Cooler temperatures, on the other hand, strongly enhance persistent currents of Nav1.3. The impaired fast inactivation of the pain-linked Nav1.7/I1461T mutation is further impaired by cooling, mimicking clinical findings.Kriegeskorte et al. show that activation of the sodium channel subtypes Na v) are key players in excitable tissues with the capability to generate and propagate action potentials. Mutations in the genes encoding Navs can lead to severe inherited diseases, and some of these so-called channelopathies show temperature-sensitive phenotypes, for example, paramyotonia congenita, Brugada syndrome, febrile seizure syndromes, and inherited pain syndromes like erythromelalgia (IEM) and paroxysmal extreme pain disorder (PEPD). Nevertheless, most investigations of mutation-induced gating effects have been conducted at room temperature, and thus the role of cooling or warming in channelopathies remains poorly understood. Here, we investigated the temperature sensitivity of four Nav subtypes: Nav1.3, Nav1.5, Nav1.6, and Nav1.7, and two mutations in Nav1.7 causing IEM (Nav1.7/L823R) and PEPD (Nav1.7/I1461T) expressed in cells of the human embryonic kidney cell line using an automated patch clamp system. Our experiments at 15°C, 25°C, and 35°C revealed a shift of the voltage dependence of activation to more hyperpolarized potentials with increasing temperature for all investigated subtypes. Nav1.3 exhibited strongly slowed inactivation kinetics compared with the other subtypes that resulted in enhanced persistent current, especially at 15°C, indicating a possible role in cold-induced hyperexcitability. Impaired fast inactivation of Nav1.7/I1461T was significantly enhanced by a cooling temperature of 15°C. The subtype-specific modulation as well as the intensified mutation-induced gating changes stress the importance to consider temperature as a regulator for channel gating and its impact on cellular excitability as well as disease phenotypes.Voltage-gated sodium channels (Na With their capability to activate and inactivate rapidly in response to changes in membrane voltage, they are initiating the upstroke of action potentials and are key players in excitable tissues. Nine Nav α-subunits, Nav1.1–1.9, and four auxiliary β-subunits have been identified in humans so far  compared with physiological (36°C) temperature mediated the increase in neuronal excitability in in vitro experiments with cortical tissue  are mostly slowly inactivated at the resting membrane potential, Nav1.8 remains excitable. To encode noxious heat, the TTX-resistant Nav1.9 is required in rodents and undergoes a large gain of function with increasing temperature  activation to more hyperpolarized potentials, while the PEPD mutations mainly impair steady-state fast inactivation  Navs. Furthermore, we studied two mutations of Nav1.7, causing IEM and PEPD to test whether the clinical phenotype is reflected in their gating properties. We performed high-throughput patch clamp experiments with an automated patch clamp device, the SyncroPatch 384, at 15°C, 25°C, and 35°C, which revealed a subtype-specific modulation by temperature as well as intensified mutation-induced gating changes. The voltage dependence of activation was shifted to more hyperpolarized potentials with increasing temperature for all investigated subtypes, with an enhanced left shift in the IEM mutation, while V1/2 of steady-state fast inactivation was in general less affected. Our results stress the importance to consider temperature as a regulator for channel gating and its impact on cellular excitability and disease phenotype.In this study, we investigated four different Nav subtypes  were stably expressed in cells from the human embryonic kidney cell line HEK293 and kept under standard culture conditions (37°C and 5% CO2). Cell culture media and supplements are listed in All investigated Na2.All electrophysiological experiments were performed with cells at passage numbers lower than 30 and after reaching a confluency of 70–95%. 1 d prior to the recording, the media was refreshed, and the cells were incubated at 32°C and 5% CO2+ –Ca2+” (Nanion Technologies GmbH) containing (in mM) 10 Hepes, 140 NaCl, 5 glucose, and 4 KCl was added. Subsequently, the cells were stored at 4°C for 10 min. Cells were pipetted up and down 8–10 times using fire-polished glass pipettes to break up cell clumps, External –Mg2+ –Ca2+ was added until a final volume of 30 ml and cells were transferred to the “Cell Hotel” on the SyncroPatch 384 (Nanion Technologies GmbH), where they rested at least 30 min at 10°C and shaking speed of 333 rpm before the recording was started.In order to carry out the experiments, the cells were washed twice with PBS-EDTA , then treated with Trypsin/EDTA (Sigma-Aldrich), and then incubated at 37°C for 6 min to detach, before “External –MgPreprint] using LJPcalc software [https://swharden.com/LJPcalc]). Because of its small size, this shift was not corrected during the experiments. Detailed information regarding series resistance and voltage error are shown in Whole-cell voltage-clamp recordings at 15°C, 25°C, and 35°C were performed using the high-throughput patch clamp robot SyncroPatch 384 with “NPC-384T 1× S-Type” chips  and, for data acquisition, PatchControl384 Version 1.9.7 (Nanion Technologies GmbH). Whole-cell recordings were conducted according to Nanion’s procedure, including initialization, cell-catch, sealing, whole-cell formation, liquid application, and recordings. After achieving the whole-cell configuration, capacitive transients were canceled, series resistance compensation was set to 65%, leak currents were subtracted online using a P/4 procedure, and signals were sampled at 20 kHz. With an increase in temperature of 10°C, the liquid junction potential is shifted by 0.81 mV  on the SyncroPatch was performed. Experiments at different temperatures were performed individually so that each cell was only recorded at one temperature. Cells of the same preparation were measured at several temperatures on the same day to increase comparability.v1.6) contained (in mM) 140 NaCl, 10 Hepes, 5 glucose, 4 KCl, 2 CaCl2, and 1 MgCl2, while the extracellular solution “External NMDG 60”  contained (in mM) 80 NaCl, 60 NMDG, 10 Hepes, 5 glucose, 4 KCl, 2 CaCl2, and 1 MgCl2 .The intracellular solution “Internal CsF110” contained (in mM) 10 EGTA, 10 Hepes, 10 CsCl, 10 NaCl, and 110 CsF. The extracellular solution “External Standard”  from the resting membrane potential of −120 in 5 mV steps with an interval of 5 s at resting membrane potential between each sweep. The conductance–voltage (G-V) relationship, the inactivation time constant τ with a mono-exponential fit on the decayed part of these traces. To quantitatively determine the dependence of the inactivation process on temperature, the Arrhenius equation was described as follows:α is the forward rate constant in the transition between open and inactivated state, z is a proportionality constant, Ea is the activation energy, R is the gas constant, T is the absolute temperature, and V is the applied voltage. β describes the backward rate constant in the transition between open and inactivated state. Since τ ≈ 1/α at depolarized potentials (where Ea(α) ≈ Ea(β)), it was possible to use the Arrhenius plot to estimate Ea by plotting ln(τ) as a function of 1/T because the slope corresponds to Ea/R in this case. The plot of Ea as a function of voltage will be parallel to the voltage axis. At more negative potentials (where Ea(α) ≠ Ea(β)), τ = 1/(α + β), and therefore the simple Arrhenius analysis is generally not applicable. The plot will deviate from a horizontal line.The population average of all included traces was used to determine the inactivation time constant INa,max is the peak sodium current elicited after the most hyperpolarized prepulse, Vm is the preconditioning pulse potential, V1/2 is the half-maximal sodium current, and k is the slope factor.The voltage dependence of steady-state fast inactivation was measured using a series of 500-ms prepulses from −130 to −20 mV in 10-mV steps followed by a 40-ms test pulse to 0 mV that assessed the non-inactivated transient current. The normalized peak inward currents were fitted using a Boltzmann distribution equation:Irecov was normalized to the maximum inward current of the prepulse Ipre and plotted against the duration of the recovery pulse. The following double-exponential equation was used:I is the current amplitude, Iplateau is the amplitude at recovery time t = 1 ms, αfast and αslow are the amplitudes for time constants τfast and τslow, and t is time. %fast is the fraction of the overall recovery that is accounted for by the faster-recovering component following the equation:The recovery from fast inactivation was measured using a 500-ms prepulse to 0 mV followed by a hyperpolarizing recovery pulse to −100 mV of varying duration . Due to the amplifier settings, it was not possible to measure time intervals shorter than 1 ms. After that, another depolarizing test pulse to 0 mV was applied to assess the rate of recovered channels. The maximum inward current of the test pulse Iramp was normalized to the maximum inward current elicited in the G-V-relationship IAct.To evoke ramp currents, slow depolarizing pulses were applied from a holding potential of −120 to +5 mV or to +20 mV. Rates of 1.4, 2.5, and 5 mV/ms were investigated and the maximum inward current I10th was normalized to the peak inward current of the first action potential I1st.5-ms voltage pulses to 0 mV from the holding potential of −120 mV were applied at frequencies of 20, 50, and 100 Hz to assess the use-dependent current decay. The peak inward current of the 10th action potential The recorded data were analyzed using DataControl384 version 2.0.0 (Nanion Technologies GmbH), IgorPro (WaveMetrics), and Prism version 9 (GraphPad Software).t test or a Mann–Whitney test in case of nonparametric testing. Comparisons between more than two experimental groups were done by one-way ANOVA followed by Sidak’s multiple comparison test for parametric testing and Kruskal–Wallis test with Dunn’s multiple comparisons in case of non-parametric testing. Statistical significance was defined as P < 0.05. All values are mentioned as mean ± SEM unless otherwise stated.For statistical testing, comparisons between two experimental groups were done by Student’s v subtype. Activation G-V curves and steady-state fast inactivation I-V curves of Nav1.3, Nav1.5, and Nav1.7 are shown in Detailed information concerning the cell culture media and supplements for each cell line individually are listed in vs is sensitive to temperature. Here, we investigated four WT isoforms: Nav1.3, Nav1.5, Nav1.6, and Nav1.7, each at 15°C, 25°C, and 35°C. The gating of Nav activation. Without exception, we found hyperpolarizing shifts of V1/2 in the range between 5.4 mV (Nav1.6) and 9.5 mV (Nav1.3) with increasing temperature from 15°C to 35°C  and 2.6 times (Nav1.3) faster at 25°C compared with 15°C. At 35°C, the activation became too fast to be accurately dissolved and no reliable analysis was possible. Taken together, the hyperpolarized V1/2 as well as the accelerated opening suggests a strong impact of temperature on the overall excitability of Navs.We analyzed the time to peak and the time from the onset of the voltage step until the maximum inward sodium current is reached. The channel opening was between 1.5 times  was about four times higher at 15°C (21.0% ± 0.6%) compared to 25°C (5.2% ± 0.2%), while at 35°C nearly no persistent current was detectable  that of Nav1.3 was 4.2 times larger (4.80 ms). This as well as the enhanced persistent current may indicate a special sensitivity of this channel’s inactivation kinetics toward cooling.We determined the inactivation time constant τ by a single exponential fit to further quantify the effect of temperature on inactivation kinetics. Rising temperature led to an acceleration of the inactivation kinetics with declining τ-values. Compared to 25°C, τ was ∼2.1–2.5 times larger at 15°C and ∼1.7–1.9 times smaller at 35°C for Na to 0 mV . For Navase of τ . Comparev1.5). It revealed roughly flat plots only for Nav1.6, with an estimated Ea of ≈50 kJ/mol  for potentials less negative than −10 mV was ≈52 kJ/mol for Nav1.5 and ≈51 kJ/mol for Nav1.7. For Nav1.3, it was 1.5 times larger with ≈79 kJ/mol. This indicates that for Nav1.3 more energy is needed to achieve fast inactivation, which may explain the strongly slowed inactivation of this subtype at 15°C.The averaged EBecause it is an important determinant of channel availability in high-frequency firing neurons, we investigated the channel’s use-dependent current decay using the voltage protocol shown in fast values  was close to the V1/2 of the WT at 35°C (V1/2 = −35.4 ± 0.5 mV), suggesting that the voltage dependence of the channel’s activation is closer to the physiological (WT) conditions at colder temperature. Additionally, the L823R mutation renders the conduction curves less steep, with a slope factor k of 10.6 ± 0.3 mV at 15°C, compared with the slope factor of 6.3 ± 0.2 mV for WT at 15°C (t tests), which was not detectable at 15°C.We showed that temperature has profound effects on Naperature . Compare at 15°C . CompariV1/2 of the mutation vs. WT at 35°C or 25°C , but at 15°C steady-state, fast inactivation, V1/2, was shifted by 4.8 mV to more hyperpolarized potentials for the mutation , than for the WT . At 25°C, the recovery curves were almost overlapping with similar τfast values . Furthermore, the voltage at which the peak ramp current occurs was shifted by 9–12 mV to more hyperpolarized potentials for the mutation compared with WT. Comparing 35°C and 15°C, we observed an ∼10 mV shift to more depolarized potentials when lowering the temperature to 15°C. These results may indicate higher sensitivity of the channels to slow subthreshold stimuli at temperatures between 15°C and 35°C.It was described before that other mutations in Nag pulses . Here, wsubtypes . Compariv1.7/I1461T, affecting the highly conserved inactivation motif in the DIII/IV-linker , 8.6 mV (25°C), and 11.8 mV (15°C) to more depolarized potentials compared with WT  decreased from 78.4% at 15°C to 51.3% at 25°C –S5 linker of DIII, Nav1.7/V1298F, and V1299F, as well as the mutation Nav1.7/T1464I showed shifts of 6.3, 4.5, and 6.8 mV, respectively, to more depolarized potentials  or cooling temperature (like for Nav1.3) or both, making it a potentially general mechanistic basis for cold aggravated symptoms and a possible explanation for the cold-induced hyperexcitability as potentially observed in patients suffering from PEPD. Cold-sensitive mutations in Nav1.4 causing PMC also support this hypothesis. PMC induces muscle stiffness in response to lowered ambient temperature. Several PMC mutations were shown to shift V1/2 of fast inactivation to more depolarized potentials, slow the current decay time constant, and accelerate the recovery from fast inactivation, similar to PEPD mutations , is likely to add important information for understanding a potential role in cold thermosensation.Taken together, a strongly slowed inactivation and thus enhanced persistent current can be induced by either a mutation , and the fast recovering proportion of channels (% τfast) decreases . At 35°C, we sometimes observed higher leak currents that weren’t subtracted properly  and largFor drug testing and safety pharmacology, it is crucial to gain deeper insight into gating mechanisms at the physiological temperature of different ion channels. The state dependence of drug binding is likely to be coupled with the occupancy of different conformational states of the channel , and thivs at different temperatures is also essential to understand the properties and to gain a deeper insight into the complex gating mechanisms of Navs at physiological temperature, otherwise important effects might be missed. Of course, in the physiological context, the excitability of a tissue does not only depend on one single channel but also the interplay between many different channels. Furthermore, passive membrane properties have been shown to be temperature sensitive and alter excitability  toward the spinal cord (∼37°C) could be addressed.Detailed electrophysiological investigation of Natability . Howeverpendence . In the vs in their physiological context.Thus, it is of enormous importance to collect experimental data of different ion channels also at physiological temperatures to improve the reliability of action potential simulations and neuronal models, draw (patho-)physiological relevant conclusions, and thus understand NaTable S1provides a ummary of cell culture media and supplements which were used for cultivation of HEK293 rNav1.3, hNav1.5, mNav1.6, hNav1.7/WT, hNav1.7/L823R, and hNav1.7/I1461T.Click here for additional data file.Table S2shows voltage error and series resistance for all investigated Nav-subtypes.Click here for additional data file.


The restacked films demonstrated an exceptionally high electrical conductivity of 2,666 S cm−1, electromagnetic interference shielding effectiveness (EMI SE) of 41.8 dB, and absolute EMI SE (SSE/t) of 27,859 dB cm2 g−1, which is the highest value reported for TMD-based materials. The bond-free vdW interactions between the adjacent 2H-TaS2 nanosheets provide a natural interfacial strain relaxation, achieving excellent flexibility without rupture after 1,000 bends. In addition, the TaS2 nanosheets are further combined with the polymer fibers of bacterial cellulose and aramid nanofibers via electrostatic interactions to significantly enhance the tensile strength and flexibility of the films while maintaining their high electrical conductivity and EMI SE.This work provides promising alternatives for conventional materials used in EMI shielding and nanodevices.Low-dimensional transition metal dichalcogenides (TMDs) have unique electronic structure, vibration modes, and physicochemical properties, making them suitable for fundamental studies and cutting-edge applications such as silicon electronics, optoelectronics, and bioelectronics. However, the brittleness, low toughness, and poor mechanical and electrical stabilities of TMD-based films limit their application. Herein, a TaSThe online version contains supplementary material available at 10.1007/s40820-023-01061-1. High performance electromagnetic interference (EMI) shielding materials with lightweight, ultrathin thickness, and mechanical flexibility have been an important research field because the development of high-speed communication technology and new wearable electronic devices causes considerable EMI harmful effects on the equipment and human health –6. Two-d2, 1T-TaS2), semi-metallic  and metallic  properties [Transition metal dichalcogenides (TMDs) are a class of layered 2D materials composed of transition metals and chalcogen elements that interact through van der Waals (vdW) forces –17. Diffoperties . TMDs exoperties –21, sensoperties , supercaoperties , supercooperties –27, theroperties , 29, eleoperties –33 and soperties , 35.2) is one of the popular TMDs materials, in which 2H-TaS2 exhibits metallic behavior involving a charge density wave phase transition and superconductivity [2 can be exfoliated using electrochemical [2 nanosheets for freestanding film preparation. Bulk TaS2 crystals are rigid and brittle [2 based films. Additionally, 2H-TaS2 flakes have poor electron transport along the stacking direction with an out-of-plane and in-plane electrical conductivities of approximately 0.125 and 33,300 S cm−1, respectively [2 films remains a huge challenge. Despite considerable pioneering efforts devoted to improving both mechanical properties and electrical conductivity of TaS2-based films, the tensile strength is often about 10 MPa [−1 [Tantalum disulfide  of 27,859 dB cm2 g−1, a high tensile strength of 23.3 ± 4.8 MPa, and excellent flexibility withstand 1000 bends without rupture. In addition, TaS2/fiber composite films were also fabricated to further improve the flexibility and strength of the nanosheets while maintaining a high EMI SE. The composite film can be readily folded into a complex shape and unfolded without structural disintegration while effectively shielding against the practical application of 2.4 GHz Bluetooth.Herein, the metallic 2H-TaS2, 99.99%) powder was received from Nanjing NXNANO Tech. Co., Ltd. Dimethyl sulfoxide , anhydrous lithium hydroxide  was received from Shanghai Aladdin Biochemical Tech. Co., Ltd. Hydrochloric acid  was purchased from DONGJIANG Reagent. Poly-p-phenylene terephthamide (PPTA) fibers were obtained from Dupont. Bacterial cellulose dispersion (1 wt%) was obtained from FEYNMAN NANO. Deionized water  powder (0.1 g) was mixed with lithium hydroxide solution  in centrifugal tube for 9 h at room temperature. The final mixture was then washed 3 times using DI water by centrifugation at 12,000 rpm for 15 min until the pH of supernatant is about 7. Subsequently, the resulting swelled sediment was diluted with 80 mL DI water and sonicated for 1 h in an ice bath. Finally, the TaS2 nanosheets aqueous dispersion (~ 1.25 mg mL−1) was obtained without centrifugation.2H-TaS2 freestanding films, the fresh 2H-TaS2 nanosheets aqueous dispersions (~ 1.25 mg mL−1) was subjected to vacuum filtration using polycarbonate micro-porous membrane (Whatman) as substrates, followed by drying at 50 °C for 10 h. The freestanding TaS2 ultrathin films with thickness of about 3.1 μm were finally obtained by peeling off from the substrates.To prepare pristine TaS2 nanosheets dispersion was mixed with BC/HCl dispersion. The resulting suspension was then vacuum filtered after hand shaking 1 min using polyethersulfone membranes as substrates. Subsequently, the pre-preparation TaS2/BC composite film was washed with 10 mL of DI water by continuing vacuum filtration and dried at 50 °C for 10 h to form a final TaS2/BC composite film. Based on the addition of BC, the following five types of TaS2/BC composite films with various mass ratios were prepared: TaS2/BC (10:1), TaS2/BC (10:2), TaS2/BC (10:3), TaS2/BC (10:4), and TaS2/BC (10:5).The bacterial cellulose (BC) dispersion (1 wt%) was added to HCl solution (0.01 M) protonation for 30 min, and ultrasonication for 1 min before use. Then the freshly synthesized TaS2/ANFs composite films with the same weight ratios as TaS2/BC composite films above were prepared: TaS2/ANFs (10:1), TaS2/ANFs (10:2), TaS2/ANFs (10:3), TaS2/ANFs (10:4), and TaS2/ANFs (10:5).The ANFs fibers were fabricated by proton donor-assisted deprotonation . Five ty2 and the films were characterized by X-ray diffraction . The elemental composition was characterized by ICP–OES (Agilent 7700) and NMR (Bruker Avance III 500 MHz WB). High-angle annular dark field scanning transmission electron microscopy  was used to characterize the TaS2 nanosheets. The thickness of the exfoliated nanosheets was measured using atomic force microscopy . The zeta potential of dispersions was measured on Malvern Zetasizer Nano ZS90. Hydrophilicity of the films was analyzed at 298 K using a contact angle analyzer . The mechanical properties of films were investigated by using dynamic mechanical analysis .The structures of bulk 2H-TaS2 freestanding films and TaS2 composite films, a layer of tungsten was deposited on the upper surface of the films, then cut by a focused ion beam (FIB) to provide cross-sections using a FEI Helios NanoLab 600i (using an acceleration voltage of 30 kV and a current of 2.4 nA). Due to the difference in height between the voids and cross-section, and the difference in atomic number between the polymer and TaS2, the contrast is different. The serial backscattering electron section images of TaS2 freestanding films and TaS2 composite films were obtained by FIB/SEM tomography (FIB/SEMT) with a constant separation of 30 nm (using an acceleration voltage of 5 kV and a current of 0.8 nA). Finally, the software (Thermo Scientific Auto Slice&View 4 and Avizo) was used to reconstruct the corresponding three-dimensional (3D) microstructure and automatically calculate all data.To detect the 3D reconstruction microstructure of the TaS2 powder was measured using a powder resistivity system . Electrical conductivity of all TaS2 films were measured using a non-contact resistivity tester . The electrical conductivity of all TaS2 films were calculated by the Eq.  using PNA network analyzer  in X-band frequency range (8.2–12.4 GHz).R), absorption (A), and transmission (T) coefficients were calculated by scattering parameters  as:The reflection (T), microwave reflection (SER), and microwave absorption (SEA) can be calculated from R and T coefficients as:Furthermore, the total EMI SE (SEThe absolute effectiveness (SSE/t) were calculated by the Eq.  7, 47, , 47, 48:2 nanosheets were produced by Li-ion intercalation, which involves immersing 2H-TaS2 crystals in a highly concentrated lithium hydroxide solution at room temperature followed by a mild sonication and exfoliation processes , in which the Tyndall effect is observed, appears yellow  [2 nanosheets (FWHM = 1.50°) was broader than that of the 2H-TaS2 crystals, which indicated the successful exfoliation of the sample and the presence of vdW restacking effects between the TaS2 nanosheets. Raman spectroscopy  and the in-plane vibration (E2g) modes, respectively, of 2H-TaS2 at room temperature, and a broad second-order peak, which is attributed to a two-phonon scattering process, was observed at approximately 196.2 cm−1. The E2g peak of the lyophilized TaS2 nanosheets was slightly blue shifted to 3.8 cm−1 compared to that of the bulk 2H-TaS2, indicating the effective exfoliation of TaS2 nanosheets [2g/A1g ratio of the TaS2 nanosheets is 0.107, which is very close to that of the monolayer [The metallic 2H phase of the lyophilized TaS domain) . The  and 162.0 eV (2p1/2)  was further characterized using an inductively coupled plasma optical emission spectrometer (ICP–OES), and the chemical compositions of the 2H-TaS2 nanosheets is found to be Li0.18TaS2. The presence of Li ions was also confirmed by nuclear magnetic resonance  analysis , which verified the homogeneous distribution of Ta and S elements throughout the material. The morphology of the 2H-TaS2 flakes was characterized using SEM, transmission electron microscopy (TEM), and AFM. Figure 2 nanosheets, which indicates that they are flexible with large lateral dimensions. SEM statistical analysis  shows the values of the lateral dimensions of the nanosheets (0.5–11 μm), mainly distributed at values < 5 μm . Figure 2 nanosheet (thicknesses ≈ 1.75 nm). The figure indicates that few porous defects are observed on the surface of the 2H-TaS2 nanosheets. The AFM statistical analysis of the thickness of the 2H-TaS2 nanosheets  shows that the 2H-TaS2 nanosheets mainly consist of few layers , and Ta2 nanosheets  and EDS images of the 2H-TaS2 nanosheets confirm the presence of the Ta and S elements  was prepared . Figure S7 shows the volume distribution of the 3D reconstructed voids. The FIB/SEMT results indicate that the ultralow porosity of the TaS2 films (6.01%) is significantly lower than that of the MXene films (15.4 ± 0.6%) [The TaSred Fig. a. Figuretructure , 58. The2. Figures 2/bacterial cellulose (BC) and TaS2/aramid nanofibers (ANFs) composite films. The TaS2 freestanding film exhibits a smoother surface than that of the TaS2/BC (10:5) composite film and a rougher surface than that of the TaS2/ANFs (10:5) composite film. This is attributed to the larger roughness of the pure BC film and the very smaller roughness of the pure ANFs film . Therefore, the surface roughness of composite film will be between pure fiber film and TaS2 film. The real content of BC and ANFs in the TaS2 composite films was examined by thermogravimetric analysis , and the results are presented in Table S2. An acid pretreatment of BC and ANFs allows for the protonation of the fiber, which promotes the aggregation of the TaS2 nanosheets around the fiber surface by electrostatic interaction . This effectively prevents the delamination of the composite films  due to the huge density difference between the TaS2 nanosheets and fibers. In addition, the preparation efficiency of the composite films is greatly accelerated by acid treatment (Movie S2). The cross-sectional SEM and FIB/SEMT images  and TaS2/ANFs (10:5) composite films were analyzed by Raman microscope (the size of observation area is 50 μm × 50 μm). As shown in Fig. S14, the shifts at 350–450, 1092, and 1648 cm−1 correspond to TaS2 [2 has the local high content area in TaS2/BC (10:5) or TaS2/ANFs (10:5) composite films  and aramid nanofibers (ANFs) are considered promising matrix materials because of their high stability, good film-forming properties, and high mechanical properties , 59–61.  to TaS2 , C–O of  to TaS2 , and C−N to TaS2 , respect2 freestanding film and the TaS2/fiber composite films. The tensile strength, Young’s modulus, and toughness of the TaS2 freestanding film are 23.3 ± 4.8 MPa, 14.9 ± 6.2 GPa, and 0.033 ± 0.018 MJ m−3, respectively . The tensile strength of the TaS2 freestanding film is much higher than that of the previously reported TaS2HA0.371NMF0.135 foil (9.16 MPa) [2 (0.5 wt%) film (11.27 MPa) [2 nanosheets improved the mechanical properties of the TaS2 freestanding film. The TaS2/BC (10:5) and TaS2/ANFs (10:5) composite films exhibit superior mechanical properties, i.e., their tensile strength is 87.9 ± 8.1 and 134.13 ± 1.4 MPa, respectively, and their toughness is 3.25 ± 0.45 and 4.52 ± 0.07 MJ m−3, respectively, which are the highest values reported for TaS2 composite films  composite film can easily withstand a tensile force of 1 kg. The large sized TaS2/BC (10:5) composite film (diameter ≈ 90 mm) can be rapidly prepared by our facile method , which can be attributed to the better anti-wettability of the composite films and stronger interaction between the TaS2 nanosheets and fibers compared to those in case of the pure TaS2 films . Figure 2 freestanding film, TaS2/BC (10:5) composite films, and TaS2/ANFs (10:5) composite films only decreased by 7.4%, 8.6%, and 3.0%, respectively, after 1,000 bending cycles at a speed of 500 mm min−1 and a bending radius of approximately 2.5 mm, indicating the good mechanical flexibility and electrical stabilities of the TaS2 freestanding and composite films.Figures .27 MPa) , which i2 freestanding film exhibits a well-ordered lamellar structure  values listed in Table S5. The TaS2 freestanding film shows a strong XRD 2θ peak at 9.47° and a weak peak at 18.77°, corresponding to d of 0.933 and 0.472 nm, respectively. The d of the restacked TaS2 freestanding film increased compared with that in the TaS2 crystal, resulting in a significant reduction in the interlayer coupling. The fracture morphology of the TaS2 freestanding film shows a smooth curvature of 2H-TaS2 nanosheets  and reduce the strain-induced cracks and fractures without breaking the broad-area vdW interfaces and conduction pathways.  and TaS2/ANFs (10:5) composite films show strong XRD peaks at 2θ of 14.78° and 14.66°, which correspond to d of 0.599 and 0.604 nm, respectively. These d values are smaller than that of the pure TaS2 freestanding film. The introduction of BC and ANFs nanofibers act as stiffeners because they are tightly embedded between the TaS2 nanosheets interlayer  between the fibers, van der Waals interaction between the TaS2 nanosheets, electrostatic interaction between fibers and bond-free TaS2 nanosheets, and mechanical entanglement. When the stretching procedure starts, the TaS2 nanosheets first slide past each other because of the weak vdW interactions. Meanwhile, the nanofibers crosslinking with TaS2 nanosheets via electrostatic interactions are stretched and further arrest crack propagation for accommodating large deformation before complete fracture of the sheets.Compared with the TaSS2/BC 10: and TaS2S2/BC 10: and TaS22 freestanding film is 2666 S cm−1  and the reported TaS2 based films (Table S7). This result indicates that the broad-area dangling-bond-free plane-to-plane contacts in the TaS2 nanosheets along with their minimum interfacial trapping states and low voids can facilitate the in-plane and intersheet electron transport properties of the thin film [2 nanosheets provides vertical paths for electron transmission and a passage for the adsorbed Li-ions, resulting in a high conductivity of the film [2 composite films  and larger volume distribution . However, the conductivity of the TaS2 composite films are superior to those of most reported TaS2-based films (Table S7). The alternating (between TaS2 and the fibers) multilayer stack structures of the TaS2 composite films ensured their optimal conductivity and mechanical properties.The electrical conductivity of the 3.1-μm-thick TaShin film , 63–65. the film . The con2 freestanding film resulted in an excellent EMI SE of 41.8 dB at the X-band (8.2–12.4 GHz), which is much higher than the commercialization benchmark (20 dB) in electronic equipment of civil telecom. Moreover, the EMI SE of the individual TaS2 films with a thickness of 7.5, 17.5, and 40 μm is 49.1, 54.8, and 72.5 dB, respectively, and hence, EMI SE increases with the increase in the thickness. In addition, the multi-level superimposed TaS2 films show superior EMI SE . Thus, EMI SE increases with the increase in the number of layers and the film thickness. The multi-level superimposed TaS2 films  provides a superior EMI SE of 105.2 dB. Clearly, the superimposed film has better EMI SE performance than the independent film, which may be due to the synergistic effect of multiple internal reflections between the adjacent TaS2 films and multiple-wave interference between the TaS2 nanosheets [2/BC and TaS2/ANFs composite films were measured at different component ratios at the X-band. With the increase in the BC content, the EMI SE does not significantly decrease  composite film, the average EMI SE still reaches 46.8 dB, which can effectively shield against a 2.4 GHz Bluetooth signal  achieved an excellent EMI SE of 46.8 dB. With the increase in the ANFs content, the EMI SE of the TaS2/ANFs (10:5) decreases to 39.2 dB, which can be mainly attributed to the significant decrease in its conductivity due to the ANFs. The main electromagnetic shielding mechanism of these films is reflection . The excellent EMI SE performance of the TaS2 freestanding and composite films are ascribed to the stacked structure and defects of the TaS2 nanosheets and the multi-interfaces created by BC or ANFs, which synergistically contributed to a strong interfacial polarization as well as multiple reflections and increased the dielectric loss of the incident electromagnetic waves Supplementary file2 (MP4 16198 KB)Supplementary file3 (MP4 2462 KB)Supplementary file4 (MP4 15097 KB)Supplementary file5 (MP4 14082 KB)Below is the link to the electronic supplementary material.


HIV-1 integration introduces ectopic transcription factor binding sites into host chromatin. We postulate that the integrated provirus serves as an ectopic enhancer that recruits additional transcription factors to the integration locus, increases chromatin accessibility, changes 3D chromatin interactions, and enhances both retroviral and host gene expression. We used four well-characterized HIV-1-infected cell line clones having unique integration sites and low to high levels of HIV-1 expression. Using single-cell DOGMA-seq, which captured the heterogeneity of HIV-1 expression and host chromatin accessibility, we found that HIV-1 transcription correlated with HIV-1 accessibility and host chromatin accessibility. HIV-1 integration increased local host chromatin accessibility within an ∼5- to 30-kb distance. CRISPRa- and CRISPRi-mediated HIV-1 promoter activation and inhibition confirmed integration site–dependent HIV-1-driven changes of host chromatin accessibility. HIV-1 did not drive chromatin confirmation changes at the genomic level (by Hi-C) or the enhancer connectome (by H3K27ac HiChIP). Using 4C-seq to interrogate HIV-1–chromatin interactions, we found that HIV-1 interacted with host chromatin ∼100–300 kb from the integration site. By identifying chromatin regions having both increased transcription factor activity (by ATAC-seq) and HIV-1–chromatin interaction (by 4C-seq), we identified enrichment of ETS, RUNT, and ZNF-family transcription factor binding that may mediate HIV-1–host chromatin interactions. Our study has found that HIV-1 promoter activity increases host chromatin accessibility, and HIV-1 interacted with host chromatin within the existing chromatin boundaries in an integration site–dependent manner. Chromatin structural proteins, such as CCCTC-binding factor (CTCF) and cohesion, form insulating boundaries and facilitate chromatin interactions within the same topological associating domains (TADs) and chromatin loops. As retroviruses integrate into the human genome, we postulate that the integrated provirus serves as an ectopic enhancer that recruits additional transcription factors to the integration locus, increases chromatin accessibility, alters 3D chromatin interactions, and enhances both retroviral and host gene expression. We wonder whether retrovirus-induced ectopic chromatin interactions can extend beyond chromatin boundaries or are restricted locally.Three-dimensional (3D) spatial chromatin organization brings remote enhancers in contact with promoters and regulates gene expression in In human chromatin, existing retroviral promoter long terminal repeat (LTR) elements provide transcription factor binding sites that regulate gene expression across development, tissues, and cell types . These L+ individuals under suppressive ART, HIV-1 integration sites are enriched in cancer genes  stops new rounds of HIV-1 infection, the HIV-1 reservoir is established early during infection and persists lifelong . In HIV-er genes . HIV-1 ier genes , causinger genes  and evener genes . On the er genes  and in per genes , presumacis-regulatory elements that recruit transcription factors. HIV-1 proviral accessibility has been well characterized, with three regions of DNase I hypersensitivity on the USF1 and NF-kB binding sites in the LTR and within the coding sequence, which are remodeled upon transcriptional activation  HIV-1 proviruses can persist despite effective ART and can contribute to viral rebound during treatment interruptions . Here, w+ individuals under suppressive ART  , enhanceTo further analyze these data and make statistical comparisons, we used a pairwise normalization and comparison approach in HiCcompare . In cont4 = 256 bp). Therefore, distal chromatin regions in close physical contact were ligated to form chimeric DNA regions. After reverse cross-linking, the DNA fragments were further digested by another restriction enzyme, MluCl (cutting AATT), into smaller fragments and circularized by intramolecular ligation in dilute conditions. These circularized chromatin contacts were amplified by HIV-1-specific primers targeting the HIV-1 core promoter and enhancer (HXB2 coordinates 249–728), where most transcription factors bind. Of note, the two restriction enzymes NlaIII and MluCI were carefully chosen because they preserve a large portion of the HIV-1 promoter region of interest in NL4-3-d6-dsGFP reporter virus in the cell line clones. To quantify chromatin interactions, we used the percentage of possible fragments detected (as opposed to direct read numbers) because the restriction enzyme digestion method, as opposed to shearing-based methods, lacks a unique molecular identifier (UMI)  tiled in increments of five fragments (256 bp per fragment × five fragments = 1280 bp on average) across the whole genome. The capture of multiple independent fragments within a given region allows us to approximate the interaction likelihood without relying on read count. We then plotted that number across the genome.We postulate that the impact of the HIV-1 proviral genome on chromatin structure is at the HIV-1 integration site locally. Circularized chromatin conformation capture sequencing (4C-seq)  enables er (UMI) . Specifier (UMI) . A thresDespite differences in the magnitude of signal between replicates, we observed a correlation between each of our replicates (Pearson's correlation 0.44–0.74) A–E. UninAlthough only semiquantitative in nature, 4C-seq signal is expected to decay with distance from the region of interest. This property can be used to fit a monotonic model, which the observed data can then be tested against . This apWe postulate that transcription factors mediate chromatin binding, looping, and 3D interactions between HIV-1 promoter and local host chromatin. We compared ATAC-seq peaks and 4C-seq at the integration site to identify transcription factors mediating local HIV-1–chromatin interactions. We identified genome regions having both increased chromatin accessibility (from ATAC-seq) and increased chromatin interactions (from 4C-seq), focusing on ATAC-seq peaks (size <1 kb) around the HIV-1 integration site that overlapped windows of HIV-1–host chromatin interaction as determined by 4C-seq (size ∼10 kb) A–E. EachCTCFL, a paralog of CTCF)  , ZFX (knof CTCF) , PLAGL1 of CTCF) , and KLFof CTCF) . ZNF652 ividuals . Recent ividuals . Our stucis-regulatory elements (such as transcription factor binding sites) on the HIV-1 genome, not mediated by viral proteins. Unlike HTLV-1, which has a CTCF binding site in the proviral genome, which changes chromatin looping, HIV-1 does not contain CTCF binding sites in the proviral genome  and transcription factor binding motifs (by ATAC-seq), we identified transcription factors that may mediate HIV-1–host chromatin looping, such as ZF transcription factors and ETS and RUNX family transcription factors. These factors are of great significance as many of them are known to bind HIV-1 and enhance or suppress HIV-1 transcription. Furthermore, recent studies have found an enrichment for HIV-1 integrations into ZNF genes after long-term suppressive therapy . AlthougDespite consistent interaction profiles observed by targeted amplification (4C-seq), we were not able to detect statistically robust changes in chromatin conformation (by Hi-C) or enhancer–promoter interactions (by H3K27ac HiChIP) around the integration site. It is possible that HIV-1 promoter activity and interactions are subjugated to existing host architecture or that the tendency of HIV-1 to form tight transcription and splicing linked loops  prevents+ individuals, these integration sites serve as biological replicates but not as a precise reflection of HIV-1 integration into enriched integration sites, such as BACH2, MRTFB, and STAT3 , 5F9 (harboring an HIV-1-d6-GFP provirus in INPPL1), 1D7 (harboring an HIV-1-d6-GFP provirus in SPECC1), and 8B10 —were selected for this study. HIV-1 integration sites were examined by inverse PCR  with pMDLg/pRRE (Addgene 12251), pRSV-Rev (Addgene 12253), and pCMV-VSV-G (Addgene 8454) at an equal molecular ratio. Media was changed 6 h after transfection, and pseudotyped lentivirus was harvested 48–72 h after transfection. Viruses were concentrated using Lenti-X concentrator (Takara Bio 631231) or by ultracentrifugation using a 20% sucrose cushion.For CRISPRi cell lines, Jurkat clones were transduced with pseudotyped lentiviruses encoding Lenti-dCas9-KRAB-blast (Addgene 89567) by spinoculation. Two days posttransduction, 8 µg/mL blasticidin was added to the culture medium to select transduced cells. After 1 wk of blasticidin selection, cells were transduced with pseudotyped viruses encoding gRNAs targeting the HIV-1 LTR (CTACAAGGGACTTTCCGCTG) or nontargeting (GTGCACCCGGCTAGGACCGG), cloned into the parental plasmid pCRISPRia-v2 (Addgene 84832). Two days after transduction, 1.5 µg/mL puromycin was added to the culture medium to select transduced cells.For CRISPRa cell lines, Jurkat clones were transduced with pseudotyped lentiviruses encoding Lenti-MPH v2 (Addgene 89308) by spinoculation. Two days posttransduction, 600 µg/mL of hygromycin was added to the culture medium to select transduced cells. After 1 wk of hygromycin selection, cells were transduced with pseudotyped viruses encoding gRNAs targeting the HIV-1 LTR (CTACAAGGGACTTTCCGCTG) or nontargeting (GTGCACCCGGCTAGGACCGG), cloned into the parental plasmid lentiSAMv2 (Addgene 75112). Two days post-transduction, 1.5 µg/mL puromycin was added to the culture medium to select transduced cells. Cells were cultured for at least 7 d under selection before genomic assays.At least 2 million cells were harvested, pelleted, and washed once using wash media. Cells were then fixed by resuspending them in 4% paraformaldehyde. Cells were then analyzed on a FACSAria cell sorter. Gating and quantification were performed using FlowJo 2, 0.01% digitonin, 2 U/µL RNase inhibitor) for 5 min before the addition of DIG wash buffer . Cells were washed once with DIG wash buffer and then loaded onto the 10x Chromium controller (10x Genomics) for the single-cell multiome ATAC + Gene Expression assay according to the manufacturer's protocol (10x Genomics document CG000338).Cells were prepared for DOGMA-seq as previously described . Dead ceLibraries were prepared according to the same protocol with the following modifications. During the barcoding reaction (step 4.1), 1 µL of 0.2 µM HTO oligo (GTGACTGGAGTTCAGACGTGTGC*T*C) was added. During the following purification (step 4.3.k), beads were eluted in 100 µL. A barcoding PCR was added to construct the index protein tags using 35 µL of the eluted sample, 2.5 µL of 10 µM SI-PCR, 2.5 µL of 10 µM D7xx, and 50 µL 2× KAPA mix (Roche KK2601). The reaction was run on a thermocycler starting with 3 min at 95°C; 12 cycles of 20 sec at 95°C, 30 sec at 60°C, and 20 sec at 72°C; with a final extension of 5 min at 72°C. Libraries for ATAC-seq, transcriptome, and surface protein tags were sequenced on a NovaSeq 6000 in manufacturer-suggested read configurations.2, 0.1% Tween 20, 0.1% NP-40, 0.01% digitonin) before the addition of 1 mL of ATAC-wash buffer . Cells were pelleted at 4°C and then transposed with ATAC-transposition mix  for 30 min at 37°C. DNA was isolated using a MinElute PCR cleanup kit (Qiagen 28004) and eluted in 10 µL twice. The resulting DNA was amplified using NEBNext ultra II Q5 master mix (New England Biolabs M0544X) and Nextera indexing primers for 5 min at 72°C and 1 min at 98°C; followed by five cycles of 15 sec at 98°C, 30 sec at 63°C, and 1 min at 72°C; with a final extension of 5 min at 72°C. Additional cycles were added based on qPCR measurements of library amplification for a total of eight to 20 cycles per library. Libraries were then sequenced on a NovaSeq 6000 in 150-bp paired-end mode.ATAC-seq was performed as previously described . BrieflyFive million to 10 million cells were cross-linked in 2% formaldehyde for 10 min at room temperature with shaking. The reaction was quenched by adding glycine to a final concentration of 0.125 M and shaking for 10 min at room temperature. Cross-linked cells were washed with ice-cold PBS twice before being incubated in Hi-C lysis buffer  on ice for 30 min with protease inhibitors (Sigma-Aldrich 11873580001). The resulting nuclei were washed once with Hi-C lysis buffer without protease inhibitors and then resuspended in 50 µL 0.5% SDS for 7 min at 62°C. To quench SDS, 145 µL of water and 25 µL 10% Triton X-100 were added, and samples were incubated with 500 rpm shaking for 15 min at 37°C. After SDS quenching, samples were incubated in restriction enzyme 1 mix  with 500 rpm shaking for 4 h at 37°C. Two hundred units of NlaIII was added, and nuclei were incubated with 500 rpm shaking overnight at 37°C. The following morning, nuclei were heated for 20 min to 62°C to inactivate the restriction enzyme. One thousand microliters of T4 ligase mix  and 2000 units T4 DNA ligase (New England Biolabs M0202S) were added, and nuclei were incubated with 500 rpm shaking for 6 h at 25°C. Nuclei were pelleted and resuspended in 300 µL reverse cross-linking mix . Nuclei were incubated for 1 h with 500 rpm shaking at 55°C, and then the temperature was raised to 68°C for overnight incubation.The following day, DNA was isolated using a phenol–chloroform extraction and ethanol precipitation. All DNA was then incubated with 500 rpm shaking in 200 µL restriction enzyme mix 2  for 4 h at 37°C. An additional 200 U of restriction enzyme was added, and the incubation continued overnight. DNA was ethanol precipitated and resuspended in 14 mL T4 sparse ligation mix  and incubated overnight at 16°C. The following day, DNA was ethanol precipitated. DNA was initially amplified using NEBNEXT ultra II Q5 master mix in eight 25-µL reactions with ∼1 µg DNA with a program of 2 min at 98°C; followed by 30 cycles of 10 sec at 98°C, 30 sec at 68°C, and 1 min at 72°C, reducing the annealing temperature by 1°C per cycle until 63°C was reached; with a final extension time of 5 min at 72°C. DNA was pooled across reactions and isolated using a PCR cleanup kit. Fifty nanograms of resulting PCR product was then barcoded using Nextera primers for 5 min at 72°C and 1 min at 98°C; followed by five cycles of 15 sec at 98°C, 30 sec at 63°C, and 1 min at 72°C; with a final extension for 5 min at 72°C. The resulting libraries were sequenced using a NovaSeq 6000 or HiSeq 4000 in 100-bp pair-end or 150-bp pair-end modes.g for 5 min at 4°C and washed once with Hi-C lysis buffer. The supernatant was removed, and the pellet was resuspended in 100 µL 0.5% SDS and incubated for 10 min at 62°C without shaking or rotation. Then, 285 µL of water and 50 µL 10% Triton X-100 were added to the sample. The sample was incubated at 37°C with 500 rpm shaking for 15 min. Fifty microliters of 10× CutSmart buffer (New England Biolabs B6004S) and 375 U of MboI (New England Biolabs R0147M) were added before the sample was shaken at 500 rpm for 2 h at 37°C. Samples were incubated for 20 min at 62°C to inactivate the restriction enzyme without rotation. Ends were filled in and marked with biotin with rotation for 1 h at 37°C after the addition of biotin master mix ; 4.5 µL each of 10 mM dCTP, dGTP, dTTP; and 50 U of DNA Polymerase I, large (Klenow) fragment (New England Biolabs M0210). Nine hundred forty-eight microliters of ligation master mix was then added . The reaction was then incubated with rotation for 4 h at room temperature before pelleting at 2500g for 5 min at room temperature. The supernatant was removed.Hi-C and HiChIP libraries were produced as previously described , 2017 wig at 4°C. Clarified supernatant was diluted with an equal volume of 2× ChIP dilution buffer  in a new tube.Nuclei were pelleted and brought up to 880 µL in nuclear lysis buffer  and transferred to a Covaris millitube (Covaris 520135). Samples were sheared on a Covaris S220 for 2 min with a fill level of 10, duty cycle of five, PIP of 140, and cycles/burst of 200. Sheared samples were clarified by centrifugation for 15 min at 16,000For H3K27ac HiChIP only, the supernatant was then incubated with 60 µL of Protein A beads (New England Biolabs S1425S) for 1 h with rotation at 4°C. Samples were placed on a magnet, transferred to a new tube, and incubated with 7.5 µg of H3K27ac antibody (Abcam ab4729) with rotation overnight at 4°C. Sixty microliters of Protein A beads was added the following morning and incubated with rotation for 1 h at 4°C. Three washes each of low-salt wash buffer , high-salt wash buffer , and LiCl wash buffer  were performed at room temperature. Beads were eluted in 50 mM sodium bicarbonate (pH 8.0), 1% SDS for 10 min with rotation followed by 3 min at 37°C with shaking. Beads were placed on a magnet, and the supernatant was transferred to a new tube. Two total elusions were performed. The 300 µL of eluted sample was incubated with 15 µL of Proteinase K (New England Biolabs P8107S) with 500 rpm shaking for 45 min at 55°C to promote protein digestion. The temperature was increased to 67°C for 1.5 h to reverse cross-links, and samples were purified using a DNA clean and concentrator kit (Zymo Research D4003) and eluted in 10 µL of water.2, 30% dimethylformamide). The resulting beads were resuspended with 4 µL of TDE1 enzyme (Illumina 20034197) in TD buffer for a total of 50 µL. Samples were incubated for 10 min at 55°C and incubated with 50 mM EDTA for 30 min at 50°C, and then the supernatant was removed. Beads were washed twice with 50 mM EDTA and then Tween wash buffer for 2 min at 55°C. Beads were washed once with 10 mM Tris (pH 7.5) before PCR.For all samples, 5 µL of Streptavidin C-1 beads (Thermo Fisher Scientific 65001) were washed with Tween wash buffer  and then resuspended in binding buffer . Beads were incubated for 15 min with shaking and then washed twice with Tween wash buffer. Beads were incubated for 2 min at 55°C with 500 rpm shaking and then washed with 100 µL of 1× TD buffer , a peak caller purpose-built to handle ATAC-seq data including accounting for duplicate and multimapping reads with the options “-j -y -r -e chrM -v.” Peaks were merged across all ATAC-seq samples. Overlapping peaks were merged to generate a consensus set of peaks using BEDTools merge (version 2.30.0)  (Reads were trimmed to remove adapter sequences using cutadapt (version 3.5) . Trimmed 2.30.0) . BAM fil 2.30.0)  to compan 3.5.1) . In the n 3.5.1) . Plottinn 3.5.1)  and Inten 3.5.1) . Differen 3.5.1) .Reads were trimmed in paired-end mode to remove primer sequences from the 5′ end of reads associated with the target window  and filtered to select only reads with the correct primer sequence using the cutadapt option “‐‐discard-untrimmed.” Trimmed reads were then mapped to hg38 using Bowtie 2. The genome was segmented into restriction enzyme fragment windows corresponding to the first restriction enzyme (NlaIII) using the HiC-Pro utility script “digest_genome.py.” The resulting BED file was used to quantify fragment coverage across the genome and in each sample using BEDTools multicov. In parallel, BAM files were used to generate bigWig files using deepTools. Results were visualized using R (version 4.2)  and IGV.Binarization and fragment quantification were performed as follows in R. The quantification file was loaded into R, and the quantification for each sample was binarized by setting the data point to “one” if at least 10 reads mapped to that fragment or to zero otherwise. A threshold of 10 reads was chosen to overcome observed index hopping in the NovaSeq platform based on the number of reads present in the uninfected Jurkat sample. Index hopping is a known sequencing artifact impacting patterned flow cell platforms . Index hopping  occurs bP-value was below 0.05. HOMER was used to output motif images and to annotate significantly interacting peaks with top enriched motifs for visualization in IGV.Overlapping regions of 4C-seq signal and ATAC-seq peaks were extracted using R, selecting only ATAC-seq peaks which were on (1) the same chromosome as the integration of interest and (2) overlapped with 4C-seq windows in which at least one replicate detected at least 10 of the possible 40 fragments. Peaks were filtered to remove regions overlapping encode blacklist regions , a set oReads from transcriptome and ATAC-seq libraries were aligned to a concatenated hg38 and NL43-d6-GFP HIV genome using Cell Ranger ARC . The resulting matrices were analyzed using Seurat (version 4.1.1)  and SignAfter clustering, peaks were recalled using MACS2 (version 2.2.7.1)  through Reads were mapped to a concatenated hg38 and NL43 genome, filtered, quantified into contact matrices, and quality controlled using HiC-Pro (version 3.1.0)  with thehttps://www.ncbi.nlm.nih.gov/geo/) under accession number GSE230314.All raw and processed sequencing data generated in this study have been submitted to the NCBI Gene Expression Omnibus (GEO; Supplement 1Supplement 2Supplement 3Supplement 4


Lewis et al. reveal that herpes simplex virus infection alters the heterochromatin landscape of the host to support the egress of progeny out of the nucleus. This work elegantly demonstrates that host chromatin architecture, beyond transcriptional regulation, is a critical structural component in virus-host interactions. Herpes simplex virus (HSV-1) progeny form in the nucleus and exit to successfully infect other cells. Newly formed capsids navigate complex chromatin architecture to reach the inner nuclear membrane (INM) and egress. Here, we demonstrate by transmission electron microscopy (TEM) that HSV-1 capsids traverse heterochromatin associated with trimethylation on histone H3 lysine 27 (H3K27me3) and the histone variant macroH2A1. Through chromatin profiling during infection, we revealed global redistribution of these marks whereby massive host genomic regions bound by macroH2A1 and H3K27me3 correlate with decreased host transcription in active compartments. We found that the loss of these markers resulted in significantly lower viral titers but did not impact viral genome or protein accumulation. Strikingly, we discovered that loss of macroH2A1 or H3K27me3 resulted in nuclear trapping of capsids. Finally, by live-capsid tracking, we quantified this decreased capsid movement. Thus, our work demonstrates that HSV-1 takes advantage of the dynamic nature of host heterochromatin formation during infection for efficient nuclear egress. Nuclear-replicating viruses must contend with host chromatin to establish a successful infection. Like most DNA viruses, herpes simplex virus (HSV-1) takes advantage of host chromatin factors both by incorporating histones onto its genome to promote gene expression  and by rHeterochromatin density and subnuclear localization are affected by the presence and ratio of specific histone modifications. In uninfected cells, histone modifications such as trimethylation of histone H3 lysine 27 (H3K27me3) and the histone variant macroH2A1, among others, delineate heterochromatin regions that are largely localized to the nuclear periphery. H3K27me3 is deposited by the EZH2 enzyme , a membeIn this study, we visualized HSV-1 nuclear egress by transmission electron microscopy (TEM) and found that capsids reach the inner nuclear membrane in regions of less densely stained chromatin. Therefore, we hypothesized that HSV-1 exploits host heterochromatin dynamics to successfully egress from the nuclear compartment. We examined chromatin structure by TEM in the absence of heterochromatin markers macroH2A1 and H3K27me3 and discovered that peripheral heterochromatin is largely dependent on these marks. We used chromatin profiling of macroH2A1 and H3K27me3 during HSV-1 infection to define the specific host genomic regions bound by these markers and found that they demarcate broad regions of heterochromatin that form in transcriptionally active compartments. Importantly, we found that the loss of macroH2A1 results in significantly lower viral titers but does not impair viral transcription, protein production, or replication in both lab-adapted and clinical isolates of HSV-1. Furthermore, by inhibiting EZH2 deposition of H3K27me3, we found that reduction of H3K27me3 also leads to a significant decrease in viral titers but did not affect viral protein or genome accumulation. Finally, we determined by TEM that loss of macroH2A1 or H3K27me3 results in significantly more viral capsids trapped in the nuclear compartment, pinpointing the importance of heterochromatin dynamics in viral nuclear egress. Our study is the first to demonstrate that HSV-1 infection takes advantage of heterochromatin changes to successfully egress from the nuclear compartment.To investigate the journey of HSV-1 capsids to the inner nuclear membrane, we used TEM to image nuclei and examined heterochromatin formation in primary and diploid human foreskin fibroblast (HFF) cells. In uninfected cells, we observed dark staining, characteristic of dense heterochromatin, in the nuclear periphery , arrows.macroH2A1 gene in hTERT-immortalized HFF cells (HFF-T) cells, which resulted in the loss of total macroH2A1 (We used CRISPR-Cas9 to knock-out (KO) expression of the acroH2A1 , termed acroH2A1  using a acroH2A1 . We treaacroH2A1 . We founacroH2A1 . We quanacroH2A1 . It is iWe hypothesized that macroH2A1 and H3K27me3 are deposited at specific genomic loci on the host genome during infection to promote the formation of heterochromatin. To test this hypothesis, we used CUT&Tag  to profiTo determine whether H3K27me3 deposition was dependent on macroH2A1, we also examined H3K27me3 enrichment in domains in the absence of macroH2A1. We plotted the H3K27me3 changes at the WT clusters for macroH2A1 KO cells and found that the trends of H3K27me3 were similar between WT and macroH2A1 KO cells . This reIn contrast, the macroH2A1 CUT&Tag on the viral genome showed similar enrichments in both WT and macroH2A1 KO cells. Further, the H3K27me3 signal on the viral genomes also mirrored the IgG control. These results indicated that there was a significant background signal from the viral genome that could not be accounted for . FurtherTo investigate the transcriptional output of newly formed macroH2A1 and H3K27me3-bound regions, we performed RNA-seq on WT and macroH2A1 KO cells over the course of HSV-1 infection and calculated fold changes in RNA levels at each time point over the mock control. We identified genes that are contained within domains in each cluster defined in To determine whether these changes in RNA reflected changes in transcription, we analyzed published 4sU-RNA-labeling data during HSV-1 infection . The timFinally, we asked if the gain or loss of macroH2A1 happened across previously defined genome compartments . We deteTo determine the impact of macroH2A1 in HSV-1 infection, we infected WT and macroH2A1 KO HFF-T cells with HSV-1. First, we used RNA-seq to compare viral transcripts in WT and macroH2A1 KO cells. We found no significant change in viral transcripts at 4, 8, or 12 hpi between the two cell types . Next, wWe next examined HSV-1 replication under the same conditions and measured viral genome accumulation by droplet digital PCR (ddPCR). We observed no significant difference in viral genome accumulation between WT and macroH2A1 KO cells , 4 µg VSVg, and 2.5 µg pREV  using Attractene transfection reagent . Lentivirus was harvested at 24-, 48-, and 72-h posttransfection, filtered through 0.2-µm filters, aliquoted, flash-frozen with liquid nitrogen, and stored at −80°C. 2 ml of thawed filtered lentivirus was used to transduce ∼2.25 × 106 HFF-T or 3 × 106 RPE cells in 10-cm plates with 8 µg/ml polybrene . Cells were allowed to reach confluence before selection with 1 µg/ml puromycin for 3 d. Cells were then sorted by GFP positivity or counted by hemocytometer, plated at one cell per well into 96-well plates, and selected through serial expansion of colonies. Selected cells were screened for macroH2A1 knockout by Western blot.MacroH2A1 guide RNA  in DMSO for 3 d prior to infection. Cells were then infected at an MOI of 3, and after 1 h of incubation with the virus, fresh media with 10 µM Taz was added to previously treated cells. Control samples were treated with equivalent volumes of DMSO. Samples were harvested as above.Western blotting was performed as previously described . Brieflyg for 2 min. Pellets were flash-frozen in liquid nitrogen and stored at −80°C until processed. HSV-1 DNA within cells was isolated from frozen pellets using QIAamp DNAMini Kit .Cells were harvested at the indicated times after infection by trypsinization, washed with 1× PBS, and centrifuged at 5,000 × g, and filtered through 40-µm sterile syringe filters. DNA on the exterior of filtered capsids was digested for 1 h at 25°C with 20.3 units DNase  supplemented with 10 mM MgCl2. DNase was inactivated at 75°C for 10 min followed by vortexing. Capsids were digested with 3 mg/ml proteinase K  in 100 mM KCl, 25 mM EDTA, 10 mM Tris-HCl pH 7.4, and 1% Igepal for 1 h at 50°C. HSV-1 genomes from digested capsids were isolated using QIAamp DNAMini Kit.Supernatants were harvested at the indicated times after infection, centrifuged at >3,500 × A duplexed droplet digital PCR was performed to measure the levels of cellular or supernatant HSV-1 genomes on the QX100 droplet digital PCR system (Bio-Rad Laboratories) using a primer/probe set specific to HSV-1 gB. Cell numbers were determined using a primer/probe set specific to human Beta-globin, a reference gene that exists at two copies per cell. See g for 3 min, and counted using a hemocytometer. Nuclei from 600,000 cells were isolated by hypotonic lysis in 1 ml buffer NE1  for 10 min on ice followed by centrifugation at 1,300 × g for 4 min. Nuclei were resuspended in Wash buffer  and counted using a hemocytometer. BioMag Plus Concanavalin A-coated beads  were equilibrated with Binding buffer . Beads (5 µl) were mixed with aliquots of 75,000 nuclei and incubated at 25°C for 10 min followed by magnetic separation of beads. Beads were resuspended in 50 µl primary antibody , anti-H3K27me3 , or anti-mouse IgG  in Wash buffer supplemented with 2 mM EDTA and 0.1% bovine serum albumin (BSA) and incubated on a nutator at 4°C overnight. The beads were decanted on a magnet stand and then resuspended in 50 µl secondary antibody (Guinea pig anti-rabbit IgG [Antibodies-Online: ABIN101961] 1:100) in Wash buffer supplemented with 2 mM EDTA and 0.1% BSA and incubated on a nutator at room temperature for 1 h. The beads were decanted on a magnet stand and washed with 200 µl Wash buffer, resuspended in 50 µl pA-Tn5 (1:200) in 300-Wash buffer (Wash buffer containing 300 mM NaCl), and incubated on a nutator at room temperature for 1 h. The beads were washed twice with 200 µl 300-Wash buffer, resuspended in 50 µl Tagmentation buffer (300-Wash buffer supplemented with 10 mM MgCl2), and incubated at 37°C for 1 h. Beads were then washed with 50 µl TAPS wash buffer , resuspended in 5 µl TAPS wash buffer supplemented with 0.1% SDS, and incubated at 58°C for 1 h. SDS was neutralized on ice with 15 µl 0.67% Triton-X100. 2 µl of 10 mM indexed P5 and P7 primer solutions and 25 µl NEBnext High-Fidelity 2× Master Mix  were added. Gap-filling and 15 cycles of PCR were performed using an MJ PTC-200 Thermocycler. Library clean-up was performed by incubating beads with 65 µl SPRI bead slurry for 5–10 min, then magnetization and two washes with 200 µl 80% ethanol. Libraries were eluted with 22 µl Tris-HCl pH 8.0 and 2 µl was used for Agilent 4200 Tapestation analysis. The barcoded libraries were mixed to achieve equimolar representation as desired aiming for a final concentration as recommended by the manufacturer for sequencing on an Illumina HiSeq 2500 2-lane Turbo flow cell.Two biological replicates per time point were obtained from independent infections. The protocol was adapted from the established CUT&Tag methods . Cells wEscherichia coli . We performed alignments using bowtie2.CUT&Tag raw sequencing data were aligned to a custom genome made by concatenating human (hg38), HSV (JN555585.1), and N:Coverage at 100-bp windows of the human hg38 reference genome was calculated as the number of reads of a given length  that mapped at that window normalized by the factor E. coli DNA as a spike-in to normalize all datasets. We partitioned all chromosomes into domains of macroH2A1: domains had an enrichment that was two times the genome-wide median and at least fourfold higher than the IgG control. The normalized coverage at each base-pair from each replicate was averaged when combining multiple replicates. The normalized read density in 100-bp bins was then smoothed with a running average over the bins spanning ±1,000 bp around each bin. We then calculated the genome-wide distribution of normalized read density and medians that were plotted in Here, 10,000 is an arbitrarily chosen number. We used 2 ratio of macroH2A1 enrichment over IgG for the non-overlapping regions for the mock, 4, 8, and 12 hpi. The 4, 8, and 12 hpi enrichments were then divided by the enrichment of the mock dataset to obtain a change in macroH2A1 over time course at the non-overlapping regions. k-means clustering (k = 6) using https://doi.org/10.5281/zenodo.6783949.To measure changes in macroH2A1 across time points where the domain boundaries are not the same, we first concatenated domain definitions from all macroH2A1 datasets and then defined a set of non-overlapping intervals using the “disjoin” method of GenomicRanges R package . We theng for 2 min. Pellets were lysed with TRIzol  and total RNA was harvested according to the manufacturer’s instructions. RNAs were then treated with DNase  on RNeasy columns  per the manufacturer’s instructions. RNA was precipitated with three volumes of ice-cold 96% ethanol, one volume of 3 M sodium acetate pH 5.5, and 1 µl glycogen  overnight at −80°C. Precipitated RNAs were pelleted at 15,000 × g and 4°C for 30 min, washed with ice-cold 75% ethanol, and spun as above for 10 min. RNA was resuspended in nuclease-free water.Three biological replicates per time point were obtained from independent infections. Cells were harvested at the indicated times after infection by trypsinization, washed with PBS, and centrifuged at 5,000 × RNA was quantified by Nanodrop and integrity was analyzed with the 4200 Tapestation Bioanalyzer system (Agilent). 500 ng of total RNA with an RNA Integrity Number (RIN) >9.5 were used to prepare sequencing libraries with the TruSeq Stranded mRNA Library Prep Kit . Library concentrations were measured with Qubit dsDNA HS Assay Kit  and then analyzed with Agilent High Sensitivity D5000 ScreenTape System and pooled. Libraries were sequenced with 100-bp paired-end reads on an Illumina NovaSeq 6000 SP sequencer at the Fred Hutch Genomics Core Facility.https://www.ncbi.nlm.nih.gov/nuccore/JN555585.1/). FastQC 0.11.9 (https://www.bioinformatics.babraham.ac.uk/projects/fastqc/) and RseQC 4.0.0  and GCA_000859985.2_ViralProj15217 (https://www.ncbi.nlm.nih.gov/data-hub/genome/GCF_000859985.2/). For HSV1, annotated genes were collapsed into non-overlapping transcribed regions, e.g., X indicating a transcribed region unique to gene X, X:Y indicating an overlapping transcribed region for genes X and Y, and so on.Gene annotations were based on GENCODE V31 . The filtered expression matrix was normalized by the TMM method  and subject to significance testing using quasi-likelihood pipeline implemented in edgeR. Genes were deemed differentially expressed if fold changes were >2 in either direction, and Benjamini-Hochberg adjusted P values were <0.01.Bioconductor package edgeR 3.26.8  was usedGSE59717) and converted to transcripts per million (TPM). The TPM values across two 4sU-RNA replicates for each condition were averaged, and box plots were generated similarly to total RNA.The intervals representing the start and end of each gene were intersected with clustered macroH2A1 domains to obtain cluster assignments for genes. The intersection was performed using “intersect” function of bedtools . Genes thttps://4dn-open-data-public.s3.amazonaws.com/fourfront-webprod/wfoutput/b543cbf4-ce54-4d2d-8960-281528ff18a6/4DNFI342UZP1.bw.For calculating eigenvector distributions, the compartment wig file was downloaded from the following link: Regions from compartment file intersecting with each domain cluster were extracted, and the values were used for generating box plots shown in Cells were plated on poly-L coated glass coverslips the day prior to infection. Cells were then infected with HSV-1 at an MOI of 3 and collected at mock, 4, 8, and 12 hpi. For harvest, cells were fixed with cold 4% PFA in 1× PBS for 15 min. Cells were permeabilized with 0.5% Triton-X in 1× PBS for 10 min, then blocked in 10% goat serum in 1× PBS for 1 h, incubated with primary antibody (diluted as noted) in 3% BSA in 1× PBS for 1 h. Slides were incubated with secondary antibodies at a dilution of 1:300 in 3% BSA in 1× PBS for 1 h. Coverslips were fixed to microscope slides with Invitrogen ProLong Gold Antifade Mountant. Images were taken on Leica Stellaris Confocal with 63× oil objective at room temperature.Cells were fixed in 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.3) at 4°C. Fixed cells were rinsed briefly in 1% sucrose in 50 mM cacodylate (pH 7.2), then postfixed on ice for 30 min in a solution of 1% osmium  and 0.8% potassium ferricyanide in 50 mM cacodylate (pH 7.2). Cell pellets were washed twice briefly at 25°C in 1% sucrose in 50 mM cacodylate (pH 7.2) and then washed in three changes of 50 mM cacodylate (pH 7.2) for 5 min each. Cell pellets were treated with 0.2% tannic acid  in 50 mM cacodylate (pH 7.2) for 15 min at 25°C and then rinsed several times in water. Cells were dehydrated through a graded ethanol series and embedded in Epon 12 resin . 70-nm thin sections were cut using an Ultracut UC7 ultramicrotome (Leica Mikrosysteme) and collected on 200 mesh formvar/carbon copper grids . Sections were stained with 2% aqueous uranyl acetate and Reynolds lead citrate. Cell pellet sections were imaged using a Talos L120C microscope operated at 120 kV with a Ceta-16 M  camera (Thermo Fisher Scientific).All data were collected at spot size 5 with a 100-µm C2 aperture and 70-µm objective aperture. For quantification, nuclei were targeted at 1,250× and manually circled. Autofocus was set to −2.0 µm. Nuclei were then imaged at 11,000× as a montage and stitched together automatically using SerialEM (Nexperion Inc). Stitched maps were exported as uncompressed 16-bit tif files for further analysis. For qualitative analysis, images were manually focused to −2.0 µm defocus and then 20 1-s frames were collected and drift-corrected using Velox software (Thermo Fisher Scientific). Final summed images were exported as 16-bit tif files and cropped using ImageJ.Image analysis pipelines for counting capsids and for measuring chromatin density at the nuclear envelope were deployed in MATLAB R2020b. Scripts are available on the Fred Hutch GitHub repository. Capsid pipeline follows three steps: (1) nuclear boundaries identification, (2) capsid detection, (3) capsid classification. Nuclear boundaries were identified with user input by outlining a freehand contour of the nucleus of interest from which a binary mask is extracted, and its area and perimeter were calculated. Capsid detection was performed on the median-filtered complement of the original image using a Circular Hough Transform-based algorithm, with phase coding for radii estimation , and seaThe width of heterochromatin abutting the nuclear envelope was quantified by measuring the length of the binarized chromatin from 1D intensity profiles along the normal of the nuclear perimeter, sampled at every 10-perimeter pixels. Dense heterochromatin was binarized using global Otsu’s thresholding method applied to the background-corrected complement of the contrast-adjusted original image. Noise from the binarized image was further reduced by applying a 2D order statistic filter using the minimum value of a varying domain interactively defined by the user, with a default value of 8-by-8 pixels. The resulting heterochromatin density distribution was normalized to the total length of the nucleus’ perimeter.2 using an environmental control system consisting of an objective heater and a humidified incubation chamber.Cells were infected with HSV-1 VP26mCherry  based on the R-package CelltrackR (Y=YM−(YM−Y0)×exp(−k×x) with GraphPad Prism v.9.5.0. Y0 represents the starting population (in [µm2]), K determines the rate constant (inverse of X [sec]), and YM represents the plateau (same units as Y). We then calculated the mean corral diameter using the following equation: (modified from corral is the corral diameter, dparticle is the estimated HSV-1 capsid size of 125 nm (Cells were continuously imaged for 40 s with a 561-nm wavelength laser in a single plane and a frame rate of 9.5 frame/sec. Single particle tracking was performed in ImageJ Fiji  with thelltrackR . To calcf 125 nm , and YM GSE209820.The data generated are available in the published article and its online supplemental material. The CUT&Tag and RNA-seq data can be found here: Series Review HistoryClick here for additional data file.SourceData F1is the source file for Fig. 1.Click here for additional data file.SourceData F3is the source file for Fig. 3.Click here for additional data file.SourceData F4is the source file for Fig. 4.Click here for additional data file.SourceData FS1is the source file for Fig. S1.Click here for additional data file.SourceData FS4is the source file for Fig. S4.Click here for additional data file.SourceData FS5is the source file for Fig. S5.Click here for additional data file.