The AAA+ (ATPases Associated with diverse cellular Activities) superfamily is comprised of ~100 proteins in humans (Erzberger and Berger, 2006; Human AAA+ protein count was obtained as follows: in supfam.org, search was performed for "Extended AAA-ATPase domain" and refined for proteins within the human genome). These ATPases are essential for many cellular processes, including DNA replication, proteostasis, membrane remodeling, and cytoskeletal organization (Hanson and Whiteheart, 2005). Extensive cell biological and biochemical studies have revealed that these enzymes couple ATP hydrolysis to substrate remodeling and directional transport, processes that can occur on the timescale of seconds or minutes (Hanson and Whiteheart, 2005). Thus, small molecule inhibitors that can modulate AAA+ activity on similarly fast timescales are likely to be valuable tools to probe their cellular functions (Lampson and Kapoor, 2006). Valosin-containing protein (Deshaies, 2014) and dynein (Firestone et al., 2012) are the only two human enzymes in this large superfamily for which well-characterized small molecule antagonists have been reported.

Dyneins are microtubule-based motor proteins in the AAA+ family that have been divided into two classes, axonemal and cytoplasmic. Axonemal dyneins are required for the beating of flagella. Cytoplasmic dyneins, of which there are two isoforms (hereafter, dynein 1 and 2), are present in metazoan cells and are required for a wide range of cellular processes (Vale, 2003; Allan, 2011; Vallee et al., 2004). Transport of cargo along microtubules requires a balance of forces directed toward opposite ends of the filament. While multiple motor proteins in the kinesin family provide the plus-end directed force to drive this motion, their activity in many contexts is opposed by one of two cytoplasmic dyneins, the primary motor proteins transporting cargos toward the minus-end of microtubules (Vale, 2003). Dynein 1 has many functions in the cytoplasm, where it moves diverse cargoes ranging from mRNA molecules to whole organelles. In contrast, dynein 2’s functions are restricted to cilia and flagella. The primary cilium is an antenna-like organelle that protrudes from the cell surface. Bidirectional transport in the cilium, known as intraflagellar transport, is required for Hedgehog signaling, a developmental pathway (Goetz and Anderson, 2010). Cargos of both dynein isoforms can move at rates of >1 µm/s in cells (Allan, 2011; Mijalkovic et al., 2017) and therefore, fast-acting, reversible chemical inhibitors are likely to be useful probes for dynamic dynein-dependent cellular processes.

Ciliobrevins were recently reported as the first selective, cell-permeable probes of dynein (Firestone et al., 2012). Although other chemical antagonists of dynein have contributed to understanding the biochemistry of dynein, their use in cell biology has been limited because they either have limited cell permeability (e.g. vanadate) or are non-selective in cells (e.g. EHNA) (Kobayashi et al., 1978; Gibbons et al., 1978; Bouchard et al., 1981). Ciliobrevins were discovered as inhibitors of Hedgehog signaling and shown to block cytoplasmic dynein 1- and 2-dependent transport in cells (Firestone et al., 2012; Hyman et al., 2009). Ciliobrevins have been used as tools to examine the role of dynein in a number of processes, including formation of the immunological synapse, transport of signaling proteins in the primary cilium, axonal transport of transcription factors, and axon extension and branching in cultured neurons (Sainath and Gallo, 2015; Liu et al., 2013; Yi et al., 2013; Ye et al., 2013). However, the use of ciliobrevins has been limited by their low potency and suboptimal chemical properties (Roossien et al., 2015), which is often noted for first-in-class compounds identified through high-throughput screening. Complete inhibition of dynein by ciliobrevins can require high doses (50–100 µM) and selective protein target inhibition can be difficult to achieve at such concentrations.

The ciliobrevins are based on a benzoylacrylonitrile-substituted quinazolinone scaffold (Figure 1A). This type of acrylonitrile has the potential to react with nucleophiles, and instability of ciliobrevins during storage has been noted (Sainath and Gallo, 2015). The benzoylacrylonitrile core is required for in vitro and cellular activity (Firestone et al., 2012; See et al., 2016). This functional group has the potential to isomerize, and the ciliobrevin scaffold may exist as either of two isomers about the benzoylacrylonitrile olefin (C2 - C9, Figure 1A). The preferred isomer of this compound has not been determined and chemical modification of the quinazolinone or acyl groups, even distal to the acrylonitrile functionality, has the potential to affect the geometry of the compound's core. Together, these factors have made activity-guided modifications to improve compound potency challenging. Further, up to ~100 fold differences in potency have been noted between biochemical and cell based assays (0.2 µM – 30 µM for ciliobrevin D), raising concerns about target specificity. Design and chemical synthesis of alternative scaffolds that address these limitations and retain activity against dynein are needed.

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Dynein is a large, ~4600 amino acid protein that contains six unique AAA ATPase domains (Carter, 2013). Many AAA+ enzymes function as homohexameric arrays of identical AAA domains, and thus all six of the ATPase sites can be biochemically equivalent (Erzberger and Berger, 2006). However, for dyneins, the six unique AAA domains reside on a single polypeptide and each can have a specialized role in motor protein function (Carter, 2013). In the case of dynein 1, four of the six AAA domains contain the residues necessary for nucleotide binding; of these, only AAA1 and AAA3 substantially contribute to ATP hydrolysis and have been shown to be required for microtubule motility (Carter, 2013). Dynein’s ATPase activity is stimulated by interactions with microtubules and is thought to occur mainly at AAA1, while hydrolysis at AAA3 plays a regulatory role (Bhabha et al., 2014; DeWitt et al., 2015; Nicholas et al., 2015a). Earlier work suggested that ciliobrevins are ATP-competitive inhibitors of dynein (Firestone et al., 2012), but it remains unclear which of the six AAA sites are modulated, adding to the challenges of inhibitor optimization.

Here, we characterize the conformation of ciliobrevins and design tricyclic pyrazoloquinazolinone derivatives that are more potent inhibitors of dynein. One derivative, which we name dynapyrazole-A inhibits dyneins 1 and 2 with similar potencies in vitro and in cellular assays. Biochemical analyses of this compound showed that while it inhibits the microtubule-stimulated ATPase activity of dynein, it does not potently block the microtubule-independent basal activity. This mode of activity is unlike that of the ciliobrevins, which inhibit both basal and microtubule-stimulated hydrolysis.

To design new analogs with improved properties, we analyzed the conformation of the ciliobrevin scaffold. For these studies, we first used X-ray crystallography. Efforts to crystallize ciliobrevin D were unsuccessful; however, compound 1, a derivative with a 2-morpholinoethyl ether substitution, was synthesized using a previously described procedure and was found to readily crystallize (Figure 1B,C,D) (See et al., 2016). The X-ray data suggest that compound 1 exists as a single isomer with an E-olefin configuration of the C2-C9 double bond. The measured bond lengths are consistent with electrons in a π-system delocalized across the benzoylacrylonitrile core (Figure 1D). The C2-C9 bond length (1.44 Å), which is longer than that of a typical olefin, indicates significant single bond character. The proximity between O11 and N1-H suggests that the ciliobrevin structure is stabilized by an intramolecular hydrogen-bond between N1 and O11 (Figure 1A and D). A closely related acylacrylonitrile-substituted quinazolinone was found to favor an alternative conformation, corresponding to a Z-olefin at C2-C9, that was also stabilized by intramolecular hydrogen-bonding (Milokhov et al., 2013). All these data together indicate it is likely that the isomeric preference of the pharmacophore is sensitive to distal substitutions.

As crystal packing may impact conformation, we turned to NMR spectroscopy to analyze the structure of the ciliobrevin scaffold in solution. A one-dimensional NMR spectrum of ciliobrevin D showed a peak at 13.5 ppm, which could be assigned to a proton at one of the quinazolinone nitrogens (Milokhov et al., 2013) (Figure 1—figure supplement 1). The presence of this broad, downfield peak suggests that one exchangeable N-H proton is stabilized by a hydrogen-bonding interaction. A NOESY spectrum of ciliobrevin D revealed all the expected resonances (Figure 1E). In addition, we detected a coupling between the proton at 13.5 ppm and the proton at the 8-position of the quinazolinone, consistent with the proton at N1 being involved in hydrogen bonding. Together, these data suggest that ciliobrevin D has similar orientation in solution to that of 1 in the crystal, and the benzoylacrylonitile functional group favors an E-isomer configuration that is likely to be stabilized by hydrogen bonding (Figure 1A and E).

We hypothesized that replacing the benzoylacrylonitrile core with a heterocyclic scaffold that maintained the observed ciliobrevin geometry could lead to improved dynein inhibitors. We reasoned that a tricyclic scaffold could replace the non-covalent N1-H-O11 interaction and maintain the overall ciliobrevin pharmacophore. We envisioned replacing the C2-C9 olefin and C10 ketone in ciliobrevin with either a pyrrole or pyrazole ring (Figure 2A–C). We adapted established procedures to synthesize 2 and 3, which differ from known compounds only in the substitution pattern of the D ring (Figure 2A and B) (Süsse and Johne, 1981; Orvieto et al., 2009). We also devised a synthetic route to a series of tricyclic analogs exemplified by 4. This synthesis relies on the condensation of a 2-fluorobenzoic acid methyl ester with a 4-cyano-aminopyrazole under basic conditions (Figure 2C). This strategy allowed convergent synthesis of the desired cyclized ciliobrevin analogs.

Figure 2

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To test if these compounds inhibit dynein, we employed a microtubule gliding assay. We first focused on human cytoplasmic dynein 2, the isoform involved in ciliary transport and Hedgehog signaling (Goetz and Anderson, 2010), as its inhibition in this assay by ciliobrevin D has not been previously demonstrated. We purified an N-terminally GFP-tagged motor-domain construct of dynein 2 using an insect cell expression system (Schmidt et al., 2015). The GFP tag in this construct (GFP-dynein 2, amino acids 1091–4307) allowed the motor protein to be immobilized on passivated glass coverslips used in microtubule gliding assays. GFP-dynein 2 was obtained as a mono-disperse peak by gel filtration (Figure 2D and E). In the presence of ATP (1 mM), this protein moved microtubules with a velocity of 128 ± 14 nm/s (mean ± SD, Figure 2G and I). This activity is readily revealed by a time-series montage, which shows a fluorescently labeled microtubule being displaced ~3 µm in 20 s (Figure 2G). This rate is comparable to previous analyses of this construct (Schmidt et al., 2015). Ciliobrevin D (20 µM) reduced the microtubule gliding velocity to 83 ± 10 nm/s (Figure 2G and I), consistent with its inhibition of dynein-2-dependent processes in cells (Firestone et al., 2012; Ye et al., 2013).

We next tested compounds 2–4 at 20 µM. Compound 2 did not substantially change microtubule gliding velocity (126 ± 17 nm/s, Figure 2H and I). Compounds 3 and 4 inhibited dynein-2-driven gliding to a velocity comparable to that observed in the presence of ciliobrevin D at this same concentration.

To improve the potency of compound 4, we compared its structure with that of ciliobrevin derivative compound 1. In particular, we superimposed the crystal structure of 1 with an energy-minimized structure of 4 computationally generated using Maestro (Schrödinger). This analysis revealed that the dichlorophenyl D ring of 4 projects from the pyrazoloquinazolinone core such that it is offset from the D ring of ciliobrevin (Figure 3A). We reasoned that adding a single carbon atom between the pyrazoloquinazolinone and the phenyl ring may lead to closer alignment with ciliobrevin and that using a cyclopropyl spacer would restrict rotation of the resulting scaffold. Using our modular synthesis strategy for pyrazoloquinazolinones, we generated a set of ciliobrevin derivatives with a cyclopropyl group separating the aromatic ring systems (5–8, Figure 3B). Three of these compounds had substitutions at the 6-position of the quinazolinone (6, 7, and 8, Figure 3B), a modification that we have previously reported to improve the potencies of ciliobrevins (Firestone et al., 2012). The crystal structure of 5 indicates that its D ring aligns with that of compound 1 better than the chlorophenyl ring of compound 4 (Figure 3A, 3C, and 3D).

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We next tested whether compounds 5–8 (20µM) inhibited dynein-2-dependent microtubule gliding (Figure 3E and F). Compound 5 reduced velocity to 58 ± 14 nm/s, while addition of a trifluoromethyl group at the 6-position of the ‘A’ ring in compound 6 led to near-complete inhibition of gliding (6 ± 2 nm/s). By contrast, compound 7, with a methoxy group at the 6-position of the A ring, did not substantially inhibit dynein-2-driven microtubule gliding (128 ± 21 nm/s, Figure 3E and F). Compound 8, with a 6-iodo substituent at the A ring, showed comparable activity to 6 (6 ± 2 nm/s). For the two most active compounds (6 and 8) as well as ciliobrevin D, we performed dose-dependent analyses. We found six- to eightfold increases in potencies of compounds 6 (IC₅₀: 2.9 ± 0.6 µM) and 8 (IC₅₀: 2.6 ± 1.3 µM) relative to ciliobrevin D (IC₅₀ of 20 ± 1.0 µM) in this assay (Figure 3G,H). To our knowledge, compound 8 is the most potent inhibitor of dynein 2 published to date. Hereafter, we designate compound 8 as dynapyrazole-A and compound 6 as dynapyrazole-B.

In order to directly test the effect of replacing the benzoylacrylonitrile-quinazolinone system by a pyrazoloquinazolinone, we also synthesized uncyclized congeners of dynapyrazoles. Due to poor solubility of a 6-iodo-substituted ciliobrevin derivative it was not possible to test its activity and compare it with that of compound 8. Comparisons between compound 6 (dynapyrazole-B) and compound 9, a 6-CF3-substituted ciliobrevin derivative (Figure 3H), show that cyclization leads to ~6-fold improvement in the potency of dynein 2 inhibition in the microtubule gliding assay. 

We next examined the inhibition of dynein 2 by dynapyrazole-A in cell-based assays. In cell culture, serum starvation results in the formation of primary cilia, which are required for Hedgehog signaling (Goetz and Anderson, 2010). Quiescent cells respond to the Hedgehog ligand or to a synthetic agonist and pathway activity can be measured using a Gli-driven luciferase reporter (Taipale et al., 2000; Chen et al., 2002). Expression of Hedgehog-driven luciferase reporter was inhibited by dynapyrazole-A (Compound 8, IC₅₀: 1.9 ± 0.6 µM) ~8 fold more potently than ciliobrevin D (IC₅₀: 15.5 ± 3 µM, Figure 3I). Cell death was observed at concentrations 10-fold above the IC₅₀ for dynapyrazole-A (20 µM) over the 28 hr time course of this experiment. In this assay, we have also observed cell death at high ciliobrevin D concentrations (200 µM,~10x above its IC₅₀). To stimulate Hedgehog pathway activity, we used the synthetic agonist of smoothened (SAG) (Chen et al., 2002), which competes with many known Hedgehog pathway inhibitors for binding to Smoothened, a key protein in this signaling pathway (Sharpe et al., 2015). As dynapyrazole-A inhibits Hedgehog signaling at high SAG concentrations (500 nM), it is likely to act downstream of Smoothened, as was previously noted for ciliobrevins and consistent with inhibition of dynein 2 (Hyman et al., 2009; Firestone et al., 2012).

The Hedgehog signaling pathway depends on dynein-2-driven retrograde intraflagellar transport in the primary cilium (Goetz and Anderson, 2010). We used time-lapse microscopy to examine the dynamics of fluorescently labeled intraflagellar transport protein-88 (mNeonGreen-IFT88, Figure 4A–C, Videos 1–4), an approach we have previously used to study the effect of ciliobrevins and derivatives (Ye et al., 2013; See et al., 2016). Time-lapse recordings of cells treated with control media revealed a steady flow of IFT88 punctae moving from cilium tip to base and vice versa, as expected (Videos 1 and 2). The wider end of the cilium was identified as its base, a convention suggested previously by others (Yang et al., 2015). We find that in cilia treated with dynapyrazole-A (compound 8, 5 µM), retrograde-directed IFT88 punctae were markedly slowed (Videos 3 and 4). In contrast, anterograde motion did not appear to be substantially altered in the presence of dynapyrazole-A.

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To quantitatively assess the effect of dynapyrazole-A treatment on intraflagellar transport, we analyzed time-lapse images of cilia using KymographDirect, an automated analysis algorithm that extracts particle velocities from kymographs (Mangeol et al., 2016). Previous analyses of the effect of dynein 2 inhibition using a temperature-sensitive mutant in Chlamydomonas revealed that dynein 2 depletion causes a ~60–70% reduction in retrograde velocities and a ~20% reduction in anterograde velocities as well as 30–60% reductions in the frequency of particle transport in both directions (Engel et al., 2012). Under control conditions (0.3% DMSO, Figure 4B), anterograde particles moved with a speed of 694 ± 117 nm/s (Figure 4D and F, mean ± S.D., 429 particles, 38 cilia) and retrograde particles moved at 421 ± 156 nm/s (Figure 4D and F, 244 particles, 38 cilia), consistent with previous studies (Ye et al., 2013). Following addition of dynapyrazole-A to cells, the speed of retrograde particles was markedly reduced at five minutes, the fastest reliable time line for this experiment on our microscopy set-up (Figure 4C,E and F; 5 µM compound 8: mean velocity 156 ± 107 nm/s, 211 particles, 52 cilia). In contrast, anterograde particle velocities were only reduced by ~18% (Figure 4C,E and F, 5 µM 8: 566 ± 116 nm/s, 443 particles, 52 cilia). After 10 min of treatment, reductions in velocities were similar to those at the 5 min time point (Figure 4—figure supplement 1). Treatment of cilia with a higher dynapyrazole-A concentration (10 µM) slowed both retrograde- and anterograde-directed motion (Figure 4—figure supplement 2). Again, retrograde motion was more strongly inhibited. Dynapyrazole-A treatment (5 µM and 10 µM) also reduced the frequency, that is, the number of particles moving across a cilium per minute, in both anterograde and retrograde directions (Figure 4G, Figure 4—figure supplement 2). We note that dynapyrazole-A, at concentrations close to the IC₅₀ for inhibiting microtubule gliding in vitro, alters intraflagellar transport in a manner similar to what has been observed following dynein 2 loss-of-function in Chlamydomonas (Engel et al., 2012).

We next examined whether inhibition of intraflagellar transport by dynapyrazole-A was reversed following washout of the compound. Ciliated cells treated with dynapyrazole-A (5 µM compound 8, 5 min) were transferred to solvent-control media with serum (0.3% DMSO, 10% FBS) and incubated for an additional 10 min. Both retrograde and anterograde velocities recovered to control levels (Figure 4F,velocities following washout: retrograde: 467 ± 136 nm/s, 173 particles, 18 cilia; anterograde: 697 ± 149 nm/s, 256 particles, 18 cilia) as did transport frequencies (Figure 4G). When media with a lower serum concentration was used in washout experiments, retrograde velocities recovered only partially, suggesting that serum may accelerate the partitioning of this compound out of cells (Figure 4—figure supplement 3). Taken together, our data suggest dynapyrazole-A is likely to be a useful reversible probe to study intraflagellar transport.

We predicted that dynapyrazole-A, like ciliobrevin D, should also inhibit cytoplasmic dynein 1 (See et al., 2016). To examine the inhibition of dynein 1 by dynapyrazole-A in vitro we generated recombinant human protein. We expressed and purified a GFP-tagged human dynein 1 (AA 1320–4646) construct similar to the one we used for GFP-dynein 2. This protein migrated with a peak elution volume of 12.6 mL in size exclusion chromatography and SDS-PAGE analysis showed >90% purity (Figure 5A,B). GFP-dynein 1 moves microtubules at 508 ± 60 nm/s (n = 5 independent experiments, 191 microtubules analyzed), a velocity expected based on studies of other mammalian dynein 1 homologs (Yamada et al., 2008). Under these conditions, ~97% of the filaments analyzed had velocities >50 nm/s (Figure 5—figure supplement 1). Montages showed that both ciliobrevin D and dynapyrazole-A slowed dynein-dependent microtubule gliding (Figure 5C). Dose-dependent analysis indicated that dynapyrazole-A blocked GFP-dynein-1-driven motility with an IC₅₀ of 2.3 ± 1.4 µM,~6-fold more potently than ciliobrevin D (15 ± 2.9 µM, Figure 5D). Inhibition of dynein-1-dependent microtubule gliding by dynapyrazole-A was reversed following washout, as is also the case for dynein-2-dependent motility (Figure 5—figure supplement 3). As has been previously noted for ciliobrevin D, the potency of dynapyrazole-A was sensitive to the protein (e.g. blocking agent, serum) concentration in solution (Figure 5—figure supplement 4), likely due to the hydrophobicity of these compounds (calculated logarithm of octanol:water partition coefficient [ClogP]: ciliobrevin D: 4.5; dynapyrazole-A: 4.2) (Firestone et al., 2012). We therefore focused our cell-based analyses of dynapyrazole-A on dynein-1-dependent processes in assays that do not require high serum concentrations.

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Lysosome transport can be observed in the neurites of CAD cells, a murine cell line that displays neuron-like properties in serum-free cell culture media (Figure 5E–M and Videos 5–7) (Qi et al., 1997). This bidirectional transport of lysosomes requires kinesins and dynein 1 and can be observed by imaging live cells stained with the LysoTracker dye (Vale, 2003; Pu et al., 2016). A coupling between anterograde and retrograde motion has been described for membrane-bound organelles, with disruption of dynein-driven motion blocking bi-directional organelle motility (Barlan et al., 2013). We imaged control solvent (0.1% DMSO) treated cells and observed bidirectional motion of LysoTracker-labelled puncta (Figure 5E,H,K, Video 5). Overlays of successive images from a time-lapse series, color-coded for displacement, reveal organelle displacements (Figure 5H–J). Quantitative analysis using kymographs shows that ~20% of particles move at speeds below 25 nm/s in both anterograde and retrograde directions under control conditions and a similar fraction of particles have velocities above 250 nm/s (Figure 5K and N). Treatment with dynapyrazole-A suppressed bidirectional motion. In particular, the percentage of retrograde-directed lysosomes moving slower than 25 nm/s increased to ~47% (3.5µM) and ~59% (5µM) while the proportion of lysosomes moving at speeds > 250 nm/s decreased to below 10% (Figure 5N). We observed equivalent effects of dynapyrazole-A on anterograde motion velocities (kymographs in Figure 5K,L and M). In control cells, lysosomes with measureable displacements had trajectories that averaged 32 ± 11 µm (Figure 5O). Dynapyrazole-A (3.5 µM, Figure 5I and L) reduced the average track length of moving particles to 15.7 ± 4.7 µm, which can be seen as white punctae that result from the overlay of multiple color-coded frames without particle translocation. A higher dose of dynapyrazole-A 5 µM) shortened the average track length of moving particles to 6.7 ± 2.7 µm (Figure 5J,M,O). We found that ATP levels in cells treated with dynapyrazole-A at concentrations that inhibited lysosome transport (5 µM) were stable for up to 3 hr (Figure 5—figure supplement 5). Higher concentrations of dynapyrazole-A (15 µM) lowered intracellular ATP levels by ~70% at 3 hr (Figure 5—figure supplement 5). Together, these data suggest that at concentrations near its in vitro IC₅₀, dynapyrazole-A suppresses lysosome transport in cells via inhibition of dynein 1.

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Dynein 1, whose biochemical activity has been extensively characterized, serves as a model for the mechano-chemistry of this motor protein family. Therefore, to dissect the mechanism of inhibition of dynein by dynapyrazole-A, we focused on this isoform. The basal ATPase activity of GFP-dynein was 0.62 ± 0.2 s⁻¹ (Figure 6A, n = 8 , mean ± S.D.). We also purified an N-terminally polyhistidine (His)-tagged dynein motor domain construct (AA 1320–4646) that was routinely obtained in five-fold higher yield than the GFP-tagged construct (Figure 6B). Following a three-step purification protocol, we obtained this protein with >80% purity. The specific activity of His-dynein 1 (0.66 ± 0.1 s⁻¹, n = 11, mean ± S.D. Figure 6C) was similar to that of GFP-dynein. The basal ATPase rates of these human dynein constructs were within the range observed by others for mammalian dyneins (Shpetner et al., 1988; Ori-McKenney et al., 2010; Nicholas et al., 2015b). We examined the inhibition of dynein 1’s ATPase activity by dynapyrazole-A. Remarkably, a high concentration of dynapyrazole-A (40 µM) did not inhibit the basal ATPase activity of GFP-dynein 1 (Figure 6A) and only partially inhibited His-dynein 1 (~30% inhibition Figure 6C). Under similar conditions, ciliobrevin D (40 µM) inhibited the ATPase activity of both GFP-dynein 1 and His-dynein 1 by ~70% (Figure 6A and C). The rather modest inhibition of dynein's ATPase activity by dynapyrazole-A was unexpected, as dynapyrazole-A can inhibit microtubule gliding at a 20-fold lower concentration.

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We next examined if dynapyrazole-A inhibits dynein 1’s microtubule-stimulated ATPase activity. We find that human dynein’s ATP hydrolysis rate is stimulated in a microtubule concentration-dependent manner (Figure 6D, 2.5 µM: 1.2 ± 0.05 s⁻¹, n = 3; 8 µM: 1.5 ± 0.3 s⁻¹, n = 4; 15 µM: 2.2 ± 0.5 s⁻¹, n = 4). Dynapyrazole-A (30 µM) inhibited the microtubule-stimulated ATPase activity to a residual rate of ~0.5 s⁻¹ (8 or 15 µM microtubules, Figure 6D). Dose-dependent analysis was carried out at a microtubule concentration (2.5 µM) that gave two-fold stimulation of dynein. At increasing concentrations of dynapyrazole-A dynein’s microtubule-stimulated ATPase activity saturated at ~0.5 s⁻¹ (Figure 6E). Fitting to this non-zero plateau value gave an IC₅₀ of dynapyrazole-A of 6.2 ± 1.6 µM. In comparison, dose-dependent inhibition of dynein 1’s microtubule-stimulated ATPase activity by ciliobrevin D did not reveal a plateau up to the highest concentrations the compound remained soluble in these assay conditions (Figure 6—figure supplement 2).

To better understand how dynapyrazole-A inhibits dynein activity, we monitored its effect on ADP-vanadate-dependent photocleavage of dynein. In this assay, an established read-out of nucleotide binding at dynein's AAA1 site (Lee-Eiford et al., 1986; Kon et al., 2004), ultraviolet irradiation leads to ADP-vanadate-dependent photocleavage of dynein at the AAA1 site, resulting in two protein fragments (Gibbons and Gibbons, 1987). Using His-dynein 1, 100 µM ATP, and 100 µM vanadate, we observed 42 ± 3% photocleavage (n = 3) of His-dynein 1 (Figure 6F and G) and fragments consistent with cleavage at AAA1. At a dynapyrazole-A concentration near its IC₅₀ for in vitro dynein inhibition (4 µM), ADP-vanadate dependent photocleavage of dynein was reduced ~2.5-fold relative to controls (17 ± 2% photocleavage, n = 3). These data are consistent with dynapyrazole-A binding at the AAA1 site.

To further dissect how dynein is inhibited by dynapyrazole, we expressed and purified a construct with a Walker A lysine-to-alanine substitution at AAA3 (K2601A), a mutation that disrupts nucleotide binding at AAA3 (Kon et al., 2004; Silvanovich et al., 2003). We found that the mutant protein eluted during size exclusion chromatography with a major peak at a similar elution volume to that for the wild-type construct (12.4 mL for the mutant, 12.2 mL for wild-type, Figure 6H and I), suggesting that the mutations do not disrupt overall protein folding. We also purified a comparable AAA1 (K1912A) mutant, but this protein had ~80–90% reduced ATPase activity, making analysis of its ATPase activity difficult (Figure 6—figure supplement 3). We found that the AAA3 mutant had basal activity that was elevated relative to wild-type (1.1 ± 0.2 s⁻¹, n = 5), as has been previously shown for yeast dynein with a similar mutation(DeWitt et al., 2015). The ATPase activity of this mutant was stimulated by microtubules to 1.5 s⁻¹ (3 µM microtubules) and 1.7 s⁻¹ (11.5 µM microtubules) (Figure 6J). The reduced microtubule stimulation of the ATPase activity is consistent with findings for comparable dynein constructs from other organisms (DeWitt et al., 2015; Kon et al., 2004).

Across the range of microtubule concentrations tested, dynapyrazole-A (30µM) reduced the AAA3 mutant dynein’s ATPase activity by 70–80% (Figure 6J). Surprisingly, unlike what we observed for the wild-type protein, strong inhibition of the basal activity of this mutant construct was also observed (~70% inhibition; 30 µM dynapyrazole-A). A dose-dependent analysis showed that dynapyrazole-A inhibits the basal activity of the AAA3 mutant with an IC₅₀ of 5.5 ± 1.6 µM, while ciliobrevin D has an IC₅₀ of 38 ± 6 µM (Figure 6K). Even in the context of the AAA3 mutant enzyme, a residual ATPase activity of ~0.2 s⁻¹ was observed at the highest dynapyrazole-A concentrations tested (Figure 6K). It is noteworthy that dynapyrazole-A inhibits basal activity of the AAA3 mutant and the microtubule-stimulated ATPase activity of the wild-type enzyme with similar potency, suggesting that a common ATPase site, likely AAA1, might be inhibited in both contexts.

In this study, we analyzed the structure of the ciliobrevins and designed tricyclic derivatives in which the benzoylacrylonitrile of ciliobrevin was replaced with a cyanopyrazole. These compounds inhibited dynein in vitro and in cells more potently than ciliobrevins and have improved chemical properties. We also find that dynapyrazole-A inhibits dynein by a mechanism that is distinct from that of ciliobrevins.

The pyrazoloquinazolinone scaffold of the dynapyrazoles addresses many of the limitations of the ciliobrevins. First, activated acrylonitriles, such as the benzoylacrylonitrile at the core of the ciliobrevin scaffold, have been shown to be susceptible to attack by intracellular nucleophiles, raising the possibility of unwanted reactivity of these dynein inhibitors (Serafimova et al., 2012). While ciliobrevins have been observed to be unstable under standard laboratory storage conditions, the pyrazoloquinazolinone of the dynapyrazoles is unlikely to be reactive or unstable but retains many of the electrostatic and steric elements present in ciliobrevin (Sainath and Gallo, 2015). Second, the potency of dynapyrazole-A is similar across in vitro and cell-based assays, consistent with dynein being its cellular target. This was not consistently observed for ciliobrevin analogs. Dynapyrazole-A and -B inhibited dynein-dependent microtubule gliding six- to eight-fold more potently than ciliobrevins; however, different mechanisms of inhibition make it difficult to relate this change in potency directly to an increased binding affinity. Third, the ciliobrevin scaffold is present in chemical screening libraries and a close derivative has annotated anti-malarial activity (Pillai et al., 2010). In contrast, to our knowledge, the aryl-substituted or cyclopropyl-phenyl-substituted cyano-pyrazoloquinazolinone structure of compounds 4-8 has not previously been reported in a bioactive compound. The hydrophobicity of the dynapyrazoles remains a limitation that needs to be addressed. We note that the high ClogP value of dynapyrazole-A, which is similar to that of ciliobrevins, may explain why this compound, like ciliobrevin D, has reduced activity in the presence of high protein concentrations (e.g. serum). Likewise, acute reversal of intraflagellar transport inhibition was most effective using cell culture medium with serum. We believe that the dynapyrazole scaffold will be valuable for further optimization of these chemical probes.

In current models, the motion of membrane-bound organelles requires a balance of motors moving toward microtubule plus- and minus-ends (Barlan et al., 2013). Studies of different organelles across several cell types have established that depletion or inhibition of dynein reduces bidirectional organelle motion (Waterman-Storer et al., 1997; Gross et al., 2002; Martin et al., 1999). Remarkably, organelle motion following dynein depletion can be restored by add-back of a minus-end directed kinesin, suggesting that micromechanical coupling is involved and an opposing force is needed to activate directional motion of the motor proteins that drive the directional motion of these cellular cargoes (Barlan et al., 2013; Ally et al., 2009). These observations provide a plausible explanation for the bi-directional inhibition of lysosome motility upon dynapyrazole-A treatment. Although the individual cargos in the cilium are not likely membrane-bound, this intracellular transport is no less complex than organelle movement in the bulk cytoplasm (Prevo et al., 2017). We believe that dynapyrazole-A will be a useful tool to dissect how the length and shape of the cilium is linked to directional transport and to tease apart the contributions of active transport and passive diffusion of proteins that participate in Hedgehog signaling.

Dynapyrazole-A blocks both dynein 1 and dynein 2. As dynein 1 plays a critical role in numerous cellular processes, its persistent inhibition is expected to be toxic to cells (Figure 5—figure supplement 5). However, acute inhibition may be useful in deciphering dynein function in different contexts. Can dynapyrazoles be chemically modified to develop inhibitors that are selective for dynein 1 or 2? Previously, we found that modest selectivity for inhibition of dynein 2 relative to dynein 1 can be achieved under certain conditions ([ATP] <1 µM) by appending tolyl ethers at the 7-position of the ciliobrevin quinazolinone (See et al., 2016). However, inhibition of dynein's ATPase activity by these ciliobrevin analogs is suppressed at higher ATP concentrations (e.g. 100 µM) in vitro. Studies of mammalian dynein 1 indicate that there are two ATP-binding sites with K_m values separated by two orders of magnitude (~2 µM and ~600 µM [Ross et al., 2006]). It is possible that the observed selectivity is achieved by targeting the high-affinity ATPase site and therefore, at higher, close to physiologic ATP concentrations, the inhibition by these ciliobrevin analogs is suppressed. In contrast, dynapyrazoles inhibit dynein at high ATP concentrations (1 mM), consistent with a different mode of inhibition. Also, substitution of this compound with a methyl ether at the 6-position abrogated dynein inhibition. These data suggest that the selectivity gains observed from appending tolyl ethers to the ciliobrevin scaffold are unlikely to be applicable to dynapyrazoles and developing a dynein isoform-selective dynapyrazole derivative will require more extensive structure-activity relationship studies.

Of the six AAA sites in dynein, only mutations at AAA1 and AAA3 have been found to block motility (Kon et al., 2004). In current models, ATPase activity at AAA1 is linked to rigid-body movements of subdomains during individual steps of the motor protein along microtubule tracks, while nucleotide state at AAA3 modulates the activity at AAA1 (Carter, 2013). Three lines of evidence indicate that dynapyrazole-A targets the AAA1 site. First, dynapyrazole-A blocks ADP-vanadate-dependent photocleavage at site 1. Second, dynapyrazole-A inhibits the activity of a dynein 1 construct with a mutation in the AAA3 domain (Walker A lysine). In this construct, the ATPase activity is expected to be mainly due to the AAA1 site. Third, sequence comparisons indicate that the AAA1 sites in dynein 1 and 2 are highly conserved, while the other AAA sites are less conserved. In fact, the residues within 4 Å of the bound nucleotide in AAA1 are identical between dynein 1 and 2 (Figure 6—figure supplement 5). This sequence similarity and our observation that dynapyrazole-A inhibits dynein 1 and 2 with comparable potency are consistent with a model in which this compound selectively inhibits hydrolysis of dynein’s AAA1 site. However, further biophysical and structural studies will be required to determine whether dynapyrazole-A binds at or near the AAA1 site.

Our finding that dynapyrazole-A does not potently inhibit ATP hydrolysis in the absence of microtubules suggests that, for dynein 1, the AAA1 site contributes to only a small fraction of this enzyme’s basal activity. We posit that a separate site, likely AAA3, accounts for the majority of dynein’s ATPase activity in the absence of microtubules. This site likely remains active even when ATPase activity at AAA1 is inhibited by dynapyrazoles. Currently, we cannot exclude the possibility that additional ATPase sites, such as AAA4, may also contribute partially to the basal ATPase activity of dynein. The nucleotide state at AAA3 is known to regulate the ATPase cycle of AAA1 when microtubules are present, with the apo- and ATP-bound states of AAA3 slowing overall hydrolysis and the ADP-bound state of AAA3 leading to rapid hydrolysis at AAA1 in wild-type dynein (Bhabha et al., 2014; DeWitt et al., 2015). Our finding that the AAA3 mutant has elevated basal activity relative to the wild-type is consistent with the protein being in a state that mimics one in which hydrolysis at AAA1 is activated. Although additional biophysical studies are needed to analyze this further, dynapyrazole-A is likely to be useful as tool for chemical ‘separation of function’ to distinguish between dynein’s microtubule stimulated and basal activities in different contexts.

While restricting conformational flexibility of ligands is a well-established strategy for improving potency, its successful application typically depends on structural data that reveal the compound conformation required to bind its target protein (Babine and Bender, 1997). In the case of the ciliobrevins, the geometric constraints we designed were guided by conformational analysis of the compounds alone and afforded ~6–8 fold improvement in potency. We note that replacement of a scaffold prone to isomerization with a fixed tricyclic pyrazoloquinazolinone did not lead directly to improved potency. Rather, incorporation of a cyclopropyl group into the dynapyrazole scaffold was required, likely by better matching the shape of the ciliobrevin pharmacophore. Our unexpected finding that conformational restriction led to changes both in potency and mechanism of inhibition raises the question of whether alternative scaffolds with different conformational constrains could reveal still other modes of dynein inhibition. More broadly, conformational restriction may be an effective strategy for improving the few AAA+ inhibitors described to date. Compounds with distinct mechanisms of action, which may result from these efforts, will be especially valuable for these multi-site enzymes whose activity is often regulated by intricate allosteric communication.

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Author details

1. Jonathan B Steinman

   Laboratory of Chemistry and Cell Biology, Rockefeller University, New York, United States

   Contribution

   JBS, Conceptualization, Supervision, Validation, Investigation, Visualization, Writing—original draft, Writing—review and editing

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0001-9492-4746

2. Cristina C Santarossa

   Laboratory of Chemistry and Cell Biology, Rockefeller University, New York, United States

   Contribution

   CCS, Formal analysis, Validation, Investigation, Visualization, Writing—review and editing, Performed and analyzed analyses of intraflagellar transport

   Competing interests

   The authors declare that no competing interests exist.

3. Rand M Miller

   Laboratory of Chemistry and Cell Biology, Rockefeller University, New York, United States

   Contribution

   RMM, Conceptualization, Validation, Investigation, Writing—original draft

   Competing interests

   The authors declare that no competing interests exist.

4. Lola S Yu

   Laboratory of Chemistry and Cell Biology, Rockefeller University, New York, United States

   Contribution

   LSY, Conceptualization, Validation, Investigation

   Competing interests

   The authors declare that no competing interests exist.

5. Anna S Serpinskaya

   Department of Cell and Molecular Biology, Feinberg School of Medicine, Northwestern University, Chicago, United States

   Contribution

   ASS, Investigation, Visualization, Performed and analyzed lysosome motility experiments

   Competing interests

   The authors declare that no competing interests exist.

6. Hideki Furukawa

   Tri-Institutitional Therapeutics Discovery Institute, New York, United States

   Contribution

   HF, Conceptualization, Formal analysis, Supervision, Validation, Investigation, Writing—original draft, Designed and executed synthesis of compounds in this manuscript, Performed analytical validation of their identity and purity, Also supervised synthetic research efforts of others

   Competing interests

   The authors declare that no competing interests exist.

7. Sachie Morimoto

   Tri-Institutitional Therapeutics Discovery Institute, New York, United States

   Contribution

   SM, Formal analysis, Validation, Investigation, Designed and executed synthesis of compounds in this manuscript, Performed analytical validation of their identity and purity

   Competing interests

   The authors declare that no competing interests exist.

8. Yuta Tanaka

   Tri-Institutitional Therapeutics Discovery Institute, New York, United States

   Contribution

   YT, Formal analysis, Validation, Investigation, Designed and executed synthesis of compounds in this manuscript, Performed analytical validation of their identity and purity

   Competing interests

   The authors declare that no competing interests exist.

9. Mitsuyoshi Nishitani

   Pharmaceutical Research Division, Takeda Pharmaceuticals Ltd, Kanagawa, Japan

   Contribution

   MN, Formal analysis, Validation, Investigation, Performed and analyzed x-ray crystallography experiments

   Competing interests

   The authors declare that no competing interests exist.

10. Moriteru Asano

    Tri-Institutitional Therapeutics Discovery Institute, New York, United States

    Contribution

    MA, Formal analysis, Validation, Investigation, Designed and executed synthesis of compounds in this manuscript, Performed analytical validation of their identity and purity

    Competing interests

    The authors declare that no competing interests exist.

11. Ruta Zalyte

    Medical Research Council Laboratory of Molecular Biology, Cambridge, United Kingdom

    Contribution

    RZ, Resources, Supervision, Investigation

    Competing interests

    The authors declare that no competing interests exist.

12. Alison E Ondrus

    Division of Chemistry and Chemical Engineering, California Institute of Technology, Pasadena, United States

    Contribution

    AEO, Formal analysis, Supervision, Investigation, Supervised and helped analyze two-dimensional NMR experiments

    Competing interests

    The authors declare that no competing interests exist.

13. Alex G Johnson

    Department of Chemical and Systems Biology, Stanford University School of Medicine, Stanford, United States

    Contribution

    AGJ, Formal analysis, Investigation, Performed and helped analyze two-dimensional NMR experiments

    Competing interests

    The authors declare that no competing interests exist.

14. Fan Ye

    Department of Molecular and Cellular Physiology, Stanford University, Stanford, United States

    Contribution

    FY, Supervision, Methodology, Writing—review and editing, Helped guide the analysis and interpretation of intraflagellar transport assays

    Competing interests

    The authors declare that no competing interests exist.

15. Maxence V Nachury

    Department of Molecular and Cellular Physiology, Stanford University, Stanford, United States

    Contribution

    MVN, Resources, Methodology, Writing—review and editing

    Competing interests

    The authors declare that no competing interests exist.

16. Yoshiyuki Fukase

    Tri-Institutitional Therapeutics Discovery Institute, New York, United States

    Contribution

    YF, Supervision, Investigation, Led chemical synthesis efforts

    Competing interests

    The authors declare that no competing interests exist.

17. Kazuyoshi Aso

    Tri-Institutitional Therapeutics Discovery Institute, New York, United States

    Contribution

    KA, Supervision, Project administration

    Competing interests

    The authors declare that no competing interests exist.

18. Michael A Foley

    Tri-Institutitional Therapeutics Discovery Institute, New York, United States

    Contribution

    MAF, Supervision, Project administration

    Competing interests

    The authors declare that no competing interests exist.

19. Vladimir I Gelfand

    Department of Cell and Molecular Biology, Feinberg School of Medicine, Northwestern University, Chicago, United States

    Contribution

    VIG, Supervision, Visualization, Writing—review and editing

    Competing interests

    The authors declare that no competing interests exist.

20. James K Chen

    Department of Chemical and Systems Biology, Stanford University School of Medicine, Stanford, United States

    Contribution

    JKC, Conceptualization, Supervision, Visualization, Writing—review and editing

    Competing interests

    The authors declare that no competing interests exist.

21. Andrew P Carter

    Medical Research Council Laboratory of Molecular Biology, Cambridge, United Kingdom

    Contribution

    APC, Resources, Supervision, Writing—review and editing

    Competing interests

    The authors declare that no competing interests exist.

22. Tarun M Kapoor

    Laboratory of Chemistry and Cell Biology, Rockefeller University, New York, United States

    Contribution

    TMK, Conceptualization, Supervision, Validation, Investigation, Writing—original draft, Project administration, Writing—review and editing

    For correspondence

    kapoor@rockefeller.edu

    Competing interests

    The authors declare that no competing interests exist.

    "This ORCID iD identifies the author of this article:" 0000-0003-0628-211X

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