1. Biochemistry and Chemical Biology
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YcgC represents a new protein deacetylase family in prokaryotes

  1. Shun Tu
  2. Shu-Juan Guo
  3. Chien-Sheng Chen
  4. Cheng-Xi Liu
  5. He-Wei Jiang
  6. Feng Ge
  7. Jiao-Yu Deng
  8. Yi-Ming Zhou
  9. Daniel M Czajkowsky
  10. Yang Li
  11. Bang-Ruo Qi
  12. Young-Hoon Ahn
  13. Philip A Cole  Is a corresponding author
  14. Heng Zhu  Is a corresponding author
  15. Sheng-Ce Tao  Is a corresponding author
  1. Shanghai Jiao Tong University, China
  2. State Key Laboratory of Oncogenes and Related Genes, China
  3. National Central University, Taiwan
  4. Chinese Academy of Sciences, China
  5. Wuhan Institute of Virology, Chinese Academy of Sciences, China
  6. National Engineering Research Center for Beijing Biochip Technology, China
  7. School of Biomedical Engineering, Shanghai Jiao Tong University, China
  8. Johns Hopkins University School of Medicine, United States
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Cite this article as: eLife 2015;4:e05322 doi: 10.7554/eLife.05322

Abstract

Reversible lysine acetylation is one of the most important protein posttranslational modifications that plays essential roles in both prokaryotes and eukaryotes. However, only a few lysine deacetylases (KDACs) have been identified in prokaryotes, perhaps in part due to their limited sequence homology. Herein, we developed a ‘clip-chip’ strategy to enable unbiased, activity-based discovery of novel KDACs in the Escherichia coli proteome. In-depth biochemical characterization confirmed that YcgC is a serine hydrolase involving Ser200 as the catalytic nucleophile for lysine deacetylation and does not use NAD+ or Zn2+ like other established KDACs. Further, in vivo characterization demonstrated that YcgC regulates transcription by catalyzing deacetylation of Lys52 and Lys62 of a transcriptional repressor RutR. Importantly, YcgC targets a distinct set of substrates from the only known E. coli KDAC CobB. Analysis of YcgC’s bacterial homologs confirmed that they also exhibit KDAC activity. YcgC thus represents a novel family of prokaryotic KDACs.

https://doi.org/10.7554/eLife.05322.001

eLife digest

After proteins have been made, they can be modified in several ways. For example, chemical tags called acetyl groups may be added to (and later removed from) the protein to regulate cell activities such as aging and metabolism. Enzymes are proteins that help catalyze the reactions that add or remove the acetyl tags on certain “substrate” proteins. In the bacteria species Escherichia coli, many enzymes that help to add acetyl groups to proteins have been discovered. However, only a single E. coli “deacetylase” enzyme that removes the acetyl group has been identified.

Now, Tu, Guo, Chen et al. have devised a technique to identify new deacetylases, called the “clip-chip” approach. In this method, thousands of proteins that are potential deacetylases are arrayed on a glass slide, and substrate proteins are immobilized on another slide. The two slides are then clipped together face-to-face, allowing the potential enzymes to transfer to the substrate slide and interact with the substrates.

Using this approach, Tu, Guo, Chen et al. identified a protein called YcgC as a deacetylase in bacteria. Further characterization experiments revealed that YcgC works in a different way to other known deacetylases, and that it targets different substrates to the previously known E. coli deacetylase.

Tu, Guo, Chen et al. found that the equivalents of YcgC in other bacteria species are also deacetylases; these enzymes therefore represent a new deacetylase family. In the future, the clip-chip approach could be used to discover new members of other enzyme families.

https://doi.org/10.7554/eLife.05322.002

Introduction

Protein (de)acetylation plays critical roles in many key biological processes, for example, transcriptional regulation, aging, and metabolism (Cohen et al., 2004; Grunstein, 1997; Lin et al., 2009; Lu et al., 2011). Recent mass spectrometry (MS) efforts have revealed that many proteins are acetylated in Escherichia coli, although only a single E. coli lysine deacetylase (KDAC), CobB, has been identified so far (Choudhary et al., 2009; Henriksen et al., 2012; Zhang et al., 2013a). The fact that induction of CobB only had a limited impact on reducing the global protein acetylation level suggests that additional KDACs may exist. However, homolog searching has failed to reveal any additional KDACs in E. coli, presumably because these enzymes emerged via convergent evolution. In contrast to bioinformatics methods, biochemical approaches have proven effective for identifying new enzymes resulting from convergent evolution (Tsukada et al., 2006; Yamane et al., 2006), though their laborious, time-consuming nature has limited their applications to high-throughput, proteome-wide screens. Herein, we established a ‘clip-chip’ approach to enable a proteome-wide, activity-based search for novel KDACs in E. coli.

Results

The clip-chip strategy

The principle behind the clip-chip approach is the delivery of thousands of purified proteins spotted on a glass slide (e.g., a proteome microarray) to a substrate of interest immobilized on another slide (i.e., the substrate slide) such that thousands of desired biochemical reactions can be carried out in parallel, in order to identify new enzymes of interest (Figure 1a and Figure 1—figure supplement 1). The substrate slide is created by immobilizing a substrate of interest onto a nitrocellulose-coated slide. After thousands of purified proteins are spotted on a plain glass slide, it is then ‘clipped’ onto the substrate slide in a face-to-face manner, resulting in the delivery of the proteins onto the substrate slide. Owing to the highly porous nature of nitrocellulose and the tiny volume of the protein droplets (0.3–0.5 nL), the delivered protein droplets are immediately absorbed and kept locally in the nitrocellulose, preventing cross-contamination. To determine which transferred proteins possess the enzymatic activity in question, the ‘clipped’ substrate slide is then incubated with an appropriate reaction buffer, followed by signal detection.

Figure 1 with 1 supplement see all
Screening the Escherichia coli proteome to discover new KDACs using the ‘clip-chip’ strategy.

(a) Schematic of the ‘clip-chip’ strategy. (b,c) Identification of YcgC as a potential protein deacetylase. E. coli proteome chips were clipped onto three substrate slides separately coated with acetylated RutR, NhoA, and YceC. After incubation in a protein deacetylase buffer, the reactions were terminated by adding wash buffers, followed by a signal detection step with a pan α-AcK antibody coupled with a Cy3-labeled secondary antibody as detection reagent to visualize the loss of signals (e.g., black holes in (b,c). To determine the identity of proteins that generated the holes, the substrate slide was subsequently probed with an α-6xHis antibody followed by a Cy5-conjugated secondary antibody. (d) Using acetylated RutR proteins purified from E. coli, of the four candidates tested, YcgC showed robust deacetylation activity in vitro. Equal amounts of RutR proteins were used in each reaction and loss of acetylation was detected with the pan α-AcK antibody.

https://doi.org/10.7554/eLife.05322.003

Screen new KDAC using the E. coli proteome microarray

To screen for new KDAC candidates in the E. coli proteome, we prepared separate substrate slides for three E. coli proteins, namely NhoA, RutR, and YceC, which were chosen because they have a rather high endogenous acetylation level and because CobB exhibits only modest ability to deacetylate them (Zhang et al., 2013b). After 4256 individually purified E. coli proteins (Chen et al., 2008) were spotted on plain glass slides, they were clipped separately onto the three substrate slides, followed by incubation with a standard deacetylase reaction buffer containing NAD+. The reactions were terminated by adding wash buffers, followed by a signal detection step with a pan α-acetyllysine (α-AcK) antibody coupled with Cy3-labeled secondary antibodies as detection reagents. Proteins that efficiently deacetylated the substrates could be readily identified as they left behind pairs of black holes in fluorescence images of the substrate slides (Figure 1b,c). To help determine the identity of the proteins with potential KDAC activity, we subsequently probed the clipped substrate slides with an α-6xHis antibody to visualize the E. coli proteins delivered onto the substrate slides. As a negative control, substrate slides were also processed in parallel without the clipping step. We identified four candidates that showed significant deacetylation activities against at least one of the three substrates tested.

To validate the KDAC activity observed above, we purified the four candidate proteins and performed solution phase deacetylation reactions against RutR. CobB was also included for comparison. By evaluating the decrease in acetylation signals using an immunoblot assay with α-AcK, we confirmed that one of the candidates, YcgC, could readily deacetylate RutR in vitro, and that CobB also deacetylated RutR. YjgD did not show any detectable deacetylation activity against RutR, while Gnd and YhbL showed slight activity (Figure 1d). As YcgC also showed KDAC activity against NhoA and YceC (data not shown), we then focused on characterizing the function of YcgC. YcgC is previously known as DhaM, a subunit of dihydroacetone kinase complex and a nonessential gene in E. coli. Because the endogenous level of YcgC is very low, YcgC was overexpressed on the wild-type background in the subsequent experiments.

In vitro characterization of YcgC’s KDAC activity

As the M subunit of the dihydroxyacetone kinase complex, the possibility that YcgC has intrinsic enzymatic activity has not been reported previously (Molin et al., 2003). Therefore, we first determined whether YcgC also requires NAD+ and/or Zn2+ to deacetylate RutR, as class III deacetylases require NAD+ as a cofactor and other classes are dependent on Zn2+ (Thiagalingam et al., 2003). We chose RutR as the substrate for YcgC, because endogenous RutR proteins are highly acetylated and because it is known to regulate genes directly or indirectly involved in the complex pathways of pyrimidine and purine metabolism (Shimada et al., 2007; Shimada et al., 2008). To reduce possible contamination from other proteins, cofactors, or metal ions as much as possible, both 6xHis-tagged YcgC and RutR proteins were affinity purified from E. coli under stringent wash conditions, followed by overnight dialysis. Immunoblotting of the deacetylation reactions clearly showed that YcgC could deacetylate RutR effectively, but this activity did not appear to be dependent on NAD+ or Zn2+ (Figure 2a, Figure 2—figure supplement 1). High-performance liquid chromatography analysis and inductively coupled plasma-mass spectrometry (ICP-MS), respectively, confirmed that there was no detectable NAD+ or Zn2+ in the reaction (data not shown). Of note, Coomassie staining of the decaetylated RutR protein product band appeared at a slightly lower molecular weight than acetylated RutR and this is explored below.

Figure 2 with 4 supplements see all
In vitro and in vivo characterization of YcgC’s KDAC activity.

(a) In vitro assays of the KDAC activity of YcgC on RutR demonstrated that its KDAC activity does not require either NAD+ or Zn2+ as cofactors. Incubation with YcgC almost completely abolished the slower migrating acetylated RutR bands (upper panel) as evidenced by immonublotting (lower panel). (b,c) LC-MS/MS analysis to determine the residues of RutR deacetylated by YcgC. RutR was treated with YcgC first and the untreated RutR used as the control. Both these two samples were resolved on a SDS-PAGE gel side by side. The upper band represents the Kac-containing starting materials and the lower band represents the K-containing product, which were then recovered from the gel and subjected for MS/MS analysis (inserts). Lys52 was identified as an acetylated site in RutR protein (b). After incubating with YcgC, acetylation on K52 was no longer detectable (c). (d) RutR is deacetylated by YcgC in Escherichia coli. A 3xFLAG tag was chromosomally inserted at the 3′-end of rutR coding sequence. Acetylation of 3xFLAG-tagged RutR was monitored upon induction of YcgC. While the protein level of RutR was unchanged (middle panel), its acetylation level was dramatically reduced as a function of YcgG induction (upper panel). YcgC’s expression was monitored using a custom-made antibody (lower panel). (e) Mutagenesis of RutR confirmed that K52 and K62 are acetylated in vivo. Two single mutants K52Q and K62Q and one double mutant K52/62Q were constructed. These mutants along with WT RutR were produced and purified in E. coli. Equal amounts of purified proteins were Western blotted with the α-AcK antibody, quantitation of acetylation level of these samples were performed. KDAC: Lysine deacetylase; LC-MS/MS: Liquid chromatography–mass spectrometry; IP: Immunoprecipitation.

https://doi.org/10.7554/eLife.05322.005

Next, we employed liquid chromatography–mass spectrometry (LC-MS/MS) to determine which acetylated lysine residues of RutR were deacetylated by YcgC. We found that Lys52 and Lys62, present in the peptide sequences LEQIAELAGVSK52TNLLYYFPSK and TNLLYYFPSK62EALYIAVLR, respectively, were acetylated in RutR expressed in wild-type (WT) cells (Figure 2b and Figure 2—figure supplement 2a). However, after RutR was incubated with YcgC, acetylation of K52 or K62 was no longer detectable (Figure 2c and Figure 2—figure supplement 2b). Therefore, YcgC effectively deacetylates RutR on residues K52 and K62 in vitro.

In vivo validation of YcgC’s KDAC activity

To determine whether YcgC could deacetylate RutR in cells, we performed immunoprecipitation (IP)-coupled immunoblotting to measure changes in the acetylation levels of RutR over the period of YcgC induction. To enable immunoprecipitation of endogenous RutR proteins, a 3xFLAG tag was chromosomally inserted into the 3′-end of the rutR coding sequence. An isopropyl-beta-D-thiogalactopyranoside (IPTG)-inducible ycgC construct was then transformed into the rutR:3xFLAG cells and induced for YcgC expression for up to 4 hr. Using IP-coupled immunoblotting analysis, we observed that acetylation levels of RutR proteins were significantly reduced in a YcgC expression level-dependent manner as detected by a custom-made α-YcgC monoclonal antibody (Figure 2d and Figure 2—figure supplement 3). In contrast, the total amount of RutR was not affected by YcgC induction (Figure 2d). These results confirmed that YcgC effectively deacetylates RutR in vivo without affecting its stability.

To examine whether K52 and K62 acetylation sites of RutR were deacetylated by YcgC in vivo, we created two single (K52Q; K62Q)- and one double (K52/62Q)-mutants of RutR. After transformation of these mutants into E. coli, subsequent IP-coupled immunoblotting demonstrated that, compared with WT RutR, mutation of either K52 or K62 resulted in a substantial loss of acetylation signals in RutR, with the K52/62Q double mutant showing the lowest acetylation signals (Figure 2e). RutR K-to-R mutants (i.e., K52R, K62R, and K52/62R) were also created and tested, and similar results were observed to those with the K-to-Q mutants (Figure 2—figure supplement 4). These results suggest that both K52 and K62 are major acetylation sites in RutR, and can be effectively deacetylated by YcgC in E. coli.

YcgC belongs to the serine hydrolase family

Because KDACs catalyze hydrolytic reactions on lysine residues, we tested a variety of hydrolase inhibitors against YcgC in in vitro deacetylation reactions as described above. We found that Halt Protease Inhibitor Cocktail (with or without ethylene glycol tetraacetic acid [EGTA]; Thermo Scientific, Rockford, IL) and Complete Protease Tablet (Roche, Mannheim, Germany) could significantly inhibit YcgC’s deacetylase activity (Figure 3—figure supplement 1a). Further analysis revealed that the active component in the Halt Protease Inhibitor Cocktail was a serine hydrolase inhibitor 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF), but not the other components. Additional assays demonstrated that YcgC’s deacetylase activity could not be inhibited by well-known deacetylase inhibitors, including trichostatin A, SAHA (suberoylanilide hydroxamic acid), and NAM (Nicotinamide), and that hydrolase inhibitors, phenylmethylsulfonyl (PMSF), leupeptin, ethylenediaminetetracetic acid and EGTA, showed no detectable inhibition of YcgC (Figure 3—figure supplement 1b,c). Thus, it is likely that YcgC belongs to the serine hydrolase family, which has no significant homology to any of the annotated KDACs to date.

To identify which Ser residue in YcgC was most critical for its hydrolase (KDAC) activity, we examined five Ser residues, namely S7, S10, S73, S77, and S200, which are highly conserved among its prokaryotic homologs on the basis of protein sequence alignment. Next, we created a quintuple Ser-to-Ala mutant (i.e., 5SA) in YcgC and tested its ability to deacetylate RutR in vitro. As compared with the WT YcgC, the 5SA mutant appeared devoid of RutR deacetylase activity (Figure 3a). To determine which of the five Ser residues was likely to be the catalytic nucleophile for hydrolase activity, we incubated WT YcgC with hydrolase inhibitor AEBSF and the following MS/MS analysis revealed that Ser200 was the only conserved residue that was covalently labeled with AEBSF (Figure 3b). To confirm its importance, a Ser200-to-Ala (S200A) mutant was created and tested in the same in vitro assay. As anticipated, the deacetylase activity was not detectable with S200A YcgC, establishing Ser200 as the likely key catalytic residue.

Figure 3 with 3 supplements see all
S200 is critical for YcgC’s deacetylation activity.

(a) Two mutants of YcgC, that is, S S8/10/73/77/200A and S200A were constructed through gene synthesis. In vitro assays of the KDAC activity of these two mutants on RutR demonstrated that their KDAC activities were completely abolished. (b) S200 on YcgC was identified as an AEBSF binding site by LC-MS/MS analysis. YcgC was incubated with AEBSF, then trypsin digested and subjected to MS/MS analysis. Upon AEBSF-mediated sulfonation, a 183 Da molecular weight increase is predicted. (c) In vitro assays of the KDAC activity of YcgC on heat-denatured RutR demonstrated that YcgC was still active (right panel), while the downshift band disappeared (left panel). Similar results were observed when heat-denatured RutR was treated with CobB. KDAC: Lysine deacetylase; LC-MS/MS: Liquid chromatography–mass spectrometry; AEBSF: 4-(2-Aminoethyl)benzenesulfonyl fluoride; WT: Wild type.

https://doi.org/10.7554/eLife.05322.010

As mentioned, RutR deacetylation by YcgC reproducibly induces faster migration on sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). This behavior was also observed with deacetylation by the sirtuin family member CobB (Figure 2a; Figure 3a). This phenomenon is not typically associated with protein deacetylation and we considered the possibility that proteolytic degradation of deacetylated RutR was occurring. N-terminal Edman sequencing of RutR after YcgC treatment showed N-terminal truncation with loss of 14 residues (N-MTQGAVKTTGKRSR) or 11 residues (N-MTQGAVKTTGK). RutR without YcgC treatment was also sequenced, no N-terminal truncation was observed. We considered the possibility that YcgC might show protease activity in addition to its unambiguous deacetylase activity. However, we believe this is unlikely since treatment of RutR with the sirtuin CobB, mechanistically and structurally unrelated to YcgC, induces a similar N-terminal truncation in RutR, visualized by SDS-PAGE and detected by N-terminal sequencing with loss of 14 residues. We suspect that this N-terminal cleavage of RutR results from autoproteolysis. To explore this possibility, intact RutR proteins were first heat-denatured and then incubated with either YcgC or CobB under the same deacetylation conditions. As shown in Figure 3, heat-denatured RutR could be partially deacetylated by both YcgC and CobB, but did not show a downshifting on SDS-PAGE. These results are consistent with the possibility that deacetylation of native acetylated RutR sparks autoproteolysis but denaturation inhibits this autoproteolytic activity.

To understand whether specific Lys residue(s) play a role in facilitating RutR autoproteolysis, K52R, K62R, K52/62R, K52Q, K62Q, and K52/62Q mutant and WT RutR protein were treated with YcgC and analyzed using Coomassie-stained SDS-PAGE (Figure 3—figure supplement 2). Measurement of the ratios of cleaved/intact RutR revealed that K52Q and K52R RutR behaved similar to WT RutR. In contrast, K62Q, K62R, and the two double mutant RutR proteins showed diminished cleavage. Therefore, we propose that the apparent autoproteolytic activity of RutR is dependent on its deacetylation and that removal of the acetyl group from K62 appears most important for its cleavage. Future studies will be needed to further understand the molecular mechanism for the apparent RutR autocleavage and its biological function.

Because the N-terminal cleavage of RutR is tightly coupled with its deacetylation by YcgC, the downshifted band of RutR in the YcgC deactylation reaction can be conveniently used as a surrogate of YcgC’s activity. We thus performed steady-state assays to monitor the enzyme kinetics of YcgC via measuring the production of the downshifted RutR band, and estimated the Km and Vmax of YcgC to be 2.13 ± 0.65 μM and 0.29 ± 0.07 μM/min/μM, respectively (Figure 3—figure supplement 3).

YcgC regulates transcription through deacetylating RutR

Originally identified as a transcriptional repressor of the rutABCDEFG operon in E. coli, RutR (YcdC) was later found to bind to 19 additional E. coil chromosomal loci (Shimada et al., 2008), including the coding regions of pmrD and gcd (Umezawa et al., 2008). However, deletion of rutR alone does not result in a significant elevation of the expression levels of most of its target genes, suggesting that RutR regulates target gene transcription via a different mechanism. To determine whether deacetylation of RutR by YcgC plays a direct role in transcription regulation of RutR’s downstream target genes, we monitored the expression levels of 15 known target genes of RutR in ycgC-induced cells. Interestingly, while induction of ycgC did not change the expression level of rutR, the expression of two rutR targeting genes, pmrD and gcd, was significantly decreased, by as much as fivefold as measured with quantitative polymerase chain reaction (PCR) over a 2-hr period of ycgC induction (Figure 4a). On the other hand, induction of cobB in parallel did not affect expression levels of rutR or any of the 15 of RutR’s target genes (data not shown), suggesting the possibility that YcgC regulates a different set of substrates from CobB.

YcgC and CobB target distinct sets of substrates.

(a) YcgC regulates gene expression via deacetylating RutR. Expression of gcd and pmrD is significantly reduced upon RutR induction over a period of 2 hr as measured by quantitative real-time PCR. Double asterisks indicate that the observed fold changes are statistically significant, p<0.01. (b) Global gene expression analysis of ycgC- and cobB-induced cells. Clustering analysis shows clearly that impact on global transcription of induction of ycgC is distinct from that of cobB. Venn diagram showing that there is no significant overlap between genes down- and up-regulated due to CobB and YcgC induction. (c) Overexpression of YcgC affects global protein acetylation levels in E. coli. After ycgC and cobB were separately induced for 1 hr, global acetylation was detected in whole lysates of Escherichia coli using two pan α-AcK antibodies. The WT E. coli strain was also included for comparison. Boxed areas indicate regions that show obviously different staining patterns in ycgC- and cobB-induced cells. PCR: Polymerase chain reaction; WT: Wild type.

https://doi.org/10.7554/eLife.05322.014

To test this hypothesis, we examined the impact of induction of ycgC and cobB on global gene expression profiles in E. coli (Allard et al., 1999; Yeung et al., 2004). Using a standard DNA microarray approach, we found that, compared with WT cells, 197 genes were significantly repressed and 93 genes were activated after 4 hr of ycgC induction. In agreement with the above observations, expression levels of pmrD and gcd were significantly reduced (Figure 4b; Supplementary file 1). A similar analysis in parallel revealed that 4 hr of cobB induction resulted in 195 and 136 up- and down-regulated genes, respectively, compared with WT cells. However, with the exception of xerC, none of the RutR’s targets was affected (Supplementary file 2). Furthermore, Venn diagram analysis did not reveal any significant overlap between either the up- or down-regulated gene groups in the ycgC and cobB induction experiments (Figure 4b). These results suggest that YcgC profoundly affects global gene expression and it probably functions via distinct biological processes from CobB.

This conclusion is further supported by evidence obtained at the protein level. Using immunoblotting with pan α-AcK, we observed that overexpression of ycgC decreases the acetylation levels of many proteins, resulting in a global change in acetylation profiles compared with those of WT cells (Figure 4c). Importantly, changes in the acetylation profile of ycgC-induced cells were different from those in cobB-induced cells. For example, in boxed areas 2 and 3 (Figure 4c), the acetylated bands are almost completely absent in ycgC-overexpressing cells, while they are essentially unchanged in cobB-overexpressing cells. On the other hand, in boxed area 1, cobB overexpression completely abolished the acetylation signals, whereas only a modest decrease in acetylation signals is observed in ycgC-overexpressing cells (Figure 4c). Taken together, the above results suggest that YcgC and CobB each target a distinct set of substrates.

YcgC’s homologs show protein deacetylase activities

To determine whether YcgC’s KDAC activity is evolutionarily conserved, we searched for its homologs in both eukaryotes and prokaryotes. Although YcgC shows limited homology to components of the phosphotransferase system (Punta et al., 2012), no statistically significant homologs were identified in eukaryotes (Molin et al., 2003). However, many prokaryotic homologs with high similarity were readily identified. Other than homologs from Escherichia strains, the closet homolog is the DhaM protein from Shigella sp. (str. 2457T), and statistically significant homologs were also identified in more remotely related bacterial species. Therefore, we selected five homologs, representing a wide range of homology, for closer scrutiny (Figure 5a). Sequence alignment of the five representative homologs with YcgC showed that the N-terminal regions (i.e., amino acids 1–2501–250) of these proteins are more conserved than the C-terminal regions (Figure 5b). To determine whether these YcgC homologs possess protein deacetylase activity, the five selected genes were synthesized, subcloned into the same IPTG-inducible expression vector as that of E. coli YcgC, and transformed into E. coli. After a 4 hr induction of each YcgC homolog, changes in global acetylation profiles were determined with two pan α-AcK antibodies and compared with that for WT cells (Figure 5c,d). Results clearly showed that induction of each of the five YcgC homologs gave rise to a unique protein deacetylation signature that is different from that of the WT and YcgC-induced strains. For example, overexpression of the Klebsiella homolog significantly reduced the acetylation levels of proteins at ~55 kDa (red box; Figure 5c). As another example, overexpression of the Pantoea homolog substantially reduced the acetylation levels of proteins around 60 and 43 kDa compared with the WT strain (wide and narrow red boxes; Figure 5d). Taken together, these results demonstrate that all five YcgC homologs possess readily detectable KDAC activity with different substrate preferences. Because YcgC and its homologs share little similarity with all the known KDACs identified so far, and because its activity does not require NAD+ or Zn2+, these results strongly suggest that this group represents a novel prokaryotic KDAC family.

YcgC represents a new family of KDACs.

(a) Five representative YcgC homologs with protein sequence homology ranging from low to high. (b) Amino acid sequence homology analysis between YcgC and five selected YcgC homologs from other bacteria. The consensus strength among the six homologous proteins at each amino acid position of YcgC is indicated with colored bars. Red, orange, green, light blue, dark blue, and blank bars represent 100, 80, 60, 40, 20, and 0% consensus strength, respectively. (c,d) Changes in global E. coli acetylation profiles upon induction of the five YcgC homologs. The five selected YcgC homologs were cloned, transformed into E. coli, and induced to overexpress. Global acetylation profiles of each induced strain were detected with a pan monoclonal antibody (Cell Signaling, #9441) and a pan polyclonal antibody (PTM-Biolabs, PTM-105), as shown in c and d, respectively. WT E. coli cells were also processed in parallel as a comparison. An antibody against myelin basic protein was used as a loading control. WT: Wild type.

https://doi.org/10.7554/eLife.05322.015

Discussion

In this study, we have applied a clip-chip approach to identify new KDAC candidates in E. coli. Our in-depth biochemical characterization revealed that the novel KDAC YcgC removes the acetyl groups on K52/62 of its substrates RutR via a previously unknown Ser hydrolyase activity. A surprising observation was that, after deacetylated by either YcgC or CobB, the RutR showed a significant downshift on SDS-PAGE, suggesting possible proteolytic activity. Further biochemical analysis established that this is likely autoproteolysis of RutR that is stimulated by deacetylation of K62. Our data suggest that acetylation of RutR may enhance its stability. Indeed, endogenous RutR purified from cells grown under standard conditions is heavily acetylated. Although a complete understanding of this phenomenon will require future study; to our knowledge, this is the first example of a protein deacetylation event driving proteolytic activation.

Our in vivo functional studies on YcgC revealed that it down-regulates the expression of several RutR target genes by catalyzing the deacetylation of two lysine residues on RutR. It has been puzzling until now how RutR represses target gene transcription by its previously reported binding to coding regions of pmrD and gcd, as the binding sites are located hundreds of base pairs downstream of the start codons, and deletion of rutR does not enhance their expression levels (Umezawa et al., 2008). Based on the results of this study, we propose that YcgC deacetylates RutR leading in turn to the recruitment of additional cofactors that enhance silencing of target gene expression. As PmrD has been demonstrated to serve as a connector between multiple two-component signal-transduction systems in Salmonella enterica, E. coli, and other bacteria, our study also suggests the possibility of crosstalk between protein acetylation and phosphorylation in E. coli, a prevalent regulatory mechanism found in eukaryotes (Eguchi and Utsumi, 2005).

This study also highlights several advantages of the clip-chip approach. First, as an activity-based screen, this method can be readily adopted to search for many other types of enzyme activities. Second, the clip-chip approach does not require any prior knowledge of the enzymes of interest as long as a robust biochemical assay is available. Third, it is a proteome-wide, high-throughput screen that does not require further deconvolution (e.g., in MS/MS) of the positive signals because of the use of a protein microarray on which each protein is physically addressable. Finally, the clip-chip approach is capable of functional annotation of enzymes using both gain- and loss-of-signal reactions. We envision that the ‘clip-chip’ strategy will proved to be of wide application for the de novo discovery of enzyme activity in biology.

Materials and methods

Chemicals and reagents

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Unless otherwise stated, all chemicals used in this study were purchased from Sigma-Aldrich (St Louis, MO), and enzymes were purchased from New England Biolabs (Ipswich, MA).

E. coli proteome chip preparation

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E. coli proteome chips were prepared as described previously (Chen et al., 2008). In brief, expression plasmid-carrying E. coli cells were cultured, induced, and harvested in 96 deep-well plates. To purify the fusion proteins, cell pellets were treated with lysozyme and incubated with Ni-NTA Superflow (QIAGEN, Valencia, CA) in Multiscreen Nylon Mesh filter plates (Millipore, Billerica, MA). After six washes, the proteins were eluted with 250 mM imidazole. To prepare the proteome chip, the purified proteins were re-arrayed from 96-well plates into 384-well plates in a cold room using an Apricot system (Apricot Designs, Covina, CA). The re-arrayed proteins were printed in duplicate onto plain glass slides.

Discovery of new protein deacetylases using an E. coli proteome chip

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A substrate slide was prepared by coating FAST slides with 200 μl acetylated protein at a protein concentration ≥0.1 μg/μL. The E. coli proteome chip was imprinted onto the substrate slide. After the removal of the proteome chip, the substrate slide was submerged in protein deacetylation buffer (50 mM Tris-HCl, 4 mM MgCl2, 50 mM NaCl, 50 mM KCl, 1 mM NAD+ , pH 8.0). The reaction was carried out at 26°C for 16 hr. The slide was washed three times with 1× Tris-buffered saline and Tween 20 (TBST), 5 min each time, and incubated with an α-AcK antibody (#9441 of Cell Signaling Technology, Danvers, MA). The incubation was carried out with a 1:1000 antibody dilution at room temperature for 1 hr. The slide was washed four times with 1× TBST, 5 min each time, followed by incubation with a Cy3-conjugated secondary antibody from Jackson ImmunoResearch (West Grove, PA). To facilitate the identification of positive spots, the substrate slide was further probed with an α-6xHis antibody followed by a Cy5 conjugated secondary antibody from Jackson ImmunoResearch. A GenePix 4200A microarray scanner was used to record the results. Since this is a loss-of-signal assay, the signal intensity of each protein spot was defined as ‘Background-Foreground’. The signal intensity of each protein was averaged from the two replicate spots. Signal-to-noise ratio (SNR), that is, signal/standard deviation of background, was set as the final signal of each protein. The cutoff to call protein deacetylase candidates was set as SNR ≥3.

Protein deacetylase assays

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Deacetylase candidates identified by clip-chip were overexpressed and purified in E. coli. In a 20 μL reaction, the three acetylated substrates, that is, 3 μg of RutR and YceC, and 0.5 μg of NhoA, were individually incubated with 5 μg of each deacetylase candidate. The reactions were carried out in protein deacetylation buffer at 37°C for 1 hr. These protein samples were then analyzed by both silver staining and Western blotting. Membranes were further probed with an IRDye 800 secondary antibody at room temperature for 1 hr and visualized with an Odyssey Infrared Imaging System from LI-COR Biosciences (Lincoln, NE).

The deacetylation assays with hydrolase inhibitors

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Hlat protease inhibitor cocktail was purchased from Thermo Scientific, cOmplete protease tablets were from Roche, AEBSF, aprotinin, bestatin, and pepstatin A were obtained from Sangon Biotech Co., Ltd (Shanghai, China). The deacetylation assays were performed as described above except for the addition of a variety of hydrolase inhibitors at appropriate concentrations. The solvents of these inhibitors, that is, dimethyl sulfoxide and ethanol, were also tested as controls.

Measure NAD+ by liquid chromatography-coupled high-resolution mass spectrometry

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Affinity purified YcgC (80 µg) was diluted in 200 µL phosphate-buffered saline (pH 7.4), and denatured at 100°C for 10 min to release any bound NAD+. After centrifugation at 12,000 rpm for 10 min, the supernatant was transferred to a new Eppendorf tube, and acetonitrile (ACN) was added to the supernatant at a ratio of 3:1 (vol/vol). The reaction was mixed well, allowed to stand for 20 min at 4°C, then centrifuged at 12,000 rpm for 10 min. The resulting supernatant was then subjected to liquid chromatography-coupled high-resolution mass spectrometry (LC-HRMS) analysis. An aliquot of pure NAD+ (50 µM) was tested to calibrate the LC-HRMS system and as a positive control. LC-HRMS was performed as described previously (Vogliardi et al., 2011) on a Waters ACQUITY UPLC system equipped with a binary solvent delivery manager and a sample manager, coupled with a Waters Micromass Q-TOF Premier Mass Spectrometer equipped with an electrospray interface (Waters Corporation, Milford, MA). Briefly, LC was performed on a Syncronis HILIC column (50 × 2.1 mm, 1.7 µm) (Thermo Scientific). The column was eluted with 200 mM ammonium formate aqueous solution and ACN in gradient mode at a flow rate of 0.30 mL/min at 30°C. MS was performed using negative polarity, 2.4 KV capillary voltage, 30 V sampling cone, 4 eV collision energy, a source temperature of 110°C, and a desolvation temperature of 350°C. The flow rate for the desolvation gas was set at 600 L/hr. Scan range was set to m/z 50–1000, scan time to 0.3 s, and interscan time to 0.02 s.

Metal analysis by ICP-MS

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ICP-MS analysis (Goullé et al., 2005) was performed according to the manufacturer’s instructions. Briefly, 0.48 mg of YcgC was prepared in 4 mL concentrated nitric acid and 2 mL deionized water. The solution was then subjected to microwave digestion with a Multiwave 3000 instrument from Anton Paar ShapeTec GmbH (Wundschuh, Austria) at 600 W power for 15 min. The digested sample was filtered through a filter paper. The sample was analyzed on an ELAN 9000 ICP-MS instrument from PerkinElmer, Inc. (Waltham, MA). Human hair GBW07601a (GSH-1a) from Institute of Geophysical and Geochemical Exploration (Hebei, China) was included as a positive control. All experiments were carried out at room temperature in a dust-free area with a relative humidity of 10–85%.

Measuring the Km and Vmax of YcgC

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YcgC (0.3 μM) was incubated with RutR at a series of concentrations, that is, 1.5, 2.3, 3, 4.5, 6, 7.5, 9, 12, and 15 μM. The reaction was carried out in protein deacetylation buffer without NAD+ at 37°C for 15 min. Protein samples were then resolved by 12% SDS-PAGE followed by silver staining. The gel was scanned with a PowerLook 2100XL from Techville, Inc. (Dallas, TX), converted to an 8-bit grayscale image and analyzed by Image J (NIH; http://rsb.info.nih.gov/ij/).

Identification of deacetylation sites by MS

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Acetylated RutR and deacetylated RutR were trypsin-digested and analyzed with a nanoflow LC-MS/MS coupled online with a Q Exactive Plus quadrupole orbitrap mass spectrometer (Thermo Scientific, San Jose, CA) equipped with a nanoelectrospray ion source. Briefly, the peptide mixtures were loaded onto a C18 column (100 mm inner diameter, 10 cm long, 5 mm resin) from Michrom Bioresources (Auburn, CA) using an autosampler. Peptides were eluted with a 0–35% gradient (Buffer A, 0.1% formic acid, and 5% ACN; Buffer B, 0.1% formic acid, and 95% ACN) over 80 min and detected online with a Q Exactive Plus quadrupole orbitrap mass spectrometer using a data-dependent TOP10 method (Haas et al., 2006).

Construction of an E. coli strain (W3110) harboring chromosomal 3xFLAG-tagged RutR

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E. coli strain (W3110) harboring chromosomal 3xFLAG-tagged RutR was constructed using the Red recombination system (Poteete, 2001). In short, the DNA cassette for recombination was composed of a 150 bp upstream flanking sequence, the rutR gene, a 3xFLAG tag before the stop codon of rutR, followed by the sequence of the kanamycin resistance gene, and a 150 bp downstream flanking sequence. This cassette was synthesized and cloned into pUC57 by GenScript (Nanjing, China). The cassette was amplified by high fidelity PCR and treated with DpnI. One microgram of the linear DNA fragment was electrotransported into E. coli W3110 cells carrying pKD46, and these recombinants were selected using kanamycin medium and verified by colony PCR.

Determination of the deacetylation activity of YcgC in vivo

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The plasmid carries ycgC from the E. coli AG1 strain that we used for the construction of the E. coli proteome chip, was extracted and transformed into the E. coli W3110 strain harboring chromosomal 3xFLAG-tagged RutR. The transformed E. coli strain was then cultured in lysogeny broth media to a OD600 of 0.6–0.8 and induced by 1 mM IPTG at 37°C for 0.5, 1, 2, and 4 hr. Cells were harvested and treated with lysis buffer (50 mM NaH2PO4, 300 mM NaCl, 20 mM imidazole, 1× CelLytic B, 50 units/mL of Benzonase proteinase inhibitor cocktail, and 1 mM PMSF, pH 8.0) at 4°C for 2 hr with vigorous shaking. The 3xFLAG-tagged RutR was then immunoprecipitated using an α-FLAG antibody and protein G conjugated agarose beads. Samples were resolved on a 10% SDS-PAGE gel followed by Western blotting with an α-AcK antibody (Cell Signaling Technology, Shanghai, China) and an α-FLAG polyclonal antibody.

Determination of the acetylation level of RutR mutants

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RutR mutants K52Q, K62Q, and double mutant K52Q/K62Q were synthesized and cloned into pET28a+ with GenScript (Supplementary file 3). The acetylation level of these RutR mutants was detected with an α-AcK antibody (Cell Signaling Technology) and compared with that of the WT E. coli strain.

Preparation of α-YcgC antibody

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Mouse α-YcgC monoclonal antibody was custom-made by Abmart, Inc. (Shanghai, China). Western blotting was applied to characterize the antibody. The sensitivity of the antibody was tested using serially diluted RutR and its specificity was tested by spiking purified RutR into a whole lysate of E. coli.

Identify AEBSF binding site on YcgC by LC-MS/MS

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At a molar ratio of 500:1, AEBSF and YcgC were incubated at 37°C for 1 hr. After SDS-PAGE and Coomassie staining, YcgC band was cut off and in-gel trypsin digestion was done according to the standard protocol. The digested YcgC was then analyzed using Ultimate 3000 Nano Pump LC system from Thermo Scientific coupled with an electrospray ionization quadrupole time-of-flight mass spectrometer from Bruker Daltonics (Bremen, Germany). The LC setup was coupled online to a Q-TOF using a nano-ESI source from Bruker Daltonics in data-dependent acquisition mode (m/z 350–1500). Tandem mass spectra were extracted, charge state was deconvoluted and deisotoped by Compass Data Analysis version 4.1 from Bruker Daltonics. Mascot version 2.4 from Matrix Science (Boston, MA) was set up to search the database (entries). Carbamidomethyl on cysteine was specified as fixed modifications, oxidation of methionine was specified as variable modifications.

Identification of the N-terminal sequence of deacetylated YcdC by YcgC and CobB

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RutR was incubated with YcgC in deacetylation buffer at 37°C for 1 hr. Protein sample was then resolved by 12% SDS-PAGE and transferred to the polyvinyl difluoride membrane. The shift RtuR band was dyed by Ponceau S and cut off, and then N-terminal sequenced by protein sequencer PPSQ-33A from Shimadzu (Kyoto, Japan). The raw data and graphs generated by PPSQ-33A were identified and exported by PPSQ-33A data processing. The N-terminal sequence RtuR was then determined.

Determination of the acetylation status of E. coli whole lysates

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E. coli cells were cultured in LB medium at 37°C. Before and after induction with 1 mM IPTG for 1 hr, cells were treated with lysis buffer at 4°C for 2 hr with vigorous shaking. Cell debris was removed by centrifugation at 4°C. The protein concentration of the whole lysate was determined using the BCA Protein Assay (Pierce, Rockford, IL); 100 μg of whole lysate was then resolved on a 10% SDS-PAGE gel followed by Western blotting with an α-AcK antibody (Cell Signaling Technology) overnight at 4°C. As a loading control, the protein lysates were Western blotted with an α-myelin basic protein antibody from Abmart, Inc. (Noinaj et al., 2013; Spanò et al., 2011). Results were recorded using an IRDye 800 secondary antibody and the Odyssey Infrared Imaging System (LI-COR Biosciences).

Quantitative real-time PCR

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Total RNA was extracted using an RNA extraction kit from TIANGEN Biotech Co., Ltd. (Beijing, China). RNA was then reverse-transcribed to cDNA using a random oligo primer from Promega (Beijing, China), according to the manufacturer’s instructions. Primers were synthesized by Sangon Biotech. (Shanghai, China) (Supplementary file 4) and validated by regular PCR and melting curve analysis. Real-time PCRs were carried out using FastStart Universal SYBR Green Master from Roche (Shanghai, China) and the ABI 7500 real-time PCR platform (Life Technologies Corporation, Shanghai, China).

Global gene expression analysis using DNA chips

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E. coli DNA chips were purchased from CapitalBio Corp. (Beijing, China). cDNA labeled with a fluorescent dye (Cy5 and Cy3-dCTP) was produced by Eberwine’s linear RNA amplification method and subsequent enzymatic reaction (Guo et al., 2005; Patterson et al., 2006). Arrays were hybridized in a CapitalBio BioMixer II Hybridization Station overnight and scanned with a LuxScan scanner and the images obtained were then analyzed using LuxScan 3.0 software from CapitalBio Corp. A space- and intensity-dependent normalization based on a LOWESS program was employed (Yang et al., 2002). To identify significantly differentially expressed genes, SAM 3.02 was used. Unsupervised hierarchical clustering was used to cluster samples or genes. The distance between single samples or genes was based on Pearson’s correlation coefficients. Distances between clusters were calculated using the ‘complete linkage’ method. Venn diagrams were drawn using the R package Vennerable. The size of each circle proportionally reflects the number of unique genes in each group.

Homology analysis, phylogenetic tree construction, and sequence alignment

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To identify YcgC’s bacterial homologs, ‘dihydroxyacetone kinase subunit M’ was used as a search term in PubMed under the ‘Protein’ category. This search showed that the top taxonomic groups, that is, Escherichia, Klebsiella, Shigella, Serratia, Citrobacter, Yersinia, Enterobacter, Salmonella, Pantoea, and Providencia, all belonged to the Enterobacteriaceae. The amino acid sequence of YcgC was then Blasted against the most significant taxonomic groups using BlastP. Bacterial strains of the closest homologs from each taxonomic group were determined, that is, Citrobacter koseri ATCC BAA-895, Enterobacter aerogenes KCTC 2190, Klebsiella oxytoca KCTC 1686, Pantoea ananatis LMG 5342, Providencia stuartii MRSN 2154, Serratia odorifera 4Rx13, Shigella flexneri 2a str. 2457T, and Yersinia enterocolitica subsp. enterocolitica WA-314, and the amino acid sequences of these homologs, along with that for YcgC from E. coli K12 W3110, were then subjected to phylogenetic tree construction and sequence alignment using LaserGene (DNAstar Inc. Madison, WI).

References

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Decision letter

  1. Leemor Joshua-Tor
    Reviewing Editor; Cold Spring Harbor Laboratory, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for sending your work entitled "YcgC Represents a New Protein Deacetylase Family in Prokaryotes" for consideration at eLife. Your article has been favorably evaluated by Michael Marletta (Senior Editor), Leemor Joshua-Tor (Reviewing Editor), and three reviewers.

The Reviewing Editor and the reviewers discussed their comments before we reached this decision, and the Reviewing Editor has assembled the following comments to help you prepare a revised submission.

The paper describes an interesting study that uses an application of proteome array "Clip-Chip" technology to the discovery of new enzymatic functions. This appears to be a new twist on the use of proteome arrays and therefore very appealing. This technology has clearly identified a new class of protein deacetylases that does not require metal ions or NAD+. This is a fortunate happenstance and highlights the utility of the method for discovery of enzymes that informatics would not have picked up by homology searching. Thus, this paper will be of considerable interest to a broad spectrum of biological scientists.

However, several issues have been raised specifically regarding the activity of YcgC as a lysine deacetylase that would have to be addressed:

1) The biochemical data supporting that YcgC has deacetylase activity was based on Western blots using pan-specific acetyl lysine antibodies. One particular concern is that YcgC might be a protease that can cleave RutR at a specific sequence. This cleavage may in turn affect the recognition of an acetyl lysine residue in RutR by the antibody, leading to a decreased acetyl lysine signal on Western blots. This is highly likely because Figure 2A shows that RutR became smaller after treatment with YcgC. In addition, further post-translational modifications adjacent to the acetylated lysine might alter the epitopes for proper readout by an antibody. To rule out this concern, more biochemical data is needed. One suggestion is to examine whether YcgC can deacetylate a synthetic acetyl peptide and monitoring products using LC-MS based methods, which can provide definitive proof for the deacetylase activity.

2) In Figure 1D legend, you state: "using acetylated RutR proteins […] YcgC showed robust deacetylation activity in vitro". It seems that the SDS-PAGE mobility of RutR doesn't change following deacetylation. However, in Figure 2D, there is at least 5KD shift to a lower molecule weight after deacetylation. Such a discrepancy should be addressed.

3) The mass spectrometry data showing the acetylated K52 peptide (Figure 2D) is problematic. Although the spectra itself is not readable, it is clear that the peptide was obtained by trypsin digestion (as stated in the experimental section), which cleaves after lysine and arginine, but not after acetyl lysine. A long peptide with Kac embedded in the center should be detected. Figure 2D shows a peptide cut after acetyl lysine, which is highly unlikely. This indicates that either the acetyl group is not on K52 or the MS data is unreliable. Although all the Y ions are marked, for the C-Kac peptide as shown, the Y ions shouldn't be visible because it lost the positively charged Lys which shouldn't fly in MS. In contrast, the C-Lys peptide should fly. However, the similar ionization efficiency for the b and y ions for the two peptides doesn't make sense. The images are at low resolution and therefore hard to examine in detail. In addition, HPLC retention times should be indicated. The same concern is noted for the MS data shown in the supplementary figures.

4) The gel shown in Figure 2F is not convincing. The authors state in the text that the double mutant provides the lowest level of acetylation of RutR, the substrate protein. This is not apparent. At least by my eye the K62Q and the double mutant appear to provide quite similar results. Some sort of quantification of these results should be provided.

5) It was not clear in the manuscript whether or not YcgC is an essential protein in E. coli. A related point is that when the authors overexpress the protein in vivo, it was not clear whether this was being done in a ∆ycgC strain or on top of the native level of YcgC. Please provide this information.

Other issues to address:

6) It is surprising that 1x or 2x-acetylation/deacetylation can cause such a dramatic mobility shift (Figure 2A). Fortunately, the authors showed that CobB also deacetylates the same substrate. This data should be included to show that deacetylation can indeed cause a 5kD mobility shift.

7) In Figure 2C, why are the band intensities of alpha-Flag and α-YcgC so different? In addition, even given that α-Flag bands have equal intensity, the decreased α-Kac bands should correspond to two α-Flag bands given the decreased mobility shift following deacetylation.

8) For the MS analyses, it is not unclear whether the authors run the SDS page gels first and then cut the band for MS analysis. In theory, there should be two bands (Kac-containing starting materials and K-containing products), which should be resolved well and analyzed in parallel.

9) In Figure 2F, interestingly, although the negative signals of α-Kac antibodies, the Coo. bands showed the similar mobility as shown in Figure 2A. It seems that the same reaction occurs when RutR was mixed with YcgC. It is another strange observation.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "YcgC Represents a New Protein Deacetylase Family in Prokaryotes" for further consideration at eLife. Your revised article has been favorably evaluated by Michael Marletta (Senior Editor), a Reviewing Editor, and two reviewers. The manuscript has been significantly improved but there are some remaining issues that need to be addressed before acceptance, as outlined below:

The reviewers all felt that this work is significant and important. Precisely because of this, it is important to make sure as much as possible that observations reported are not due to a contaminating protease activity. The MS data showing 95% sequence coverage of RutR sequence cannot prove that there is no proteolysis. The fact that YcgC does not work on a synthetic peptide is another reason for concern.

There are a couple of simple things that could be done to address this concern:

1) In Figure 2A, include a negative control with CobB but without NAD+.If there is a protease contamination, it is likely an endogenous E. coli protein that is present in both the YcgC and CobB prep. Using CobB without NAD+ should resolve this.

2) In Figure 2F, K52Q and K62Q mutants were blotted and they appear to be of similar size to the WT protein. One can argue that the K to Q mutant behaves similarly to the acetylated WT protein. However, the authors should repeat these experiments using the K52R and K62R mutants. If these mutants migrate faster as the deacetylated RutR, it will also help address these concerns. In addition, the K to R mutants may improve the Ac Western blot because Q is generally thought to mimic acetyl lysine.

It is interesting that the 5-kd mobility shift only occurs for native YcgC but not N-terminal flagged YcgC. Such an observation should be described explicitly in the text to avoid potential confusion, especially if the K52R and K62R mutants don't migrate as fast. This type of shift by 1 or 2 Ac groups is quite large.

3) Another possibility is to mutate the conserved Ser residues in YcgC and show that the mutant loses deacetylase activity. There are only five conserved Ser residues and mutating them would not be a huge effort.

4) For Figure 1D, the experimental details provided in the main text and response letter are still not clear. Did the authors run "Coo. Stain" on the samples and then use the same sample for the deacelylation reaction or did they run "Coo. Stain" and deacelylation with equal amounts of the two aliquots? If the latter is the case, the samples after the deacelylation reaction should be subsequently stain by a Western-compatible dye followed by anti-acetyllysine antibody.

https://doi.org/10.7554/eLife.05322.020

Author response

[…] However, several issues have been raised specifically regarding the activity of YcgC as a lysine deacetylase that would have to be addressed: 1) The biochemical data supporting that YcgC has deacetylase activity was based on Western blots using pan-specific acetyl lysine antibodies. One particular concern is that YcgC might be a protease that can cleave RutR at a specific sequence. This cleavage may in turn affect the recognition of an acetyl lysine residue in RutR by the antibody, leading to a decreased acetyl lysine signal on Western blots. This is highly likely because Figure 2A shows that RutR became smaller after treatment with YcgC. In addition, further post-translational modifications adjacent to the acetylated lysine might alter the epitopes for proper readout by an antibody. To rule out this concern, more biochemical data is needed. One suggestion is to examine whether YcgC can deacetylate a synthetic acetyl peptide and monitoring products using LC-MS based methods, which can provide definitive proof for the deacetylase activity.

We thank the reviewers for giving us this opportunity to further clarify this concern. In this study, the acetylated lysine residues were determined by comparing the MS/MS peaks obtained from the upper and lower (i.e., down-shifted) RutR protein bands after PAGE separation. To make this point clearer, we have now added an insert to Figures 2D and 2E, as well as to Figure 2–figure supplement 3. To demonstrate that the down-shifted RutR was not caused by protease activity of YcgC, we repeated the MS/MS analysis several more times against the down-shifted RutR bands recovered from PAGE gels. (Please also see our response to Point 3 below.) To our satisfaction, we obtained a peptide-coverage for up to 95% and with both the N- and C- terminal of the full-length RutR, indicating that the down-shifted RutR was very unlikely to be caused by protease cleavage. Moreover, we observed that CobB-deacetylated RutR protein also appeared as a similar down-shifted band on SDS-PAGE, providing further confirmation of our interpretation of the data. Taken together, these additional results help establish that YcgC is a bona fide deacetylase and not a protease. We have now clarified this in the second paragraph of the subsection “In vitro characterization of YcgC's KDAC activity”, and added the CobB treated RutR to Figure 2.

In addition, we have also treated RutR-derived synthetic peptides in acetylated and unacetylated forms with YcgC as recommended by the reviewers. There was no evidence that the RutR acetylated peptide was deacetylated or otherwise modified by YcgC treatment as analyzed by HPLC. Our interpretation of these results is that a folded substrate conformation or long-range recognition elements within RutR are critical for YcgC processing.

2) In Figure 1D legend, you state: "using acetylated RutR proteins […] YcgC showed robust deacetylation activity in vitro". It seems that the SDS-PAGE mobility of RutR doesn't change following deacetylation. However, in Figure 2D, there is at least 5KD shift to a lower molecule weight after deacetylation. Such a discrepancy should be addressed.

We thank the reviewers for pointing out this “discrepancy.” The purpose of Coomassie staining in the lower panel of Figure 1D is to show that an equal amount of RutR proteins was used for each reaction. Since they were not deacetylated by YcgC, no down-shift would be expected and none as observed. We have now clarified this in the Figure 1 legend.

3) The mass spectrometry data showing the acetylated K52 peptide (Figure 2D) is problematic. Although the spectra itself is not readable, it is clear that the peptide was obtained by trypsin digestion (as stated in the experimental section), which cleaves after lysine and arginine, but not after acetyl lysine. A long peptide with Kac embedded in the center should be detected. Figure 2D shows a peptide cut after acetyl lysine, which is highly unlikely. This indicates that either the acetyl group is not on K52 or the MS data is unreliable. Although all the Y ions are marked, for the C-Kac peptide as shown, the Y ions shouldn't be visible because it lost the positively charged Lys which shouldn't fly in MS. In contrast, the C-Lys peptide should fly. However, the similar ionization efficiency for the b and y ions for the two peptides doesn't make sense. The images are at low resolution and therefore hard to examine in detail. In addition, HPLC retention times should be indicated. The same concern is noted for the MS data shown in the supplementary figures.

We have consulted several mass spectrometry experts, and re-performed the MS/MS analysis with a different MS spectrometer (Q ExactiveTM Plus quadrupole orbitrap mass spectrometer) of a much higher mass resolution. Long peptides with Kac embedded in the center for both K52 and K62 were clearly observed for untreated RutR (-YcgC), while not for treated RutR (+YcgC). We have now updated Figure 2 and Figure 2–figure supplement 3 with the new MS/MS data, and provided clear images of HPLC retention times. We have also modified the manuscript accordingly (in the second paragraph of the subsection “In vivo validation of YcgC's KDAC activity”, and in the subsection “Identification of deacetylation sites by mass spectrometry").

4) The gel shown in Figure 2F is not convincing. The authors state in the text that the double mutant provides the lowest level of acetylation of RutR, the substrate protein. This is not apparent. At least by my eye the K62Q and the double mutant appear to provide quite similar results. Some sort of quantification of these results should be provided.

To clearly demonstrate the difference of acetylation level between the wild-type and mutant RutR proteins, we further optimized the amount of proteins and repeated the experiments for several times. We now provide quantification and statistic analyses to demonstrate the significant differences of their acetylation levels in the lower panel of Figure 2F. The figure legend and main text have been updated accordingly.

5) It was not clear in the manuscript whether or not YcgC is an essential protein in E. coli. A related point is that when the authors overexpress the protein in vivo, it was not clear whether this was being done in a ∆ycgC strain or on top of the native level of YcgC. Please provide this information.

YcgC is also known as DhaM, a subunit of the dihydroacetone kinase complex. It is not essential in E. coli. Because the endogenous level of YcgC is very low, YcgC was overexpressed on the wild type background. We have now clarified this point in the last paragraph of the subsection “Screen new KDAC using the E. coli proteome microarray”.

Other issues to address: 6) It is surprising that 1x or 2x-acetylation/deacetylation can cause such a dramatic mobility shift (Figure 2A). Fortunately, the authors showed that CobB also deacetylates the same substrate. This data should be included to show that deacetylation can indeed cause a 5kD mobility shift.

We thank the reviewers for pointing this out. We have now added the CobB reaction to Figure 2A, and modified the manuscript accordingly (subsection “In vitro characterization of YcgC's KDAC activity”).

7) In Figure 2C, why are the band intensities of alpha-Flag and α-YcgC so different? In addition, even given that α-Flag bands have equal intensity, the decreased α-Kac bands should correspond to two α-Flag bands given the decreased mobility shift following deacetylation.

In Figure 2C the anti-FLAG antibodies were used to detect the expression level of endogenous RutR, because we chromosomally tagged RutR gene with FLAG (subsection “In vivo validation of YcgC's KDAC activity”, first paragraph), whereas anti-YcgC mAb (created during this study) was for detecting YcgC during a time course of induction. It is evident that the endogenous level of YcgC is very low as shown in the control lane in Figure 2C. The down-shifted RutR bands were only observed in our in vitro deacetylation reactions, when RutR was almost completely decetylated on both K52 and K62 (as shown by MS/MS analysis). While the evidence of Figure 2C indicates that YcgC can deacetylate endogenous RutR in vivo, it is likely to be far from complete compared with in vitro assays (Figure 2C), potentially accounting for the lack of a shift. The FLAG tag may also affect the mobility, as well as other PTMs.

8) For the MS analyses, it is not unclear whether the authors run the SDS page gels first and then cut the band for MS analysis. In theory, there should be two bands (Kac-containing starting materials and K-containing products), which should be resolved well and analyzed in parallel.

Yes, we treated RutR with YcgC first and then ran it together with untreated RutR on a PAGE gel. After Coomassie staining, we recovered both the upper band in the untreated RutR lane and the down-shifted band in the YcgC-treated lane. Both recovered proteins were then subjected to MS analysis. Please also see our response to Point 1 above. We have now added an insert to Figures 2D and E and Figure 2–figure supplement 3, and have clarified this in the manuscript (Figure 2 legend; Figure 2–figure supplement 3 legend).

9) In Figure 2F, interestingly, although the negative signals of α-Kac antibodies, the Coo. bands showed the similar mobility as shown in Figure 2A. It seems that the same reaction occurs when RutR was mixed with YcgC. It is another strange observation.

In Figure 2F, lysine-to-glutamine mutations were used to demonstrate that K52 and K62 were the two dominant acetylated residues. Because K-to-Q mutation has been commonly used to generate an acetylated lysine mimic due to chemical similarity of an acetamide and the Gln side chain (e.g., PMID: 23904479; 19303850; 21906795), such mutation may not generate enough structural change of the protein to cause a down-shifting of the band. To better demonstrate the differences in their acetylation level, we have repeated this assay and added statistical analyses to Figure 2F. Please also see our response to Point 4 above. We have now updated Figure 2F and modified the manuscript accordingly (Figure 2 legend).

[Editors' note: further revisions were requested prior to acceptance, as described below.]

The reviewers all felt that this work is significant and important. Precisely because of this, it is important to make sure as much as possible that observations reported are not due to a contaminating protease activity. The MS data showing 95% sequence coverage of RutR sequence cannot prove that there is no proteolysis. The fact that YcgC does not work on a synthetic peptide is another reason for concern. There are a couple of simple things that could be done to address this concern: 1) In Figure 2A, include a negative control with CobB but without NAD+.If there is a protease contamination, it is likely an endogenous E. coli protein that is present in both the YcgC and CobB prep. Using CobB without NAD+ should resolve this.

We have now included the negative control with CobB but without NAD+. As expected, no deacetylase activity was observed for this reaction (Figure 2–figure supplement 1), suggesting that it was unlikely due to contamination by an endogenous protease. We have added this result and modified the manuscript accordingly (subsection “In vitro characterization of YcgC's KDAC activity”, first paragraph).

2) In Figure 2F, K52Q and K62Q mutants were blotted and they appear to be of similar size to the WT protein. One can argue that the K to Q mutant behaves similarly to the acetylated WT protein. However, the authors should repeat these experiments using the K52R and K62R mutants. If these mutants migrate faster as the deacetylated RutR, it will also help address these concerns. In addition, the K to R mutants may improve the Ac Western blot because Q is generally thought to mimic acetyl lysine.

It is interesting that the 5-kd mobility shift only occurs for native YcgC but not N-terminal flagged YcgC. Such an observation should be described explicitly in the text to avoid potential confusion, especially if the K52R and K62R mutants don't migrate as fast. This type of shift by 1 or 2 Ac groups is quite large.

We thank the reviewers for their insightful suggestions. The major concern was the observation of the 5-kDa downshift of RutR after the treatment of either YcgC or CobB, as it raised the possibility that YcgC might act as a protease that cleaved off the acetylated lysines of RutR. To explicitly address this concern, we have performed a series of assays. First, we recovered the downshifted RutR after YcgC treatment (e.g., Figure 2A) and performed N-terminal Edman sequencing. We identified two versions of cleaved RutR: the major species involves deletion of 14-aa (N-MTQGAVKTTGKRSR), and the minor species involves deletion of 11 aa (N-MTQGAVKTTGK) from the N-terminus of RutR. Because the identified acetylated K52 and K62 are quite distant from the N-terminus, this N-terminal cleavage cannot account for YcgC’s KDAC activity, although it still did not rule out YcgC’s protease activity. Second, we sequenced the downshifted RutR after CobB treatment and found the same cleaved RutR missing the 14-aa at the N-terminus. Since CobB is a NAD+-dependent Class III sirtuin without any known protease activity, these results suggest that the RutR cleavage is intrinsic to RutR autoproteolysis. Third, to test whether RutR carries any protease activity, intact RutR proteins were heat-denatured and then incubated with either YcgC or CobB under the same deacetylation conditions. It became clear that denatured RutR could no longer downshift, while YcgC and CobB could still significantly deacetylate denatured RutR as compared with the untreated RutR. Finally, we examined the role of specific lysines in RutR’s proteolytic activity. As suggested by the reviewers, we created K52R, K62R and K52/62R mutants in RutR and tested them together with all the K-to-Q mutants in YcgC deacetylation reactions. On the basis of the ratios of cleaved/intact RutR in each reaction, we observed that when K62 was deacetylated in either K52Q or K52R, the proteolytic activity of these mutated RutR was almost as strong as WT RutR. In contrast, K62Q, K62R, and the two double mutants showed significantly lower proteolytic activities. Taken together, the above results confirmed YcgC’s KDAC activity and revealed that the autoproteolytic activity of RutR is dependent on deacetylation of K52/62 with K62 playing a major role. Because these new results are both important and interesting, we added a new Figure 3 and supplemental figures (Figure 2–figure supplement 4, Figure 3–figure supplement 2) to fully describe our new observations.

3) Another possibility is to mutate the conserved Ser residues in YcgC and show that the mutant loses deacetylase activity. There are only five conserved Ser residues and mutating them would not be a huge effort.

We thank the reviewers for this insightful suggestion. As the reviewers expected, once these five conserved Ser residues were mutated in YcgC, the S8/10/73/77/200A mutant could no longer deacetylate RutR (see new Figure 3A). To further pinpoint the essential Ser for the KDAC activity, we incubated WT YcgC with hydrolase inhibitor AEBSF and the following MS/MS analysis revealed that Ser200 was the only residue covalently labeled with AEBSF (See new Figure 3B). This result was further validated by the observation that S200A mutation completely abolished YcgC’s KDAC activity (Left panel; new Figure 3A). We have now added these new results to the main text and to the new Figure 3.

4) For Figure 1D, the experimental details provided in the main text and response letter are still not clear. Did the authors run "Coo. Stain" on the samples and then use the same sample for the deacelylation reaction or did they run "Coo. Stain" and deacelylation with equal amounts of the two aliquots? If the latter is the case, the samples after the deacelylation reaction should be subsequently stain by a Western-compatible dye followed by anti-acetyllysine antibody.

In the previous figure, one aliquot was Coomassie stained as loading control to show that equal amounts of RutR were loaded to each lane. Another aliquot was deacetylated and then WB by anti-acetyllysine antibody. To further clarify this, after the deacetylation reaction, RutR was divided into two aliquots of equal amounts: one aliquot was then subjected for Coomassie stain, and the other was western blotted by anti-acetyllysine antibody. We have also clarified this point in the manuscript accordingly (subsection “Screen new KDAC using the E. coli proteome microarray”, second paragraph).

https://doi.org/10.7554/eLife.05322.021

Article and author information

Author details

  1. Shun Tu

    1. Shanghai Center for Systems Biomedicine, Key Laboratory of Systems Biomedicine, Shanghai Jiao Tong University, Shanghai, China
    2. State Key Laboratory of Oncogenes and Related Genes, Shanghai, China
    Contribution
    ST, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Contributed equally with
    Shu-Juan Guo and Chien-Sheng Chen
    Competing interests
    No competing interests declared.
  2. Shu-Juan Guo

    1. Shanghai Center for Systems Biomedicine, Key Laboratory of Systems Biomedicine, Shanghai Jiao Tong University, Shanghai, China
    2. State Key Laboratory of Oncogenes and Related Genes, Shanghai, China
    Contribution
    S-JG, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Contributed equally with
    Shun Tu and Chien-Sheng Chen
    Competing interests
    No competing interests declared.
  3. Chien-Sheng Chen

    Graduate Institute of Systems Biology and Bioinformatics, National Central University, Jhongli, Taiwan
    Contribution
    C-SC, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Contributed equally with
    Shun Tu and Shu-Juan Guo
    Competing interests
    No competing interests declared.
  4. Cheng-Xi Liu

    1. Shanghai Center for Systems Biomedicine, Key Laboratory of Systems Biomedicine, Shanghai Jiao Tong University, Shanghai, China
    2. State Key Laboratory of Oncogenes and Related Genes, Shanghai, China
    Contribution
    C-XL, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  5. He-Wei Jiang

    1. Shanghai Center for Systems Biomedicine, Key Laboratory of Systems Biomedicine, Shanghai Jiao Tong University, Shanghai, China
    2. State Key Laboratory of Oncogenes and Related Genes, Shanghai, China
    Contribution
    H-WJ, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  6. Feng Ge

    Key Laboratory of Algal Biology, Institute of Hydrobiology, Chinese Academy of Sciences, Wuhan, Hubei, China
    Contribution
    FG, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  7. Jiao-Yu Deng

    State Key Laboratory of Virology, Wuhan Institute of Virology, Chinese Academy of Sciences, Wuhan, China
    Contribution
    J-YD, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  8. Yi-Ming Zhou

    National Engineering Research Center for Beijing Biochip Technology, Beijing, China
    Contribution
    Y-MZ, Analyzed the microarray data, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  9. Daniel M Czajkowsky

    Bio-ID Center, School of Biomedical Engineering, Shanghai Jiao Tong University, Shanghai, China
    Contribution
    DMC, Analyzed the microarray data, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  10. Yang Li

    1. Shanghai Center for Systems Biomedicine, Key Laboratory of Systems Biomedicine, Shanghai Jiao Tong University, Shanghai, China
    2. State Key Laboratory of Oncogenes and Related Genes, Shanghai, China
    Contribution
    YL, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  11. Bang-Ruo Qi

    1. Shanghai Center for Systems Biomedicine, Key Laboratory of Systems Biomedicine, Shanghai Jiao Tong University, Shanghai, China
    2. State Key Laboratory of Oncogenes and Related Genes, Shanghai, China
    Contribution
    B-RQ, Performed the experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  12. Young-Hoon Ahn

    Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Baltimore, United States
    Contribution
    Y-HA, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  13. Philip A Cole

    Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Baltimore, United States
    Contribution
    PAC, Conceived and designed the study with assistance from YHA, Analyzed the data, Wrote the manuscript, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    pcole@jhmi.edu
    Competing interests
    PAC: Reviewing editor, eLife
  14. Heng Zhu

    1. Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Baltimore, United States
    2. The High-Throughput Biology Center, Johns Hopkins University School of Medicine, Baltimore, United States
    Contribution
    HZ, Conceived and designed the study with assistance from YHA, Analyzed the data, Wrote the manuscript, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    hzhu4@jhmi.edu
    Competing interests
    No competing interests declared.
  15. Sheng-Ce Tao

    1. Shanghai Center for Systems Biomedicine, Key Laboratory of Systems Biomedicine, Shanghai Jiao Tong University, Shanghai, China
    2. State Key Laboratory of Oncogenes and Related Genes, Shanghai, China
    3. Bio-ID Center, School of Biomedical Engineering, Shanghai Jiao Tong University, Shanghai, China
    Contribution
    S-CT, Conceived and designed the study with assistance from YHA, Analyzed the data, Wrote the manuscript, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    taosc@sjtu.edu.cn
    Competing interests
    No competing interests declared.

Funding

National Natural Science Foundation of China (31370750)

  • Daniel M. Czajkowsky

National Institutes of Health (GM62437)

  • Philip A Cole

Flight Attendant Medical Research Institute

  • Philip A Cole

National Institutes of Health (RR020839)

  • Heng Zhu

National Institutes of Health (GM076102)

  • Heng Zhu

National Institutes of Health (HG006434)

  • Heng Zhu

Ministry of Science and Technology of the People's Republic of China (2010CB529205)

  • Sheng-Ce Tao

National Natural Science Foundation of China (31370813)

  • Sheng-Ce Tao

National Natural Science Foundation of China (31000388)

  • Sheng-Ce Tao

Ministry of Science and Technology of the People's Republic of China (2012AA020103)

  • Sheng-Ce Tao

Ministry of Science and Technology of the People's Republic of China (2012AA020203)

  • Sheng-Ce Tao

Ministry of Health of the People's Republic of China (2013ZX10003006)

  • Sheng-Ce Tao

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

The authors are grateful to W Yan, YK Wan, PY Yang, and MJ Tan for their expert research assistance and comments and J Fleming for editing the manuscript. This study was supported in part by grants from the National Natural Science Foundation of China (No. 31370813 and 31000388), and the National High Technology Research and Development Program of China (No. 2012AA020103 and 2012AA020203) to SCT, grant from National Natural Science Foundation of China (No. 31370750) to DMC, grants from the NIH (RR020839, GM076102, and HG006434) to HZ, and grant NIH GM62437 to PAC. We thank the FAMRI (Flight Attendant Medical Research Institute) Foundation for support of this work.

Reviewing Editor

  1. Leemor Joshua-Tor, Cold Spring Harbor Laboratory, United States

Publication history

  1. Received: October 25, 2014
  2. Accepted: October 28, 2015
  3. Version of Record published: December 30, 2015 (version 1)
  4. Version of Record updated: August 30, 2016 (version 2)

Copyright

© 2015, Tu et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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