 associated with maturation depends on intact filaments, and is sensitive to inhibitors of actin 318 polymerizing molecules. While it remains to be determined exactly which actin regulatory 319 pathways control cortical stiffness in mature DCs, our data show that both Arp2/3 complex and 320 formins are involved. Moreover, we find that DCs lacking the Arp2/3 activator WASp are 321 abnormally soft. In keeping with these findings, DC maturation is known to induce changes in 322 the activation state and localization of Rho family GTPases, especially Cdc42, a molecule that 323 can activate both WASp and formins (Garrett et al., 2000;Vargas et al., 2016;West et al., 2000). 324 Since the overall levels of active Cdc42 are diminished during DC activation, it seems likely that 325 the observed increase in cortical stiffness results from redistribution of the active pool. 326 We show that DC cortical stiffness is a cell-intrinsic property that is unaffected by substrate 327 stiffness. In this respect, DCs are different from other cell types that adapt their stiffness to 328 showing that DCs rapidly change their method of locomotion in order to maintain consistent 331 migration speed and shape while crossing over different surfaces (Renkawitz et al., 2009). This 332 behavior has been proposed to allow DCs to pass through tissues with widely different 333 mechanical properties. In the same way, we propose that the ability of DCs to regulate cortical 334 stiffness as a function of maturation state in spite of environmental cues reflects the importance 335 of this property for priming an appropriate T cell response. 336 A central finding of this paper is that changes in DC stiffness serves as a costimulatory signal for 337 T cell priming. By using a matrix of different hydrogels spanning the biologically relevant range 338 defined for immature and mature DCs (2 -8 kPa), coated with increasing pMHC concentrations, 339 we found that stimulatory substrates with lower stiffness required higher concentrations of 340 pMHC to achieve T cell activation. Similarly, when compared to WT DCs, softer WASp knockout 341 DCs required higher concentrations of OVA peptide to induce the same level of proliferation. 342 Our results indicate that increases in cortical stiffness, together with diminished ligand mobility 343 Importantly, it appears that the 378 TCR's ability to sense stiffness is closely related to its ability to transduce force-dependent 379 signals during T cell-APC interaction. Indeed, there is evidence that signaling downstream of TCR 380 engagement is increased on stiffer substrates (Judokusumo et al., 2012) and that the location of 381 early tyrosine phosphorylation events corresponds to sites of maximum traction force (Bashour 382 et al., 2014). We propose that stiffer substrates allow T cells to exert more force through TCR 383 interactions, and consequently induce more effective signaling. This accounts for the co-384 stimulatory property of substrate stiffness on T cell activation. 385 The mechanism by which force application on the TCR is translated into biochemical signals 386 remains controversial. Nevertheless, there is evidence to suggest that force applied on the TCR 387 complex induces conformational changes within TCRαβ that exposes ITAM sites on the CD3 and 388 TCRζ chains for phosphorylation and downstream signaling 57,58 . Importantly The IS is often described as a platform for information exchange between the T cell and APC. 413 Together with our recent work on ligand mobility, the findings presented here indicate that the 414 mechanical properties of the APC side of the IS influence T cell priming, likely because they 415 augment force-dependent conformational changes in TCRs, integrins, and potentially other 416 molecules. Going forward, it will be important to determine how these properties are 417 modulated during DC maturation, and whether there are also local changes induced by signaling 418 events taking place at the IS. In addition, it will be important to tease apart the molecular events 419 through which T cells sense and respond to these mechanical cues. 420

MATERIALS AND METHODS 421
Key Resource

Inhibitors and antibodies 424
Cytochalasin-D and Latrunculin-B were from EMD Millipore, (S)-nitro-Blebbistatin was from 425 Cayman Chemical, CK666 was from Calbiochem, and Y27632 and SMIFH2 were from Sigma- Roche Inc (Lahm & Stein, 1985)), to give a final IL-2 concentration of 100 units/ml. Cells were 489 cultured at 37°C and 10% CO2, and passaged as needed to be kept at 0.8x10 6 cells/ml for 7 more 490 days. CTLs were used at days 8 or 9 after activation. assays, ruling out the possibility that differences in T cell activation are due to differential ligand 529 binding. Importantly, initial experiments included a 1 kPa hydrogel surface that yielded no 530 response across all assays. Therefore, data from this condition is not shown and was not included 531 in repeated experiments. For experiments where exogenous IL-2 was added, media was 532 supplemented with IL-2 to a final concentration of 25 U/ml. To measure IL-2 secretion, 533 supernatants were harvested 22-24 hours post stimulation, and IL-2 concentration was measured 534 using a mouse IL-2 ELISA kit (Invitrogen). For early activation marker expression assays, cells were 535 plated immediately after isolation, and harvested 22-24 hours post stimulation for flow cytometry 536 analysis. For CFSE dilution assays, purified cells were washed once with PBS and stained for 3 min 537 with 2.5 µM CFSE (ThermoFisher). After quenching the excess CFSE by addition of 1 ml FBS for 30 538 seconds, cells were washed and plated. Cells were harvested 68-72 hours post stimulation for 539 flow cytometry analysis. For ligand comparison assays, surfaces were first coated with 10 µg/ml 540 of NutrAvidin (ThermoFisher) and 2 µg/ml anti-CD28 (PV1) overnight at 4°C. Surfaces were then 541 washed twice with 200 µL of PBS and coated with varying concentrations of biotinylated ligands 542 (anti-TCRVβ5.1/5.2, anti-CD3ε, or pMHC-II monomers) for 2 hours at 37°C. 543

Cytotoxic T cell degranulation assays 544
Assays were conducted on day 8 or 9 of culture. 2x10 5 CTLs were plated onto surfaces coated with 545 various concentrations of pMHC-I in the presence of 2 µg/ml PE-conjugated anti-CD107a. After 3 546 hours of re-stimulation, CD107a labeling was quantified by flow cytometry analysis. Cells were 547 gated based on size, live cells, and expression of CD8 + . CD107a mean fluorescence intensity (MFI) 548 was extracted using FlowJo. 549

T cell priming assays 550
Priming assays were carried out in round bottom 96 well plates. 5x10 4 LPS-matured BMDCs were 551 plated in each well and pulsed with OVA323-339 peptide at various concentrations (0.1 -1 µg/ml). 552 1.5x10 5 CFSE stained, OT-II CD4 + T cells were added to each well and incubated for 68-72 hours. 553 Cells were then harvested and analyzed using flow cytometry. 554

Atomic force microscopy (AFM) 555
All experiments were carried out at room temperature using a Bruker Bioscope Catalyst AFM 556 mounted on a Nikon TE200 inverted microscope. Micro-indentation measurements were made 557 with a spherical tip from Novascan. The tip was comprised of a 1 µm silicon dioxide particle 558 mounted on a silicon nitride cantilever with a nominal spring constant of 0.06 N/m; each 559 cantilever was calibrated using the thermal fluctuation method. The AFM was operated in fluid 560 contact mode, with 2 Hz acquisition. Total vertical cantilever displacement was set to 5 µm, 561 producing a maximal approach/retraction speed of ~20 µm/sec. Maximal deflection (Trigger 562 threshold) was adjusted for each cantilever to apply a maximal force of 6 nN on the measured cell 563 (e.g. for a 0.06 N/m cantilever, the trigger threshold was set to 100 nm). The actual indentation 564 depth was ~1.5 µm depending on the measured cell stiffness (Figure 1 -Supplement Figure 2). 565 Analysis of force-distance curves was carried out using the Nanoscope Analysis software (Bruker). 566 The Young's modulus was extracted using the Hertzian model for spherical tips with a contact 567 point-based fitting on the extend curve data. For each individual cell, two separate measurements 568 were conducted at different locations near, but not directly over the nucleus. The reported cell 569 stiffness value represents the average between these independent measurements. Note that 570 when measurements of cortical stiffness were made over the nucleus, no significant differences 571 in Young's modulus values were found (not shown). To measure BMDC stiffness, 1x10 5 cells 572 (untreated or LPS matured) were seeded onto Poly L-lysine coated coverslips and allowed to 573 spread for 4 hours at 37°C, 5% CO2. Prior to data acquisition, cells were incubated for 10 min with 574 the Fc blocking antibody 2.4G2, washed and stained for CD86 for 20 min, then washed and 575 mounted on the AFM. All antibody incubations and data acquisition steps were performed in L-576 15 media (Gibco) supplemented with 2mg/ml glucose. For treated cell measurements, drugs 577 [Latrunculin-B (10 µM), Cytochalasin-D (10 µM), s-nitro-Blebbistatin (50 µM), Y27632 (25 µM), 578 CK666 (100 µM), or SMIFH2 (10 µM)] were pre-incubated with the cells at 37°C, 5% CO2 for 30 min 579 prior to Fc blocking and maintained in the cultures throughout staining and data acquisition. 580

Statistical Methods 581
All datasets were subjected to outlier analysis prior to execution of statistical testing. Outliers 582 were defined as data points with values outside the range of mean +/-2.5xStDev, and were 583 deleted from the dataset. Testing for a statistically significant difference between experimental 584 groups was done using an unpaired one-way ANOVA test with a post-hoc Tukey correction for 585 multiple comparisons. 586 Throughout the paper, data shown represents biological, and not technical, replicates.