SNS-032 and AT-7519 were initially identified as potent CDK inhibitors (Davis et al., 2011; Vasta et al., 2018). We profiled our extensive library of >50 SNS-032 and >100 AT-7519 analogs using the CDKL5 NanoBRET assay. In parallel, the CDKL5 actives (IC₅₀ < 1 µM) were profiled using the GSK3β NanoBRET assay. NanoBRET is a cellular target engagement assay that relies upon bioluminescence resonance energy transfer (BRET) between a tracer with a red-shifted fluorophore attached and a protein of interest with NanoLuciferase appended. Test compounds, which compete with the tracer for binding to the ATP-binding site, are introduced in a dose-dependent manner and BRET plotted versus concentration, allowing calculation of a target engagement IC₅₀ value (Vasta et al., 2018). Based on the data generated, a plate of 25 compounds was assembled. The criteria for inclusion on this plate included an IC₅₀ < 400 nM in the CDKL5 NanoBRET assay and >20fold difference between the CDKL5 NanoBRET and the GSK3β NanoBRET IC₅₀ values (Table 1).

A set of 20 CDKL5 inhibitors were selected based on their selectivity properties for CDKL5 versus GSK3α/β for testing their inhibition of CDKL5 in rat primary cortical neuron cultures. We tested whether we could detect a dose-dependent reduction of EB2 pSer222 following 1 hr incubation in culture media. We normalized pS222 EB2 with total EB2 levels to assess a specific reduction in phosphorylation as opposed to a loss of total protein levels. Almost all inhibitors showed a pSer222 EB2 reduction at 500 nM; most inhibitors showed a reduction at 50 nM (Figure 1). Three inhibitors caused a significant reduction in pSer222 EB2 at 5 nM without a change in total EB2 levels: CAF-382 (B1), HW2-013 (B4), and LY-213 (B12) (Figure 1). These represented different chemical parent backbones, so we decided to study these inhibitors further.

As shown in Figure 2, B1 is an SNS-032 analog, while B4 and B12 are analogs of AT-7519. SNS-032 and AT-7519 are published inhibitors of several members of the cyclin-dependent kinase (CDK) and cyclin-dependent kinase-like (CDKL) families (Davis et al., 2011). Before using these compounds as tools for dissecting CDKL5 biology, we needed to first understand their kinome-wide selectivity. We analyzed B1, B4, and B12 at 1 µM using the Eurofins DiscoverX scanMAX panel, which assesses binding to 403 wild-type (WT) human as well as several nonhuman and mutant kinases (Figure 3). These cell-free assays employ active site competition to quantitatively measure interactions between a test compound and a kinase. The percent of control (PoC) values are generated for each kinase contained within the panel (Davis et al., 2011).

Orthogonal binding or enzymatic assays were employed to confirm the single concentration DiscoverX scanMAX panel results above a specified PoC threshold for B1, B4, and B12. A binding assay that employs the Luceome KinaseSeeker technology was run for CDKL5 since no commercial enzymatic assay is available. Radiometric enzymatic assays that measure phosphorylation of a validated substrate in the presence of [gamma-33P]-ATP were executed for all other kinases at Eurofins. Finally, NanoBRET assays corresponding with GSK3α, GSK3β, and CDKL5 were run for all three compounds. Additional NanoBRET profiling was carried out for B1.

For B1, an analog of SNS-032, pan CDK inhibition was maintained (Figure 3A). All kinases with PoC ≤ 20 in the DiscoverX scanMAX panel plus GSK3β were profiled using orthogonal binding or enzymatic assays. While B1 has a good selectivity score (S₁₀ [1 µM] = 0.017; 7 kinases with PoC < 10 at 1 µM), several CDKs (CDK9, PCTK1/CDK16, PCTK2/CDK17, PCTK3/CDK18) are potently inhibited (IC₅₀ ≤ 100 nM) (colored green in the nested table in Figure 3A). Weaker inhibition of CDK7 as well as both GSK3α and GSK3β was noted. Analysis of biochemical data reveals potent binding/inhibition of CDKL5, CDK9, CDK16, CDK17, and CDK18, and a 44-fold selectivity window between CDKL5 and the next most potently inhibited kinase (CDK7). NanoBRET data reflects a window for potent engagement of CDKL5 at which CDKs are only weakly bound by B1 (≤100 nM). Curves corresponding to the affinity data for CDKL5, GSK3, and several CDKs are included in Figure 3—figure supplements 1 and 2A. As the binding pockets share a high degree of similarly, the selectivity of B1 within the CDKL family was further profiled via thermal shift and enzymatic assays (Figure 3—figure supplement 3). The highest change in melting temperature (ΔTm, >5°C) was noted for CDKL5, while a modest ΔTm of 3–4°C was observed for CDKL2. Radiometric enzyme assays were run for CDKL1–4 to corroborate and expand on this in-family selectivity assessment. When comparing the binding and enzymatic assay results, nearly 100-fold selectivity was observed for CDKL5 versus the most potently inhibited kinase, CDKL2. Inhibition of CDKL3 and CDKL4 by B1 was modest (IC₅₀ = 2.1–2.7 µM), and this compound did not inhibit CDKL1. We propose that only at higher concentrations (>500 nM in a biochemical assay) would inhibition of these kinases become significant, and, based on the data in Figure 3A, that an even larger window may exist between CDKL5 and CDKL2 when cells are dosed with B1. Importantly, these data support that when cells are dosed with B1 at ≤100 nM, no CDKL family member other than CDKL5 will be inhibited.

Compounds B4 and B12, which are analogs of AT-7519, also demonstrated good kinome-wide selectivity with S₁₀(1 µM) scores of 0.02 and 0.01, respectively (Figure 3B and C). These selectivity scores correspond with high-affinity binding (PoC < 10) to eight kinases for B4 and four kinases for B12 when profiled at 1 µM. Kinases with PoC < 10 in the DiscoverX scanMAX panel when B4 was screened were profiled using orthogonal binding (CDKL5) or enzymatic (remaining kinases) assays. This orthogonal profiling of B4 confirmed high-affinity binding to CDKL5, GSK3α, and GSK3β (green in nested table in Figure 3B, curves in Figure 3—figure supplements 1 and 2B), with slightly weaker affinity for GSK3α/β than CDKL5 evaluated via the respective cellular target engagement (NanoBRET) assays (Figure 3, curves in Figure 3—figure supplements 1 and 2). Orthogonal profiling of B12 confirmed high-affinity binding to CDKL5, GSK3α, and GSK3β (green in nested table in Figure 3C, curves in Figure 3—figure supplements 1 and 2C). This compound is a more effective inhibitor of GSK3α when compared to GSK3β. As was the case for B4, B12 bound with greater affinity to CDKL5 than GSK3α/β in the respective NanoBRET assays (Figure 3, curves in Figure 3—figure supplements 1 and 2C). Aside from GSK3α/β, a large (>100-fold) window exists between CDKL5 and the next most potently inhibited kinase, DYRK2, when considering the enzymatic and/or binding data.

Binding and orthogonal assay results in combination with western blots confirmed that in cells B1 is a CDKL5 and multi-CDK inhibitor with much weaker GSK3α/β affinity (>1.8 µM) and inhibitory activity. Compounds B4 and B12 exhibited similar inhibition profiles and are potent inhibitors of CDKL5, GSK3α, and GSK3β. Despite its potency, compound B4 demonstrated suboptimal selectivity when compared with B1 and B12. This was determined based on the number of WT human kinases with PoC < 35 in the DiscoverX scanMAX panel when profiled at 1 µM: B1 = 12, B4 = 17, and B12 = 9.

We employed in vitro kinase assays to evaluate the impact of B1 on CDKL5 activity (Figure 4). After preparing human WT and kinase dead (KD, CDKL5 K42R; Lin et al., 2005; Kim et al., 2020b) recombinant proteins, equal amounts were employed in CDKL5 activity assays (Figure 4B). We observed that, in the presence of ATP (50 µM, Figure 4A), WT CDKL5 but not KD CDKL5 elicited kinase activity, as measured by turnover of ATP to ADP via a luminescent reagent (ADP-Glo assay). Next, the activity of untreated WT CDKL5 was compared with WT CDKL5 treated with vehicle (DMSO) or kinase inhibitors (10 or 100 nM). B1, AST-487, and lapatinib were the inhibitors chosen. AST-487 is a confirmed, but less selective inhibitor of CDKL5 (positive control) (Canning et al., 2018). Lapatinib is a relatively selective kinase inhibitor that does not inhibit CDKL5 (negative control). The difference between DMSO, lapatinib, and untreated CDKL5 was not significant at 10 or 100 nM (Figure 4C and D). B1 and AST-487, however, robustly inhibited CDKL5 in a dose-dependent manner (Figure 4C and D). B1 was confirmed to be a potent inhibitor of CDKL5 kinase activity using this assay.

Kinase domains of GSK3β and CDKL5 have high similarity. To test the cross-talk of these inhibitors with GSK3β, we measured phospho-β-catenin (Ser33/37/Thr41), which is directly phosphorylated by GSK3β (Baltussen et al., 2018). We measured pSer222 EB2 levels normalized to total EB2 at increasing concentrations of inhibitors in primary neurons and compared this to phospho-β-catenin normalized to total β-catenin (Figure 5). We found that CAF-382 (B1) showed a significant reduction in pSer222 EB2/total EB2 at 500 nM concentration (Figure 5A). There was a trend of pSer222 EB2/total EB2 reduction from 5 nM, which was not significant (Figure 5A). Phospho-β-catenin/total β-catenin was not changed over the entire concentration range, indicating that CAF-382 (B1) does not inhibit GSK3b (Figure 5A). HW2-013 (B4) was effective in reducing EB2 pSer222 at 500 nM but also inhibited GSK3β significantly at 500 nM with a trend of inhibition from 5 nM, indicating that this inhibitor affects GSK3b (Figure 5B). Finally, LY-213 (B12) reduced pSer222 EB2 at 500 nM but also seemed to decrease β-catenin phosphorylation at this concentration. LY-213 (B12) showed comparable trends of inhibition for GSK3βand CDKL5 (Figure 5C). There were no significant differences in total CDKL5, EB2 or β-catenin levels during these treatments (Figure 5—figure supplement 1). In support of this assay, phospho-β-catenin (Ser33/37/Thr41) is reduced upon exposure to GSK3β inhibitor CHIR 99021 while EB2 pSer222 is not affected by GSK3β inhibition (Figure 5—figure supplement 2). We concluded that CAF-382 (B1) can inhibit CDKL5 activity without affecting GSK3 activity in neurons.

To determine whether CAF-382 (B1) could inhibit CDKL5 activity within intact brain tissue, we prepared acute hippocampal slices from P20–30 rats and incubated them with CAF-382 (B1) for 2 hr as described. As for primary cultures, CAF-382 (B1) reduced EB2 phosphorylation in a dose-dependent fashion (Figure 6). Similarly, SNS-032, the parent compound of CAF-382 (B1), also reduced EB2 phosphorylation in hippocampal slices (Figure 6—figure supplement 1).

Previous studies found that genetic knock-down of Cdkl5 in primary rodent brain cultures reduced glutamatergic excitatory synaptic transmission associated with loss of dendritic spines (Ricciardi et al., 2012). In this approach, the time course of loss of CDKL5 function is expected to occur over several days. To clarify the more acute role of CDKL5 in regulating excitatory synaptic transmission, we applied CAF-382 (B1) (10–100 nM) to hippocampal slices while measuring field excitatory postsynaptic potentials (fEPSPs) in the CA1 hippocampus. While varying the stimulation strength applied to presynaptic Schaffer-collateral pathway inputs, we were able to measure the input–output responsiveness through measuring the input (fiber volley size) versus output (fEPSP slope). The slope of this relationship was reduced in a dose–response relationship by CAF-382 (B1) within 30 min of application (Figure 7A). In other words, for a similar input (fiber volley amplitude), the postsynaptic responsiveness (fEPSP slope) was reduced (Figure 7B) in a time- and dose-dependent fashion (Figure 7C). This suggests that the effect of acute CDKL5 inhibition by CAF-382 (B1) on glutamatergic synaptic transmission is primarily postsynaptic. Determination that amphiphysin-1, a presynaptic protein, is phosphorylated by CDKL5 (Sekiguchi et al., 2013) suggested a role of CDKL5 in presynaptic function. However, Cdkl5 knock-out studies showed that amphiphysin-1 was not phosphorylated by CDKL5 in vivo, suggesting an alternative amphiphysin-1-independent mechanism is responsible for observed presynaptic changes (Kontaxi et al., 2023). To investigate the presynaptic role of CDKL5 kinase function, the paired-pulse ratio across several paired-pulse intervals was measured (Figure 8). CAF-382 (B1) (100 nM × 1 hr) had no effect on the paired-pulse ratio, confirming the acute effects of CAF-382 (B1) are restricted to the postsynapse. fEPSP slopes are mediated by AMPA-type glutamate receptors with NMDA-type glutamate receptors largely silent under these conditions. In order to visualize the impact of CAF-382 (B1) on NMDA-type glutamate receptor mediated responses, extracellular Mg²⁺ was removed from the recording medium (Coan and Collingridge, 1985). Under these conditions, an NMDA-receptor-mediated field potential (population spike 2) becomes apparent; this was not affected by CAF-382 (B1) (Figure 9) but was completely blocked by the NMDA-type glutamate receptor antagonist D-APV (50 μM). This result suggests that the acute effect of CAF-382 (B1)-mediated CDKL5 inhibition is to selectively reduce AMPA-type glutamate receptor-mediated responses postsynaptically.

A previous study found that germ-line knock-out of Cdkl5 in the mouse enhanced LTP at 7–13-week-old (wo) hippocampal Schaffer Collateral to CA1 synapses without alteration of the input–output relationship (Okuda et al., 2017). In another study, germ-line knock-out of Cdkl5 in a mouse at 4–5 wo did not affect stable LTP at this synapse (Yennawar et al., 2019). In a rat Cdkl5 knock-out, LTP at this synapse was selectively increased at 3–4 wo but not at later ages (de Oliveira et al., 2022). To clarify the more acute role of CDKL5 in regulating LTP at this synapse, we incubated 3–4 wo hippocampal slices with CAF-382 (B1) (100 nM) for at least 30 min and, following stable baseline measurements of fEPSP slope, induced LTP via theta burst to Schaffer Collateral to CA1 synapses in the continuous presence of CAF-382 (B1). Compared to interleaved recordings from control slices, CAF-382 (B1) significantly reduced LTP at 60 min post theta-burst (Figure 10). To investigate this further, LTP at these synapses of isolated AMPA-type glutamate receptors via a pairing protocol was measured using whole-cell patch-clamp. In this case, hippocampal slices were incubated with CAF-382 (B1) (100 nM) for only 1 hr prior to, but not during, recording. Compared to interleaved recordings from control slices, CAF-382 (B1) significantly reduced LTP at 30 min post pairing (Figure 11). The induction and expression of LTP include the contributions of presynaptic and postsynaptic mechanisms at both excitatory and inhibitory synapses (Collingridge and Abraham, 2022; Nicoll, 2017). Since we do not see a presynaptic effect of CAF-382 (B1) (Figure 8), and the approach (Figure 11) removed the contribution of inhibitory synapses, this supports the direct role of CDKL5 in the postsynaptic mediation of LTP at this synapse at this age.

As our studies were all executed in vitro, the suitability of B1 for use in vivo was unknown. To gauge whether B1 could be dosed in animals using a water-based formulation, we first analyzed its aqueous kinetic solubility (Table 2). B1 demonstrated good solubility (196 µM), supporting that it readily dissolves in aqueous buffer without observed precipitation. Mouse liver microsomal stability data was also collected (Table 2). B1 proved to be very tolerant to microsomal exposure, demonstrated excellent stability (>85%) after 30 min, and was not prone to non-NADPH-dependent enzymatic degradation. This stability is not true for other kinase inhibitor scaffolds, which are significantly degraded after just 30 min (Yang et al., 2023). Since compound B1 was found to have the required aqueous solubility and mouse liver microsomal stability, it was submitted for in vivo characterization. Pharmacokinetic analysis and brain exposure of mice following intraperitoneal administration with B1 were next examined. Plasma was sampled 0.5, 1, 2, and 4 hr post-dose with 2.29 mg/kg of a salt form of B1 (Figure 12). No abnormal clinical symptoms were observed in treated animals. The half-life of B1 was found to be <1 hr, with the highest concentration of drug in the plasma found 0.5 hr after dosing (Cmax = 508 ng/mL) and total system exposure (AUCinf) of 646 h*ng/mL (Figure 12). The same dose (2.29 mg/kg) as well as a 7.63 mg/kg dose were used to analyze the brain penetration of B1 in mice 1 hr after administration. As shown in Table 3, brain and plasma sampling indicated that brain penetration of B1 was generally low.

Reagents were purchased from reputable commercial vendors, including Sigma-Aldrich, Combi Blocks, Enamine, and others, and used in accordance with safety data sheets. A rotary evaporator was employed to remove solvent(s) under reduced pressure (in vacuo). Thin-layer chromatography (TLC) as well as LC–MS were used to monitor reaction progress. The following abbreviations are used in experimental procedures: mmol (millimoles), μmol (micromoles), μL (microliters), mg (milligrams), equiv (equivalent(s)), min (minutes), and hr (hours). Compound B1 was prepared as a TFA salt with >95% purity (by HPLC) according to Scheme 1 (Figure 13) via a reductive coupling reaction in the presence of sodium borohydride and ethanol followed by amide coupling and deprotection. Compounds B4 and B12 were prepared as an HCl salt with >95% purity (by NMR) according to Scheme 2 (Figure 14). An amide coupling reaction in the presence of propylphosphonic anhydride (T3P) followed by nitro group reduction generated a common intermediate. Subsequent amide coupling reaction followed by deprotection yielded B4 and B12. Full characterization of the intermediates and final products (Figure 2) is included as (Figure 13—figure supplement 1 and Figure 14—figure supplements 1 and 2). ¹H NMR and ¹³C NMR spectra as well as microanalytical data were collected for a key intermediate and final compound B1 to confirm their identity and evaluate their purity prior to initiating cell- and animal-based studies. ¹H and ¹³C NMR spectra were collected in DMSO-d₆ using Bruker spectrometers. Magnet strength is indicated in each corresponding line listing. Peak positions are included in parts per million (ppm) and calibrated versus the shift of DMSO-d₆; coupling constants (J values) are noted in hertz (Hz); and multiplicities are included as follows: singlet (s), doublet (d), pentet of doublets (pd), and multiplet (m).

Human embryonic kidney (HEK293) cells obtained from ATCC (Manassas, VA) were cultured in Dulbecco’s Modified Eagle’s medium (DMEM, Gibco) supplemented with 10% (v/v) fetal bovine serum (FBS, Corning). These cells were incubated at 37°C in 5% CO₂ and passaged every 72 hr with trypsin (Gibco) so that they never reached confluency. Promega (Madison, WI) kindly provided full-length, human constructs for NanoBRET measurements of CDKL5 (NLuc-CDKL5; hCDKL5_1), GSK3α (NLuc-GSK3α), GSK3β (NLuc-GSK3β), CDK9 (NLuc-CDK9), CDK16 (NLuc-CDK16), CDK17 (NLuc-CDK17), and CDK18 (NLuc-CDK18) included in Table 1 as well as in Figure 3, Figure 3—figure supplement 1, and Figure 3—figure supplement 2. The fusion vector sequences for each can be found on the Promega website. hCDKL5 and rCdkl5 isoforms are homologous through the kinase domain (Hector et al., 2017b; Hector et al., 2016; Hector et al., 2017a). N-terminal NLuc orientations were used for all seven kinases. NanoBRET assays were executed in dose–response (12-pt curves) format in HEK293 cells as previously reported (Wells et al., 2021; Drewry et al., 2022b; Wells et al., 2020). Assays were executed as recommended by Promega, using 0.31 μM of tracer K11 for CDKL5, 0.13 μM of tracer K8 for GSK3α, 0.063 μM of tracer K8 for GSK3β, 0.063 μM of tracer K8 for CDK9, 0.25 μM of tracer K12 for CDK16, 0.13 μM of tracer K12 for CDK17, and 0.13 μM of tracer K12 for CDK18. Cyclin K was transfected into the cells for the CDK9 NanoBRET assay, while cyclin Y was transfected into the cells for the CDK16, CDK17, and CDK18 NanoBRET assays. Where shown, error bars represent SD.

The broad selectivity of compounds B1, B4, and B12 at 1 µM was evaluated via the Eurofins DiscoverX Corporation (Fremont, CA) scanMAX assay platform, the largest commercial kinase panel available. The scanMAX assays are cell-free, ATP-free, and employ proprietary active site competition to quantitatively measure binding between a test compound and a kinase. Different kinase constructs, truncates or full-length, are used to enable different assays for mutant, lipid, and atypical human kinases as well as pathogen kinases. Since these assays do not require ATP, values generated are reflective of thermodynamic interaction affinities and span a large dynamic affinity range. In total, 403 wild-type (WT) human kinases were included in these analyses. Percent of control (PoC) values were generated, which allowed for the calculation of selectivity scores (S₁₀ [1 µM]) noted in Figure 3 (Davis et al., 2011). Selectivity scores (S₁₀ [1 µM]) were calculated using the PoC values for only WT human kinases in the DiscoverX scanMAX panel. The S₁₀ score expresses selectivity and corresponds with the percent of the kinases screened that bind with a PoC value <10. A lower S₁₀ score indicates enhanced kinome-wide selectivity as a lower percentage of kinases bind with high affinity (PoC values closer to zero). Binding to a single kinase would generate an S₁₀ score of 0.002, while binding to 10 kinases yields an S₁₀ score of 0.025. WT human kinases in the scanMAX panel with PoC ≤ 20 (and GSK3β) for B1, with PoC < 10 for B4, and PoC < 35 for B12 are also included in Figure 3. The kinome tree diagrams in Figure 3 were created based on WT kinases with PoC < 10 for each compound.

A homogeneous competition binding assay for CDKL5 was executed at Luceome Biotechnologies, LLC (Tuscon, AZ) using their KinaseSeeker technology (Jester et al., 2010). Briefly, this is a luminescence-based assay that relies on displacement of an active site-dependent probe by an ATP-competitive test compound. A change in luminescence signal indicates binding. These assays are robust and highly sensitive with low background and minimal interference from test compounds. Test compounds were screened at 11 different concentrations in duplicate and percentage of activity remaining plotting against compound concentration to calculate the corresponding IC₅₀ value. Compounds B1, B4, and B12 were evaluated in dose–response (12-pt curve) format in duplicate. Curves with error bars showing standard deviation are included in Figure 3—figure supplements 1 and 2, and IC₅₀ values generated are embedded in tables within Figure 3.

Radiometric enzymatic assays were executed at Eurofins DiscoverX Corporation (Fremont) at the K_m value for ATP to generate dose–response (9-pt) curves for all kinases listed in Figure 3 and Figure 3—figure supplements 1 and 2 (except for CDKL5). A detailed protocol for these assays, which includes the protein constructs, substrate, and controls employed, is part of the Eurofins website: https://www.eurofinsdiscoveryservices.com. Briefly, either full-length or a truncate of a human kinase was incubated with [gamma-33P]-ATP in the presence of buffer, salts (magnesium source), and kinase-specific additives (such as cyclins for CDKs), where necessary, in the presence of a known substrate. Scintillation counting after quenching and washing with phosphoric acid allowed plotting of the data versus concentration and calculation of an IC₅₀ value. Inclusion of a positive control ensured assay performance.

The kinetic solubility of compound B1 was evaluated using an aliquot of a 10 mM DMSO stock solution dissolved in phosphate-buffered saline solution (PBS) at pH 7.4. This analysis was done by Analiza, Inc (Cleveland, OH) as described previously (Drewry et al., 2022a). The reported solubility values in Table 2 have been corrected for background nitrogen present in DMSO and the media.

Mouse liver microsomal stability of compound B1 was evaluated by Analiza, Inc as described previously (Drewry et al., 2022b). The assay was run in the presence and absence of NADPH. NADPH is required for the cytochrome P450 enzyme system, which is a common metabolism pathway of compounds. Any metabolism that occurs in the absence of NADPH would be due to non-NADPH-dependent enzymatic degradation and would be indicative of alternative metabolic pathways contributing to compound breaking down. Monoamine oxidase and carboxylesterases are examples of non-NADPH-dependent enzymes. This data generated by Analiza is included in Table 2.

Kinase domains of human CDKL1, CDKL2, CDKL3, and CDKL5 proteins were produced using the construct boundaries previously used to generate the X-ray crystal structures with PDB codes 4AGU, 4AAA, 3ZDU, and 4BGQ, respectively. 4 µM of the kinase domain of CDKL1, CDKL2, CDKL3, or CDLK5 in 10 mM HEPES-NaOH pH 7.4 and 500 mM NaCl was incubated with B1 (12.5, 25, or 50 µM) in the presence of 5× SyPRO orange dye (Invitrogen). Next, fluorescence was evaluated using a Real-Time PCR Mx3005p machine (Stratagene). A previously reported protocol was followed to run the Tm shift assays and assess melting temperatures (Fedorov et al., 2007). Data is included in Figure 3—figure supplement 2. Error bars represent SD.

These assays were executed as previously described (Lin et al., 2005; Kim et al., 2020b). Briefly, full-length FLAG-tagged WT (transcript variant II, NCBI Reference Sequence: NM_001037343.2) or kinase dead (KD, CDKL5 K42R) constructs of full-length human CDKL5 were subcloned into a pT7CFE1-CHis plasmid (Thermo Fisher). A HeLa cell lysate-based Kit (1-Step Human Coupled IVT Kit—DNA, 88881, Life Technologies) enabled in vitro translation of these constructs. His Pur cobalt spin columns (Thermo Scientific) were used to purify the in vitro-translated proteins. To execute the in vitro kinase assays, myelin basic protein (Active Motif, 31314) was employed as a substrate for recombinant CDKL5. Myelin basic protein is used as a substrate for multiple kinases, both serine/threonine and tyrosine kinases, to enable in vitro kinase assays due to the presence of multiple sites for phosphorylation. As such, we and others have used this protein as a kinase substrate for evaluating kinase activity (Kim et al., 2020b; Kameshita et al., 2014). These two components were incubated in kinase buffer (Cell Signaling, 9802) supplemented with or without ATP (50 µM) for 30 min at 30°C, followed by kinase assays run using the ADP-Glo Kinase Assay kit (Promega). After the termination of kinase assay, NuPAGE LDS Sample Buffer (4×) was added to the reaction mixture and the samples were heated at 100°C for 10 min. The protein samples were run on Invitrogen Bis-tris gradient midi-gels and transferred to PVDF membrane for western blot analysis. Primary antibody used for western blot analysis was from Cell Signaling: FLAG (#14793) and was used at 1:1,000 dilution. Anti-rabbit HRP-conjugated secondary antibody was from Jackson ImmunoResearch (#111-035-144) and used at 1:5000 dilution. Immunoblot signals were detected using SignalFire ECL Reagent (#6883) and X-ray films (Bioland #A03-02). Canon LiDE400 scanner was used for scanning the films. Precision Plus Protein Dual Color Standard prestained protein marker (Bio-Rad) was used to estimate the molecular weight of sample proteins.

The pharmacokinetics of compound B1 was evaluated by Pharmaron (San Diego, CA) following a single intraperitoneal administration to CD1 mice (two males). A dose of 2.29 mg/kg of the TFA salt of compound B1 in NMP/Solutol/PEG-400/normal saline (v/v/v/v, 10:5:30:55) was prepared just before use. This dose was calibrated due to the compound being a salt form. Plasma was sampled 0.5, 1, 2, and 4 hr post-dose. Mean values from the two animals without dispersion are shown in Figure 12. Tmax (Figure 12) was defaulted to the earliest time point and likely occurred earlier than 0.5 hr. Similarly, Cmax (Figure 12) was recorded as the concentration after 0.5 hr, the first sampling time point. Bioanalytical assays were run using Prominence HPLC and AB Sciex Triple Quan 5500 LC/MS/MS instruments, and a HALO column (90A, C18, 2.7 µm, 2.1 × 50 mm). Snapshot PK results are included in Figure 12. PK parameters were estimated by non-compartmental model using WinNonlin 8.3 (Certara, Princeton, NJ). The lower limit of quantification for plasma sampling was 1.53 ng/mL.

The brain/plasma concentration of compound B1 was evaluated by Pharmaron following a single intraperitoneal administration to CD1 mice (three males per dose). Two doses (2.29 mg/kg and 7.63 mg/kg) of the TFA salt of compound B1 in NMP/Solutol/PEG-400/normal saline (v/v/v/v, 10:5:30:55) were prepared just before use. These doses were calibrated due to the compound being a salt form. The same dose as was used for Snapshot PK (2.29 mg/kg) was chosen to have a more comprehensive profile, including brain penetrance, at this concentration. Since brain penetration was anticipated to be low, a higher concentration was also selected. Plasma and brain samples were collected 1 hr post-dose. Brain samples were homogenized at a ratio of 1:3 with PBS (W/V, 1:3) and then final brain concentrations corrected. Bioanalytical assays were run using Prominence HPLC and AB Sciex Triple Quan 5500 LC/MS/MS instruments, and a HALO column (90A, C18, 2.7 µm, 2.1 × 50 mm). Brain/plasma concentration results are included in Table 3. The lower limit of quantification for plasma sampling was 1.53 ng/mL and 0.763 ng/mL for brain.

Animals were housed in a controlled environment with 12 hr light/dark cycle. They were fed ad libitum and used in accordance with the Animals (Scientific Procedures) Act 1986 of the United Kingdom. Rat handling and housing was performed according to the regulations of the Animal (Scientific Procedures) Act 1986. Animal studies were approved by the Francis Crick Institute ethical committee and performed under U.K. Home Office project license (PPL P5E6B5A4B).

Primary cortical cultures were prepared from embryonic day (E) 18.5 embryos of Long–Evans rats as described (Roşianu et al., 2023). Pregnant females were culled using cervical dislocation, embryos were removed from the uterus, and the brains were taken out. Cortices were dissected out, pooled from multiple animals, and washed three times with HBSS. An incubation with 0.25% trypsin for 15 min at 37°C was followed by four times washing with HBSS. Cells were dissociated and then counted using a hemocytometer. Neurons were plated on 12-well culture plates at a density of 300,000 cells per well. The wells were coated with 0.1 M borate buffer containing 60 µg/mL poly-D-lysine and 2.5 µg/mL laminin and placed in the incubator overnight. Neurons were plated with minimum essential medium (MEM) containing 10% FBS, 0.5% dextrose, 0.11 mg/mL sodium pyruvate, 2 mM glutamine, and penicillin/streptomycin. After 4 hr, cultures were transferred to neurobasal medium containing 1 mL of B27 (Gibco), 0.5 mM GlutaMAX, 0.5 mM glutamine, 12.5 µM glutamate, and penicillin/streptomycin. Primary neuronal cultures were kept at 37°C and 5% CO₂. Every 3–4 d, 20–30% of the maintenance media was refreshed. Between DIV14-16, neurons were treated with 5 nM, 50 nM, 500 nM, and 5 µM of the different compounds for 1 hr. The compounds were added directly to the media and the plates were placed at 37°C for the time of the treatment. DMSO was added to the well for the control condition.

After treatment, neuronal cultures were lysed in 300 µL of 1× sample buffer (Invitrogen) containing 0.1 M DTT. Lysates were sonicated briefly twice and denatured at 70°C for 10 min. The samples were centrifuged at 13,300 rpm for 10 min and ran on NuPage 4–12% Bis-Tris polyacrylamide gels (Invitrogen). Proteins were transferred onto a Immobilon PVDF membrane (Millipore), which was then blocked in 4% milk in Tris-buffered saline containing 0.1% Tween-20 (TBST) for 30 min. Primary antibodies were incubated at 4°C overnight, and HRP-conjugated secondary antibodies at room temperature (RT) for 2 hr. The following primary antibodies were used: rabbit anti-CDKL5 (1:1000; Atlas HPA002847), rabbit anti-pS222 EB2 (1:2000; Covalab, from Baltussen et al., 2018), rat anti-EB2 (1:2000; Abcam ab45767), mouse anti-tubulin (1:100,000; Sigma T9026), rabbit anti-β-catenin (1:1000; Cell Signaling 9562), and rabbit anti phospho-β-catenin (Ser33/37 Thr41) (1:250, Cell Signaling 9561). The following secondary antibodies were used at a concentration of 1:10,000: HRP-conjugated anti-rabbit (Jackson 711-035-152), HRP-conjugated anti-mouse (Jackson 715-035-151), and HRP-conjugated anti-rat (Jackson 712-035-153). The membrane was developed using ECL reagent (Cytiva) and was visualized with an Amersham Imager 600 (GE Healthcare). Quantification of western blots was manually performed using Image Studio Lite Software (version 5.2). EB2 phosphorylation was measured relative to total EB2 and catenin phosphorylation to total catenin. The other proteins were normalized to tubulin, if not indicated otherwise.

All studies conformed to the requirements of the National Institutes of Health Guide for the Care and Use of Laboratory Rats and were approved by the Institutional Animal Care and Use subcommittee of the University of Colorado Anschutz Medical Campus (protocol 00411). Timed-pregnant Sprague–Dawley rats (Charles Rivers Labs, Wilmington, MA) gave birth in-house. All rodents were housed in micro-isolator cages with water and chow available ad libitum.

As done previously (Cornejo et al., 2007; Bernard et al., 2014; Bernard et al., 2013), following rapid decapitation and removal of the rat brain at postnatal day (P) 20–30, sagittal hippocampal slices (400 µm) were made using a vibratome (Leica VT 1200, Buffalo Grove, IL) in ice-cold sucrose artificial cerebral spinal fluid (sACSF: 206 mM sucrose, 2.8 mM KCl, 1 mM CaCl₂, 3 mM MgSO₄, 1.25 mM NaH₂PO₄, 26 mM NaHCO₃, 10 mM D-glucose, and bubbled with 95%/5% O₂/CO₂). Following removal of CA3, slices were recovered in a submersion-type chamber perfused with oxygenated artificial cerebral spinal fluid (ACSF: 124 mM NaCl, 3 mM KCl, 1 mM MgSO₄, 2 mM CaCl₂, 1.2 mM NaH₂PO₄, 26 mM NaHCO3, 10 mM D-glucose and bubbled with 95%/5% O₂/CO₂) at 28°C for at least 90 min and then submerged in a recording chamber perfused with recirculated ACSF. All electrophysiology was performed in the CA1 region at 28°C. Drugs were added to ACSF. Two twisted-tungsten bipolar-stimulating electrodes were offset in the CA1 to stimulate one or two independent Schaffer collateral-commissural pathways using a constant current source (WPI, Sarasota, FL) with a fixed duration (20 µs), each at a rate of 0.033 Hz. fEPSPs were recorded from the stratum radiatum (or stratum pyramidale region where indicated) region of CA1 using a borosilicate glass (WPI) microelectrode (pulled [Sutter, Novato, CA] to 6–9 MΩ when filled with ACSF), amplified 1000× (WPI and Warner, Hamden, CT), and digitized (National Instruments, Austin, TX) at 20 kHz using winLTP-version 2.4 (Anderson and Collingridge, 2001) to follow fEPSP slope (averaged over four EPSPs), measured using 20–80% rise times, expressed as percent of baseline, during the course of an experiment. To be sure only ‘healthy’ slices were included in our studies, responses had to meet several criteria: fiber volleys less than 1/3 of response amplitude and peak responses larger than 0.5 mV; responses and fiber volley must be stable (<5% drift). Following baseline stabilization of fEPSP slope at ~50% of maximal slope for at least 30 min, measurements of synaptic physiology were performed. Stimulation was adjusted to make 4–6 measurements from ~10 to 90% maximal slope to measure input-output (I/O) curves; recordings that deviated from linear slope were rejected. Paired-pulse ratios of slopes and peaks were measured at ~50% of maximal slope. For measurements of NMDA-receptor-dependent population spikes (Coan and Collingridge, 1985), Mg²⁺ was omitted from recording ACSF, the recording electrode was placed in stratum pyramidale, stimulation was adjusted to evoke a first population spike >2 mV, and stabilization of the second NMDA-receptor-dependent population spike (measured from the coastline of the fEPSP) for >30 min was allowed prior to measuring the effect of antagonists. Theta burst LTP was induced at test strength of 2 trains, separated by 20 s, of 10 trains of 4 stimuli at 100 Hz, separated by 200 ms. Potentiation was statistically compared over the last 5 min.

For whole-cell, voltage-clamp electrophysiological recordings of excitatory postsynaptic currents (EPSCs), 350 μm horizontal hippocampal slices were prepared from rats at P 20–30 similar to previously (Sanderson et al., 2021; Sanderson et al., 2016). After 15 min recovery at 34°C, slices were maintained at RT until recording in modified ACSF containing (in mM) 126 NaCl, 5 KCl, 2 CaCl₂, 1.25 NaH₂PO₄, 1 MgSO₄, 26 NaHCO₃, 10 glucose, and 2 N-acetyl cysteine. Slices were incubated in either CAF-382 (B1) (100 nM) or vehicle (water) in modified ACSF for at least 1 hr prior to recording. Using infrared–differential interference contrast microscopy for visualization, whole-cell recordings (Axopatch 200B amplifier and pClamp software [Molecular Devices]) from CA1 pyramidal neurons (series resistance between 3 and 6 MΩ) were performed at 29–30°C with an intracellular solution containing (in mM) 130 Cs gluconate, 1 CsCl, 1 MgSO₄, 10 HEPES, 1 EGTA, 4 MgATP, 0.4 MgGTP, spermine (0.01), and 2 QX-314, pH 7.3. AMPA-type EPSCs, evoked using a bipolar tungsten-stimulating electrode as above, at a holding potential of −65 mV were pharmacologically isolated using picrotoxin (50 μM; Tocris). EPSC peaks were evoked every 20 s. After establishing whole-cell access, a 3 min baseline recording at −65 mV was followed by LTP induction using a pairing protocol: cells were depolarized to 0 mV for 90 s and stimulated every 20 s then followed by delivery of 1 s × 100 Hz high-frequency stimulation. Evoked EPSC amplitudes were subsequently monitored at –65 mV. Potentiation was statistically compared over the last 5 min.

Hippocampal slices were prepared and recovered as for electrophysiology with the addition of cuts to isolate CA1 from the remaining hippocampus; after recovery, the slices from a given rat were incubated in inhibitor or vehicle (water) in ACSF for 2 hr. Following this, slices were suspended in STE buffer, sonicated, boiled for 5 min, and then frozen until further use. To minimize the effects of slice preparation (Osterweil et al., 2010) and control for slice quality, only slices that came from a preparation that met electrophysiological criteria were used. All concentrations were quantified with BCA and then loaded in duplicate on 10% polyacrylamide gel and a five-point dilution series of naive rat hippocampal homogenate was included on each gel as a standard curve for quantification of immunoreactivity/μg loaded protein (Grosshans and Browning, 2001). Following transfer to PolyScreen PVDF transfer membrane (Genie, Idea Scientific Company, Minneapolis, MN), blots were blocked in BSA or Carnation nonfat dry milk for 1 hr and incubated either 1 hr at RT or overnight at 4°C with antibodies. The following primary antibodies were used: sheep anti-CDKL5 (1:2000; University of Dundee, anti-CDKL5 [350-650]), rabbit anti-pS222 EB2 (1:1000; Covalab, from Baltussen et al., 2018), and rat anti-EB2 (1:1000; Abcam ab45767). Blots were then subjected to three 10 min washings in Tris-buffered saline (140 mm NaCl, 20 mm Tris pH 7.6) plus 0.1% Tween 20 (TTBS), before being incubated with anti-sheep, anti-rat, or anti-rabbit secondary antibody (1:5000) in 1% BSA or milk for 1 hr at RT, followed by three additional 10 min washes preformed with TTBS. Immunodetection was accomplished using a chemiluminescent substrate kit (SuperSignal West Femto Maximum Sensitivity Substrate; Pierce) and the Alpha Innotech (Alpha Innotech, San Leandro, CA) imaging system. Quantification was performed using AlphaEase software (Alpha Innotech) and Excel (Microsoft, Redmond, WA). Immunoreactivity was reported as the density of sample bands relative to the standard curve. For phospho-proteins, ratios of immunoreactivity/μg to totals are reported without standardization. Only values falling within the standard curve generated from the dilution series included on each gel were incorporated into the final analysis. Some of the blots were then stripped (Restore PLUS Western Blot Stripping Buffer; Thermo Scientific, Rockford, IL) and reblotted if needed.

Data were analyzed via GraphPad Prism 9. Each assay was run in triplicate, and mean values are graphed in Figure 4 with error bars showing standard deviation (SD). In Figure 4, statistical analysis was done using one-way ANOVA with Dunnett’s multiple-comparisons test. Thresholds for significance (p-values) were set at ***p<0.0001 and nonsignificant statistics were not included.

Data were analyzed using GraphPad Prism 9. Exact values of n and statistical methods are mentioned in the figure legends. Each concentration was compared to the control using a one-way ANOVA test. A p-value >0.05 was not considered statistically significant. Thresholds for significance were placed at *p≤0.05, **p≤0.01, ***p≤0.001, and ****p≤0.0001. All errors bars in the figures are SD where indicated. Nonsignificant statistics were not included.

Data are expressed as mean ± SEM with n = number of rats for a given treatment, unless otherwise stated. Data are plotted using SigmaPlot 12.5 (Systat, Chicago, IL). Mann–Whitney rank-sum, one-way, two-way, and repeated measures (RM) ANOVA (Holm–Sidak post hoc testing) and Student’s t-tests were used, where indicated and appropriate, for statistical comparisons for electrophysiological and biochemical using SigmaPlot 12.5 (Systat). Significance is reported at p<0.05 unless otherwise stated. All errors bars in the figures are standard error (SE) where indicated. Nonsignificant statistics were not included.