s2fieldsofstudy: Biology Chemistry

text:
A conformational switch in response to Chi converts RecBCD from phage destruction to DNA repair

The RecBCD complex plays key roles in phage DNA degradation, CRISPR array acquisition (adaptation) and host DNA repair. The switch between these roles is regulated by a DNA sequence called Chi. We report cryo-EM structures of the Escherichia coli RecBCD complex bound to several different DNA forks containing a Chi sequence, including one in which Chi is recognised and others in which it is not. The Chi-recognised structure shows conformational changes in regions of the protein that contact Chi and reveals a tortuous path taken by the DNA. Sequence specificity arises from interactions with both the RecC subunit and the sequence itself. These structures provide molecular details for how Chi is recognised and insights into the changes that occur in response to Chi binding that switch RecBCD from bacteriophage destruction and CRISPR spacer acquisition, to constructive host DNA repair.

Introduction
The RecBCD complex, and its close relatives AddAB and AdnAB, plays a critical role in bacterial DNA double-strand break repair (1,2). These complexes load onto DNA ends resulting from a DNA break and are highly processive helicase/nucleases that digest the duplex until they encounter a special sequence called Chi (Crossover hotspot instigator) (3) at which they load RecA protein to initiate repair by homologous recombination (1,2). In E.coli, approximately 75% of Chi sequences are present in the leading strand with an orientation bias in the direction of DNA replication (4). This arrangement has been proposed to favour activity towards the replication origin (4).
However, the complexes are voracious DNA degrading machines and a major defence system against bacteriophages (as their genomes typically lack Chi sequences), digesting the viral DNA starting from breaks created by restriction enzyme cleavage (5,6). Indeed, the default mode of action of RecBCD and AddAB is DNA destruction; that is only modified to a benign DNA repair mode upon interaction with Chi. Furthermore, phage DNA fragments that are produced by both RecBCD and AddAB digestion are incorporated into CRISPR arrays to recognise and respond rapidly to subsequent phage infection, thus providing a mechanism to avoid incorporation of "self" DNA into these arrays via Chi recognition (7,8). Although structural, genetic and biochemical studies of RecBCD have contributed considerably to our understanding of the role of RecBCD in the repair of DNA doublestrand breaks (1), how Chi regulates the switch between bacteriophage digestion mode and DNA repair has remained enigmatic for almost half a century.
Here, using gel mobility shift assays, we probed specificity of RecBCD for unwound DNA forks containing a Chi sequence in the 3'-tail to determine the requirements of the complex for Chi recognition. In this way, we identified a DNA fork with the Chi sequence spaced appropriately for recognition and response. Using this substrate, we were able to obtain high-resolution structural information of the complex by cryo-EM. The structure revealed a surprisingly compact and twisted conformation of the Chi sequence, which is bound and recognised quite differently compared to an equivalent structure of AddAB bound to its Chi sequence (9). This different mode of binding explains a conundrum, in that the final cleavage of the 3'-tail after Chi recognition by each enzyme occurs at a different distance from the Chi sequence. Furthermore, the structure reveals an on-enzyme equilibrium between Chi recognised and unrecognised states that explains a mechanism to prevent phage escape from these destructive enzymes.

DNA substrate optimisation and cryo-EM analyses
The recognition of Chi by E. coli RecBCD requires the sequence to enter the enzyme complex as a DNA duplex, which is then unwound to present the Chi sequence (GCTGGTGG) in a single-stranded DNA context in the 3'-terminated strand (10). Previous studies on both RecBCD and the related AddAB system have shown that the RecBCD complex is more highly regulated than AddAB and that this control emanates from the RecD subunit (11)(12)(13)(14)(15). Indeed, the nuclease activity of the RecB subunit1 (14) is controlled by the RecD subunit (11,13) by a mechanism that involves binding of the 5'-tail to RecD to induce a conformational change in RecB, releasing an α-helix that blocks the nuclease active site (15,16). Interactions of RecD with the 5'-tail also alter the conformation of domains of the RecC subunit, the subunit proposed to interact with Chi (15)(16)(17)(18)(19)(20)(21)(22), indicating that a 5'-tail of around 15 bases or more would be required to activate the enzyme. an encounter with Chi on a physiological substrate (Figure 1a). A Chi site was positioned along the 3'-tail at increasing distance in bases from the fork junction to mimic steps in unwinding a fork. At a spacing of eight bases from the junction, an altered gel mobility shift was observed suggesting a specific response to the Chi sequence at that position ( Figure 1a). We thus designed a DNA fork substrate with 25 base pair duplex, a 15 base 5'-tail and a 20 base 3'-tail comprising eight T bases, followed by the eight base Chi sequence (GCTGGTGG), and terminated with four T residues (Figure 1a).
Cryo-EM analysis of RecBCD in complex with this DNA fork substrate resulted in three main structures -a 3.7Å structure of the Chi-recognised complex, a 3.9Å structure with a DNA fork bound but in which Chi has not been recognised, and an intermediate structure at 4 The structure in which Chi is not recognised is indistinguishable from our previous structure with a long 5'-tail (16). Density corresponding to the bound DNA fork is visible for the duplex region, the 5'-tail, and for the 3'-tail across the RecB subunit, although it is disordered beyond the sixth base so the Chi sequence is not visible (Figure 2a). Consequently, this structure provides a high resolution internal control to delineate which conformational changes are specific to the Chi response. As further confirmation, the structure of a complex with an equivalent substrate that lacks a Chi sequence (Chi-minus) and does not induce an altered gel mobility shift (Figure 1a), was determined at 3.8Å resolution and shows no density in the Chi-binding site ( Table 1, Supplementary Figures 2 & 3). Similarly, structures of complexes with forks in which the 3'-tail contained either six (Chi-minus 2, 3.6Å resolution) or ten (Chi-plus 2, 3.8Å resolution) bases between the junction and the Chi site also revealed no density in the Chi site (Table 1, Extended Data Figures 3 -5). Consequently, we can be confident that the changes we observe in the Chi-bound complexes are a result of specific interactions with that sequence and that a spacer of eight residues is required between the fork junction and the start of the Chi site.

Conformational changes in response to Chi recognition
In the Chi-recognised complex, density for the bound DNA is visible for all twenty bases of the 3'-tail, including the Chi sequence (  (Figures 2 -4). The conformational changes in RecC that are induced by the 5'-tail binding to RecD (15) appear to facilitate Chi recognition and bring several residues into proximity of the Chi-binding site prior to Chi binding, switching the subunit into a Chi-recognition mode. Binding to Chi induces a rotation of the 1A and 1B domains of RecC. The 1A domain of RecC forms a significant part of the interface with the RecB nuclease domain and the largest Chidependent movement is of the nuclease domain that rotates towards the Chi-binding site as part of a rigid-body movement with domain 1A of RecC (Supplementary Movie 2). The active site residues of the nuclease domain are now located close to the terminal T residue of the 3'-tail ( Figure 5). The final cleavage of the 3'-tail is located 4-6 bases after the Chi sequence (23), suggesting the structure is that after the final cleavage event following Chi recognition.
Details of the interactions between RecBCD and the bound Chi sequence (Figures 2 -4, Supplementary Movies 1 & 3) reveal multiple side chain interactions with residues in RecC, many of which have been identified from mutational studies (19,20,22). Interactions with each of the eight bases of the Chi sequence provide specificity while interactions with the phosphodiester backbone (mainly arginine side chains) help to stabilise the twisted conformation of the DNA, allowing the first and fourth residues (both guanines) of the Chi sequence to stack upon each other which provides additional specificity. It has been determined that mutation of any one of the eight bases in Chi is sufficient to reduce affinity to a level that is not functional in vivo (20).
In addition to the particles we assigned to the Chi-recognised structure, a further group were isolated in a class that also showed good density for Chi (Extended Data Figures 1 & 3). The majority of the structure was the same as the Chi-recognised complex, but the nuclease domain was in a slightly altered location, intermediate between the positions of the recognised and unrecognised complexes, that was slightly misaligned for interaction with the 3'-terminus. The density for the nuclease was also less well ordered (Extended Data Figure 1) suggesting the domain is more mobile. This class appears to have recognised Chi but is not optimally aligned for cleavage so we suggest this may be an intermediate in which Chi has been recognised but the nuclease is not aligned for cleavage and during active translocation may not have time to cleave before the Chi sequence is pushed out. We, therefore, refer to this as the Chi-intermediate structure.

Comparison with AddAB/Chi recognition complex
RecBCD shares many aspects of its activity with AddAB (1,2) and the structure of B.subtilis AddAB bound to its Chi sequence (AGCGG) has been determined (9). However, comparison of the structures reveals that interactions with their respective Chi sequences are very different, even though the structures and conformations of the protein components, and their interactions with the DNA fork, are all very similar ( Figure 4). In particular, the conformation of the Chi sequence itself is completely different despite the two sequences sharing some apparent similarity (9) ( Figure 4). However, the different interaction modes for Chi with RecC and AddB observed in the structures explains why the final cleavage event for RecBCD is 4-6 bases after Chi (23) but only a single base away for AddAB (24). In each case, the bound ssDNA is presented to the nuclease active site in a manner that is consistent with these differing cleavage sites (Figures 4 & 5). Consistent with our structural data, residues beyond the canonical eight base Chi site may also contribute to specificity, particularly in the 4-7 bases on the 3' side of the Chi sequence (21,22), albeit with reduced specificity. Furthermore, a RecBCD complex that carries a mutation in the RecC protein (17) recognises an extended eleven base Chi sequence (Chi* = GCTGGTGCTCG) that includes this region (25).

Mechanism of RecBCD stalling
Single-molecule studies have shown that, upon encountering Chi, RecBCD pauses for several seconds before recommencing translocation at (usually) a reduced speed (26,27). It is also known that an active RecB motor is required for Chi recognition and response (28). The RecB motor precedes the Chi-binding site in RecC so reactivation would push DNA towards the bound Chi sequence. However, the twisted and compacted Chi sequence would act as a physical block to prevent translocation and the DNA would become compressed within the space at the RecB/RecC interface. The conformational changes that provide additional interactions between RecC and the Chi sequence would tighten this grip even further and prevent translocation across and through the RecC subunit. As described previously (19,20), once translocation by the RecB motor recommences, the only exit for this DNA would be through a protein gate between the RecB and RecC subunits that is operated by an ion pair "latch".
Such a mechanism has similarities to the "DNA scrunching" mechanism observed for RNA polymerase during initiation of transcription that forces the RNA product out of the enzyme complex (29). Blocking the 3'-channel would also explain why nuclease digestion of the 3'strand is attenuated after Chi binding (30). Although the structure explains why RecBCD stalls at Chi sites (26,27) it does not provide details of the subsequent steps that precede full conversion to repair mode such as reactivation of translocation or how the RecD subunit becomes inactivated (27).

A mechanism to prevent phage evasion
It is known that RecBCD only cleaves at a Chi sequence in 25-40% of encounters (30,31). There are two explanations for this counter intuitive behaviour. The first is that the rate of translocation by RecBCD is so rapid that the Chi sequence passes over the RecC binding site before the complex has a chance to recognise the sequence. A second alternative is that there is an intrinsic equilibrium between bound and unbound states that is reached at every step of translocation. For this to be the case, at a translocation speed of around 1000 bases per second, this equilibrium would need to be established within 1 ms. For the particles with bound DNA substrate, around 40% had the Chi sequence bound specifically and were in the Chi-recognised state with the nuclease correctly positioned (Extended Data Figure 1), suggesting that the latter explanation is correct. Although an additional 20% have also bound Chi (Chi-intermediate), the nuclease domain does not appear to be correctly positioned for cleavage. The remaining 40% of molecules have failed to recognise Chi. Since singlemolecule studies of RecBCD indicate that translocation speeds vary considerably (26,27), this would ensure that Chi is recognised and cleaved with similar frequency, regardless of the translocation speed on the 3'-strand (RecB motor). Furthermore, bacteriophages that infect E. coli, such as lambda, typically lack Chi sequences or have them at very low frequency. Consequently, the reduced recognition frequency is an adaptation mechanism to ensure that even bacteriophages that acquire a Chi sequence will likely fail to escape degradation by RecBCD. By contrast, the elevated frequency of Chi sequences in the E. coli genome (4) ensures DNA repair is initiated within a few kilobases of the break site. Thus, these contradictory requirements for Chi recognition can be achieved simultaneously; allowing RecBCD to make the switch between being friend to the host, yet foe to bacteriophages.

Discussion
The twisted conformation of the Chi sequence when bound to RecBCD was unanticipated in light of the related AddAB/Chi structure reported previously (9). In AddAB, the Chi sequence is extended with each base sitting in a pocket on the surface of the AddB subunit, making side chain contacts with the protein that explain the specificity for the sequence. By contrast, RecC contacts Chi in a quite different manner with fewer base-specific contacts. Instead, the specificity appears to come from an overall interaction with a specific shape of the bound ssDNA that is stabilised, at least in part, by contacts such as the intra-sequence contacts between the first and fourth guanine residues in the sequence. Nonetheless, many side chains have been shown to be essential for Chi recognition (19,20,22) so these are, presumably, assisting in the shape recognition that leads to sequence specificity.
RecBCD has been validated as a target for antibiotic development (32). The differing modes of Chi interaction adopted by AddAB and RecBCD have important implications for drug discovery. Recognition of Chi by AddAB provides opportunities for the design of specific drugs that might interact at any or each of the base specificity pockets to prevent Chi binding. Such interactions could be different in organisms with different Chi sequences creating the potential for narrow spectrum targeting of groups of organisms with similar Chi sequences. By contrast, blocking of Chi recognition in RecBCD may require a different approach to find compounds that interact with a part of the pocket in a different manner. Nevertheless, the twisted conformation of the Chi sequence provides multiple surfaces and pockets that might provide specificity for drug design.
In E.coli, 75% of Chi sequences are located in the leading strand and oriented with a bias to favour activity towards the replication origin (4). Such an arrangement has been proposed to promote DNA repair processes during replication (4,33). Given this property, one would imagine that determination of Chi sequences would be straightforward in different organisms. However, that has proved not to be the case and analysis of bacterial genomic data has had limited success in identifying Chi sequences (34). Indeed, even the notion that Chi sequences have evolved to facilitate repair has been questioned. Instead, it was shown that G-rich sequences are naturally more prevalent and the orientation of G-rich sequences other than Chi also showed an apparent directional bias in E.coli but less so in other organisms (such as H.influenzae) that do not show an intrinsic sequence bias (35). This suggests that, rather than Chi sequences evolving as a regulator of Chi, an alternative is that RecBCD adopted a naturally occurring sequence with appropriate characteristics as a regulator, possibly initially with a higher redundancy but later to be more specific as a coevolution with the abundance of sequence of Chi itself. Such a proposal might explain why Chi sites have proved to be so hard to identify from bioinformatics analysis alone because under these circumstances there need be no conservation of sequences between organisms. To date, Chi sequences that bind to RecBCD have only been identified in two other organisms (H.influenzae and P.synringae) and these are both very similar to the E.coli sequence (E.coli -GCTGGTGG, H.influenzae -G(G/C)TGGTGG and P.syringae -GCTGGCGC).
The structures presented here help us to explain the long-standing observation of Chiregulation and its effects on phage biology. They help explain the switch from destructive to repair mode but several questions remain unanswered. For example, the structures show how the enzyme is stopped in its tracks during degradation but they do not explain how the complex changes in order to recommence translocation after pausing at the Chi site. Several events take place after Chi recognition including inactivation of the RecD motor (27) and loading of RecA protein onto the 3'-tail. Hopefully, future structures will shed light on those processes.

Purification of RecBCD
To prevent digestion of the single-stranded tails of the DNA substrate, a nuclease-deficient E. coli RecBCD mutant (D1080A in RecB) was used in this work. The complex was produced from three plasmids, pETduet-His6-TEVsite-recBD1080A, pRSFduet-recC and pCDFduet-recD in a ΔrecBD E. coli strain as described previously (12). His-tagged RecBCD complex was expressed and purified as described previously (16,36) with the Histag removed by TEV protease. The purified protein was flash-frozen with 15% glycerol and stored at -80 °C until use.

Preparation of forked DNA substrates
The forked DNA substrates used during this work were made by annealing equal molar quantities of two respective ssDNA oligonucleotides to generate forks with 25 bp duplex, 15 base single-stranded 5'-tail and a 20 base single-stranded 3'-tail. The same upper strand, responsible for generating the 5'-tail, was used in all cases and had the sequence: 5' TTTTTTTTTTTTTTTgagcgactgcactacaacagaacca 3' (lower case represents the bases that form the duplex region). The variable lower strand, responsible for presenting Chi (GCTGGTGG), contained the sequence: 5' tggttctgttgtagtgcagtcgctc(Y)GCTGGTGG(Z) 3'. Y and Z represent the variable number of T bases used to alter the position of the Chi sequence whilst keeping the total number of unpaired bases to 20 (see Figure 1). In the Chiminus negative control substrate, the Chi sequence was replaced by eight T bases to give a 3'-tail of twenty unpaired T bases.

Native gel mobility assays
For native gel mobility assays, all DNA substrates contained the upper strand labelled with fluorophore (Cy5) at the 5'-terminus and for the lower strand Y values of 1-10 with corresponding Z values of 11-2 were used. Typically, 500 nM RecB D1080A CD was incubated with 500 nM DNA substrate in a 5μl binding solution containing 50 mM Tris-HCl (pH 8.0), 100 mM NaCl, 0.5 mM TCEP and 5% glycerol, which was incubated on ice for 10 min. Samples were separated on 5% native polyacrylamide gels in 1 × TB buffer. Gels were scanned in fluorescence mode (Cy5) on a BIO-RAD ChemiDocTM MP imaging system.

Cryo-electron microscopy grid preparation and data collection
Prior to preparing grids, RecBCD was thawed and desalted into 20 mM Tris-HCl (pH 8.0), 50mM NaCl, 0.5 mM TCEP using Sephadex G25 spintrap columns (GE Healthcare). The protein was mixed with a 1.5 fold excess of DNA substrate for 10 min at room temperature. Final concentrations were 1 μM RecBCD and 1.5 μM DNA substrate. A RecBCD-Chi complex was prepared using a Chi-containing DNA substrate with a Y spacing of 8 and a Z spacing of 4 (see Figure 1). For the Chi-minus dataset, a complex was prepared using the negative control substrate lacking Chi. For the Chi-minus 2 dataset, a complex was prepared using a Chi-containing DNA substrate with a Y spacing of 6 and a Z spacing of 4 ( Figure 1). For the Chi-plus 2 dataset, a complex was prepared using a Chi-containing DNA substrate with a Y spacing of 10 and a Z spacing of 4 ( Figure 1). Quantifoil R2/2 μm holey carbon film grids (300 mesh) were treated by plasma cleaning for 30 s before being covered with graphene oxide sheets. In order to improve grid preparation reproducibility and hydrophilicity of the graphene oxide, the following method was used: 5μl of graphene oxide solution (Aldrich 763705, 2mg/ml) were mixed with 5μl of 1.5% w/v solution of nonionic detergent n-Dodecyl-β-D-Maltoside. The mixture was diluted 100 times with water and 5ul was applied to the carbon side of the grid. A sharp edge of a piece of filter paper was applied to the centre of the opposite surface of the grid to pull the graphene oxide solution through the grid. Grids were used within 30 minutes of graphene oxide application. Sample (4 μL) was evenly applied to the graphene oxide-coated side of the grid, followed by a 5 s wait time, 1 s blot time, and freezing in liquid ethane using a Vitrobot Mark IV (FEI). The Vitrobot chamber was maintained at close to 100% humidity at 4°C.
The dataset with the Chi substrate was collected using a Titan Krios microscope operated at 300 KV at eBIC, Diamond, UK. Zero loss energy images were collected automatically using EPU (FEI) on a Gatan K2-Summit detector in counting mode with a pixel size of 1.047 Å. A total of 3,721 images were collected with a nominal defocus range of −1.3 to −2.5 μm in 0.3 μm increments. Each image consisted of a movie stack of 40 frames with a total dose of 45 e-/Å 2 over 10 s corresponding to a dose rate of 5 e-/pixel/s. The dataset with the Chi-minus control substrate was collected using a similar collection strategy again with a Gatan K2-Summit detector and Titan Krios microscope at eBIC, Diamond, UK. The pixel size was 1.048 Å and a total of 788 images were collected with a similar defocus range to the above. A total dose of 45 e-/Å 2 was split into 40 frames over 7 s, corresponding to a dose rate of 7 e-/pixel/s. The datasets with the Chi-minus 2 and Chi-plus 2 were collected using a Titan Krios microscope operated at 300 KV at eBIC, Diamond, UK. Zero loss energy images were collected automatically using EPU (FEI) on a Falcon3 detector in integrating mode with a pixel size of 1.085 Å. A total of 4,013 images (Chi-minus 2 substrate) and 3,613 images (Chi-plus 2 substrate) were collected with a nominal defocus range of −1.2 to −2.7 μm in 0.3 μm increments. Each image consisted of a movie stack of 39 frames with a total dose of 76 e -/Å 2 over 1 s corresponding to a dose rate of 89 e -/pixel/s.

Data processing -The Chi-containing dataset
Movie stacks were aligned and summed using Motioncor2 (37). Template-free particle picking was done in Gautomatch, using a circular diameter of 150 Å and CTF parameters were estimated for each micrograph using Gctf (38). A total of 1,053,451 picked particles were extracted 2x binned from 3,721 images for two rounds of 2D classification in RELION3 (39) in which 373,476 real RecBCD particles were kept and picking artefacts/ noise discarded. A consensus refinement was run on all the particles followed by unmasked 3D classification without alignment. From this, 204,633 particles with complete density for the complex were selected and sub-stoichiometric classes discarded (no DNA density, no RecB nuclease domain density or weaker RecD density). These 204,633 particles were reextracted unbinned, 3D refined and Bayesian polished prior to 3D classification without alignment using a mask of RecC around the prospective Chi-binding site. This separated 127,268 particles with ordered density for the complete ssDNA 3' tail, including the Chi sequence, from 77,365 particles with no ssDNA density beyond the RecB helicase domains.
Each class was further refined and classified without alignment, this time using a mask around both RecD and the RecB nuclease. The classifications removed 8,452 and 3,092 particles from the Chi and no Chi classes respectively, which had weaker density for the RecD helicase domains. This left 74,273 homogenous particles from the no Chi class, that were refined to give a 3.9 Å resolution (0.143 FSC cutoff, RELION3) map representing a Chi-unrecognised complex. This structure was very similar a previous RecBCD structure with a forked DNA substrate containing long ssDNA tails (PDB: 5LD2) (16). From the Chi class, two significant conformations were identified. The major state, with 74,496 particles, refined to a resolution of 3.7 Å (0.143 FSC cutoff, RELION3) and represented a Chirecognised complex with ordered Chi density and significant local conformational changes involving RecC and the RecB nuclease. The second Chi state contained 44,320 particles, refined to 4.1 Å (0.143 FSC cutoff, RELION3) and similarly showed ordered Chi density and the associated local conformational changes in RecC. However, this third state did not show such a substantial movement in the RecB nuclease domain, therefore may represent an intermediate between the Chi-unrecognised and Chi-recognised states, which we have called the Chi-intermediate complex. Work flows for data processing for this and all other structures are presented in Extended Data Figures 1 and 2. An analysis of particle orientations for all structures is presented in Extended Data Figure 6.

Data processing -The Chi-minus control dataset
The Chi-minus movie stacks were initially processed similarly to the Chi dataset. A total of 271,210 particles were extracted 2x binned from 788 images for two rounds of 2D classification in RELION3, resulting in 99,545 RecBCD particles. A consensus 3D refinement was run followed by unmasked 3D classification without alignment from which a homogeneous subset of 87,229 particles with complete density for the complex was selected. The particles were re-extracted unbinned, refined and Bayesian polished prior to 3D classification without alignment using the mask of RecC around the Chi binding site. This time there was only one class, with no DNA density observed in the Chi-binding channel. 3D classification with a mask of RecD plus the RecB nuclease domain separated a class of sub-stoichiometric particles (24,417 particles). The remaining major class (62,812 particles) was refined to produce a 3.8 Å resolution map (0.143 FSC cutoff, RELION3). This represents the Chi-minus complex, which resembles the Chi-unrecognised complex and similarly does not contain ordered 3' ssDNA beyond the RecB helicase.

Data processing -The Chi-minus 2 dataset
The movie stacks were initially processed similarly to the Chi-containing dataset. A total of 1,694,205 particles were extracted 2x binned from 4,013 images for two rounds of 2D classification in RELION3, resulting in 754,150 RecBCD particles. A consensus 3D refinement was run followed by unmasked 3D classification without alignment from which a homogeneous subset of 687,825 particles with complete density for the complex was selected. The particles were re-extracted unbinned, refined and Bayesian polished prior to 3D classification without alignment using the mask of RecC around the Chi binding site. Only one class, with no DNA density observed in the Chi-binding channel was obtained. 3D classification with a mask of RecD plus the RecB nuclease domain separated a class of substoichiometric particles (308,035 particles). The remaining major class (379,790 particles) was refined to produce a 3.6 Å resolution map (0.143 FSC cutoff, RELION3). This structure resembles the Chi-unrecognised complex and similarly does not contain ordered 3' ssDNA beyond the RecB helicase.

Data processing -The Chi-plus 2 dataset
The movie stacks were initially processed similarly to the Chi-containing dataset. A total of 1,809,768 particles were extracted 2x binned from 3,613 images for two rounds of 2D classification in RELION3, resulting in 795,810 RecBCD particles. A consensus 3D refinement was run followed by unmasked 3D classification without alignment from which a homogeneous subset of 776, 381 particles with complete density for the complex was selected. The particles were re-extracted unbinned, refined and Bayesian polished prior to 3D classification without alignment using the mask of RecC around the Chi binding site. Only one class, with no DNA density observed in the Chi-binding channel was obtained. 3D classification with a mask of RecD plus the RecB nuclease domain separated a class of substoichiometric particles (395,469 particles). The remaining class (380,912 particles) was refined to produce a 3.8 Å resolution map (0.143 FSC cutoff, RELION3). This structure resembles the Chi-unrecognised complex and similarly does not contain ordered 3' ssDNA beyond the RecB helicase.

Model building and refinement
The structure of RecBCD in a complex with a DNA fork (PDB 5LD2) (16) was used as a starting model for global docking in Chimera (40) for the Chi-recognised, Chi-unrecognised, Chi-minus 2, and Chi-plus 2 complexes. Once finalised, the Chi-recognised model was then used as a starting template for the Chi-intermediate complex. The Chi-unrecognised model was used a starting template for the Chi minus complex. Each of the six models was edited initially by jelly-body refinement with Refmac (41) in CCP-EM (42) followed by cycles of manual rebuilding in Coot (43) and real-space refinement with PHENIX (44). A 3.7 Å resolution (0.143 FSC cutoff, RELION3) map from a masked focused refinement of the Chi region for the 127,268 Chi-bound particles was used to help build the novel Chi bases and surrounding protein contacts. The six models were finally checked for consistency with one another, apart from where obvious density differences occurred in the maps. A final run of PHENIX real-space refinement with group B-factor refinement was run to generate the final models and model statistics (Table 1).  a, Density corresponding to the bound DNA fork substrate in the Chi-unrecognised state with the protein complex shown as a faint cartoon. Strong density is observed for the duplex and 5'-tail regions. Density is also observed for the first six bases of the 3'-tail, but is disordered beyond that. b, Density corresponding to the bound DNA fork substrate in the Chi-recognised state. The density is similar to that observed in the Chi-unrecognised state except that additional density is observed for the entire 20 bases of the 3'-tail, including the Chi sequence. Inset, Expanded view of the density (contoured at 2.5 σ) for the eight Chi bases with the Chi bases individually coloured as in panel (c) demonstrating the highly twisted conformation of the DNA. c, Details of the interactions with each base of the Chi sequence. Hydrogen bonds are shown as dotted lines. A cross-eye stereo view of the Chi-binding site and interacting residues is shown. Interacting residues are shown as sticks and coloured according to previously published point mutation studies (19,20). Mutations at residues that failed to recognize Chi are in violet, those that showed altered/relaxed Chi recognition specificity are in green, and those that showed little or no influence after mutation are grey. Additional residues which failed to recognize Chi after mutation (22) are coloured in yellow. a, Transparent surfaces of RecBCD (above) and AddAB (below) coloured by subunit with the DNA fork backbones overlaid as a cartoon in grey with their respective Chi sequences depicted in red and the bases between Chi and the nuclease active site in yellow. b, Expanded cartoon view of the Chi-binding region in each structure with the RecC domains labelled to highlight the different Chi-binding surfaces utilised by the two enzymes. c, Cartoon representation detailing the contacts between the proteins and DNA. The nuclease domains of RecBCD and AddAB are shown as cartoons. Key residues participating in DNA binding and catalysis are shown as sticks. Hydrogen bonds are shown as dotted lines. The DNA forks used for both enzymes represent substrates after the final cleavage, with four bases after Chi for RecBCD but just a single base for AddAB (in each case depicted in yellow). The difference in the final cleavage site relates to the distance between the end of the Chi-binding site and the nuclease even though the relative position of the nuclease, relative to RecC or AddB, are similar. The normal cleavage of DNA after Chi in RecBCD would result in a terminal 3'-phosphate but this was not added to the substrate synthesised for the structure determination. Cheng


extfieldsofstudy: Chemistry Medicine

s2fieldsofstudy: Biology Chemistry Medicine

text:
TNF-α enhances Th9 cell differentiation and antitumor immunity via TNFR2-dependent pathways

Tumor specific Th9 cells are potential effector cells for adoptive therapy of human cancers. TNF family members OX40L, TL1A and GITRL have been shown to promote the induction of Th9 cells and antitumor immunity. However, the role of TNF-α, the prototype of the TNF superfamily cytokines, in Th9 cell differentiation and their antitumor efficacy is not defined. Here, we showed that TNF-α potently promoted naïve CD4+ T cells to differentiate into Th9 cells in vitro. Furthermore, the addition of TNF-α during Th9 cell differentiation increased T cell survival and proliferation. More importantly, the adoptive transfer of TNF-α-treated Th9 cells induced more potent antitumor effects than regular Th9 cells in mouse tumor model. TNF-α signals via two cell surface receptors, TNFR1 and TNFR2. Mechanistic studies revealed that TNF-α drove Th9 cell differentiation through TNFR2 but not TNFR1. In addition, under Th9 polarizing condition, TNF-α activated STAT5 and NF-κB pathways in T cells in a TNFR2-dependent manner. Inhibition of STAT5 and NF-κB pathways by their specific inhibitors impaired TNF-α-induced Th9 cell differentiation. Our results identified TNF-α as a new powerful inducer of Th9 cells and clarified the molecular mechanisms underlying TNF-α-induced Th9 cell differentiation.

Introduction
Adoptive T-cell therapy (ACT) has shown encouraging results in some cancer types; however, most tumors are refractory to ACT [1]. The efficacy of ACT of cancers relies on the cytolytic activity and the in vivo persistence of the transferred effector T cells [1][2][3][4]. Tumor-specific CD8 + T cells have powerful cytolytic activities against tumor cells [1]. However, CD8 + T cells used in ACT of cancers are often terminally differentiated and have a disappointing lack of persistence in vivo [2]. CD4 + T helper (Th) cells are another option of effectors in ACT of cancers. Among Th cell subsets, the cytotoxic Th1 cells are a useful T cell lineage for ACT of cancers [3]. However, Th1 cells display an exhausted phenotype, and have a short-term persistence in vivo [3]. Compared to Th1 cells, the "stem cell-like" memory Th17 cells have reduced cytolytic function in vitro, but persist significantly longer in vivo, resulting in better antitumor efficacy in ACT of cancers [4,5]. Th9 cells are a unique Th cell subset with a mature T cell phenotype, highly cytolytic activity and prolonged persistence in vivo based on their hyperproliferation, suggesting an excellent effector for ACT of human cancers [6].
TNF-α is a potent proinflammatory cytokine, which is implicated in the immunopathology of various inflammatory diseases [24]. TNF-α stimulates inflammatory cytokine production, cell growth, cell survival and paradoxically, cell death [25][26][27]. In cancer immunology, TNF-α activates antigen-presenting cells and promotes the activation and proliferation of effector T cells [28,29]. TNF-α impairs the function of regulatory T (Treg) cells [30], which contributes to antitumor immunity. TNF-α has two cell surface receptors TNFR1 and TNFR2 [24]. TNFR1 is widely expressed, whereas the expression of TNFR2 is limited to immune and endothelial cells [24,31]. The cytoplasmic regions of TNFR1 contain a conserved 'death' domain which is essential for triggering cell apoptosis and subsequent activation of NF-κB [27,31]. In contrast, TNFR2 lacks the cytoplasmic 'death' domain and has mainly been linked to cell survival and proinflammatory reactions [31,32].
In this study we found that TNF-α profoundly stimulates Th9 cell differentiation. And the adoptive transfer of TNF-α-treated Th9 cells induces more potent inhibition on melanoma tumor growth than regular Th9 cells in mouse models. In addition, we clarified the TNFR2-dependent signaling pathways of TNF-α-induced Th9 cell differentiation.

Materials and methods
Mice and cell lines C57BL/6 (H-2 b ), TNFR1 −/− (B6.129-Tnfrsf1a tm1Mak /J) and TNFR2 −/− (B6.129S2-Tn frsf1b tm1Mwm /J) mice were purchased from the Jackson Laboratory. Mice were housed in specific pathogen-free conditions at the First Hospital Animal Center of Jilin University. Mice at 6-8 weeks of age were used in experiments. All animal experiments were approved by the Animal Ethical Committee of First Hospital of Jilin University.
Anti-TNFR1 and anti-TNFR2 blocking mAbs and control IgG were purchased from Biolegend. STAT5 inhibitor and Bortezomib (a NF-κB inhibitor) were purchased from Santa Cruz and Selleckchem respectively.
To examine the role of TNFR1 and TNFR2 in TNF-α-induced Th9 cell differentiation, naïve CD4 + T cells were cultured under the Th9-polarizing conditions with or without addition of TNF-α (50 ng/mL). Cell cultures were added with anti-TNFR1 (50 μg/mL), anti-T NFR2 (50 μg/mL) mAbs or control IgG (50 μg/mL). After 3 days of culture, the cells and culture supernatants were harvested and analyzed.
To explore the signaling pathways involved in TNF-α-induced Th9 cell differentiation, naïve CD4 + T cells were cultured under the Th9-polarizing conditions in the presence or absence of TNF-α. In some cell cultures, STAT5 Inhibitor (10 μg/mL) or Bortezomib (1 nM) was added. After 3 days of culture, the cells and culture supernatants were harvested and analyzed.

Enzyme-linked immunosorbent assay (ELISA)
Concentrations of IL-9 in culture supernatant were detected by ELISA as previously described [23]. IL-9 Capture/detection antibodies were purchased from BD Biosciences. Recombinant mouse IL-9 used as the standards in ELISA were purchased from R&D Systems. Avidin-HRP was purchased from Biolegend.

RNA sequencing (RNA-Seq)
Mouse naïve CD4 + T cells were cultured under Th9 polarizing conditions with or without addition of TNF-α for 6 h and cells were collected for RNA extraction. Total RNA was extracted with the Trizol (Thermo-Fisher) and RNA-Seq was done by the genomics core of Lerner Research Institute in Cleveland Clinic with Illumina HiSeq2500.

Co-immunoprecipitation (co-IP) and mass spectrometry (MS)
Naïve CD4 + T cells were cultured under Th9 polarizing conditions with or without addition of TNF-α for 3 h. Cells were collected and cell lysates were prepared in non-denaturing lysis buffer. Cell lysates were incubated with anti-TNFR2 antibody for 2 h at 4°C, and subjected to immunoprecipitation (IP) using protein A-sepharose beads. The beads were washed with IP buffer. The proteins were eluted with SDS sample buffer and heated at 98°C for 5 min. IP samples were then separated using SDS-PAGE and visualized by Coomassie Blue staining. The immunoreactive bands were excised from stained gels and digested overnight with trypsin (10 ng/μL) at room temperature. Peptides in the digested sample were analyzed using liquid chromatography mass spectrometry (LC-MS) provided by Mass Spectrometry Laboratory for Protein Sequencing (Lerner Research Institute, Cleveland Clinic, Cleveland, Ohio, USA).
Adoptive tumor immunotherapy 2 × 10 5 B16-OVA cells were injected subcutaneously into C57BL/6 mice. To generate Th9 cells, naïve CD4 + T cells from OT-II mice were cultured under Th9 polarizing conditions in the presence or absence of TNF-α for 2 days. On Day 2 after tumor injection, the mice were randomly divided into groups and transfused with Th9 or TNF-α-treated Th9 cells (1 × 10 6 ) via tail vein injection. Mice treated with PBS served as controls. Tumor development was monitored over time. The mice were killed when the tumor diameter reached between the range of 1.5 and 2 cm. Tumor volume was calculated by the formula: 3.14 × (mean diameter) 3 /6.

Statistical analysis
The Student t test (2 groups) and one-way ANOVA (> = 3 groups) were used to compare various experimental groups. A P value of less than 0.05 was considered significant.

TNF-α promotes Th9 cell differentiation in vitro
To examine the role of TNF-α in Th9 cell differentiation, naïve CD4 + T cells were cultured in the presence of anti-CD3/28 antibodies plus TGF-β, IL-4 and/or TNF-α for 3 days. The addition of TNF-α combined with Th9 polarizing cytokines TGF-β and IL-4 increased Th cell expression of IL-9 mRNA and protein ( Fig. 1a, b), and the frequency of Th9 cells (Fig. 1c). However, TNF-α alone or TNF-α plus TGF-β or IL-4 could not induce Th9 cell differentiation (Fig. 1a-c). Interestingly, TNF-α did not increase the expression of Spi1 or Irf4 in Th9 cells (Fig. 1d), suggesting that TNF-α may drive Th9 cell differentiation through other Th9-related transcription factors. We also examined the expression of the other Th cell-related cytokines and transcription factors and found that TNF-α-treated Th9 cells did not express most of Th1-, Th2-, Th17-and Treg-related cytokines and transcription factors, such as Ifng, Il4, Il17, Tbx21, Gata3, Rorc and Foxp3 (Fig. 1d, e), although Il5 and Il13 were increased (Fig. 1e) in TNF-α-treated Th9 cells compared to regular Th9 cells. We also examined the effects of TNF-α on the expression of Il9 in Th9 cells at different time points. We found that the expression of Il9 in TNF-α-treated Th9 cells increased on Day 1, reached the highest level on Day 2 or Day 3, and then slightly decreased from the highest level on Day 4 (Fig. 1f).
Together, these results demonstrated that TNF-α promotes Th9 cell differentiation in vitro.

TNF-α increases the survival and proliferation of Th9 cells in vitro
The survival and proliferation of tumor-specific T cells are crucial for their persistence and therapeutic potential in vivo. We next examined the effects of TNF-α treatment on Th9 cell survival and proliferation. Naïve CD4 + T cells were cultured under Th9-polarizing conditions in the presence or absence of TNF-α for 6 h. RNA-seq analysis of the cultured T cells revealed that the addition of TNF-α decreased T cell expression of Casp9, Tradd, Tnfsf10, Dffb, Bid, Traf2, Cflar and Casp8 (Fig. 2a), the genes related to cell apoptosis, and increased the expression of Bcl2, Birc2, Birc3 and Nfkbia (Fig. 2a), the genes related to cell survival, suggesting that the addition of TNF-α during Th9 cell differentiation inhibits T cell apoptosis and promotes T cell survival.
To further exploit the role of TNF-α in Th9 cell survival and proliferation, Th9 cells with or without TNF-α-treatment were generated and cell death was assessed by Annexin V staining. While Th9 cells showed less cell apoptosis than Th0 cells (Fig. 2b), TNF-α treatment further reduced the apoptosis of Th9 cells (Fig. 2b). In addition, TNF-α treatment increased Th9 cell proliferation (Fig. 2c). These results demonstrated that TNF-α enhances T cell survival and proliferation during Th9 cell differentiation.
PD-1 is an immune-checkpoint and PD-1 up-regulation promotes CD4 + T cell apoptosis [33]. We next examined the role of TNF-α on the expression of PD-1 by Th9 cells. While Th9 cells expressed lower mRNA levels of Pdcd1 than Th0 cells (Fig. 2d), TNF-α treatment further decreased the expression of Pdcd1 in Th9 cells (Fig. 2d). Furthermore, TNF-α treatment increased Th9 cell expression of CD44 (Fig. 2e), a marker for effector and memory T cells which contribute to T cell survival and proliferation [34]. Together, these results demonstrated that TNF-α enhances Th9 cell survival and proliferation in vitro.

TNF-α treatment improves the antitumor efficacy of Th9 cells in vivo
To assess the antitumor efficacy of TNF-α-treated Th9 cells, naïve CD4 + T cells from OT-II mice were differentiated into Th9 cells in the presence or absence of TNF-α. Cells were used to treat B16-OVA-bearing C57BL/6 mice. While Th9 cells mediated a higher inhibition on melanoma tumor growth than PBS control (Fig. 3a), the addition of TNF-α during Th9 cell differentiation further improved their antitumor efficacy (Fig. 3a), demonstrating that TNF-α improves the antitumor efficacy of Th9 cells.
The infiltration of effector T cells into tumor sites is associated with their antitumor effects. We next examined the tumor-infiltrating capability of TNF-α-treated Th9 cells. OT-II Th9 cells and TNF-α-treated Th9 cells were transfused to B16-OVA-bearing C57BL/6 mice, and cells from tumor-draining lymph nodes (TDLNs) were collected and analyzed. While mice transfused with Th9 cells had higher frequencies of IL-9 + CD4 + T cells in TDLNs than PBS control mice (Fig. 3b), the percentages of IL-9 + CD4 + T cells in TDLNs further increased in mice receiving TNF-α-treated Th9 cells compared to regular Th9 cells (Fig. 3b). In addition, TDLN CD4 + T cells from mice receiving TNF-α-treated Th9 cells expressed higher levels of Il9 (Fig. 3c), Il5 and Il13 (Fig.  3d) than mice transfused with Th9 cells or PBS control. We also examined CD4 + T cells isolated from pulmonary tumor tissues. As shown in Fig. 3e, cells from mice receiving TNF-α-treated Th9 cells expressed higher levels of Il9 than mice transfused with Th9 cells or PBS controls. These results indicated that TNF-α treatment increases the tumor-infiltrating capability of Th9 cells.

TNF-α enhances Th9 cell differentiation through TNFR2
There are two cell surface receptors for TNF-α, TNFR1 and TNFR2 [24]. CD4 + T cells expressed both TNFR1 and TNFR2 (Fig. 4a). We next explored the contribution of TNFR1 and TNFR2 to TNF-α-induced Th9 cell differentiation. TNFR1 (αR1) or TNFR2 (αR2) blocking antibodies were used during the in vitro differentiation of Th9 cells with or without the addition of TNF-α. The addition of αR1 did not affect the expression of IL-9 mRNA and protein in TNF-α-treated Th9 cells as compared to control IgG (Fig. 4b, c), whereas the addition of αR2 abolished TNF-α-induced up-regulation of IL-9 expression in Th9 cells (Fig. 4b, c). In addition, the addition of αR2 but not αR1 abolished the up-regulation of Il5 and Il13 expression induced by TNF-α in Th9 cells (Fig. 4d). These results indicated that TNFR2 but not TNFR1 mediates the stimulatory activity of TNF-α in Th9 cell differentiation.

TNF-α enhances Th9 cell differentiation through TNFR2-STAT5 signaling
We next exploited the downstream signaling pathways of TNFR2 that are responsible for TNF-α-induced Th9 cell differentiation. RNA-seq assay showed that the addition of TNF-α during Th9 cell differentiation decreased the expression of Stat1 and increased the expression of Nfkb2, Traf6, Irf1 and Irf4 (Fig. 5a), genes related to cytokine-induced signaling pathways. To exploit the proteins that may interact with TNFR2, naïve CD4 + T cells were cultured under Th9 polarizing conditions in the presence or absence of TNF-α and immunoprecipitation (IP) of the cell lysates was performed by using an anti-TNFR2 antibody, followed by mass spectrometry (MS) analysis of the immune precipitates. Interestingly, higher levels of STAT1 were detected in TNF-α-treated cells as compared to untreated control cells (Fig. 5b), suggesting that STAT signaling pathways may be involved TNF-α-induced Th9 cell differentiation.
We next explored the effects of TNF-α on the activation of STAT family members in T cells cultured under Th9 polarizing condition. Western-blots detected increased levels of phosphorylated (p) STAT1 (pSTAT1), independent experiments or presented as mean ± SD of three (b-i) independent experiments. NS, non-significant; *P < 0.05; **P < 0.01 pSTAT3 and pSTAT5 in T cells at Hour 3 compared to Hour 1 (Fig. 5c); whereas only pSTAT5 increased further in TNF-α-treated T cells compared to untreated controls at Hour 3 (Fig. 5c). These results indicated that TNF-α enhances STAT5 activation during Th9 cell differentiation.
To determine the role of TNFR1 and TNFR2 in TNF-α-induced activation of STAT5, naïve CD4 + T cells were isolated from WT, R1 −/− and R2 −/− mice and cultured under Th9 polarizing conditions in the presence or absence of TNF-α. The addition of TNF-α remarkably increased pSTAT5 in R1 −/− T cells as compared to the untreated control (Fig. 5d); however, TNF-α treatment exhibited minor effects on the protein levels of pSTAT5 in R2 −/− T cells as compared to the untreated control (Fig. 5d). These results demonstrated that TNF-α activates STAT5 via TNFR2 but not TNFR1.
To investigate the role of STAT5 signaling in TNF-α-induced Th9 cell differentiation, a STAT5 specific inhibitor (STAT5i) was used during Th9 cell differentiation. The inhibition of STAT5 significantly decreased IL-9 mRNA and protein expression in both TNF-α-treated Th9 cells and regular Th9 cells (Fig. 5e, f), indicating that TNF-α-induced Th9 cell differentiation relies on STAT5 signaling pathways. Collectively, these results demonstrated that TNF-α induces Th9 cell differentiation through TNFR2-mediated activation of STAT5.

TNF-α enhances Th9 cell differentiation through NF-κB pathway
RNA-seq assay showed that TNF-α increased the expression of Nfkb2 and Traf6 in T cells (Fig. 5a), genes related to NF-κB signaling pathway. To further determine the role of TNF-α in the activation of NF-κB pathway in T cells, naïve CD4 + T cells were treated with TNF-α under Th9 polarizing conditions and cells were analyzed by Western-blots. TNF-α treatment increased the protein levels of p-IKKα/β in T cells (Fig. 6a), indicating that TNF-α activates NF-κB pathway during Th9 cell differentiation.
We next explored the role of NF-κB pathway in TNF-α-induced Th9 cell differentiation. We first performed luciferase reporter assays to examine whether these NF-κB molecules could bind directly to Il9 promoters and stimulate its expression. We found that p50, c-Rel-p50, p50-RelB and p52-RelB dimmers could bind to and activate Il9 promoter (Fig. 6b). Bortezomib was used as an inhibitor of NF-κB pathway [11]. To further determine the role of NF-κB pathway in TNF-α-induced Th9 cell differentiation, Bortezomib was used during TNF-α-induced Th9 cell differentiation. Th9 cells treated by TNF-α plus bortezomib expressed lower levels of IL-9 mRNA and protein than those treated by TNF-α alone (Fig. 6c, d). These results indicated that TNF-α induces Th9 cell differentiation through NF-κB signaling pathway.

Discussion
Tumor-specific Th9 cells are potential effector cells for adoptive therapy of human cancers [6]. Therefore, identifying factors that can stimulate Th9 cell development may have important clinical significance. Recently, TNF family cytokines OX40L, TL1A, and GITRL have been shown to promote Th9 formation and their antitumor effects [20][21][22][23]. However, the role of TNF-α in Th9 cell differentiation and their antitumor functions remains unknown. In this study, we found that TNF-α potently promotes Th9 cell differentiation and IL-9 production. In addition, TNF-α stimulates IL-9 expression in T cells at multiple time-points between Day 1 and Day 4 during Th9 cell differentiation. More importantly, the adoptive transfer of TNF-α-treated Th9 cells induces more potent antitumor effects than regular Th9 cells in mouse tumor models. Multiple mechanisms may be involved in the increased antitumor capability of TNF-α-treated tumor specific Th9 cells. The addition of TNF-α increases Th9 cell expression of IL-9, which is a mediator of antitumor immunity [6,16]. TNF-α increases the expression of IL-2 in T cells [35] and the survival and proliferation of Th9 cells, which may prolong their persistence in vivo [6]. Furthermore, the high tumor-infiltrating capability of TNF-α-treated tumor-specific Th9 cells may also contribute to their antitumor efficacy. Thus, our data identify TNF-α as a new powerful inducer of Th9 cells.
TNF-α has two cell surface receptors TNFR1 and TNFR2 [24]. And T cells express both TNFR1 and TNFR2. In this study, we discovered that the blockade or depletion of TNFR2 in T cells abrogates the stimulatory effects of TNF-α on Th9 cell formation, indicating that TNFR2 downstream signaling is required for TNF-α-induced stimulation of Th9 cell formation. Compared to TNF-α/TNFR1 signaling, which has been shown to activate NF-κB, MAPK and caspase 8-mediated cell apoptotic pathways [36][37][38], the TNF-α/ TNFR2 downstream signaling is not well characterized. Recent studies indicated that TNFR2 can interact with Fig. 6 TNF-α enhances Th9 cell differentiation through NF-κB signaling pathway. (a) Naïve CD4 + T cells were cultured under Th9 polarizing conditions with or without addition of TNF-α for 3 h. Western-blots examined the protein levels of phosphorylated IKKα/β (p-IKKα/β) and IκB-α in cells. β-actin was used as a loading control. (b) 293 T cells were transiently transfected with vectors contained Il9 promoter or empty vector, followed by transfecting with vectors expressing the indicated NF-κB molecules. Luciferase reporter assay showed NF-κB-dependent activation of Il9 promoter in 293 T cells. (c, d) Naïve CD4 + T cells were cultured under Th9 polarizing conditions in the presence or absence of TNF-α with or without (DMSO) the addition of NF-κB inhibitor bortizomib (Bor) for 3 days. IL-9 expression was examined by qPCR (c) and ELISA (d). Data are representative of three (a) independent experiments or presented as mean ± SD of three (b-d) independent experiments. *P < 0.05 TRAF proteins TRAF1, 2 and 3 [39] and activate both the canonical and the noncanonical NF-κB signaling pathways [40,41]. In this study, we found that TNF-α activates NF-κB signaling by increasing the expression of TRAF6 and p-IKKα/β in T cells. In addition, we discovered that TNF-α/TNFR2 signaling activates STAT5 pathway in T cells. And, blocking NF-κB or STAT5 by their specific inhibitors partially abrogates TNF-α-induced stimulation of Th9 cell formation. Therefore, TNF-α/ TNFR2 signaling contributes to Th9 differentiation via two different mechanisms: the activation of (i) the transcription factor STAT5 and (ii) the TRAF6-NF-κB pathway. These two mechanisms may act synergistically to promote Th9 cell differentiation. Interestingly, published studies showed that STAT5 and NF-κB pathways are also involved in the stimulation of Th9 cell differentiation by other TNF family cytokines OX40L, TL1A, and GITRL [20][21][22], indicating that the TNF family cytokines may promote Th9 cell differentiation via some common signaling pathways.
IL-5 and IL-13 are type-2 cytokines expressed primarily in Th2 and mast cells [45]. GATA3 is the master transcription factor for Th2 cells and controls the expression of Th2-derived cytokines, including IL-5 and IL-13 [45]. In this study, we found that TNF-α increases the expression of IL-5 and IL-13 in Th9 cells. However, TNF-α/TNFR2 signaling does not increase GATA3 expression, but activates the STAT5 and NF-κB pathways in T cells during Th9 cell differentiation. Interestingly, a previous study showed that STAT5 and NF-κB stimulate the expression of IL-5 and IL-13 in Th cells [46]. These observations suggest that TNF-α/TNFR2 signaling enhances the expression of IL-5 and IL-13 in Th9 cells through STAT5 and NF-κB pathways.
Cytokine milieu is the major determinant for Th cell differentiation [47]. In this study, we observed that TNF-α/TNFR2 signaling promotes Th9 cell differentiation and increases the antitumor capability of tumor-specific Th9 cells. However, previous studies also showed that TNFR2 signaling enhances the differentiation, expansion and function of Treg cells [48][49][50], suggesting inhibitory effects on antitumor immunity. These observations imply that the TNFR2 signaling promotes the differentiation and functions of not only Th9 cells but also Treg cells, which suggests that TNF-α/TNFR2 signaling may exert both beneficial and detrimental effects on tumor immunotherapy. Detailed mechanisms underlying the different role of TNF-α/TNFR2 signaling in the induction of Th9 cells and Treg cells need to be defined. Further studies will be necessary to investigate strategies of converting the TNF-α/TNFR2 signaling to the induction of Th9 cells but not Treg cells in tumor immunotherapy.
In summary, our study demonstrates that TNF-α potently promotes the induction of Th9 cells in vitro. And, the addition of TNF-α during Th9 cell differentiation increases T cell survival and proliferation. The adoptive transfer of TNF-α-treated Th9 cells induces potent therapeutic antitumor effects in mouse models. TNF-α drives Th9 cell differentiation via TNFR2 but not TNFR1, and TNF-α/TNFR2 signaling activates STAT5 and NF-κB pathways, which are required for TNF-α-induced Th9 cell differentiation. Our results identified TNF-α as a new powerful inducer of Th9 cells and clarified the molecular mechanisms underlying TNF-α-induced Th9 cell differentiation.


Role of Pyroptosis in Cardiovascular Diseases and its Therapeutic Implications

Pyroptotic cell death or pyroptosis is characterized by caspase-1-dependent formation of plasma membrane pores, leading to the release of pro-inflammatory cytokines and cell lysis. Pyroptosis tightly controls the inflammatory responses and coordinates antimicrobial host defenses by releasing pro-inflammatory cellular contents, such as interleukin (IL)-1β and IL-18, and consequently expands or sustains inflammation. It is recognized as an important innate immune effector mechanism against intracellular pathogens. The induction of pyroptosis is closely associated with the activation of the NOD-like receptor 3 (NLRP3) inflammasome which has been linked to key cardiovascular risk factors including hyperlipidemia, diabetes, hypertension, obesity, and hyperhomocysteinemia. Emerging evidence has indicated pyroptosis as an important trigger and endogenous regulator of cardiovascular inflammation. Thus, pyroptosis may play an important role in the pathogenesis of cardiovascular diseases. Design of therapeutic strategies targeting the activation of NLRP3 inflammasome and pyroptosis holds promise for the treatment of cardiovascular diseases.

Introduction
Cell death is one of the ancient processes with fundamental biological importance in both normal physiology and pathological states. Recently, caspase-1 in both human and mouse, caspase-4/5 in human, and caspase-11 in mouse, are recognized to mediate a novel form of programmed cell death termed as pyroptosis. Pyroptosis is characterized by the formation of membrane pores, cell lysis and the release of pro-inflammatory cytokines and intracellular content. This event is predicted to be inherently inflammatory and coincides with the activation of inflammasome, a molecular platform resulting in caspase-1 activation and interleukin (IL)-1β and IL-18 secretion upon cellular infection or stress. The active caspase-1 triggered by inflammasome activation cleaves gasdermin D (GSDMD) to generate an N-terminal GSDMD fragment (GSDMD-NT), which further induces the formation of membrane pores and subsequent inflammatory responses. As an innate immune effector mechanism, pyroptosis has the ability to defense against infection. Emerging evidence has indicated that pyroptosis and related inflammasome activation play important roles in the progression of vascular inflammation and cardiovascular diseases. In this review, we summarize the molecular pathways of pyroptosis and the implications in cardiovascular research.

Pyroptosis: a form of pro-inflammatory programmed cell death.
In 1992, morphological features of pyroptosis were firstly observed in macrophages infected with the Gram-negative bacterial pathogen Shigella flexneri, which were mistakenly considered as morphological Ivyspring International Publisher changes of apoptosis at that time [1]. A similar phenotype was observed in Salmonella-infected macrophages in 1999 [2]. Further studies revealed that both selective caspase-1 inhibitor and caspase-1 knockdown efficiently blocked the shigella flexneriinduced cell death, whereas caspase-3 inhibitor did not, suggesting a novel form of cell death, depending on the activation of caspase-1, but not classical apoptotic caspase-3 [3][4][5]. Until 2001, pyroptotic cell death (pyroptosis) was firstly defined as a novel form of caspase-1-dependent programmed cell death by Cookson BT and Brennan MA [6]. In addition to microbial signaling, endogenous contents released by cells in overwhelming stress were also identified to induce macrophage pyroptosis. Pyroptosis has been found not only in monocyte/macrophages, but also in other cells including dendritic cells [7], hepatic cells [8], endothelial cells [9] and myocardial cells [10].
Macrophages undergoing pyroptosis show several morphological features of apoptosis, and also exhibit some characters which are similar to necrosis. Indeed, pyroptosis is characterized by rapid formation of membrane pores with a diameter of 10-15 nm [11]. Cellular ionic gradients are dissipated by these pores, which allow water influx, cell swelling and osmotic lysis with the release of intracellular pro-inflammatory contents including IL-1β, IL-18, high mobility group box-1 protein (HMGB-1) and heat shock protein (HSP). These are similar to oncosis but remarkably opposite to apoptosis which is characterized with the formation and noninflammatory phagocytic uptake of apoptotic bodies. Several features of pyroptotic cells seem to overlap with apoptotic cells. Both pyroptosis and apoptosis share the feature of chromatin condensation, but the nucleus remains intact and karyorrhexis does not occur in pyroptosis [12]. Another feature that is shared between pyroptosis and apoptosis is annexin V positive staining. During the early stages of apoptosis, phosphatidylserine is translocated to the outer leaflet, leading to positive cell surface staining with annexin V [13]. As cell membrane is ruptured during pyroptosis, annexin V is permited to enter the cell and stains the inner leaflet of the membrane. In contast, 7-aminoactinomycin or propidium iodide, as alive cell membrane impermeant dyes, can stain the nucleus of pyroptotic cell through the membrane pores, but not the early-stage apoptotic cell [14]. Thus, these dyes are applied to differentiate between apoptosis and pyroptosis. Moreover, pore formation causes cell swelling in pyroptotic cell, whereas cell shrinks in apoptosis [14]. Caspases play central role in initiating both apoptosis and pyroptosis. The characterized effectors of apoptosis are caspase-3, -5 and -7, whereas pyroptosis is induced by pro-inflammatory caspases (especially for caspase-1).

Mechanisms of Pyroptosis
The host can sense intracellular and extracellular 'danger' signals generated by invading microorganisms or by the host in response to tissue injury. The innate immune responses rely on specific hostreceptors which are termed as pattern-recognition receptors (PRRs) to detect pathogen-associated molecular patterns (PAMPs) and damage-associated molecular patterns (DAMPs) [15][16][17][18]. Based on their location, the PRRs can be differentiated into membrane-bound PRRs that recognize signals of infection in the cellular milieu or endosomes, such as Toll-like receptors (TLRs) and C-type lectin-like receptors, and cytoplasmic PRRs that recognize invasive pathogens, such as nucleotide-binding oligomerization domain (NOD)-like receptors (NLRs), retinoic acid-inducible gene I-like receptors, absent in melanoma 2 (AIM2)-like receptors (ALRs), cyclic GMP-AMP synthase, and stimulator of interferon gene [18][19][20][21]. Detection of PAMPs and DAMPs by NLRs and ALRs triggers the assembly of a caspase-1activating complex that was firstly termed inflammasome by Tschopp et al [22]. Active caspase-1 further processes pro-inflammatory cytokines (IL-1β and IL-18), and initiates pyroptosis by disruption of plasma membrane integrity [18,23]. In addition, mouse caspase-11 and human caspase-4 and -5 can directly bind to cytosolic lipopolysaccharide (LPS), leading to non-canonical inflammasome activation [18]. Non-canonical inflammasome can also induce pyroptosis, but cannot directly process pro-inflammatory IL-1β and IL-18. However, non-canonical inflammasome is capable of inducing the processing of pro-inflammatory cytokines indirectly by activating the canonical inflammasome through a not well-defined mechanism.

Pyroptotic pathway by caspase-1 dependent model
It is critical for pyroptosis to activate pro-inflammatory caspases. Thus, how inflammatory caspases are activated is a key question to elucidate the mechanism of pyroptosis. In 2002, inflammasome was firstly identified as the molecular platform triggering the activation of inflammatory caspases and consequently processing of pro-IL-1β and pro-IL-18 by Tschopp et al [22]. Inflammasome is multiprotein complex, composed of sensor, the adaptor protein ASC (apoptosis-associated speck-like protein containing a caspase recruitment domain (CARD)) and caspase-1. To date, several types of inflammasomes have been identified, including NLRP1, NLRP3, NLRC4, NLRP6, and AIM2 inflammasomes. Among these different types of inflammasomes, the NLRP3 inflammasome has been extensively studied in a variety of mammalian cells, and has been linked with various autoimmune and inflammatory diseases. Therefore, the role of NLRP3 inflammasome and related pyroptosis in cardiovascular disease will be discussed in this review.
The canonical inflammasome activation has been proposed to be a two-step process involving priming followed by assembly [30]. The activation of NLRP3 inflammasome requires 'priming' with TLRs agonists (such as LPS) to initiate signaling cascades (primarily nuclear factor-κB (NF-κB)-dependent pathway) that ultimately promote a transcriptional response to up-regulate pro-inflammatory mediators (such as pro-IL-1β) (Figure 1). After this priming phase, inflammasome assembles in response to a diverse range of PAMPs or DAMPs to form a large molecular platform triggering activation of inflammatory caspases and processing of pro-IL-1β.
In addition, three models about the assembly of NLRP3 inflammasome have been proposed (Figure 1). Extracellular ATP stimulates the purinergic receptor P2X, ligand-gated ion channel 7 (P2X7) on cell membrane [31], triggering K + efflux and inducing recruitment of pannexin 1 to form large membrane pores [32]. Therefore, extracellular agonists are able to penetrate the cell membrane and activate NLRP3 inflammasome. A second model was presented for particulate or crystalline agonists, such as cholesterol crystal, monosodium urate, silica and asbestos. Uptake of such substances leads to lysosomal damage, resulting in the leakage of lysosomal cathepsins B and L that are sensed by the NLRP3 inflammasomes [33]. Under the third model, agonists trigger the generation of reactive oxygen species (ROS), which is the common pathway engaging the NLRP3 inflammasome. Intriguingly, all three models are not functionally independent but rather interactive.
The agonist of NLRP3 inflammasome serves as potential inducer for cell pyroptosis. The detailed molecular basis for pyroptosis was unclear until very recently. Several studies simultaneously identified GSDMD as a critical executor of pyroptosis.
GSDMD belongs to a gasdermin family that shares the pore-forming domain [34]. Both caspase-1 and caspase-4/ 5/11 can cleave GSDMD to release GSDMD-NT that perforates the plasma membrane to induce cell swelling and The NLRP3 oligomerization and ASC recruitment trigger pro-caspase-1 autocleavage, leading to autocatalytic activation of caspase-1, which in turn converts inactive pro-IL-1β and pro-IL-18 into their bioactive and secreted forms (IL-1β and IL-18). The active caspase-1 also cleaves GSDMD to generate GSDMD-NT, which forms plasma membrane pores to induce pyroptosis. Three models about the assembly of NLRP3 inflammasome have been proposed: (1) The NLRP3 inflammasome is activated by extracellular ATP through one of the following mechanisms: the P2X7-activated pannexin-1 pore allows cytoplasmic entry of extracellular factors that are direct NLRP3 ligands, or NLRP3 senses either K + efflux or the loss of membrane integrity. (2) Crystalline or particulate agonists are phagocytosed, leading to the leakage of lysosomal cathepsins B and L, which are sensed by the NLRP3 inflammasome. (3) NLRP3 agonists including DAMPs and PAMPs trigger the production of ROS, which leads to the activation of the NLRP3 inflammasome. In general, the three models discussed above are not functionally independent but rather interactive. (4) Besides above three models, the activation of NLRP3 inflammasome also requires 'priming' with TLRs agonist (such as LPS). osmotic lysis. Other gasdermin family members also show pore-forming activity, but they are not substrates of the inflammatory caspases [28]. GSDMD is cleaved at the conserved residue D276, which separates GSDMD into a GSDMD-NT domain (p30) and a GSDMD C-terminal domain (p20) [11]. GSDMD-NT can combine with lipid in the plasma membrane and form large oligomeric pores, leading to the disruption of cell membrane integrity, the increase of intracellular osmotic pressure, and the release of inflammatory intracellular contents, such as HMGB-1 and HSP (Figure 1) [11,28,35,36]. This caspase-1dependent GSDMD cleavage is defined as canonical pyroptotic pathway [37].

Pyroptotic pathway by caspase-1 independent model
The other inflammatory caspases, mouse caspase-11 and human caspase-4 and -5, are required for non-canonical inflammasome activation, thus are major mediators of non-canonical pyroptosis ( Figure  2). In 2011, Kayagaki et al reported that gram-negative bacteria (Escherichia coli, Vibrio cholerae, and Citrobacter rodentium)-induced IL-1β maturation and secretion were blocked in caspase-11 -/macrophages rather than caspase-1 -/macrophages, whereas ATP and monosodium urate-induced canonical inflammasome activation was intact in caspase-11 -/macrophages, indicating caspase-11 but not caspase-1 was required for non-canonical inflammasome activation and related cell death [38]. Follow-up studies identified intracellular LPS as the trigger of TLR4-independent non-canonical inflammasome [39,40]. Recent study demonstrated that the high-affinity interaction between the lipid A component of LPS and the CARD domain of cytoplasmic caspase-11/4/5 led to direct binding of them, and resulted in inflammatory caspase oligomerization and activation [41,42]. Active caspase-4/5/11 ultimately cleaved GSDMD to induce cell pyroptosis similar to canonical pathway ( Figure  2) [41]. In addition, non-canonical inflammasome activation led to secondary activation of the canonical inflammasome and caspase-1, and subsequent maturation and secretion of proinflammatory cytokines [38], which may be an indirect consequence of GSDMD-NT [37]. Although a study reported that caspase-4 could cleave pro-IL-1β directly [43], the inability of caspase-4/5/11 to cleave the precursors of proinflammatory cytokines was widely accepted and considered as a major difference between canonical and non-canonical pyroptotic pathways.
In addition, gasdermin E (GSMDE) can shift caspase-3-mediated classical apoptosis induced by tumor necrosis factor alpha (TNFα) or chemotherapy drug to pyroptosis. GSDME is specifically cleaved by non-proinflammatory caspase-3, producing a GSDME N-terminal cleavage product that perforates membrane and thereby induces cell pyroptosis [44]. These findings reveal novel insights into mechanisms underlying pyroptosis.

Pyroptosis and Atherosclerosis
Atherosclerosis is complicated by a number of events: endothelial dysfunction; accumulation and oxidation of low-density lipoprotein (LDL); recruitment of monocytes and lymphocytes; migration and proliferation of smooth muscle cells; activation of proinflammatory cytokines; adherence of platelets. In these processes, inflammatory reaction was recognized in the atherosclerotic lesions from the early stage of fatty streak to culmination into an acute cardiovascular event caused by plaque erosion or rupture. However, how the inflammation contributes to atherosclerosis is still largely unclear.
More and more studies revealed that risk factors of atherosclerosis could activate NLRP3 inflammasomes in both endothelial cells and macrophages. In addition, NLRP3 inflammasomemediated pyroptosis has been observed in the atherosclerotic plaques and is positively correlated with the plaque rupture and vascular inflammation, indicating that NLRP3 inflammasome and related pyroptosis play an important role in the progression of atherosclerosis.

Pyroptosis and Dyslipidemia
Cholesterol crystals have been identified as a potential biomarker for atherosclerosis and a major risk factor for plaque vulnerability [45]. Duewell et al firstly reported that cholesterol crystals activated NLRP3 inflammasomes by inducing lysosomal damage, and led to the release of IL-1β [46]. Combined confocal-reflection microscopy revealed that a large load of cholesterol crystals were deposited in the aortic wall and coincided with increased recruitment of macrophages in high cholesterol diet-fed low-density lipoprotein receptor-null (LDLR -/-) mice. In contrast, a significant decrease of atherosclerotic lesion size was also observed at the aortic sinus of high fat diet-fed LDLR -/mice reconstituted with NLRP3-, ASC-or IL-1α/β-knockout bone marrow cells [46]. Consistent with these in vivo findings, cholesterol crystals activated NLRP3 inflammasomes in human macrophages, which was suppressed by silencing of NLRP3 [47]. These studies confirmed that NLRP3 inflammasomes were activated by cholesterol crystals and required for atherogenesis. Given that caspase-1 activation induced by NLRP3 inflammasome is a major mediator for pyroptosis, these work further suggested that pyroptotic macrophages might take part in the progression of atheroslerosis.
In addition, oxidized LDL (ox-LDL), a major risk factor for atherosclerosis, triggered the assembly of NLRP3 inflammasome, and subsquently activated caspase-1 and processed IL-1β in macrophages [46,48]. Immunohistochemistry results showed that caspase-1 was activated in advanced atherosclerotic lesion and co-located with macrophages and TUNEL positive area, suggesting that pyroptosis was implicated in advanced plaque. Another in vitro experiment further confirmed that ox-LDL induced pyroptosis in macrophages [49], which was inhibited by the depletion of mitochondrial DNA [50].
In addition to ox-LDL, NLRP3 inflammasome was also activated by high level of triglyceride, a marker for several types of atherogenic lipoproteins [51,52]. Yin et al also reported that early hyperlipidemia induced caspase-1 activation along with enhanced expression of intercellular adhension molecule, vascular cell adhension molecule and E-selectin in the aorta, and promoted monocyte recruitment in ApoE -/mice, whereas the knockout of caspase-1 obviously suppressed these effects and decreased the area of atheroclerotic plaque [53]. They further performed in vitro expriments and found that ox-LDL induced cell pyroptosis through ROSdependent pathway in endothelial cells. The pyroptosis of vascular endothilal cells was firstly linked to atherosclerotic progression in this work.
As same as ox-LDL, palmitic acid, a saturated fatty acid, has been reported to induce pyroptosis in ROS-dependent manner in human umbilical venin endothelial cells (HUVECs), while pretreatment with dihydromyricetin inhibited palmitic acid-induced pyroptosis by activating Nrf2-antioxidant response element signaling pathway [54]. Another study also pointed out that cholesterol crystal obviously induced NLRP3 inflammasome-dependent pyroptosis in primary coronary endothelial cells [55], indicating that inflammasome-mediated pyroptosis might play an important role in cholesterol crystal-induced atherosclerosis. As endothelial cells cover the inner surface of vessel wall and are directly exposed to the atherosclerosis-related danger signals in the circulation, endothelial dysfunction is an initial event responsible for monocyte recruitment in atherogenesis [56]. It is very likely that cell pyroptosis can switch endothelial cells to the proinflammatory state which subsequently contributes to atherosclerosis.

Pyroptosis and Hyperhomocysteinemia
Hyperhomocysteinemia (HHcy) was defined as elevated plasma level of homocysteine (Hcy) (>15 μmol/L). HHcy is an important and independent risk factor involved in several disorders including atherosclerosis [57]. Earlier reports have shown that plasma levels of proinflammatory cytokines including IL-6 and TNFα were increased in HHcy individuals, which played a crucial role in endothelial damage and atherosclerosis [58]. Since Hcy can be autooxidized with anthor Hcy molecule to generate ROS and the disulfide [59], Hcy has the potential to activate NLRP3 inflammasome. Recently, a study from our laboratory demonstrated that HHcy-induced activation of NLRP3 inflammasome in macrophages contributed to vascular inflammation and atherosclerosis [60]. ROS scavenger N-acetyl-L-cysteine suppressed the activation of NLRP3 inflammasomes and alleviated atherosclerosis induced by HHcy [60]. In fact, there was an evidence showing that the conbination of Hcy and LPS synergistically induced endothelial cell pyroptosis via caspase-1-dependent inflammasome activation [61]. Further study found that intracellular ROS levels determined Hcy/LPS-induced death destiny in endothelial cells: relatively high ROS levels directly induced apoptosis, while moderate ROS generation preferred to trigger pyroptosis [61]. These phenomena indicate that HHcy-aggravated atheroclerosis may be closely associated with not only apoptosis of endothelial cells but also the pyroptosis-mediated inflammation.
Evidence shows that certain heavy metals may regulate NLRP3 inflammasome activation. Cadmium (Cd), a toxic and nonessential heavy metal, has been linked to cardiovascular diseases including atherosclerosis [62]. Chen et al recently found that Cd induced NLRP3 inflammasome activation and subsequent pyroptosis via mitochondrial ROSdependent pathway in vascular endothelial cells [9]. Whether NLRP3 inflammasome and related pyroptosis contribute to Cd-induced cardiovascular diseases needs further exploration in animal models.

Pyroptosis in Obesity and Diabetes: Reminding of NLRP3 Inflammasome
Obesity and related metabolic syndrome pose huge threats to human health. Active inflammation in obese adipose tissue is increasingly recognized as a key event underlying metabolic diseases. Obesity provides a proinflammatory condition which contributes to the increase of circulating proinflammatory cytokines [63]. Studies showed that the expression of caspase-1, ASC and NLRP3 was upregulated in high fat diet-induced obesity mice, while the knockout of NLRP3 or caspase-1 suppressed obesity-induced inflammasome activation in fat depots and liver [64]. Interestingly, obese db/db mice exhibited ultrastructural abnormalities similar to pyroptotic cells with ROS increment and caspase-1 activation in the subcutaneous and visceral hypertrophic adipocytes [65]. In addition, Konene et al found that the expression of caspase-1 in the abdominal adipose tissue was markedly higher than in the subcutaneous fat, indicating that pyroptotic cell death was probably presented in the abdominal adipose tissue [66].
A study showed that peripheral blood-derived macrophages from drug-naïve patients with type 2 diabetes displayed enhanced expression of NLRP3 and ASC together with caspase-1 activation and IL-1β maturation [67]. It was reported that high glucose stimulated IL-1β secretion in pancreatic β cells [68]. Further study demostrated that high glucose induced NLRP3 inflammasomes activation and IL-1β secretion, leading to pancreatic β cell death [68]. In normal condition, the thioredoxin (TRX)-interacting protein (TXNIP) was bound to TRX, however, TXNIP was dissociated from TRX to interact with and activate NLRP3 inflammasome in various types of cells when incubated with high glucose [68]. In contrast, γ-tocotrienol, known to exert potent anti-inflammatory function in various cell types [69], was shown to inhibit NLRP3 inflammasome activation and the recruitment of macrophages to adipose tissues, thus ameliorated insulin resistance [70]. In fact, γ-tocotrienol not only inhibited the tumor necrosis factor receptor-associated factor 6/NF-κB pathway, but also activated AMP-activated protein kinase/autophagy axis, leading to reduced cleavage of caspase-1. These results suggested that NLRP3 inflammasome and related pyroptosis might be potentially promising targets for new drug research in diabetes.

Pyroptosis in Hypertension: Reminding of NLRP3 Inflammasome
Hypertension is a well-known risk factor for atherogenesis-related cardiovascular and cerebrovascular diseases. Recently, the role of inflammation in the pathologenesis of hypertension has attracted great interests. Although there was no direct proof that cell pyroptosis contributed to pathological process of hypertension, studies showed that circulating levels of IL-1β and IL-18 were elevated in hypertensive mice and rats [71,72]. Dalekos et al found that serum level of IL-1β in patients with essential hypertension was higher than in normotensive controls [73]. Given that IL-1β maturation is dependent on caspase-1, there is a possibility that hypertension may be assiociated with the activation of caspase-1. Another study showed that mRNA expression of pro-caspase-1 in the aorta and renal artery was increased in spontaneously hypertensive rats, which indirectly supported this possibility [74]. Moreover, high salt increased the expression of NLRP3, ASC and pro-caspase-1, the activation of caspase-1, and the maturation of IL-1β in mouse kidney [71]. Interestingly, NF-κB knockdown was associated with reduced expression of NLRP3 inflammasome components in hypothalamic paraventricular nucleus, and thus delayed the progression of hypertension in Dahl salt-sensitive hypertensive rats [75]. Therefore, targeting NLRP3 inflammasome may be a therapeutic strategy to treat hypertension. Based on this hypothesis, Tang et al found that ellagic acid, a promising treatment candidate for pulmonary artery hypertension (PAH), inhibited NLRP3 inflammasomes activation primarily by exerting its antioxidant effect [76].

Pyroptosis and Myocardial Ischemia/Reperfusion Injury
Therapeutic strategies aimed at rescuing ischemic myocardium have been studied extensively. Reperfusion remains the definitive treatment for acute myocardial infarction (MI). However, restoring blood flow carries the potential to exacerbate lethal tissue injury, a process termed as "ischemia/reperfusion injury". The major contributory factors of reperfusion injury include oxidative stress [34], calcium overload [41], opening of mitochondrial permeability transition pores [42], and in particular inflammation [18]. Myocardial ischemia/reperfusion injury (MIRI) may cause MI and cardiac arrhythmias as well as cardiac dysfunction [77]. MI is defined as myocardial cell death due to prolonged ischemia and usually caused by the sudden thrombotic occlusion of coronary artery [78]. Recent studies found that NLRP3 inflammasome and related pyroptosis were strongly associated with pathological process of MIRI and MI. As previously mentioned, NLRP3 inflammasome activation and pyroptosis contributed to plaque rupture, and the release of IL-1β was an early and prominent event of inflammatory responses in MI [49]. Investigators have also found that inflammatory responses and subsequent injuries, including myocardial dysfunction and fibrosis, were significantly alleviated in ASC or caspase-1 deficient mice [79]. Further studies showed that NLRP3 inflammasome was activated in the ischemic heart as evidenced by increased NLRP3 expression and IL-1β and IL-18 production with caspase-1 activation, whereas NLRP3 siRNA, treated via intramyocardial injection, inhibited these effects and decreased TUNEL positive areas. Another study showed that pyroptosis was induced by ROS-dependent NLRP3 inflammasome activation, and inhibition of NLRP3 inflammasome activation alleviated MIRI in diabetic rats, suggesting that NLRP3 inflammasome-mediated pyroptosis played a critical role in inflammatory responses and MIRI in diabetic rats [80]. In addition, silence of cryopyrin (also known as NLRP3) and P2X7 prevented the formation of NLRP3 inflammasome, limited myocardial infarct size and ameliorated cardiac remodeling after acute MI [81]. These results indicated that NLRP3 inflammasome activation and related pyroptosis are potential novel therapeutic targets for MIRI.

Pyroptosis and Diabetic Cardiomyopathy
Diabetic cardiomyopathy (DCM) is a diabetesinduced cardiomyopathy that occurs in diabetic subjects without coronary artery disease and hypertension [82,83]. DCM is characterized by functional and structural abnormalities, including myocardial dysfunction [84], myocardial cell death [85], myocardial fibroblast activation [86], and metabolic deregulation [87]. Among them, myocardial cell death is a key factor in the progression of DCM, leading to a loss of contractile units, conduction disturbances, compensatory hypertrophy of myo-cardial cells, and myocardial fibrosis [88]. Apoptosis has been widely recognized in the pathogenesis of DCM. In addition, recent evidence showed pyroptosis could occur in DCM heart. Electron microscopy studies demonstrated that most of the dying cells had characteristics of swollen fibril and mitochondria similar to the features of pyroptosis in the myocardium of diabetic mice and rats [89][90][91]. Intriguingly, Luo et al reported that NLRP3 inflammasome-caspase-1mediated pyroptosis was presented in myocardium of diabetic rats, while silencing of NLRP3 ameliorated cardiac inflammation and pyroptosis, and improved myocardial function [92].
Numerous studies have shown that microRNAs (miRNAs) are involved in the regulation of cardiomyocyte pyroptosis in DCM. For example, Li et al demonstrated that miR-30d was upregulated in high glucose-treated cardiomyocytes and streptozotocin (STZ)-induced diabetic rat hearts, which regulated cardiomyocyte pyroptosis in DCM [10]. In an effort to explore the pro-pyroptotic mechanisms of miR-30d, they found that miR-30d directly repressed the expression of Forkhead box O3 and its downstream factor, apoptosis repressor with caspase recruitment domain [10]. Moreover, miR-9 inhibited hyperglycemia-induced pyroptosis in human ventricular cardiomyocytes [93], suggesting that miR-9 may have a protective effect on DCM.

Pyroptosis in Viral Myocarditis: Reminding of NLRP3 Inflammasome
About 20 common types of viruses have been related to myocarditis in human [94]. The dominant etiological agent of viral myocarditis is coxsackievirus B3 (CVB3) [95]. Wang et al firstly reported that the activation of NLRP3 inflammasome was involved in CVB3-induced myocarditis [96]. Both in vivo and in vitro results showed that CVB3 infection activated NLRP3 inflammasome. Conversely, once inflammasome activation was inhibited, CVB3-induced myocarditis manifested by weight loss, increased serological indexes of creatine kinase and CK-MB activities, was obviously suppressed. Caspase-1 inhibitor also improved left ventricular ejection fraction and left ventricular fractional shortening in CVB3-induced myocarditis [96]. Further study showed that ROS generation and K + efflux were involved in the activation of NLRP3 inflammasome caused by CVB3 infection [96]. Recent study demonstrated that NOD2 mRNA expression was higher in endomyocardial biopsy tissue from CVB3-positive patients as compared with myocarditis patients without persistent CVB3 infection [97]. Interestingly, mesenchymal stromal cells suppressed NOD2 expression and NLRP3 inflammasome activation, which was associated with improved expression of markers for cardiac contractility and fibrosis in the heart of CVB3-infected mice, suggesting that NOD2 might coordinate NLRP3 inflammasome activation in CVB3-infected myocardium [98]. In addition, Wang et al. found that CVB3-induced myocarditis was aggravated by cathepsin B-induced activation of NLRP3 inflammasome and pyroptosis [99]. It is still not clear whether other types of viruses regulate NLRP3 inflammasome activation in viral myocarditis.

Pyroptosis and Heart Failure
Heart failure (HF) means that the heart is unable to pump sufficiently to sustain blood flow to meet the needs of the body [100]. HF is the end stage of many cardiovascular diseases. Common causes of HF include myocarditis, MI, cardiomyopathy, hypertension, atrial fibrillation (AF), valvular heart disease, alcohol abuse, and infection. Recently, elevated mRNA of NLRP3 and enhanced cleavage of caspase-1 were observed along with cardiac hypertrophy and ventricular dilatation in calcineurin transgene (CNTg) mice, an established mouse model for chronic HF [101]. As expected, they also discovered that genetic ablation of NLRP3 or administration of IL-1 receptor antagonist attenuated cardiac inflammation and systolic dysfunction [101]. Similarly, the expression of NLRP3 inflammasome components was significantly increased, and associated with elevated inflammatory mediators and profibrotic factors and impaired cardiac function in a mouse model of transverse aortic constriction (TAC)-induced ventricular hypertrophy [102] [103]. Interestingly, the inhibition of NLRP3 by triptolide [103] or pirfenidone [102] attenuated myocardial remodeling and improved cardiac function in TAC mice. In addition, the activation of NLRP3 inflammasome was observed in LPSstimulated cardiac fibroblasts [104] and myofibroblasts [105], suggesting that NLRP3 inflammasome and related pyroptosis may contribute to myocardial dysfunction in sepsis.
AF, a common kind of arrhythmia, is associated with an increased risk of worsening HF [106,107]. A strong inflammatory response is frequently associated with AF progression [108,109]. Increased serum levels of IL-1β and IL-18 were positively correlated with AF progression from paroxysmal AF (pAF) to long-lasting persistent AF (perAF) [109,110]. Recently, Yao et al firstly showed that NLRP3 inflammasome was activated in the atrial cardiomyocytes from patients with pAF and perAF [111]. Cardiomyocytespecific activation of NLRP3 induced abnormal Ca 2+ release from sarcoplasmic reticulum and promoted inducible AF, which was attenuated by MCC950, a specific NLRP3 inflammasome inhibitor, and genetic silencing of NLRP3 [111]. Genetic silencing of NLRP3 also prevented AF development in cAMP-responsive element modulator (CREM)-transgenic mice, a spontaneous AF mouse model [111].

Targeting of NLRP3 inflammasome or cell pyroptosis as a therapeutic strategy
Based on experimental data, inhibiting NLRP3 inflammasome or cell pyroptosis may supply a new therapeutic strategy for cardiovascular disease. The complicated signaling of NLRP3 inflammasome and related pyroptosis offers diverse targets for inhibiting their activation, including suppression of upstream signals of NLRP3 inflammasome, prevention of NLRP3 inflammasome assembly, inhibition of caspase-1 activation and GSDMD cleavage, and neutralizing antibodies targeting inflammasomederived inflammatory cytokines. Preventing the assembly of NLRP3 inflammasome could be established by inhibition of K + efflux with glyburide [112] and β-hydroxybutyrate [113], blockage of P2X7 signaling with human cathelicidin LL-37 [114], as well as the application of ROS scavenger [115]. It was reported that MCC950 disrupted the interaction between NLRP3 and ASC, and showed strong inhibitory effect on NLRP3 inflammasome and beneficial effect on inflammatory diseases [116]. In addition, parthenolide, BAY 11-7082, INF39, and 3,4methylenedioxy-β-nitrostyrene (MNS) were reported to directly inhibit the ATPase activity of NLRP3, but these inhibitors have unspecific effects limiting their use [117][118][119]. Recently, Jiang et al. showed that CY-09 specifically blocked NLRP3 inflammasome activation by binding to the ATP-binding motif of NLRP3 NACHT domain and thus inhibiting the ATPase activity of NLRP3. Treatment with CY-09 showed therapeutic effects on cryopyrin-associated autoinflammatory syndrome and type 2 diabetes in mouse models [120]. The suppression of NLRP3 inflammasome components could be achieved by genetic silencing of NLRP3, ASC or caspase-1 using specific shRNAs [111] or clustered regularly interspaced short palindromic repeat (CRISPR)-associated system 9 (Cas9) [121]. Xu et al screened an optimized cationic lipid-assisted nanoparticle to deliver Cas9 mRNA and guide RNA targeting NLRP3 into macrophages, which improved insulin sensitivity and reduced adipose inflammation in high fat diet-induced diabetic mice [122]. Moreover, caspase-1 inhibitors are currently under development for inflammatory diseases [123,124]. However, further studies are needed to determine whether these inhibitors are appropriate for cardiovascular diseases.
To date, most drug candidates targeting NLRP3 inflammasome have been studied only in animal models, and need to be further translated into clinical practices. The current available clinical treatments for NLRP3 inflammasome-related diseases are the agents that block IL-1β, including the recombinant IL-1 receptor antagonist anakinra, the IL-1β-neutralizing antibody canakinumab, and the soluble decoy IL-1β receptor rilonacept [125]. Interestingly, the recently completed large-scale clinical trial CANTOS showed that canakinumab targeting the IL-1β innate immunity pathway significantly decreased the rate of recurrent cardiovascular events [126], and reduced incident lung cancer and lung cancer mortality in patients with atherosclerosis [127]. However, IL-1β is not the only biological production of NLRP3 inflammasome and pyroptosis, other inflammatory cytokines, such as IL-18 and HMGB-1, also involved in the pathogenesis of cardiovascular diseases. Therefore, inhibition of IL-1β signaling may have limitations in clinical treatments.

Conclusion
Emerging evidence has demonstrated that NLRP3 inflammasome activation is a critical regulator for pyroptosis involved in the progression of various cardiovascular diseases (Figure 3). Pyroptosis has been observed in different types of cells and tissues in in vivo and in vitro models of cardiovascular diseases ( Table 1). Despite of growing studies focused on the role of pyroptosis in cardiovascular diseases, the detailed molecular mechanisms remain elusive. Although to what extent does NLRP3 inflammasome activation lead to pyroptosis remains unclear, NLRP3 inflammasome activation indeed mediates pyroptosis in most situations. Therefore, blocking the inflammasome pathway has been predicated to have clinical benefits in delaying the progression of cardiovascular diseases. Several types of NLRP3 inflammasome inhibitors have been developed and validated in cell culture studies and animal models of inflammationrelated diseases. However, prospective clinical trials are needed for their potential translation into clinical practice. Also other potential candidates are required for further development, such as autophagy-inducer, antioxidant and miRNA agents. Future studies are needed to answer many unresolved questions. For example, can drugs or antibodies targeting GSDMD pyroptotic pathway enhance the therapeutic potential of single NLRP3 inflammasome inhibitor on cardiovascular diseases? Further studies exploring novel signaling pathways of pyroptosis will be important for the development of new therapeutic drugs.   [104] * indicated this study with cells or tissue from human.