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PHF6 regulates phenotypic plasticity through chromatin organization within lineage-specific genes

In this study, Soto-Feliciano et al. describe the function of the plant homeodomain finger 6 (PHF6) protein in leukemia and define its role in regulating chromatin accessibility to lineage-specific transcription factors. Their findings suggest that active maintenance of a precise chromatin landscape is essential for sustaining proper leukemia cell identity and that loss of a single factor (PHF6) can cause focal changes in chromatin accessibility and nucleosome positioning that render cells susceptible to lineage transition.

Developmental and lineage plasticity have been observed in numerous malignancies and have been correlated with tumor progression and drug resistance. However, little is known about the molecular mechanisms that enable such plasticity to occur. Here, we describe the function of the plant homeodomain finger protein 6 (PHF6) in leukemia and define its role in regulating chromatin accessibility to lineage-specific transcription factors. We show that loss of Phf6 in B-cell leukemia results in systematic changes in gene expression via alteration of the chromatin landscape at the transcriptional start sites of B-cell-and T-cell-specific factors. Additionally, Phf6 KO cells show significant downregulation of genes involved in the development and function of normal B cells, show up-regulation of genes involved in T-cell signaling, and give rise to mixed-lineage lymphoma in vivo. Engagement of divergent transcriptional programs results in phenotypic plasticity that leads to altered disease presentation in vivo, tolerance of aberrant oncogenic signaling, and differential sensitivity to frontline and targeted therapies. These findings suggest that active maintenance of a precise chromatin landscape is essential for sustaining proper leukemia cell identity and that loss of a single factor (PHF6) can cause focal changes in chromatin accessibility and nucleosome positioning that render cells susceptible to lineage transition. Lymphopoiesis is a carefully orchestrated process in which hematopoietic stem cells differentiate into B cells and T cells through the activation of precise gene expression programs governed by well-described transcription factors. During the process of lymphocyte differentiation, gene expression must be activated, silenced, or maintained. In B-cell development, the accessibility of specific genomic loci for binding of transcription factors (e.g., PU.1, E2A, EBF1, and PAX5) is vital for transcriptional regulation. However, the critical mechanisms regulating the chromatin landscape and transcription factor access have yet to be elucidated (Schebesta et al. 2002).
Lymphoid cells are terminally differentiated but still possess the capacity to transdifferentiate into distinct lineages when subjected to specific genetic perturbations.
For example, loss of Pax5 in mature B cells can produce functional T cells in immune-deficient mice (Cobaleda et al. 2007). Similarly, Pax5 deletion in pro-B cells allows for transdifferentiation into macrophages, granulocytes, osteoclasts, dendritic cells, and natural killer cells (Nutt et al. 1999). Overexpression of CEBPα/β can transform mature B and T cells into macrophages (Xie et al. 2004;Laiosa et al. 2006). In addition, Ebf1 loss converts pro-B cells into innate lymphoid cells and T cells (Nechanitzky et al. 2013). Interestingly, these lineage-specific transcription factors are often found to be altered in B-cell acute lymphoblastic leukemia (B-ALL). These findings highlight the plasticity of leukemia cells and how aberrant lymphoid developmental programs can favor leukemogenesis (Horcher et al. 2001;Rathert et al. 2015;Somasundaram and Sigvardsson 2015).
Plant homeodomain finger protein 6 (PHF6) was first described as the single gene mutated in the X-linked intellectual disability called Börjeson-Forssman-Lehmann syndrome (Lower et al. 2002). Inactivating mutations in PHF6 were subsequently identified in human leukemia of the T and myeloid lineages, highlighting its role as a tumor suppressor gene in these malignancies (Van Vlierberghe et al. 2010, 2011. Our group recently de-scribed a tumor-promoting role for Phf6 in a murine model of BCR-ABL1 + B-ALL (Williams et al. 2006;Meacham et al. 2015). In this study, we found that hairpin-mediated knockdown of Phf6 leads to impaired growth of B-ALL cells in vivo. Altogether, these observations suggest that PHF6 can act as a tumor suppressor or an oncogene in a lineage-dependent manner. However, the molecular mechanisms underlying PHF6's function in hematological malignancies remain entirely unknown (Fig. 1A). . Kaplan-Meier survival analysis of mice injected with either 10 3 (dotted) or 10 6 (solid) Phf6 WT (blue) and Phf6 KO (red) B-ALL cells. The number (n) of mice per genotype analyzed is shown. Statistical analysis (log-rank test, Mantel-Cox) was performed for the different groups in comparison with mice injected with Phf6 WT cells. P-values are shown for the comparisons. (C) Representative hematoxylin and eosin (H&E) (top) and immunohistochemistry (bottom) staining of serial sections from lymph nodes (LNs) and lymphoma (mass) of recipient mice injected with Phf6 WT , shPhf6, and Phf6 KO cells. mCherry immunochemistry demarcates tumor cells. Bars, 600 µm. (D, top) Size comparison of representative LNs from Phf6 WT (left) and Phf6 KO (right) recipient mice. (Bottom) Quantification of combined LN weight of Phf6 WT (blue; n = 5) and Phf6 KO (red; n = 5) recipients. (E) Tumor burden in the blood of Phf6 WT (blue; n = 7) and Phf6 KO (red; n = 8) recipient mice. mCherry demarcates tumor cells. (F ) Bar graphs showing the percentage of the CD4 + fraction among mCherry + cells isolated from Phf6 WT (blue; n = 9) and Phf6 KO (red; n = 5) tumors in bone marrow (left) and LNs (right). Data represent the mean ± standard deviation (SD) in D-F. Statistics were calculated with two-sided Student's t-test. ( * * * ) P < 0.001; ( * * * * ) P < 0.0001. Despite the knowledge obtained through sequencing studies, only a handful of functions have been described for PHF6. The protein contains two atypical PHD-like zinc finger domains, implying the capacity to bind modified histones similar to canonical PHD domains (Wysocka et al. 2006). However, PHF6 has only been shown to bind dsDNA in vitro (Liu et al. 2014). In addition, it has been shown to interact with transcriptional regulatory factors such as the nucleosome remodeling and deacetylation (NuRD) complex, the RNA polymerase II-associated factor 1 (PAF1) transcription elongation complex, and the rRNA transcriptional activator UBF (Todd and Picketts 2012;Wang et al. 2013;Zhang et al. 2013). To better understand the function of PHF6 as a potential chromatin regulator and examine its lineage-specific roles in hematological malignancies, we decided to thoroughly investigate its role in B-ALL. Here, through integrated genomics and in vivo studies, we show that PHF6 regulates the chromatin landscape of B-ALL cells, where it is responsible for maintaining a chromatin state that enables a transformed pre-B-cell identity. PHF6 controls the transcription of target genes by supporting a chromatin configuration that permits or blocks the binding of lineage-specific transcription factors. Furthermore, we show that the associated transcriptional and chromatin state changes that occur in the absence of PHF6 contribute to an emerging mechanism of drug resistance, termed pathway indifference (Cooley et al. 2015). Loss of PHF6 results in chromatin instability and genomic plasticity, which allows malignant cells to reprogram transcriptional outputs and tolerate aberrant lineage signaling.

Loss of Phf6 decreases the leukemogenic potential of B-ALL cells and results in the development of mixedlineage lymphoma in vivo
Recent studies suggest that PHF6 can act as a lineage-specific regulator of tumor growth. However, the molecular mechanisms underlying PHF6's function in hematological malignancies remain widely unclear (Fig. 1A). To evaluate the effects of complete loss of Phf6 on B-ALL growth, we engineered isogenic Phf6 knockout (Phf6 KO ) B-ALL cells using CRISPR-Cas9 (Supplemental Fig. S1A-D; Ran et al. 2013;Sánchez-Rivera and Jacks 2015). To maximize the chance of functionally inactivating the PHF6 protein, we used a domain-focused targeting strategy  by using a single-guide RNA (sgRNA) designed to target the second conserved plant homeodomain (ePHD2), which has been identified as a mutational hot spot in T-cell acute lymphoblastic leukemia (T-ALL) (Van Vlierberghe et al. 2010) and acute myeloid leukemia (AML) (Van Vlierberghe et al. 2011) (Supplemental Fig. S1B-D). The Phf6 KO cells were characterized extensively with regard to their in vitro growth properties, showing no significant differences in their proliferation rates and cell cycle profiles when compared with their wild-type counterparts (Supplemental Fig. S1E,F).
To determine the effects of complete loss of Phf6 on B-ALL growth in vivo, we performed syngeneic trans-plants into immunocompetent recipient mice . Tumor formation in mice injected with 10 6 Phf6 KO cells was significantly delayed compared with mice transplanted with (1) Phf6 WT cells, (2) cells expressing an shRNA targeting Phf6 (shPhf6), and (3) Phf6 KO cells rescued with the Phf6 cDNA (Fig. 1B;Supplemental Figs. S1G,H,S2A). When the number of transplanted cells was reduced 1000-fold, Phf6-deficient cells failed to develop detectable leukemia in recipient animals. Thus, complete loss of Phf6 reduces the fitness and leukemiainitiating capacity of B-ALL cells in vivo.
Notably, mice transplanted with Phf6 WT and Phf6 KO cells developed malignancies with pronounced differences. BCR-ABL1 + B-cell leukemia reproducibly presents with an enlarged spleen and disseminated disease that infiltrates the bone marrow (BM) and peripheral blood, as can be seen upon transplantation with Phf6 WT cells (Williams et al. 2006). However, tumors that developed from Phf6 KO cells pathologically resembled lymphoma, showing prominent lymphadenopathy with marked blood vessels rooting from and to the lymph nodes (LNs), which completely effaced the normal LN histology in recipient mice (Fig. 1C,D). These tumors resulted in size and combined organ weights that were >200% greater than those of wild-type B-ALL recipients (Fig. 1D). Conversely, tumor burden in the blood and the spleen size of mice injected with knockout cells were significantly reduced relative to those from mice injected with wild-type cells ( Fig. 1E; Supplemental Fig. S2B). These changes were confirmed in a second isogenic Phf6 KO B-ALL cell line (Supplemental Fig. S3A,B). Moreover, reintroduction of Phf6 cDNA into Phf6 KO B-ALL cells resulted in leukemia that was indistinguishable in latency from that produced by Phf6 WT B-ALL cells, ruling out off-target effects (Supplemental Figs. S1G,H, S2A). Altogether, the complete loss of Phf6 in B-ALL results in a drastic alteration of the natural ontology of the malignancy (changing from leukemic/disseminated disease to lymphoma) and a consequential extension in survival of recipient mice.
To further investigate the marked differences in disease presentation between Phf6 KO and Phf6 WT cells, we performed immunophenotypic analyses on tumor cells isolated from the LNs and BM of recipient mice. As expected, Phf6 WT tumor cells isolated from the BM and LNs expressed the B-cell markers CD19 and B220 and lacked IgM (Williams et al. 2006). In contrast, tumor cells isolated from Phf6 KO B-ALL recipient mice showed a modest but significant reduction of CD19 and B220 levels (Supplemental Fig. S2C-F). Strikingly, these tumor cells also had high levels of CD4 on the cell surface, indicating that a significant proportion of Phf6 KO B-ALL cells aberrantly expressed a surface marker characteristic of the T-cell lineage (Fig. 1F;Supplemental Fig. S3C;Janeway et al. 1988;Zhu et al. 2010;Rothenberg et al. 2011). Additional immunophenotypic profiling demonstrated that leukemia cells from both genotypes did not express markers of additional lineages, including surface or intracellular CD3 (Borst et al. 1987) or the myeloid/dendritic cell marker CD11b (Supplemental Fig. S4A,B; Bell and Bhandoola 2008;Wada et al. 2008). As observed previously, loss of Phf6 expression in leukemia cells is detrimental exclusively in the in vivo setting (Meacham et al. 2015). In addition, Phf6 KO cells cultured in vitro did not express CD4 (Supplemental Fig. S4C). Importantly, the changes in disease presentation and aberrant expression of CD4 require the complete loss of function of Phf6, as they cannot be recapitulated via potent RNAi-mediated suppression of Phf6 (Supplemental Figs. S2A, S4D-F; Meacham et al. 2015).
To rule out potential effects associated with single-cell cloning, we generated a clonal line from the Phf6 WT population and performed similar survival and immunophenotyping experiments (Supplemental Fig. S4G,H). As expected, mice transplanted with the Phf6 WT clone did not show a significantly different median survival compared with Phf6 WT nonclonal cells. In addition, we did not observe CD4 overexpression in cells isolated from the LNs and BM of mice injected with the wild-type clone, mirroring the disease established by Phf6 WT nonclonal cells. Altogether, these results indicate that PHF6 is critical for growth in vivo as a disseminated disease and suggest a role for PHF6 in maintaining a stable B-cell identity in B-ALL.

Phf6-deficient B-ALL cells have altered expression of genes involved in lineage specificity
Intrigued by the underlying phenotypic plasticity observed in Phf6 KO B-ALL tumors, we sought to explore any potential transcriptional changes behind this phenotype by performing RNA sequencing (RNA-seq) ( Fig. 2; Supplemental Fig. S5). Pairwise differential expression analysis between wild-type and knockout transcriptome profiles showed that Phf6 KO cells had a markedly distinct Quantitative PCR (qPCR) analysis of Phf6 WT (blue) and Phf6 KO (red) cells transduced with empty vector (EV; solid) or a vector expressing Phf6 cDNA (cDNA; dotted). Relative mRNA levels for B-cell-associated genes are shown: Phf6, Cd22, Cd74, Il4ra, Lyn, Ly86, and Blk. (E) Schematic representation of ICA used to identify differential expression signatures (independent components [ICs]) in the integrated RNA-seq data set comprised of Phf6 WT , shPhf6, and Phf6 KO cells. Hinton diagram representation of ICA-derived signatures. Columns denote signatures, and rows denote samples. Colors denote relative directionality of gene expression ([red] upregulation; [green] down-regulation), and the size of each square represents the magnitude of the contribution of each sample to the respective IC. Each signature is two-sided. Vertical boxes denote statistically significant (P = 0.01, Mann-Whitney test) independent components. IC2 identified a Phf6 KO -specific gene signature. (F ) GSEA plot depicting (P = 0.08) enrichment in T-cell signal transduction signature upon Phf6 deletion, as compared with Phf6 WT cells. (NES) Normalized enrichment score. Data represent the mean ± SD. Statistics for these data were calculated with two-sided Student's t-test. ( * * ) P < 0.01; ( * * * ) P < 0.001; ( * * * * ) P < 0.0001; (n.s.) not significant. gene expression pattern when compared with Phf6 WT cells ( Fig Table  2). Differentially expressed genes between Phf6 KO and Phf6 WT cells were significantly enriched in pathways involved in cellular and developmental processes of lymphocytes (Fig. 2B), suggesting that complete loss of Phf6 in B-ALL cells selectively affects the expression of genes involved in lymphoid differentiation and function. Gene set enrichment analysis (GSEA) (Subramanian et al. 2005) of pairwise comparisons revealed that Phf6 KO cells had a significant decrease in the expression of genes associated with B-cell development and function (  Table 2; Hoffmann et al. 2002;Mori et al. 2008). Importantly, the expression of many of the genes that formed the leading-edge cluster of this B-cell signature (Cd74, IL4ra, Lyn, Ly86, and Blk) was restored by introduction of the Phf6 cDNA, indicating that these changes in gene expression are caused by the absence of Phf6 in the knockout cells and not an offtarget effect (Fig. 2D). Using independent component analysis (ICA), we identified a statistically significant gene signature specific to Phf6 KO replicate samples (  Table 2; Hahtola et al. 2006;Jeffrey et al. 2006;Hutcheson et al. 2008;Abbas et al. 2009;Tokoyoda et al. 2009). These analyses suggest that Phf6 KO B-ALL cells adopt a transcriptional program similar to CD4 + T cells upon complete loss of Phf6.
To directly compare murine CD4 + T cells and Phf6 KO B-ALL cells, we applied ICA signature analysis to RNA-seq data from Phf6 WT , shPhf6, and Phf6 KO B-ALL cells and murine CD4 + -single-positive (SP) T cells (Supplemental Fig. S5C; Miyazaki et al. 2015). This analysis unveiled a comparable expression profile between Phf6 KO B-ALL cells and CD4 + T cells that was markedly distinct from shPhf6 and Phf6 WT cells (Supplemental Fig. S5C, D). These results suggest that complete loss of Phf6 in B-ALL cells promotes a transcriptional program that partially resembles CD4 + T cells. It should be noted that the shPhf6 used in our previous study affects in vivo proliferation of B-ALL cells but is unable to recapitulate the breadth of transcriptional changes observed upon complete loss of Phf6 (Supplemental Figs. S4D,E, S5A,D; Meacham et al. 2015). Altogether, these findings further suggest that PHF6 is critical for the maintenance of B-cell identity through the positive transcriptional regulation of genes important for the B-cell state.

Global genomic binding profiles suggest that PHF6 interacts with chromatin
We next sought to determine whether the marked differences in gene expression profiles between knockout and wild-type cells were due to chromatin regulation by PHF6. To gain insight into the distribution of PHF6-bound sites across the B-ALL genome, we performed ChIP-seq (chromatin immunoprecipitation [ChIP] combined with high-throughput sequencing) (Lee et al. 2006) with an antibody against PHF6 (Fig. 3A). We found that PHF6 binds to both gene bodies and proximal promoter/enhancer regions of many annotated genes ( Fig. 3A; Supplemental Table 3). We observed that genes that are differentially expressed in knockout cells had more than twofold enrichment of PHF6 binding at their transcriptional start sites (TSSs) (Supplemental Fig. S6A). In addition, we found that there is enrichment of PHF6 binding within the gene body of lowly expressed genes in Phf6 WT cells (Supplemental Fig. S6B). Together, these data suggest that the genomic location of PHF6 binding might influence the transcriptional output in B-ALL cells.
Previous studies have proposed that PHF6 has the ability to bind both DNA and histone proteins (Todd and Picketts 2012;Liu et al. 2014). Given these contrasting ideas, we decided to examine both modalities, first asking whether PHF6 has any DNA sequence-specific binding properties in B-ALL cells. To do this, we performed motif enrichment analysis (Bailey and Elkan 1994) of the DNA sequences surrounding the PHF6-binding peak summits near promoters of all genes and differentially expressed genes ( Fig. 3B; Supplemental Fig. S6C). We found that no significant de novo motifs could be identified for PHF6 itself (Fig. 3B, left column). The motifs that passed the significance threshold bear a resemblance to motifs bound by known transcription factors (Fig. 3B, right column). Similar results were obtained from a motif search of genome-wide PHF6-binding peaks (Supplemental Fig.  S6C). Since PHF6 lacked any sequence-specific binding properties, we next investigated whether PHF6 is part of a transcription factor complex that could potentially cobind and coregulate gene expression in B-ALL cells.
To do this, we obtained mouse sequences for the ±1-kb regions adjacent to the TSSs of differentially expressed genes and assessed the enrichment of known motifs (those with P-value <0.01 that also have a strong PHF6 signal in ChIP-seq) (https://rdrr.io/bioc/PWMEnrich). Ultimately, 28 motifs (corresponding to 24 unique transcription factors) were identified as binding sites of putative interacting partners of PHF6 (Supplemental Table 3). These motifs were enriched for transcription factor-binding sites important during hematopoiesis and lymphopoiesis, including sites bound by PU.1, NF-kB, EGR1, EBF1, TCF3 (E2A), and TCF12 (HEB) (Supplemental Fig. S6D; Supplemental Table 3; Schebesta et al. 2002;Luo et al. 2004;Carlson et al. 2006;Wildt et al. 2007, Wang et al. 2008Arenzana et al. 2009). Through endogenous coimmunoprecipitation (co-IP) experiments, we confirmed that PHF6 interacts with TCF12 and NF-kB in B-ALL cells (Fig. 3C, top). However, these interactions are mediated in most part via binding to DNA rather than protein-protein interactions, as demonstrated by a notable reduction in immunoprecipitated protein upon addition of the DNA-intercalating agent ethidium bromide (EtBr) (Fig. 3C, bottom). These results show that PHF6 is unable to recognize specific DNA-binding motifs/sequences and therefore is not acting as a canonical transcription factor in B-ALL cells.

PHF6 regulates chromatin organization
We then tested whether PHF6 can regulate gene expression by interacting with histones and subsequently altering the chromatin state. In fact, we observed that the PHF6-binding profile is positively correlated with gene expression (Fig. 3D, left). Specifically, at highly expressed genes, PHF6 binding is enriched at regions flanking the TSS, similar to H3K27ac signal location (Fig. 3D). However, PHF6 signals tend to be more centered at the TSSs of genes with lower expression (Fig. 3D, left). In addition, we found that PHF6-binding enrichment is positively correlated with the presence of activating histone posttranslational modifications, including H3K27ac and H3K4me3, whereas low correlation is observed with the repressive mark H3K27me3 (Fig. 3D [right], E). It had been demonstrated previously that PHF6 can interact with components of the NuRD complex as well several histone proteins (H1.2, H2B.1, H2A.Z, and H3.1) (Todd and Picketts 2012). Therefore, we decided to investigate whether these interactions also occur in B-ALL cells. Through endogenous co-IP experiments, we could not confirm interactions between PHF6 and components of the NuRD complex-RBBP4, HDAC1, or HDAC2-in B-ALL cells (data not shown). However, we were able to confirm a strong interaction between PHF6 and histone H3 (Fig. 3F, top). Importantly, this interaction is independent of the presence of DNA, as treatment of cell lysates with EtBr did not interfere with it (Fig. 3F, bottom). Altogether, these results show that PHF6 exerts transcriptional control of target genes via protein-protein interactions with histones. We conclude that the large transcriptional changes and the associated phenotypic plasticity observed in vivo are caused by loss of the PHF6-chromatin interaction that occurs upon genetic deletion of Phf6 in B-ALL cells.
PHF6 is necessary for the maintenance of chromatin organization at lineage-specific genes Given the lineage promiscuity observed upon loss of Phf6 in vivo, we decided to further compare the transcriptional states of Phf6 WT and Phf6 KO cells with those of B and T cells. We used precurated lists of genes whose expression is unique and distinguishing to CD19 + B cells and CD4 + T cells (Supplemental Table 2; Painter et al. 2011). As Lineage-specific transcriptional programs are often characterized by differential chromatin accessibility (Lara-Astiaso et al. 2014). To determine whether the changes observed in the B and T gene sets are the result of altered chromatin organization, we performed ATACseq (assay for transposase-accessible chromatin [ATAC] with high-throughput sequencing) (Buenrostro et al. 2013) on Phf6 WT and Phf6 KO B-ALL cells (Supplemental Fig.  S7A-F). Phf6 loss results in focal changes in chromatin accessibility, with ∼700 regions in the genome showing statistically significant differences (Supplemental Fig. S7B). These changes are even more pronounced in the promoter regions of genes associated with B and T cells (Supplemental Fig. S7C,E,F). This analysis also demonstrated that changes in chromatin accessibility at promoter regions are predictive of changes in gene expression (Supplemental Fig. S7D,E). Consistent with our phenotypic data, differential chromatin accessibility analysis revealed that regions that become less accessible upon Phf6 loss are enriched for binding motifs of transcription factors required for B-cell development (Fig. 4B, top;Schebesta et al. 2002;Dai et al. 2007;Niebuhr et al. 2013) . Conversely, regions that became more accessible upon Phf6 loss are enriched for motifs of transcription factors associated with the development of T-ALL and AML (Fig. 4B, bottom; Eyquem et al. 2004;Thoms et al. 2011;Kiaii et al. 2013;Smeets et al. 2013). Thus, genomic regions important for B and T lineage genes as well as their regulatory factor-binding sites undergo significant changes in chromatin accessibility upon complete loss of Phf6 in B-ALL cells.
We next asked whether nucleosome positions, in addition to chromatin accessibility, were undergoing significant changes dependent on Phf6 status. To determine nucleosome positioning with high precision and resolution, we applied the NucleoATAC algorithm to our data set ( Fig. 4C; Schep et al. 2015). Globally, there are few differences in nucleosome occupancy between Phf6 WT (Fig. 4C, left, solid line) and Phf6 KO (Fig. 4C, left, dashed line) cells. However, drastic changes in nucleosome occupancy are observed in the lineage-specific gene sets. Focusing on the CD19 + B-cell gene set, we observed an increase in nucleosome occupancy at the TSS and within the gene body upon Phf6 loss (Fig. 4C, middle). When aligned with the promoter-TSS-binding profile of PHF6 in Phf6 WT cells, we observed a large overlap between regions with enriched PHF6 binding and major shifts in nucleosome occupancy (Fig. 4C, gray bar). This finding suggests that PHF6 coordinates nucleosome architecture at B-cell genes in a way that is favorable for transcriptional activity. When PHF6 is lost, nucleosome occupancy is enriched and gene expression decreases (Fig. 4A,C, middle). Focusing on the CD4 + T-cell gene subset, we observed decreased nucleosome occupancy surrounding the TSS in Phf6 KO cells (Fig. 4C, right, dashed line), with preferential depletion at the −1 nucleosome (Fig. 4C, right). Our data are consistent with promoters undergoing increased chromatin accessibility, where the −1 nucleosome undergoes more pronounced depletion (Schep et al. 2015). Furthermore, PHF6 binding is enriched at sites that are undergoing global shifts in nucleosome occupancy (Fig.  4C, right). The chromatin landscape surrounding the TSSs of genes important for CD4 + T cells demonstrates fluidity in Phf6 KO cells, leading to increased gene expression in a subset of T-cell genes (Fig. 4A) and presentation as a mixed-lineage lymphoma in vivo (Fig. 1). The shifts in nucleosome occupancy that we observed upon complete loss of Phf6 indicate that B-ALL cells are undergoing lasting and stable transcriptional changes resulting from alterations of the chromatin architecture, allowing for genomic (as well as phenotypic) plasticity. Specific genes undergoing changes in chromatin accessibility at regulatory elements include B-cell-specific genes Btg2 and Cd74 (both of which become less accessible upon Phf6 loss) and T-cell lineage genes Cd4 (distal enhancer) and Gata3 (which become more accessible upon Phf6 loss) (Supplemental Fig. S8A,B). Importantly, we confirmed enrichment of PHF6 binding at these target genes through ChIP combined with quantitative PCR (ChIP-qPCR) analysis (Supplemental Fig. S8C,D), suggesting that the observed shifts in chromatin accessibility are dependent on the presence of PHF6. In conclusion, we show that PHF6 is necessary for the maintenance of a chromatin state that is instrumental to B-cell identity and that Phf6 loss leads to the acquisition of a permissive/open state at loci of alternate lineages, allowing for the engagement of aberrant transcriptional programs that direct the resulting phenotypic plasticity.

Instability in the chromatin landscape allows for aberrant lineage signaling
Given the genomic plasticity observed in our Phf6 KO B-ALL cells and the changes in chromatin accessibility detected at regions of T-cell transcription factor-binding sites upon Phf6 loss, we hypothesized that the genome of Phf6 KO cells may now be permissive to aberrant oncogenic signaling emanating from T-cell lineage factors, such as NOTCH1. Activating mutations in NOTCH1 are well documented in >60% of cases of T-ALL (Belver and Ferrando 2016), where PHF6 is often found to be comutated with NOTCH1 (Wang et al. 2011;Li et al. 2016). Therefore, we tested the effects of overexpression of the NOTCH1 intracellular domain (NICD) in Phf6 WT and Phf6 KO B-ALL cells. As expected, we observed that NOTCH1 signaling dramatically inhibits the growth of Phf6 WT B-ALL cells ( Fig. 5A; Rothenberg 2014). However, Phf6 KO cells were viable upon NICD overexpression, with growth rates equivalent to Phf6 WT and Phf6 KO vector control cell lines (Fig. 5A). Transplantation of Phf6 KO + NICD cells into immunocompetent syngeneic mice gives rise to a latent lymphoma with extensive thymic infiltration and elevated levels of CD4 expression characteristic of T-cell malignancies ( Fig. 5B-D). In conclusion, PHF6 deficiency in B-ALL cells creates a permissive chromatin state that allows transcription factors from alternate lineages to bind to their respective genomic loci, which are normally in a closed conformation in wild-type cells. Enforced expression of NOTCH1 (T-cell transcription factor) takes advantage of the chromatin fluidity characteristic of knockout cells, where Phf6 KO tumors are driven more toward a "T-celllike" state instead of a mixed-lineage phenotype.
Phenotypic plasticity underlies differential responses to anti-cancer treatments in vivo Clinically, patients with B-cell and T-cell leukemia and lymphoma undergo different treatment strategies.
BCR-ABL1-driven B-cell leukemia is often treated with targeted kinase inhibitors, whereas T-cell leukemia and lymphoma are treated with intensive chemotherapy regimens (Schultz et al. 2009;Litzow and Ferrando 2015). To further understand the shift from a disseminated leukemia to a mixed-phenotype solid lymphoma that results from complete loss of Phf6 in B-ALL cells, we examined the response of these tumors to targeted therapy (ponatinib) and frontline chemotherapy (doxorubicin).
Unbiased transcriptome analysis shows decreased dependency on BCR-ABL1 signaling pathways, the driving oncogene in this B-ALL model ( Fig. 6A; Williams et al. 2006;Shamroe and Comeau 2013). Consistent with this, we observed a marked difference in the response to ponatinib (third-generation BCR-ABL1 inhibitor) between Phf6 WT and Phf6 KO tumors in vivo but not in vitro ( Fig. 6B; Supplemental Fig. S9A). Mice transplanted with Phf6 KO cells and treated with ponatinib have a median survival of 15 d, while their wild-type counterparts survived >60 d after treatment (Fig. 6B). Thus, the lineage plasticity that arises in vivo upon Phf6 loss drives a significantly decreased response to ponatinib. Furthermore, we examined the response of Phf6 WT and Phf6 KO tumors to the chemotherapeutic agent doxorubicin. We found that mice transplanted with Phf6 KO cells are exquisitely sensitive to doxorubicin treatment in vivo, as demonstrated by an additional 13-d extension in median survival (essentially doubling their life span) when compared with mice injected with Phf6 WT cells ( Fig. 6C; Supplemental  Fig. S9B,C). In conclusion, we found that complete loss of Phf6 in B-ALL cells changes not only the chromatin landscape of these tumors but also the phenotypic state, which is reflected in the formation of a different disease with vastly distinct responses to anti-cancer therapies. These findings support a possible underlying mechanism for the examples of lineage plasticity and drug resistance observed in a variety of malignancies and demonstrate that modulation of the chromatin landscape through the loss of regulation by a single factor (such as PHF6) may play a crucial role in acquired resistance.

Discussion
The transcriptional regulation of lymphocyte development via lineage-specific factors is well characterized, and the perturbation of this process can promote leukemogenesis (Mullighan et al. 2007). However, the dynamic regulation of chromatin states underlying transcriptional activity and its contribution to cancer are yet to be elucidated. In this study, we show that PHF6 is a novel chromatin state regulator important for the maintenance of a chromatin landscape conducive to B-cell identity in B-ALL. Through integrated genomics (RNA-seq, ChIP-seq, and ATAC-seq) and in vivo studies, we demonstrate the crucial role that PHF6-mediated chromatin modulation plays in genomic plasticity, cell identity, leukemogenesis, and response to therapy. Our data convincingly show that Phf6 loss leads to a chromatin environment that tolerates aberrant lineage and oncogenic signaling. Moreover, we demonstrate how malignant cells can hijack chromatin instability to reprogram transcriptional output and cell identity. These findings propose the first mechanistic insights into PHF6's role in a lymphoid malignancy. We show that PHF6 binds to nucleosomes at specific genomic loci and orchestrates distinct transcriptional programs. It stabilizes chromatin accessibility at B-cellspecific genes, while T-cell transcription factor-binding sites are concealed in compacted chromatin (Fig. 7A). Upon loss of this single factor, chromatin architecture Phf6 WT + NICD cDNA (blue; down triangle), and Phf6 KO + NICD cDNA (red; down triangle) cells. n = 3. (B) Kaplan-Meier survival analysis of mice injected with 10 6 B-ALL cells of the indicated genotypes infected with control vector or activated NICD vector. The number (n) of mice per genotype analyzed is shown. Statistical analysis (log-rank test, Mantel-Cox) was performed for the different groups in comparison with mice injected with Phf6 WT + control vector cells. P-values are shown for the comparisons. (C ) Quantification of the combined thymus weight of Phf6 WT + control vector (blue; n = 4), Phf6 KO + control vector (red unfilled; n = 6), and Phf6 KO + NICD cDNA (red patterned; n = 7) recipients. (D) Bar graphs showing the percentage of the CD4 + fraction among mCherry + cells isolated from Phf6 WT + control vector (blue; n = 5), Phf6 KO + control vector (red unfilled; n = 4), and Phf6 KO + NICD cDNA vector (red patterned; n = 6) tumors from LNs. Data represent the mean ± SD in C and D. Statistics were calculated with two-sided Student's t-test. ( * ) P < 0.05; ( * * ) P < 0.01; ( * * * * ) P < 0.0001. around B-cell-and T-cell-specific genes undergoes dramatic remodeling, resulting in focal genomic plasticity and acquisition of T-cell lineage markers in B-ALL cells (Fig. 7B). Given that PHF6 is frequently inactivated in T-ALL (∼30%) (Van Vlierberghe et al. 2010), we speculate that T-cell malignancies benefit from the permissive chromatin landscape that results upon PHF6 loss. Our data further support the possibility that a significant portion of T-ALLs (∼30%) (Van Vlierberghe et al. 2010) may have once started as a B-cell progenitor/precursor that acquired a mutation in PHF6 during early stages of leukemogenesis and consequently forced the cell of origin into a "pseudo-T-cell state" that then presented as a T-cell malignancy.
Lineage switching is a rapidly emerging mechanism of resistance to targeted therapies. A variety of cancers (including lung cancer, prostate cancer, and B-cell leukemia) can acquire resistance to therapy by relapsing as a histologically distinct malignancy (Sequist et al. 2011;Yu et al. 2013;Oser et al. 2015;Gardner et al. 2016;Jacoby et al. 2016;Ku et al. 2017;Mu et al. 2017). The first functional studies investigating this process as a driver of drug resistance in prostate cancer identified epigenetic reprogramming proteins (SOX2 and EZH2) as key effectors of this phenomenon Mu et al. 2017). In our study, we further demonstrate that active maintenance of chromatin architecture (by PHF6, a novel chromatin regulator) is also critical in the response to targeted therapy in B-cell leukemia. We postulate that the chromatin landscape is a pivotal component in the response of many cancer types to targeted treatments.
Here we identifed the function of PHF6 in B-ALL and ascribed its role in regulating chromatin accessibility to lineage-specific transcription factors. We showed that loss of Phf6 results in disruption of lineage differentiation via gross changes in chromatin accessibility, engagement of alternative transcriptional programs, altered disease presentation in vivo, and tolerance of aberrant oncogenic signaling. We established that the proper chromatin landscape is essential for maintaining cell identity and that loss of Phf6 can cause changes in chromatin accessibility and nucleosome positioning that result in lineage promiscuity. These findings highlight a mechanism of gene expression maintenance via chromatin landscape modulation that underlies the developmental plasticity characteristic of hematopoietic malignancies.

Cell culture
Murine BCR-ABL1 + ; p19 −/− ; mCherry + B-ALL cells (Williams et al. 2006) were cultured in RPMI-1640 (Corning) supplemented with 10% FBS, 4 mM L-glutamine, 50 µM 2-mercaptoethanol, 100 U/mL penicillin, and 100 µg/mL streptomycin. Retroviral particles were generated and packaged using the Phoenix cell system (G. Nolan, Stanford University). Briefly, a calcium phosphate transfection of 10 µg of plasmid DNA and 5 µg of helper plasmid (pCMV-Gag-Pol) was used. Transduction efficiencies were assessed 48 h after infection by flow cytometry. To generate Phf6 KO cells, sgPhf6-pX458 plasmid was transfected (Lipofectamine 3000, Life Technologies) into B-ALL cells. Transfected cells were single-cell-sorted into 96-well plates based on GFP expression. Cell proliferation assay was performed by plating B-ALL cells in triplicate (1 million cells per well in 3 mL of medium in a six-well plate), counting every 24 h by Trypan blue exclusion, and replating the initial density. This was performed for 7 d.

Drug response analysis
Cells (10,000 per well) were plated in 96-well plates, and medium containing ponatinib (LC Laboratories, AP24534) or doxorubicin (Sigma-Aldrich, D1515) was added to achieve the indicated final concentrations. Cell viability was assessed 48 h after treatment by propidium iodide (Sigma-Aldrich, 81845) exclusion and flow cytometry. Inhibitor dilutions were made in cell medium immediately before use. The relative viability compared with controls diluted in equivalent medium + DMSO is shown.

Co-IP
Antibodies were precleaned before attaching to magnetic beads using the Pierce antibody cleanup kit (ThermoFisher Scientific, 44600). A total of 10 µg of antibody was covalently attached to magnetic beads (ThermoFisher Scientific, 88828) and incubated with 1 mg of protein for 2 h at room temperature while rotating (or overnight at 4°C). Immunoprecipitation was performed according to the manufacturer's directions (Pierce Direct magnetic immunoprecipitation/co-IP kit, Thermo Fisher Scientific). For DNA-dependent interactions, 50 µg/mL EtBr was added, and lysates were incubated for 30 min on ice. Samples were centrifuged, and the supernatant was used for immunoprecipitations. qPCR RNA from 2 × 10 6 B-ALL cells was prepared using RNeasy minikit (Qiagen). Synthesis of cDNA was performed using M-MLV reverse transcriptase (Life Technologies, 28025) with oligo(dT) 20 primer. qPCR was done in an Applied Biosystems StepOnePlus machine with TaqMan fast universal PCR master mix (Life Technologies, 4352042). Data were analyzed using the comparative ΔC T method and normalized to the levels of Gapdh. The TaqMan gene expression assays (Life Technologies) used were Phf6 (Mm00804415_m1), Blk (Mm00432077_m1), Lyn (Mm01217488_m1), Gapdh (Mm99999915_g1), and Cd22 (Mm00515432_m1).

Genomic DNA isolation and sequencing
Genomic DNA from B-ALL cells was isolated using the blood and cell culture DNA minikit (Qiagen) following the manufacturer's guidelines. PCR products for sequencing were amplified using HerculaseII Fusion DNA polymerase (Agilent) (see Supplemental  Table 1 for primers used), gel-purified, cloned into pCR4-TOPO TA vector (Life Technologies), and subsequently sequenced.

Animal experiments
All animal studies described in this study were approved by the Massachusetts Institute of Technology Institutional Animal Care and Use Committee. For syngeneic transplants, 10 3 or 10 6 murine BCR-ABL1 + p19 −/− B-ALL cells (Williams et al. 2006) were injected via tail vein into syngeneic C57BL/6J (The Jackson Laboratory, 000664) recipient mice at the age of 6-8 wk. Treatment with 30 mg/kg ponatinib (administered via oral gavage, treated once daily for four consecutive days), 10 mg/kg doxorubicin (administered via immunoprecipitation, treated once; Nova-Plus), or vehicle (25 mM citric acid buffer) was initiated upon significant disease presentation. Mice were observed daily and sacrificed when moribund (dehydration, ruffled fur, respiratory distress, and poor motility). Survival curves were generated using GraphPad Prism version 6.0 software, and the Mantel-Cox test was applied to pairwise comparisons of survival data. No statistical methods were used to predetermine sample size. Investigators were not blinded to group allocation during experiments and outcome assessment. Randomization was not used because all animals used in this study were of similar sex, age, and strain background.
Organ processing and cell preparation for flow cytometry When mice reached morbidity, they were sacrificed, and lymphoid organs were collected. Briefly, cells derived from the BM were obtained by flushing the femora and tibiae with B-ALL medium. LNs, spleens, and thymi were isolated; weighed for mass measurements; ground; and filtered to obtain single-cell suspension. Subsequently, red blood cells were lysed with ACK lysing buffer (Life Technologies, A10492-01).

Immunostaining
For cell surface immunophenotyping, leukemia cell suspensions (cultured and isolated from mice) were collected and treated with the indicated antibodies. Briefly, 1 × 10 6 to 2 × 10 6 cells in 100 µL of 10% FBS in PBS were incubated with antibody for 1 h at room temperature in the dark. Antibodies were used at a 1:100 dilution. Stained samples were analyzed by flow cytometry.
For intracellular immunophenotyping, leukemia cell suspensions (cultured and isolated from mice) were collected, resuspended in PBS, and fixed with 4% formaldehyde (Tousimis, 1008A) solution for 10 min at 37°C. Cells were permeabilized with icecold methanol for 30 min on ice.
Cell cycle analysis FITC BrdU flow kit (BD Biosciences, 559619) was used for cell cycle analysis. Briefly, 1 × 10 6 B-ALL cells plated in triplicate were labeled with 10 µM BrdU for 30 min. Cells were subsequently fixed and stained with anti-BrdU and 7-AAD and analyzed by flow cytometry (LSRFortessa, BD Biosciences).

Flow cytometry
FACSAria (BD Biosciences) was used for fluorescence-activated cell sorting, and LSRFortessa (BD Biosciences) was used for flow cytometry analysis. Data were analyzed with FlowJo software (TreeStar version 10).

Histology and immunohistochemistry
For histological and immunohistochemistry analyses, mice were euthanized by carbon dioxide asphyxiation. Lymphoid tissues were harvested, fixed overnight with 10% neutral buffered formalin (VWR), transferred to 70% ethanol solution, and subsequently embedded in parafilm. Sections were cut at a thickness of 10 µm and stained with hematoxylin and eosin (H&E) for pathological assessment. Immunohistochemistry was performed on a Thermo Autostainer 360 machine. Slides were antigen-retrieved using Thermo citrate buffer (pH 6.0) in the pretreatment module. Sections were treated with Biocare rodent block, primary antibody, and anti-rabbit HRP-polymer (Vector Laboratories). The slides were developed with Thermo Ultra DAB and counterstained with hematoxylin in a Thermo Gemini stainer, and coverslips were added using the Thermo Consul coverslipper.

RNA-seq library preparation
cDNA for RNA-seq libraries was prepared using TruSeq mRNA library preparation kit (Illumina). Illumina libraries were produced using SPRIworks (Beckman-Coulter Genomics) with a 200-to 400-base-pair (bp) size selection and enriched with 14 cycles of PCR. Library quality was determined by qPCR and on the fragment analyzer (Advanced Analytical) and loaded onto two lanes of 40-nucleotide (nt) single-end sequencing on a HiSeq2000 system (Illumina).

RNA-seq data analysis
Illumina HiSeq2000 40-nt single-end reads were mapped to the University of California at Santa Cruz (UCSC) mm9 mouse genome build (http://genome.ucsc.edu) using RSEM (Li and Dewey 2011). Raw estimated expression counts were upper-quartilenormalized to a count of 1000 (Bullard et al. 2010). Given the complexity of the data set in terms of a mixture of different conditions, a high-resolution signature discovery approach was used to characterize global gene expression profiles. ICA, an unsupervised blind source separation technique, was used on this discrete count-based expression data set to elucidate statistically independent and biologically relevant signatures. ICA is a signal processing and multivariate data analysis technique in the category of unsupervised matrix factorization methods. Conceptually, ICA decomposes the overall expression data set into independent signals (gene expression patterns) that represent distinct signatures. High-ranking positively and negatively correlated genes in each signature represent gene sets that drive the corresponding expression pattern (in either direction). Signatures were visualized using the sample to signature correspondence schematic afforded by Hinton plots, where colors represent directionality of gene expression (red indicates up-regulated, and green indicates down-regulated), and the size of each rectangle quantifies the strength of a signature (column) in a given sample (row). Each signature is two-sided, allowing for identification of up-regulated and downregulated genes for each signature within each sample. Using input data consisting of a gene-sample matrix, ICA uses higher-order moments to characterize the data set as a linear combination of statistically independent latent variables. These latent variables represent independent components based on maximizing non-Gaussianity and can be interpreted as independent source signals that have been mixed together to form the data set under consideration. Each component includes a weight assignment to each gene that quantifies its contribution to that component. Additionally, ICA derives a mixing matrix that describes the contribution of each sample toward the signal embodied in each component. This mixing matrix can be used to select signatures among components with distinct gene expression profiles across the set of samples. The R implementation of the core JADE (Joint Approximate Diagonalization of Eigenmatrices) algorithm (Rutledge and Bouveresse 2013; https://cran.r-project.org/ web/packages/JADE/index.htm; https://www.bioconductor.org/ packages/release/bioc/html/MineICA.html) was used along with custom R utilities. The P-value for the knockout-specific (IC2) signature was P = 0.0119 (Mann-Whitney U-test directional P-value). Targeted pairwise differential expression analyses were conducted using EBSeq version 1.4.0 (Leng et al. 2013). Differentially expressed genes from the pairwise comparison between Phf6 KO and Phf6 WT B-ALL cells were determined using a significance level of false discovery rate (FDR) 0.05 and fold change >1.5. All RNA-seq analyses were conducted in the R statistical programming language (http://www.r-project.org). GSEA was carried out using the preranked mode with default settings (Subramanian et al. 2005). Heat maps were generated using the Heatplus package in R. Integrated RNA-seq signature analysis of B-ALL cells and a data set from CD4 + SP cells (Gene Expression Omnibus [GEO] accession code GSE64779, samples GSM1580493-GSM1580497) (Edgar et al. 2002;Barrett et al. 2012;Miyazaki et al. 2015) was performed using ICA. All gene lists, signatures, and sets used in this study are in Supplemental Table S2. A two-dimensional expression principal component (principal component analysis [PCA]) plot for each group of cells was generated by plotting the first two principal components of expression (explaining 75% and 11% of expression variance, respectively). Gene ontology (GO) analysis was done using DAVID, and top GO and PANTHER terms were plotted against their corresponding P-values (log-scale).

ChIP-seq
ChIP was performed as described before (Lee et al. 2006) with a few adaptations. A total of 150 × 10 6 cells, grown to a density of 1 × 10 6 cells per milliliter, was cross-linked for 10 min at room temperature by the addition of one-tenth of the volume of 11% formaldehyde solution (11% formaldehyde, 50 mM HEPES at pH 7.3, 100 mM NaCl, 1 mM EDTA at pH 8.0, 0.5 mM EGTA at pH 8.0) to the growth medium followed by 5 min of quenching with 125 mM glycine. Cells were washed twice with PBS, the supernatant was aspirated, and the cell pellet was flash-frozen in liquid nitrogen. Frozen cross-linked cells were stored at −80°C. Onehundred microliters of Protein G Dynabeads (Life Technologies) was blocked with 0.5% (w/v) BSA in PBS. Magnetic beads were bound with 10 µg of anti-H3K27ac (Abcam, ab4729), anti-H3K27me3 (Abcam, ab6002), and anti-PHF6 (Bethyl Laboratories, A301-451A). Nuclei were isolated as described previously (Lee et al. 2006) and sonicated in lysis buffer (20 mM Tris-HCl at pH 8.0, 150 mM NaCl, 2 mM EDTA at pH 8.0, 0.1% SDS, 1% Triton X-100) on a Misonix 3000 sonicator for 10 cycles of 30 sec each on ice (18-21 W) with 60 sec on ice between cycles. Sonicated lysates were cleared once by centrifugation and incubated overnight at 4°C with magnetic beads bound with antibody to enrich for DNA fragments bound by the indicated factor. Beads were washed with wash buffer A (50 mM HEPES-KOH at pH 7.9, 140 mM NaCl, 1 mM EDTA at pH 8.0, 0.1% sodium deoxycholate, 1% Triton X-100, 0.1% SDS), buffer B (50 mM HEPES-KOH at pH 7.9, 500 mM NaCl, 1 mM EDTA at pH 8.0, 0.1% sodium deoxycholate, 1% Triton X-100, 0.1% SDS), buffer C (20 mM Tris-HCl at pH 8.0, 250 mM LiCl, 1 mM EDTA at pH 8.0, 0.5% sodium deoxycholate, 0.5% IGEPAL CA-630, 0.1% SDS), and buffer D (TE with 50 mM NaCl) sequentially. DNA was eluted in elution buffer (50 mM Tris-HCl at pH 8.0, 10 mM EDTA, 1% SDS). Cross-links were reversed overnight. RNA and protein were digested using RNase A and Proteinase K, respectively, and DNA was purified with phenol-chloroform extraction and ethanol precipitation. Purified ChIP DNA was used to prepare Illumina multiplexed sequencing libraries. Libraries were prepared following the TruSeq DNA sample preparation kit version 2 (Illumina). Amplified libraries were size-selected using a 2% gel cassette in the Pippin preparation system (Sage Science) set to capture fragments between 200 and 400 bp. Libraries were quantified by qPCR using the Illumina library quantification kit (KAPA Biosystems) according to the manufacturer's guidelines. Libraries were sequenced on the Illumina HiSeq2500 for 40 nt in single-read mode.

ChIP-seq data analysis
To compare RNA-seq with ChIP-seq data sets, normalized RNAseq tag counts for each gene were used to plot a heat map showing the expression levels of these genes ordered by fold change values. Among these genes, 1127 symbols were mappable to ENSEMBL mouse gene IDs (unmappable gene symbols corresponded mostly to those that are not well characterized) and were used for mapping to genomic coordinates. Sequencing reads were mapped to the UCSC mm9 mouse genome using Bowtie2 (Langmead et al. 2009;Langmead and Salzberg 2012) with at most one mismatch allowed in seed alignments. Unmapped reads were removed, and alignments were output in BAM files for peak calling with the MACS software (Zhang et al. 2008). PHF6 and histone modification peaks were called using a P-value threshold of 1 × 10 −6 . The fraction of reads falling within the called peaks (FRiP) for PHF6 ChIP-seq data was 25.9% (8,333,082 out of 32,143,465 uniquely mappable reads), showing high ChIP quality. To examine whether PHF6-binding strength correlates with gene expression levels, RNA-seq data from Phf6 WT cells were sorted and grouped by expression into three bins corresponding to high, medium, and low expression by separating into three equally sized bins. Average log 2 fold change values over input signals were plotted as a metagene track for each gene set. Plots for H3K27ac signal enrichment were also generated to confirm promoter activity for each gene set. To determine PHF6 binding at promoter/enhancer regions of the differentially expressed genes, genomic sequences were obtained for the ±5-kb region surrounding the TSSs for these genes from the UCSC mm9 genome assembly and binned into 50bp bins. ChIP-seq RPM (reads per million) values for PHF6, H3K27ac, H3K27me3, and H3K4me3 (publicly available ChIPseq data sets were obtained as Sequence Read Archive-lite files, GEO accession code GSE66234, data sets GSM1617788-GSM1617789) (Edgar et al. 2002;Barrett et al. 2012;Wang et al. 2015) at these promoter bins were divided by the corresponding ChIP input (control) reads with a pseudocount of one and then log 2 -transformed to show fold enrichment. To show correlation between PHF6 and histone modification signals, Pearson's correlation coefficients between corresponding pairs of columns in the heat maps were calculated. The MEME suite (Bailey and Elkan 1994) was used for de novo motif discovery of sequences surrounding PHF6-binding peak summits. A 100-nt window upstream of and downstream from the peak summits was set to obtain sequences from the UCSC mm9 genome assembly. Among all 77,749 PHF6 summits called by the MACS pipeline (Zhang et al. 2008), 6037 fell within the proximal regulatory regions (±5 kb surrounding the TSS) of differentially expressed genes, covering a total of 1104 (out of 1123) unique promoters. For each promoter region, the PHF6 summit was selected with the highest average peak signal for motif analysis, resulting in 1044 unique summits and their surrounding sequences as input for the MEME motif discovery tool (Bailey and Elkan 1994). To see whether PHF6 displays any bias in sequence specificity at the differentially expressed gene promoters, the top 1044 among all PHF6 summits were selected and run in motif discovery in parallel. A motif length of eight was used and selected the top 10 significant motifs returned by MEME (Bailey and Elkan 1994). The motifs thus discovered were then tested for similarity to known motifs (as in the JASPAR CORE 2014 vertebrates [Mathelier et al. 2013], UniProbe mouse [Hume et al. 2015], and other human and mouse motifs [Jolma et al. 2013]) using the Tomtom tool (Gupta et al. 2007) with a minimum motif overlap of one and Pearson distance as the distance metric for motif similarity. To identify other transcription factors that cobind with PHF6, mm9 sequences were obtained for ±1-kb regions around the TSSs of differentially expressed genes and tested for known motif enrichment using the Bioconductor PWMEnrich package (https://bioconductor.org/packages/release/bioc/html/PWMEnri ch.html). A default Mus musculus genomic background and the MotifDb database of motifs (https://rdrr.io/bioc/MotifDb) included in the package were used, and a P-value threshold of 0.01 was set for reporting significant transcription factors whose motifs were enriched in the selected regions. Input sequences were randomly shuffled 100 times, motif enrichment was performed for each shuffled sequence set, and motifs that came up as significant (P < 0.001) in >5% of the shuffled sets were considered to be false positive enrichments. A total of 28 motifs corresponding to 24 unique transcription factors passed the filtering step and were reported as potential cobinding partners of PHF6 at the promoters of differentially expressed genes (Supplemental Table 3). Gene functions were determined by using the PANTHER classification system (Thomas et al. 2003;Mi et al. 2004).
Chromatin accessibility by ATAC-seq ATAC-seq libraries from Phf6 WT and Phf6 KO B-ALL cells were constructed as reported previously (Buenrostro et al. 2013). For the ATAC-seq data analysis, mapped ATAC reads were summarized into 500-bp bins. The DESeq2 package in R (Love et al. 2014) was then applied to the binned reads to analyze for differential chromatin accessibility between Phf6 wild-type and knockout cells. To call differentially accessible chromatin regions, we used a fold change cutoff of >2 and a FDR cutoff of <0.001. GREAT (genomic regions of enrichment annotations tool) (McLean et al. 2010) was used to analyze for enriched gene sets and other genomic features in the regions that were more or less accessible in Phf6 KO cells separately. To test enrichment of known and de novo sequence motifs within these differentially accessible chromatin regions, we used the HOMER motif analysis toolkit (Heinz et al. 2010) on these regions using all mappable ATAC regions as background and the UCSC mm9 genome, with all other parameters set to default values.
NucleoATAC software was used with default parameters to call nucleosome signals from raw ATAC-seq signals in Phf6 WT cells (Schep et al. 2015(Schep et al. , 2017. Background-corrected nucleosome signals ±1 kb from TSSs annotated in the mm9 genome were then stacked into a heat map with rows corresponding to individual TSSs ordered by gene expression in Phf6 WT cells. To visualize PHF6 occupancy at these sites, a heat map of normalized PHF6 signals in the same row order was also generated. ATAC-seq raw data and processed TSS nucleosome signals are available in the GEO database with accession number GSE98716 under superseries GSE77457.

Statistical analysis
Results are expressed as means ± standard deviation. If not stated otherwise, statistical significance was determined with Prism software (GraphPad version 6.0) by comparison of mean values by the two-tailed unpaired Student's t-test on two experimental conditions, with P < 0.05 considered statistically significant. Statistical significance levels are designated as follows: P < 0.05 ( * ), P < 0.01 ( * * ), P < 0.001 ( * * * ), P < 0.0001 ( * * * * ), and not significant (n.s.). Survival curves were generated using Prism software (GraphPad version 6.0), and the Mantel-Cox test was applied to pairwise comparisons of survival data. No statistical methods were used to predetermine sample size.

Accession numbers
All RNA-seq and ChIP-seq data sets have been deposited into the NCBI GEO repository under BioProjectID GSE74878. The ATACseq data set and processed TSS nucleosome signals are available in the GEO database with accession number GSE98716 under superseries GSE77457.


extfieldsofstudy: Biology Medicine

text:
Structure, Catalysis, and Inhibition of OfChi-h, the Lepidoptera-exclusive Insect Chitinase*

Chitinase-h (Chi-h) is of special interest among insect chitinases due to its exclusive distribution in lepidopteran insects and high sequence identity with bacterial and baculovirus homologs. Here OfChi-h, a Chi-h from Ostrinia furnacalis, was investigated. Crystal structures of both OfChi-h and its complex with chitoheptaose ((GlcN)7) reveal that OfChi-h possesses a long and asymmetric substrate binding cleft, which is a typical characteristics of a processive exo-chitinase. The structural comparison between OfChi-h and its bacterial homolog SmChiA uncovered two phenylalanine-to-tryptophan site variants in OfChi-h at subsites +2 and possibly −7. The F232W/F396W double mutant endowed SmChiA with higher hydrolytic activities toward insoluble substrates, such as insect cuticle, α-chitin, and chitin nanowhisker. An enzymatic assay demonstrated that OfChi-h outperformed OfChtI, an insect endo-chitinase, toward the insoluble substrates, but showed lower activity toward the soluble substrate ethylene glycol chitin. Furthermore, OfChi-h was found to be inhibited by N,N′,N″-trimethylglucosamine-N,N′,N″,N″′-tetraacetylchitotetraose (TMG-(GlcNAc)4), a substrate analog which can be degraded into TMG-(GlcNAc)1–2. Injection of TMG-(GlcNAc)4 into 5th-instar O. furnacalis larvae led to severe defects in pupation. This work provides insights into a molting-indispensable insect chitinase that is phylogenetically closer to bacterial chitinases than insect chitinases.

Insect chitinases belong to glycoside hydrolase family 18 (GH18) 3 and can be classified into 11 groups based on sequence similarity and domain architecture (1,2). Among them, chitinase-h (Chi-h) is noteworthy because its members are only found in lepidopteran insects, one of the most destructive crop pest (3,4). Chi-hs and their bacterial homologs share more than 70% sequence identity, suggesting that a gene horizontal transfer occurred between these two phylogenetic-distant species (4,5).
The physiological role of Chi-h in lepidopteran insects is mostly related to cuticle chitin degradation. During molting and metamorphosis, lepidopteran insects secrete molting fluid, which contains three chitinases (EC 3.2.1.14, group I chitinase (ChtI), group II chitinase (ChtII) and Chi-h), one N-acetyl-Dhexosaminidase (EC 3.2.1.52, Hex), and several kinds of proteases to degrade and shed the old cuticle (6). Chitinases degrade polymeric chitin into chitobiose and chitotriose, which are then further degraded into N-acetyl-D-glucosamine (GlcNAc) by Hex (7). Compared with the extensively studied ChtI (8 -19), there is limited information about the function of ChtII and Chi-h. RNAi of SeChi-h from Spodoptera exigua led to molting deficiency and death indicating that Chi-h is indispensable for molting (17). The spatial and temporal expression patterns of Chi-hs from Bombyx mori (4,5) and S. exigua (17) are similar to that of ChtI but different from ChtII. This suggests that Chi-h and ChtI may work synergistically throughout insect development.
Several crystal structures of GH18 chitinases have been determined from archaea (20), bacteria (21)(22)(23)(24)(25)(26)(27)(28)(29), fungi (30 -34), plants (35)(36)(37)(38)(39)(40)(41), and mammals (42,43). These structures show that although all of the GH18 chitinases use the same catalytic mechanism, they have large discrepancies in the shape of the substrate binding cleft. The crystal structure of OfChtI gave structural evidence that ChtI has a long and open-ended substrate binding cleft with symmetrically distributed subsites that is believed to be a structural characteristic of an endo-acting chitinase (44). According to a structure-based sequence alignment, we found that Chi-h does not contain such a substrate binding cleft but contains a long substrate binding cleft with asymmetrically distributed subsites, a structural characteristic of the processive exo-acting chitinase SmChiA from Serratia marcescens (45). Thus, it is unlikely that Chi-h would be able to act through the same mode of action as ChtI. Revealing the structure of Chi-h will increase our knowledge of why and how lepidopteran insects acquired Chi-h for old cuticle shedding.
In this study, OfChi-h, a Chi-h from the pest Ostrinia furnacalis, was investigated. The crystal structures of OfChi-h and OfChi-h in complex with a substrate analog (GlcN) 7 were obtained and resolved. Through structure-based comparison as well as biochemical characterization, we demonstrate that Chi-h acts synergistically with ChtI to degrade cuticle chitin. Moreover, N,NЈ,NЉ-trimethylglucosamine-N,NЈ,NЉ,NЉЈ-tetraacetylchitotetraose (TMG-(GlcNAc) 4 ), an inhibitor against chitinolytic Hexs (46), inhibits OfChi-h, providing a valuable clue for designing efficient inhibitors. Because Chi-h is absent in most beneficial insects including parasitic wasps and bees, this work will also help develop novel and eco-friendly agrochemicals to protect plants and defend economical loss.

Results
Overall Structure of OfChi-h-The structure of OfChi-h was determined by molecular replacement using the bacterial SmChiA as a search model and was refined to a resolution of 3.2 Å (Table 1). OfChi-h adopts a compact and elongated structure with two domains: domain I (residues 18 -125) and domain II (residues 151-553) (Fig. 1A). According to SCOP (Structural Classification of Proteins) classification (47), domain I is different from domain II. Domain I (fibronectin III domain) is an immunoglobulin-like ␤-sandwich domain comprised of eight ␤-strands. And domain II (catalytic domain) is a (␤/␣) 8 -barrel composed of eight ␤-strands and eight ␣-helices. A chitinase insertion domain (residues 437-509), which consists of five antiparallel ␤-strands flanked by two ␣-helices, is observed in the domain II (48). Domain I and domain II are connected via a 25-amino acid linker (residues 126 -150) and interact with each other via a motif consisting of two antiparallel ␤-strands and one short ␣-helix (residues 34 -51). Two N-glycosylation sites (Asn 391 and Asn 456 ) were observed (Fig. 1A).
One of the most striking features of OfChi-h is a number of aromatic residues lining the groove starting from the far end of domain I and ending at the far end of the substrate binding cleft of domain II (Fig. 1B). They are nine in total, including Trp 27 , Trp 63 , Trp 238 , Trp 225 , Tyr 163 , Trp 160 , Trp 532 , Trp 268 , and Trp 389 . Seven of these aromatic residues are in domain II, but the first two come from domain I. According to the catalytic mechanism (23), the crucial catalytic residues, Asp 304 , Asp 306 , and Glu 308 , are located in the middle of the substrate binding cleft.
Substrate Binding Cleft of OfChi-h-Although our attempts to obtain the structure of OfChi-h complexed to its substrate hexa-N-acetylchitohexaose ((GlcNAc) 6 ) failed, the structure of OfChi-h complexed to chitoheptaose ((GlcN) 7 ), a substrate analog, was obtained by soaking OfChi-h crystals with (GlcN) 7 . The structure was determined by molecular replacement using the unliganded form of OfChi-h as a searching model. The final structure was refined to a resolution of 2.7 Å ( Table 1). The sugar binding subsites were named according to Davies et al. (49), where subsite Ϫn represents the non-reducing end, subsite ϩn represents the reducing end, and the enzymatic cleavage happens between the Ϫ1 and the ϩ1 subsites.
The overall structure of OfChi-h-(GlcN) 7 is very similar to unliganded OfChi-h, with a root mean square deviation (r.m.s.d.) of 0.3 Å. The electron density map supports (GlcN) 7 binds along the substrate binding cleft and occupies the sub- sites from Ϫ5 to ϩ2 (Fig. 2). It is worth noting that the electron density signals from GlcN residues at Ϫ5, Ϫ4, Ϫ3, ϩ1, and ϩ2 are stronger than those at Ϫ2 and Ϫ1. This is different from the electron density map of (GlcN) 5 complexed with OfChtI in that the electron density signals of GlcN residues are strong at subsite Ϫ1 and Ϫ2 but weaker at Ϫ3, Ϫ4, to Ϫ5 (50). (GlcN) 7 binds the substrate binding pocket of OfChi-h in a bent conformation. According to the Cremer-Pople parameter calculation (51), the conformation of the GlcN residues at subsites Ϫ1 and Ϫ2 are 1 S 5 and 4 H 5 , respectively, whereas the conformation of the GlcN residues at the other subsites is 4 C 1 (Fig. 2). The (GlcN) 7 binds OfChi-h mainly via stacking interactions between sugar rings and aromatic residues, specifically Ϫ5 GlcN with Tyr 163 , Ϫ3 GlcN with Trp 160 , Ϫ1 GlcN with Trp 532 , ϩ1 GlcN with Trp 268 , and ϩ2 GlcN with Trp 389 . In addition, polar interactions were also observed between OfChi-h and (GlcN) 7 , including C3-hydroxyl group of the Ϫ1 GlcN with Glu 308 and C6-hydroxyl group of the Ϫ1 GlcN with Arg 439 , respectively (Fig. 2). Enzymatic Activities of OfChi-h-The substrate spectrum of OfChi-h was determined using various insoluble substrates including insect cuticle, ␣-chitin, chitin nanowhisker (CNW) as well as soluble substrate ethylene glycol chitin (EGC). Two chitinases, OfChtI and SmChiA, were chosen to compare with OfChi-h. Because the two tryptophans along the substrate binding cleft of OfChi-h were phenylalanines in SmChiA, the mutant SmChiA-F232W/F396W was thus constructed to test the effects of these site mutations ( Fig. 1B and see Fig. 6A). Among the four substrates, OfChi-h and the other enzymes exhibited the highest hydrolytic activity toward the soluble EGC but lower activities toward insoluble substrates ( Table 2). Table 2, OfChi-h exhibited higher activities toward insoluble substrates than OfChtI but showed lower activities toward EGC than OfChtI. SmChiA-F232W/F396W outperformed SmChiA in hydrolyzing insect cuticle, ␣-chitin, and CNW but showed lower activities toward EGC.

As shown in
The hydrolytic mode of OfChi-h was investigated using (GlcNAc) 6 as the substrate. In addition, the hydrolytic modes of OfChtI, SmChiA, SmChiB, and SmChiC were also investigated for comparison. Because chitin is a ␤-1,4-linked polymer of GlcNAc and GH18 chitinases hydrolyze chitin via a retaining mechanism, ␤-anomeric products will be left after cleavage. SmChiA has been experimentally determined with transmission electron microscopy (52) and high speed atomic force microscopy (45) to be an exo-chitinase that attacks chitin from the reducing end. Supplemental Fig. S2 showed HPLC analysis of ␣and ␤-anomeric hydrolytic products of (GlcNAc) 6 in the presence of OfChi-h, OfChtI, SmChiA, SmChiB, and SmChiC. As shown in Table 3, the percentage of hydrolytic ␤-anomeric products in the presence of OfChi-h was very similar to that by SmChiA but different from those in the presence of any of OfChtI, SmChiB, or SmChiC. Therefore, we deduce OfChi-h perhaps acted in a similar way as did SmChiA.
Because both OfChi-h and OfChtI are present in the insect molting fluid and may act through different hydrolytic modes, the synergistic effect was investigated using insect cuticle chitin as substrate (Fig. 3). The results clearly indicated a synergistic effect between OfChi-h and OfChtI because the activity of the combination of OfChi-h and OfChtI was significantly higher than the specific activity calculated from individual activities during the reaction. The synergistic coefficient between

TABLE 3
Percentage of ␤-anomers after partial hydrolysis of (GlcNAc) 6  OfChi-h and OfChtI was calculated by using the following function (53).

by insect and bacterial chitinases
It is worthy to note that the synergistic coefficient at different time points increased with the reaction time from 1.24 at 2 h to 1.98 at 6 h.
To test the in vivo activity, 0.2 g of TMG-(GlcNAc) 4 was injected into a 5th instar, day-3 O. furnacalis larva. The metamorphosis of the TMG-(GlcNAc) 4 -injected group was severely affected compared with the water-injected group (Fig. 4B). In the control group, 100% of the insects molted into normal pupa 5 days after injection compared with only 40% of the insects from the TMG-(GlcNAc) 4 -injected group. 23% of the insects in the TMG-(GlcNAc) 4 -injected group were arrested during the larva stage, whereas the other 37% molted into abnormal pupa. The abnormal pupa appeared to be prepupa trapped by undetached head capsules and thoracic legs (Fig. 4B). Eventually, most of the abnormal pupa and larvae died within 10 days after injection.

Comparison of OfChi-h with Its Bacterial Homolog Sm-ChiA-Insect
Chi-h is presumed to have been obtained from bacteria as it shares higher sequence identities with bacterial chitinases than insect chitinases (3,4). In this study we found SmChiA from S. marcescens had the highest sequence identity of 73% and the highest similarity of hydrolytic anomeric products profiles with OfChi-h and showed the highest structural similarity with OfChi-h (r.m.s.d. of 1.3 Å for 534 C ␣ atoms). Structure superimposition of OfChi-h and SmChiA (E315L) in complex with (GlcNAc) 8 (PDB code 1EHN) demonstrates that the aromatic residues for chitin binding at subsites Ϫ5, Ϫ3, Ϫ1, and ϩ1 are conserved, except the OfChi-h tryptophans at subsite ϩ2 (Trp 389 ) and the SmChiA-corresponding subsite, Ϫ6 (Trp 225 ), are substituted by phenylalanines (Phe 396 and Phe 232 ) in SmChiA (Fig. 6A). As previously shown, OfChi-h and SmChiA have similar substrate specificity (Table 2) and hydrolytic anomeric products composition (Table 3). Given their similar structural characteristics and enzymatic properties, insect Chi-hs and bacterial ChiAs may act similarly in their respective chitin degradation systems.
The mutation of Phe 232 to Ala in SmChiA has been reported to affect the hydrolytic activity but not the binding activity toward crystalline ␤-chitin. Phe 232 is thought to aid in guiding the chitin chain into the catalytic cleft (54). Similarly, the Phe 396 to Ala mutation in SmChiA was reported to decrease its hydrolytic activity toward crystalline ␤-chitin but increase its hydrolytic activity toward soluble chitosan (55). To explore the effect of the Phe to Trp substitutions in the chitin binding cleft of OfChi-h, Phe 232 and Phe 396 in SmChiA were mutated to tryptophan, and the substrate specificity of SmChiA-F232W/ F396W was tested using insect cuticle, ␣-chitin, CNW, and EGC as substrates. Compared with wild-type SmChiA, SmChiA-F232W/F396W showed higher hydrolytic activity for insoluble and crystalline substrates but lower hydrolytic activity for the soluble substrate (Table 2). Because Trp allows more aromatic interactions with chitin chains (56), we deduce that SmChiA-F232W/F396W may guide chitin chains into the substrate binding cleft more efficiently and may improve binding affinity for chitin. Given that the formation of the complex with the chitin chain is presumed to be the rate-limiting step for SmChiA (57), this may explain why F232W/F396W had a higher activity for insoluble chitin. This result also suggests that the substitution of Phe to Trp in OfChi-h increases its ability to degrade insect cuticles, which are highly insoluble and crystalline.
Structural Differences between OfChi-h and OfChtI-As do key chitinases during molting, we found that OfChi-h and OfChtI work synergistically according to their catalytic efficiency in vitro. Their differences in the architecture of substrate-binding sites were then discussed. Although the substrate binding clefts of both OfChi-h and OfChtI are long with both sides open, they have different structural characteristics. First, in OfChi-h, the distribution of aromatic residues aligned along the substrate binding cleft is highly asymmetric with regard to the enzymatic cleavage site (Fig. 6B). There are 13 solvent-exposed aromatic residues in the nonreducing end side but only two in the reducing end side. However, in OfChtI, the distribution of aromatic residues along the substrate binding cleft is symmetric (Fig. 6B); namely, five aromatic residues on both the non-reducing end side and the reducing end side of the cleavage site. Because oligosaccharide substrates binding to the enzyme rely largely onand/or hydrophobic interactions, these aromatic residues are likely crucial for substrate binding. And the asymmetric architecture is generally believed to be a feature of processive exo-chitinases (45). Second, a unique structural element in OfChi-h, but not in OfChtI (residues 188 -214), was observed on the wall of the substrate binding cleft. This structural element increases the depth of the substrate binding cleft and narrows the substrate binding cleft (the narrowest point between residue Ile 210 and Gln 466 is 6.6 Å) (Fig. 6B). This may further increase the binding affinity of OfChi-h for chitin chains and thus favor OfChi-h to hydrolyze crystalline substrate.
Taken together, we deduce that OfChi-h works synergistically with OfChtI, an endo-chitinase. Because both Chi-h and ChtI are highly conserved in lepidopteran species, this synergistic mechanism is likely generalizable.

Experimental Procedures
Gene Cloning and Construction of the Expression Plasmid-Total RNA was extracted from O. furnacalis during the prepupal state using RNAiso Reagent (TaKaRa, Japan) and was subjected to reverse transcription using the PrimeScript TM RT reagent Kit (TaKaRa). Based on the mRNA sequence of OfChi-h (GenBank TM accession number AB201281.1), two primers, 5Ј-CTGAAGCTTACGTAGAATTCGCGCCCC-CTGGCAAACCC-3Ј (forward) and 5Ј-GTGGTGGTGGTG-GTGGTGACTAGTCGCGCTGTTACCTAGACCCA-3Ј (reverse) were designed to amplify the gene fragment encoding mature OfChi-h and add a C-terminal His 6 tag. The resulting PCR products were digested with EcoRI/SpeI and then ligated into pPIC9 vector (Invitrogen) to generate the expression plasmid pPIC9-OfChi-h.
Protein Expression and Purification-The expression plasmid pPIC9-OfChi-h was linearized by PmeI (New England Biolabs) and transformed into Pichia pastoris GS115 cells by electroporation. Positive clones carrying His ϩ and Mut ϩ traits were selected on minimal methanol and minimal dextrose plates. The selected transformant was first cultured in minimal glycerol-complex medium at 30°C to an A 600 of 2.0. The cells were then collected and resuspended in 1 liter minimal methanol-complex medium and incubated at 30°C. Methanol (1%) was added at 24-h intervals. After 72 h of fermentation, the culture supernatant was harvested by centrifugation at 8000 ϫ g for 10 min.
OfChi-h was purified by ammonium sulfate precipitation and immobilized metal ion affinity chromatography (IMAC). Solid ammonium sulfate was added to the culture supernatant to 75% saturation. After incubation at 4°C for 24 h, the sample was centrifuged at 12,000 ϫ g for 30 min. Then the precipitate was dissolved in buffer A (20 mM sodium phosphate, 0.5 M sodium chloride, pH 7.4) and recentrifuged at 12,000 ϫ g for 15 min. Next, the resulting supernatant was loaded onto a HisTrap TM crude column (5 ml, GE Healthcare) pre-equilibrated with buffer A. Then the column was washed with buffer A containing 75 mM imidazole to remove nonspecific binding proteins. Finally, the recombinant OfChi-h was eluted with buffer A containing 250 mM imidazole. The protein was quantified using a BCA protein assay kit (TaKaRa) with bovine serum albumin as a standard protein, and its purity was analyzed by SDS-PAGE (supplemental Fig. S1). The molecular mass of the recombinant OfChi-h was determined to be 64.6 kDa, which was 4.4 kDa larger than the theoretical moleculare mass. Two N-glycans found in the crystal structure may account for this discrepancy.

JOURNAL OF BIOLOGICAL CHEMISTRY 2085
Enzymatic Assays-Three kinds of polymeric substrates, EGC (Wako Pure Chemicals, Osaka, Japan), CNW (prepared as described in Kuusk et al.;Ref. 57), and ␣-chitin (Sigma), were used as substrates for the chitinase activity assays. The 100-l reaction mixtures consist of 2 M enzyme and 3 mg/ml substrate in 20 mM sodium phosphate buffer, pH 6.0. After incubating at 30°C for an appropriate time, the amount of reducing sugars was determined by the potassium ferricyanide method (58).
The chitin from insect cuticle was prepared as follows: 50 of the 5th-instar day-3 larvae were dissected, and the integuments were collected. The integuments were milled into powder in liquid nitrogen and then washed twice with buffer B (20 mM sodium phosphate, 0.15 M sodium chloride, pH 7.4). To remove minerals and catechols, the powder was treated with 4 M hydrochloric acid at 75°C for 2 h and then rinsed thoroughly with buffer B. Next the powder was treated with 4 M sodium hydroxide for 20 h at 100°C to remove proteins and then rinsed thoroughly with buffer B before being placed in an oven at 60°C for 24 h to dry. At last, the insect chitin was suspended in buffer B to a concentration of 10 mg/ml. To evaluate the enzymatic activity and the synergism of OfChi-h and OfChtI, 3 mg/ml insect chitin was incubated with 2 M OfChi-h, 2 M OfChtI, or a mixture of 2 M OfChi-h and 2 M OfChtI. The reaction mixtures were incubated at 30°C, and 50-l samples were collected at different times to determine the production of reducing sugar.
Analysis of TMG-(GlcNAc) 4 Hydrolytic Products by ESI-TOF MS-Three copies of TMG-(GlcNAc) 4 at 10 M concentration were incubated with 0.1 nM OfChi-h, SmChiA, and SmChiB for 30 min. Then 20-l of hydrolysate was analyzed by ESI-TOF MS using an Agilent G6224A (Agilent) in positive-ion reflection mode.
In Vivo Bioevaluation of TMG-(GlcNAc) 4 by Injection-O. furnacalis larvae were reared using an artificial diet with 16 h of light and 8 h of darkness and a relative humidity of 70 -90% at 26 -28°C. Larvae at day 3 of the fifth instar were selected for the microinjection experiment. In the experimental group, 0.2 g of TMG-(GlcNAc) 4 (solved in water) was injected into the penultimate abdominal segment of larvae. In the control group, distilled water was injected instead. Each group contained 10 individual larvae with three independent replicates. After injection, all of the treated larvae were reared under identical conditions as described above. Mortality and developmental defects were recorded every day until eclosion.
Crystallization and Data Collection-Pure OfChi-h was spin-concentrated to 10 mg/ml in 20 mM bis-Tris (pH 6.5) containing 50 mM NaCl. Crystallization screening of recombinant OfChi-h was performed using the following commercially available screens: Index, Crystal Screen, and Crystal Screen 2 (Hampton Research). The hanging-drop vapor-diffusion crystallization experiments were set up at 4°C by mixing 1 l of OfChi-h and 1 l of reservoir solution. The protein crystallized after 1 month in 100 mM HEPES, pH 7.0, 30% (w/v) Jeffamine ED-2001.
Crystals of OfChi-h-ligand complexes were obtained by transferring native crystals to a reservoir solution consisting of 5 mM (GlcNAc) 6 , 10 mM (GlcN) 7 (Qingdao BZ Oligo Biotech Co., Ltd.), or 1 mM TMG-(GlcNAc) 4 . For (GlcNAc) 6 , the crystals were soaked for 5 min, 15 min, and 1 h at room temperature. For (GlcN) 7 or TMG-(GlcNAc) 4 , the crystals were soaked for 1 h at room temperature. Then the crystals were soaked for several minutes in a reservoir solution containing 25% (v/v) glycerol and subsequently flash-cooled in liquid nitrogen. Diffraction data were collected on the BL-18U1 at the Shanghai Synchrotron Radiation Facility in China, and the diffraction data were processed using the HKL-2000 package (60).
Structure Determination and Refinement-The structure of free OfChi-h was solved by molecular replacement with Phaser (61) using the structure of SmChiA (PDB code: 1EDQ) as a model. OfChi-h-(GlcN) 7 complexes were solved using the coordinates of free OfChi-h as a model. Structure refinement was performed using PHENIX (62). The molecular models were manually built and extended using Coot (63). The stereochemistry of the models was checked by PROCHECK (64). The data collection and structure refinement statistics are summarized in Table 1. The coordinates of OfChi-h and OfChi-h-(GlcN) 7 are deposited in the PDB with the codes 5GPR and 5GQB, respectively. All structural figures were prepared using PyMOL (DeLano Scientific LLC, San Carlos, CA).
Author Contributions-T. L. and Q. Y. designed the experiments. T. L., L. C., X. J., and Y. D. performed the experiments. T. L. and Y. Z analyzed the protein structures. T. L. and Q. Y. analyzed the data and wrote the paper.