ATP-dependent chromatin remodeling enzymes (remodelers) alter the structure and organization of nucleosomes, the basic building blocks of eukaryotic chromatin (Becker and Workman, 2013; Bracken et al., 2019; Clapier et al., 2017; Jungblut et al., 2020; Sundaramoorthy and Owen-Hughes, 2020). Core nucleosomes comprise 147 bp of DNA wrapped around a histone protein octamer formed by a H3-H4 tetramer that is flanked on either side by a H2A-H2B dimer. Remodelers modulate the accessibility of regulatory DNA elements by unwrapping or sliding nucleosomes, thereby presenting a fundamental level of control of transcription and other processes that require access to DNA (Brahma and Henikoff, 2020). Remodelers are large, multi-subunit complexes that harbor a Snf2-class ATPase motor (Clapier et al., 2017; Jungblut et al., 2020; Sundaramoorthy and Owen-Hughes, 2020). Nucleosome remodeling is the result of ATP-dependent DNA translocation by the ATPase, which is locked onto the nucleosome through binding additional DNA- and histone sites. ATPase activity is directed by accessory subunits that determine targeting to specific genomic loci, and the outcome of the remodeling reaction. Different classes of remodelers, defined by their ATPase and unique sets of accessory subunits, are dedicated to distinct chromatin regulatory functions.

Members of the SWI/SNF family of remodelers have mainly been implicated in generating open chromatin at gene regulatory DNA elements (Becker and Workman, 2013; Bracken et al., 2019; Brahma and Henikoff, 2020; Cakiroglu et al., 2019; Clapier et al., 2017; Pillidge and Bray, 2019). Recent structural studies revealed that SWI/SNF-class remodelers engage the nucleosome in a highly modular manner (Han et al., 2020; He et al., 2020; Jungblut et al., 2020; Mashtalir et al., 2020; Patel et al., 2019; Sundaramoorthy and Owen-Hughes, 2020; Wagner et al., 2020; Ye et al., 2019). The motor domain of the ATPase binds the nucleosomal DNA at superhelical position +2, while other modules contact the opposite DNA gyre, both faces of the histone octamer, and possibly the exiting DNA, and the histone H4 tail. SWI/SNF remodelers do not closely encompass the nucleosome, and the nucleosome remains mostly accessible. ATPase activity is only required for binding to a subset of genomic loci, but loss of ATPase activity can have dominant effects on chromatin binding, enhancer accessibility, and gene expression (Gelbart et al., 2005; Hodges et al., 2018; Pan et al., 2019). Mutations in key functional domains of SWI/SNF are associated with human cancer and developmental disorders (Bracken et al., 2019; Cenik and Shilatifard, 2021; Mashtalir et al., 2020; Sundaramoorthy and Owen-Hughes, 2020).

There are two main subclasses of SWI/SNF complexes that are conserved from yeast to man. The first includes yeast SWI/SNF, Drosophila BAP and mammalian BAF, while the second class includes yeast RSC, fly PBAP, and mammalian PBAF (Bracken et al., 2019; Clapier et al., 2017; Mohrmann and Verrijzer, 2005). Complexes of both classes share a common core, comprising related or identical subunits, associated with a set of signature subunits that define either SWI/SNF-BAF or RSC-PBAF. In mammalian cells, a third type of SWI/SNF complex has been described, named GBAF or non-canonical BAF (ncBAF; Alpsoy and Dykhuizen, 2018; Gatchalian et al., 2018; Mashtalir et al., 2018). Drosophila BAP and PBAP share a single ATPase, BRM, which is the ortholog of yeast Swi2/Snf2 and Sth1, and human SMARCA2 and SMARCA4. BRM associates with seven additional subunits to form the core of both BAP and PBAP. OSA (human ARID1A/B), D4/TTH (DPF1-3) and SS18 associate with the core subunits to form BAP, whereas POLYBROMO (PBRM1), BRD7, BAP170 (ARID2), and SAYP (PHF10) are specific for PBAF. The signature subunits play a major role in genomic targeting and functional selectivity of BAP and PBAP (Bracken et al., 2019; Chalkley et al., 2008; Mohrmann et al., 2004; Moshkin et al., 2007; Moshkin et al., 2012). Thus, SWI/SNF remodelers can be considered holoenzymes, in which different modules provide different functionalities that direct the remodeling activity of the ATPase. Although BAP and PBAP have shared activities, for example, both antagonize Polycomb repression, they also function in unique gene expression programs that control development, cell proliferation and differentiation (Chalkley et al., 2008; Mohrmann et al., 2004; Moshkin et al., 2007; Moshkin et al., 2012). Here, we use (P)BAP when making general statements that apply to both BAP and PBAP.

The natural amplification of Drosophila larval salivary gland polytene chromosomes allows the visualization of fluorescent-tagged transcription factors on interphase chromatin at native genetic loci in living cells. This has yielded fundamental insights in RNA polymerase II (RNAPII) recruitment and transcriptional dynamics (Yao et al., 2007; Zobeck et al., 2010). Salivary gland polytene chromosomes are the result of typically 10 rounds of DNA replication without segregation, resulting in cable-like super chromosomes comprising up to 1024 closely aligned chromatids. Interphase polytene chromosomes are visible by light microscopy and provided the first view of a physical genetic map (Bridges, 1935). The characteristic banding pattern of polytene chromosomes reflects the degree of interphase condensation, i.e., the ratio between the length of a stretched DNA molecule and the length of the corresponding chromosomal domain. The compacted polytene bands have been linked to transcriptionally repressed topologically associated domains (TADs), which are conserved in diploid cells (Eagen et al., 2015). Interbands contain gene regulatory regions, origins of replication, and are characterized by marks of active, open chromatin (Zykova et al., 2018). Polytene bands can be divided into moderately condensed gray chromatin, harboring the coding regions of active genes, and the highly compacted black chromatin that contains inactive developmental genes, and under-replicated, gene-poor intercalary heterochromatin (Eagen et al., 2015; Zykova et al., 2018). Very high levels of transcriptional activity results in puffing, the local uncoiling of individual chromosome strands due to the accumulation of the gene expression machinery and RNA. Pertinently, there is a close correspondence between chromatin structure of polytene chromosomes and that in diploid cells (Eagen et al., 2015; Zykova et al., 2018).

The steroid hormone 20-hydroxyecdysone (Ec) is the key regulator of Drosophila development and controls major transitions such as molting and metamorphosis (Hill et al., 2013). Exposure of larval salivary glands to Ec leads to the formation of early chromosomal puffs, starting a cascade of developmental gene expression and puffing. The Ec hormone mediates gene activation via binding to the Ec receptor (EcR), which associates with specific DNA elements. EcR belongs to the nuclear receptor family of transcription factors that mediate hormone-driven gene expression. Upon hormone binding, nuclear receptors cooperate with a slew of coactivators, including the SWI/SNF remodelers, to stimulate transcription (Hoffman et al., 2018; Paakinaho et al., 2017).

Previous live-cell microscopy of fluorescent-tagged remodelers in diploid cells suggested a fast exchange dynamic, similar to many transcription factors (Erdel et al., 2010; Johnson et al., 2008; Mehta et al., 2018; Paakinaho et al., 2017; Swinstead et al., 2016; Voss and Hager, 2014). However, the in vivo kinetics of remodeler interactions with specific natural genomic loci has not been studied. In particular, the role of ATP hydrolysis in remodeler kinetics remains unclear. Here, we used vital imaging to determine the spatial and temporal dynamics of (P)BAP interactions with endogenous target loci and the effect of chromatin condensation. Our results show that ATP-hydrolysis fuels a continuous probing of the genome by (P)BAP. In the absence of ATP-hydrolysis, there is increased chromatin retention and reduced turnover of (P)BAP. We discuss the implications of our in vivo findings for understanding remodeler function and chromatin dynamics.

We studied the role of ATP-hydrolysis during exploration of the genome by (P)BAP chromatin remodelers. We took advantage of the parallel amplification of the Drosophila genome in salivary gland polytene chromosomes to monitor (P)BAP interactions with endogenous targets in real time. This allowed us to distinguish between nucleoplasm and natural loci in interphase chromosomes by live-cell imaging, which is not possible in diploid cells. We found that a surprisingly small portion of (P)BAP (~5%) is associated with chromatin at any given time. This is caused by the coupling of ATP-hydrolysis during nucleosome remodeling to (P)BAP release. Consequently, (P)BAP acts through a continuous transient probing of the genome (Figure 9A). (P)BAP mainly interacts with chromatin in non-condensed interbands or puffs. Conversely, (P)BAP levels within highly condensed polytene bands are much lower than in the nucleoplasm. We note that the loss of (P)BAP does not affect chromatin condensation and the polytene banding pattern (Figure 1—figure supplement 1A). Based on these observations, we postulate that chromatin condensation excludes (P)BAP, thereby reducing its genomic search space. At the height of E74 and E75 transcription, a substantial portion of RNAPII is retained locally in what Lis and colleagues named a transcription compartment (Yao et al., 2007). On the same chromatin puffs, however, PBAP exchanges rapidly (within ~2–4 s) without a substantial immobile fraction. Thus, although essential for expression of E74 and E75, PBAP is not retained within a transcription compartment. The rapid exchange of (P)BAP indicates continuous cycles of nucleosome remodeling. Transient binding to chromatin might explain the relatively poor capture of remodelers by ChIP, as the crosslinking reaction has temporal constraints (Gelbart et al., 2005; Schmiedeberg et al., 2009). Indeed, a mutation in the ATP-binding pocket increased the residence time on chromatin and concomitantly enhanced ChIP signals of BRM-K804R. Consistent with our earlier developmental genetic analysis (Chalkley et al., 2008), we found that PBAP, but not BAP, is required for Ec-regulated gene expression. This finding expands the functional diversification between BAP and PBAP remodeling complexes, which is to a large extent determined by their signature subunits (Mohrmann et al., 2004; Moshkin et al., 2007; Chalkley et al., 2008; Moshkin et al., 2012). We detected the fly homolog of GLTSCR1 in GFP-BRM IPs from S2 cells (Figure 4B). Indeed, a recent study identified and characterized the fly BRM complex that is orthologous to mammalian GBAF/ncBAF (Barish et al., 2020). Thus, although GBAP appears to be mainly expressed in the nervous system, it might contribute to our analysis of BRM dynamics. We note that the dynamic characteristics of SNR1, D4 and BRD7, which are all absent from GBAP, is highly similar to that of BRM.

Figure 9

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Histones displayed only limited exchange, which was more than 2 orders of magnitude slower than that of (P)BAP. FRAP analysis of GFP-H2B in polytene chromosomes showed that it takes about 40 min to obtain a ~18% recovery (Figure 5H). These results agree with the notion that histone eviction is restricted to small regulatory regions of the genome and is associated with the placement of variant histones (Brahma and Henikoff, 2020; Cakiroglu et al., 2019; Deal et al., 2010; Pillidge and Bray, 2019). Indeed, it is difficult to envision how the limited amount of free canonical histones available in the nucleoplasm could support substantial turnover of chromosomal histones. A comparison of DNA accessibility and histone occupancy during transcriptional induction revealed that changes in the former predominate the latter (Mueller et al., 2017). Likewise, several recent studies indicated that promoter chromatin harbors so called fragile nucleosomes with accessible DNA, which depend on chromatin remodeler activity (Brahma and Henikoff, 2020). These observations suggest that the prevalent outcome of chromatin remodeling in vivo is restructuring or sliding of nucleosomes rather than eviction.

Vital imaging of fluorophore-tagged histones recapitulated the polytene banding pattern and local chromatin condensation. This enabled us to match the GFP-H2B density distribution in living cells with chromatin states derived from histone marks and DNA accessibility (Eagen et al., 2015; Filion et al., 2010; Kharchenko et al., 2011; Zykova et al., 2018). Our analysis indicated that DNA in gray bands is on average ~6-fold more condensed than that in interbands, whereas heterochromatic DNA can be up to ~150-fold more condensed. The level of gray band condensation measured by live-cell imaging agrees well with the value derived from HiC analysis of diploid Kc167 cells (Eagen et al., 2015), whereas our imaging-based estimate for black chromatin condensation is ~5-fold higher. We suspect that this discrepancy is caused by the very high degree of chromatin condensation at the chromocenter and telomers of polytene chromosomes. Indeed, measurements of black chromatin bands flanking the E74 and E75 loci, suggested a ~25-fold condensation compared to open chromatin, which falls in the range derived from HiC in diploid cells (Figure 3E; Eagen et al., 2015). Our real time measurements of histone density in polytene chromosomes in living cells complements other methods to estimate condensation that all depend on chromatin fixation (Eagen et al., 2015; Ou et al., 2017; Zykova et al., 2018). Collectively, these studies suggest that euchromatin and heterochromatin are formed by the same basic nucleosomal array, but at different degrees of condensation.

Although central to the function of Snf2 remodelers, our understanding of the role of ATP hydrolysis in the nucleosome remodeling cycle remains incomplete. A mutation in the ATP-binding pocket of BRM caused up to 5-fold stronger chromatin binding and a ~2-times slower turnover of BRM-K804R in S2 cells. In polytene cells, BRM-K804R clamped down onto chromatin and displayed no measurable turnover. Likewise, in permeabilized salivary gland cells that lack ATP, wt BRM accumulated onto polytene chromosome interbands. Chromatin binding persisted for over 1 hr, but following the addition of ATP, BRM released immediately. Most likely, the high local density of target loci in polytene chromosomes amplifies the effect of prolonged chromatin binding by BRM. We speculate that in the absence of ATP hydrolysis, slow release combined with rapid re-binding to a paired chromatid, might results in the local trapping of (P)BAP on polytene chromosomes. We note that under normal physiological conditions, ATP concentrations in cells would not become depleted to a level (< 1 mM) that would impair remodeler function. Finally, our results argue against a role for liquid-liquid phase separation in the clustering of (P)BAP in the absence of ATP-hydrolysis. Local (P)BAP accumulation is insensitive to the disruption of weak hydrophobic interactions by 1,6-hexanediol. Moreover, FRAP analysis showed that clustered BRM has a strongly reduced mobility, indicating it does not form a condensate. We consider prolonged chromatin-binding the most plausible cause of local accumulation of BRM-K804R, or wt BRM in the absence of ATP.

Studies in diploid cells often equate the formation of low-mobility aggregates of a fluorophore-tagged transcription factor with persistent chromatin binding and functional activity. However, given that chromatin typically only occupies about 3% of the nuclear volume, this interpretation remains speculative. E.g., in a study on human ISWI remodelers, an increase in the immobile fraction upon DNA damage was rationalized as an increase in activity on chromatin (Erdel et al., 2010). Moreover, it was concluded that loss of ATP-binding reduced chromatin association and increased mobility. In contrast, our results with BRM showed that chromatin binding is independent of ATP, but that its release is stimulated by ATP-hydrolysis. At the peak of PBAP- and EcR-dependent transcription of E74 and E75 genes, both retained a rapid exchange, indicating ceaseless cycles of nucleosome remodeling. Consistent with this notion, the use of small-molecule inhibitors recently showed that chromatin accessibility required continuous remodeler activity (Iurlaro et al., 2021). Transient remodeler dynamics enables competition at regulatory elements between epigenetic regulators with opposing effects on chromatin structure (Bracken et al., 2019; Cenik and Shilatifard, 2021; Kubik et al., 2019; Mohd-Sarip et al., 2017).

There are both parallels and differences in the roles of ATP in either chromatin remodeling or in RNA-duplex unwinding by DEAD-box helicases. ATP-hydrolysis is dispensable for RNA-binding and duplex unwinding by DEAD-box helicases, but is required for fast enzyme release from RNA (Liu et al., 2014). However, the formation of long-lived RNA-DEAD-box helicase complexes requires ATP-binding but not its hydrolysis. In contrast, chromatin binding by BRM is independent of ATP. The ATP hydrolysis-fueled transient probing of the genome by BRM we described here, dovetails well with structural observations on Swi2/Snf2 remodelers (Xia et al., 2016). When Snf2 is free in solution, the two domains that harbor the ATP-binding motifs are oriented away from one another and Snf2 cannot bind ATP. Binding to naked DNA or nucleosomal DNA induces a reorientation of the two ATPase lobes so that they now form a functional ATP-binding pocket. Combined with our results, these structural observations suggest a model in which nucleosome binding occurs independent of ATP but induces the formation of an ATP-binding pocket (Figure 9B). ATP hydrolysis can now fuel nucleosome remodeling coupled to an increased probability of remodeler release, resulting in rapid cycles of nucleosome remodeling and remodeler recycling. The absence of ATP-hydrolysis leads to prolonged chromatin binding of the remodeler and impedes nucleosome remodeling.

Our observation that ATP hydrolysis promotes remodeler release from chromatin raises the question of how many translocation steps a single remodeler takes before dissociation from a nucleosome in vivo. Previous single molecule studies on the RSC remodeler determined that an ATP concentration above 1 mM is required for optimal translocation speed and processivity (Zhang et al., 2006; Sirinakis et al., 2011). This value derived from in vitro experiments is remarkably similar to our in vivo estimates. Note that many biophysical studies on remodelers use much lower ATP concentrations to slow down the reaction speed and determine individual kinetic steps (see e.g. Harada et al., 2016). In the presence of 1 mM ATP in vitro, RSC translocated at a speed of 25 bp/s on naked DNA and ~13 bp/s on nucleosomes (Zhang et al., 2006; Sirinakis et al., 2011). Enzyme processivity in these experiments was ~35 bp on naked DNA and (albeit it with a widespread) on average ~100 bp per round of translocation on nucleosomes. Stopped-flow pre-steady state kinetic analysis of RSC translocation along naked DNA suggested a series of uniform steps yielding an average translocation distance of 20 bp before dissociation (Fischer et al., 2007). The (P)BAP turnover time of ~2–4 s in our in vivo FRAP experiments agrees well with the in vitro single molecule analyses and is compatible with multiple kinetic steps and substantial nucleosome remodeling prior to remodeler disengagement. Remodeler release might be considered as a probability during each ATPase cycle. Full remodeling, sliding or even eviction may require the cooperation of a cascade of remodelers acting subsequently.

BRM-K804R clamps to chromatin and impedes histone turnover, emphasizing that remodeling and remodeler recycling are closely integrated. Surprisingly, BRM-K804R also traps wt BRM onto polytene chromosomes. Thus, reflecting early genetic studies (Elfring et al., 1998), BRM-K804R behaves as a molecular dominant mutant. The retention of wt BRM onto chromatin by BRM-K804R suggests that BRM complexes might interact on chromatin. Consequently, remodeling might involve a nucleosome hand-off mechanism between different BRM complexes. Additionally, translocation by one BRM complex might help to push another one off the chromatin. This way, the persistent chromatin binding of BRM-K804R might create a ‘traffic jam’ that blocks the release of wt BRM.

In summary, our visualization of BRM remodelers engaging their endogenous genomic targets revealed highly transient and dynamic interactions. (P)BAP binds chromatin independent of ATP. However, ATP-hydrolysis couples nucleosome remodeling to (P)BAP release, resulting in rapid cycles of remodeling and remodeler turnover. Remodelers might interact on chromatin, as suggested by the trapping of wt BRM by the ATP-binding mutant BRM-K804R. Importantly, an accompanying report by Wu and colleagues, studying budding yeast remodelers through live-cell single molecule tracking, also revealed the key role of the ATPase in promoting dissociation from chromatin and fast remodeler kinetics (Kim et al., 2021). Collectively, these in vivo results provide a framework for understanding remodeler function in genome regulation.

The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium (http://www.proteomexchange.org/) via the PRIDE partner repository with the dataset identifier PXD025474. All data generated or analyzed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 1, 2 and Figure 2—figure supplement 1.

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Author details

1. Ben C Tilly

   Department of Biochemistry, Rotterdam, Netherlands

   Contribution

   Conceptualization, Data curation, Software, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing

   For correspondence

   b.tilly@erasmusmc.nl

   Competing interests

   No competing interests declared

2. Gillian E Chalkley

   Department of Biochemistry, Rotterdam, Netherlands

   Contribution

   Data curation, Validation, Investigation, Visualization, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

3. Jan A van der Knaap

   Department of Biochemistry, Rotterdam, Netherlands

   Contribution

   Conceptualization, Data curation, Validation, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

4. Yuri M Moshkin

   Department of Biochemistry, Rotterdam, Netherlands

   Contribution

   Data curation, Formal analysis, Supervision, Validation, Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

5. Tsung Wai Kan

   Department of Biochemistry, Rotterdam, Netherlands

   Contribution

   Investigation, Methodology

   Competing interests

   No competing interests declared

6. Dick HW Dekkers

   1. Department of Biochemistry, Rotterdam, Netherlands
   2. Proteomics Center, Erasmus University Medical Center, Rotterdam, Netherlands

   Contribution

   Data curation, Formal analysis, Investigation

   Competing interests

   No competing interests declared

7. Jeroen AA Demmers

   1. Department of Biochemistry, Rotterdam, Netherlands
   2. Proteomics Center, Erasmus University Medical Center, Rotterdam, Netherlands

   Contribution

   Data curation, Formal analysis, Supervision, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

8. C Peter Verrijzer

   Department of Biochemistry, Rotterdam, Netherlands

   Contribution

   Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing - original draft, Project administration, Writing - review and editing

   For correspondence

   c.verrijzer@erasmusmc.nl

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-6476-3264

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