A LOV (Light, Oxygen, or Voltage) domain containing blue-light photoreceptor ZEITLUPE (ZTL) directs circadian timing by degrading clock proteins in plants. Functions hinge upon allosteric differences coupled to the ZTL photocycle; however, structural and kinetic information was unavailable. Herein, we tune the ZTL photocycle over two orders of magnitude. These variants reveal that ZTL complexes with targets independent of light, but dictates enhanced protein degradation in the dark. In vivo experiments definitively show photocycle kinetics dictate the rate of clock component degradation, thereby impacting circadian period. Structural studies demonstrate that photocycle dependent activation of ZTL depends on an unusual dark-state conformation of ZTL. Crystal structures of ZTL LOV domain confirm delineation of structural and kinetic mechanisms and identify an evolutionarily selected allosteric hinge differentiating modes of PAS/LOV signal transduction. The combined biochemical, genetic and structural studies provide new mechanisms indicating how PAS/LOV proteins integrate environmental variables in complex networks.https://doi.org/10.7554/eLife.21646.001
Many living organisms track the 24-hour cycle of day and night via collections of proteins and other molecules that together act like an internal clock. These clocks, also known as circadian clocks, help these organisms to predict regular changes in their environment, like light and temperature, and adjust their activities according to the time of day.
Plants use circadian clocks to predict, for example, when dawn will occur and get ready to harness sunlight to fuel their growth. A plant called Arabidopsis thaliana has a light-sensitive protein called ZEITLUPE (or ZTL for short) that helps it keep its circadian clock in sync with the cycle of night and day. Previous studies have shown that light activates this protein causing part of it to change shape and then revert back after a period of about an hour and a half. However, it was unclear if this timing was important for ZEITLUPE to allow plants to keep track of time.
To help answer this question, Pudasaini et al. set out to identify a specific chemical event behind ZEITLUPE’s changes in shape. A chemical bond forms when light activates ZEITLUPE, and it turns out that how long this bond lasts before it breaks plays an important role in allowing plants to maintain a 24-hour circadian clock. This chemical bond controls the shape changes that guide the protein’s activities and, when Pudasaini et al. modified ZEITLUPE so that it took much longer for this bond to break, they could tune how fast the plant’s internal clocks run. In essence, the time between the bond forming and breaking breaks acts like a countdown on a stopwatch, and it must be precisely timed to keep the clock in pace with the environment.
These findings improve our understanding of how light can regulate an internal biological clock. This improved understanding could, in the future, allow researchers to manipulate how plants and other organisms respond to their environment. This in turn could change how these organisms develop, and how much they grow. As such, extending these findings into agricultural crops may one day lead to new ways to increase crop yields.https://doi.org/10.7554/eLife.21646.002
Organisms have developed signaling networks to measure and respond to daily (circadian) and seasonal (photoperiodic) alteration in environmental variables. Key to these circadian and photoperiodic responses is measurement of day length through complicated feedback loops involving sensory proteins. These sensory proteins involve members of the Period-ARNT-Singleminded and its Light, Oxygen, or Voltage (PAS/LOV respectively) subclass that couple photic-input to metabolic and stress pathways (Lokhandwala et al., 2015; Sahar and Sassone-Corsi, 2009; Taylor and Zhulin, 1999). Currently, how these signaling components are integrated is poorly understood due to a difficulty in decoupling photochemistry from allosteric protein changes and signal transduction. In plants and fungal species, LOV domain containing proteins act at signaling nodes to integrate sensory responses into circadian, reproductive and stress pathways (Lokhandwala et al., 2015; Imaizumi et al., 2003; Somers et al., 2000; Zoltowski et al., 2007). Central to their function are two key elements of the PAS family: (1) The ability to sense and respond to diverse environmental stimuli, and (2) The presence of multiple interaction surfaces that engage targets in a selective manner (Figure 1, Figure 1—figure supplement 1). The ability to trigger these elements with light has positioned LOV proteins as an allosteric model and enabled the development of LOV optogenetic tools (Pudasaini et al., 2015). However, limited understanding of how chemistry is linked to allostery and downstream signaling hampers our understanding of these systems.
Our current understanding of LOV signaling has benefited from detailed structural studies that revealed amino acid sites that tune LOV allostery without affecting LOV photocycle kinetics (Zoltowski et al., 2007; Harper et al., 2004; Jones et al., 2007; Salomon et al., 2000). A consensus mechanism is summarized. LOV domains are typified by blue-light induced formation of a C4a adduct between a conserved Cys residue and a bound flavin (FAD, FMN or riboflavin) cofactor (Figure 1—figure supplement 1D) (Zoltowski and Gardner, 2011). C4a adduct formation then drives rotation of a conserved Gln residue to initiate conformational changes within N- or C-terminal extensions (Ncap/Ccap) to the LOV core (Zoltowski and Gardner, 2011). These N/Ccap elements in turn regulate activity of effector domains or recruit proteins to Ncap, Ccap or β-sheet surfaces (Figure 1—figure supplement 1A,C,E). Upon incubation in the dark, or in the presence of UV-A light, the signaling adduct state spontaneously decays on a timescale of seconds to days (Zoltowski and Gardner, 2011; Kennis et al., 2004; Pudasaini and Zoltowski, 2013). In this manner, LOV proteins dictate transiently stable signaling states capable of switching between a distinct on and off state depending on lighting conditions (Zoltowski et al., 2007; Halavaty and Moffat, 2007; Harper et al., 2003; Zoltowski et al., 2013). These photoswitchable functions have made LOV proteins targets for optogenetic devices; however, currently we have limited understanding of the role of LOV photocycle kinetics for in vivo function. Genetic and photochemical studies of plant circadian and photoperiodic timing indicate Arabidopsis may function as a model organism for delineating the roles of photochemistry and allostery in LOV function.
In Arabidopsis thaliana, three LOV domain containing proteins ZTL, FLAVIN-BINDING, KELCH REPEAT, F-BOX 1 (FKF1), and LOV KELCH PROTEIN 2 (LKP2) act in circadian timing and seasonal flowering (Baudry et al., 2010). Among them, the genetic functions of ZTL and FKF1 are highly characterized. Recent research indicates that their divergent roles in the circadian clock and photoperiodic flowering may enable interrogation of how chemistry regulates PAS/LOV protein function to select for signaling pathways (Figure 1—figure supplement 1B) (Zoltowski and Imaizumi, 2014). These proteins retain analogous elements, where an N-terminal LOV domain regulates activity of a C-terminal E3 ubiquitin ligase to target clock proteins for degradation in a time dependent manner (Baudry et al., 2010; Más et al., 2003a). In addition, the LOV scaffold engages multiple proteins in a selective manner to regulate clock function (Zoltowski and Imaizumi, 2014). Despite conservation of domain elements, ZTL and FKF1 differ in their subcellular localization, degradation targets, and fundamental chemistry, thereby differentiating ZTL and FKF1 in circadian and photoperiodic timing (Zoltowski and Imaizumi, 2014).
Central to their function are key differences in photocycle kinetics. In FKF1, day-specific expression and a long-lived light-state species (τ = 62 hr) enable light-specific functions (Imaizumi et al., 2003; Pudasaini et al., 2015). In FKF1, photon absorption facilitates complex formation with GIGANTEA (GI) through its LOV domain (LOV-GI) to mediate degradation of CYCLING DOF FACTOR 1 (CDF1) and stabilization of CONSTANS (CO) (Imaizumi et al., 2005; Sawa et al., 2007; Song et al., 2012). Here, selection of degradation or stabilization appears to center on the domain involved in target recruitment, where the Kelch repeat domain (CDF1) specifies degradation and the LOV domain (CO) specifies stabilization (Zoltowski and Imaizumi, 2014; Song et al., 2014). Importantly, both functions require both light and GI.
In contrast, constitutive mRNA levels and a fast photocycle for ZTL (τ = 1.4 hr) suggest possible day (light) and evening (dark) functions (Pudasaini and Zoltowski, 2013). Light-state ZTL has two primary functions, formation of a LOV-GI complex to allow protein accumulation during the day via enhanced ZTL/GI stability (Kim et al., 2013, 2007), and ZTL/GI-dependent destabilization of CO in the morning (Song et al., 2014). The latter results in antagonistic roles of ZTL and FKF1 in the photoperiodic response, and may contribute to a late-flowering phenotype in some ztl mutants (Somers et al., 2004). Throughout the night, ZTL mediates rapid degradation of the clock components, TIMING OF CAB EXPRESSION 1 (TOC1) and PSEUDO RESPONSE REGULATOR 5 (PRR5) (Figure 1—figure supplement 2) (Kiba et al., 2007; Fujiwara et al., 2008; Más et al., 2003b). TOC1 protein levels contribute to the control of period length of the circadian clock (Zoltowski and Imaizumi, 2014). Impaired degradation of TOC1 by ZTL mutants lead to accumulation of TOC1 and PRR5 protein and a long-period phenotype consistent with strains harboring additional copies of TOC1 (De Caluwé et al., 2016). Interestingly, TOC1 degradation appears to be in competition with GI, occurs with an approximately 2 hr delay following dusk, and is enhanced in the dark (Zoltowski and Imaizumi, 2014; Más et al., 2003a; Song et al., 2014; Kim et al., 2007). Current models explain such behavior in TOC1/PRR5 turnover by imparting a differential function between day and night conditions (De Caluwé et al., 2016; Pokhilko et al., 2013), where TOC1/PRR5 are targeted for degradation by ZTL regardless of lighting conditions, but degradation activity is enhanced in the dark. How this is achieved at the molecular and chemical level is not well understood.
Herein, we present a comprehensive study of ZTL signaling to develop a broad understanding of blue-light regulation of circadian and photoperiodic timing and to understand the evolutionary basis for divergent functions of ZTL and FKF1. Therein, we tackle two outstanding questions in the PAS/LOV field; (1) What purpose, if any, does the LOV photocycle lifetime play in biological function? Previous research proposed divergent roles for ZTL and FKF1 in the measurement of light intensity and day length (Pudasaini and Zoltowski, 2013), however no experimental validation of such a model has been available. Further, with the exception of a fungal system (Dasgupta et al., 2015), the role of LOV lifetimes in biology remains elusive. (2) How do PAS/LOV proteins signal to multiple interaction surfaces to allow signal integration? Based on structural studies of ZTL we have developed protein variants that decouple photocycle lifetimes from signal transduction. In this manner, we show a definitive role of LOV lifetime in circadian timing. Further, we provide an allosteric model of LOV signal transduction enabling selection of diverse protein:protein interactions.
Examination of mathematical models of circadian function reveals that ZTL photocycle kinetics may impart phase specific degradation of TOC1 and PRR5 (De Caluwé et al., 2016; Pokhilko et al., 2013). Namely by replacing indistinct day and night conditions by a difference in activity between light and dark-state ZTL, where the light-state inhibits ZTL activity in regards to degradation. In this manner, dark-state reversion activates ZTL during evening. Incorporating ZTL photochemistry into existing models of PRR5 results in Equation 1 (see Mathematical model generation in methods), where cPRR5 is the concentration of PRR5 protein, k1 and k2 the light and dark-state degradation constants and k3 the rate of adduct decay for ZTL. An analogous equation, with equivalent k3 dependence, can be derived for TOC1 (Equation S12), however, the expression pattern of TOC1 and complex regulation of TOC1 mRNA complicates analysis of TOC1 levels during the circadian cycle. For these reasons, we use the simpler PRR5 degradation data to mathematically test our model and use a qualitative analysis for examining effects of k3 on TOC1 (see below).
Examination of Equation 1 provides distinct predictions on the effect of the ZTL adduct decay rate constant (k3) on PRR5 and TOC1 levels. If rate-altering variants only affect photocycle kinetics, k3 would dictate delays in PRR5/TOC1 degradation (Figure 1). Under these conditions a long photocycle would lead to increasing delays in PRR5 and TOC1 degradation leading to progressively longer circadian periods. Unfortunately, testing such predictions is complicated by difficulties in manipulating photochemical kinetics without altering allosteric regulation of ZTL function.
To delineate these aspects we focused on three residues in close proximity to the flavin active site that differ between ZTL and FKF1. These are G46 (Ser in FKF1) that lies adjacent to the active site Q154 implicated in Gln-flip signal transduction mechanisms (Zoltowski and Gardner, 2011), V48 (Ile in FKF1) that sterically interacts with C82 (Zoltowski et al., 2009), and G80 that lies in a GXNCRFLQ motif (X80=G for ZTL, X=R for FKF1) (Figure 1—figure supplement 1E). The V48 position is known to alter LOV photocycle kinetics by V/I/T substitutions (Zoltowski et al., 2009; El-Arab et al., 2015; Christie et al., 2007; Lokhandwala et al., 2016). This site has been exploited in optogenetic tools to tune signal duration under the presumption that it does not affect allosteric regulation of protein function (Strickland et al., 2012). In contrast, to the best of our knowledge, the G80 position has not been exploited for tuning LOV kinetics. This likely is due to Arabidopsis ZTL being unusual amongst LOV proteins in containing a Gly residue at this position. As a consequence of G80, ZTL expresses poorly in E. coli and is largely confined to inclusion bodies (see Materials and methods). Since, G80 is unique to Arabidopsis ZTL, and R/Q/K substitutions are permitted in other ZEITLUPE and LOV proteins (see Discussion), it is unlikely to affect allostery and may function as a site to uniquely affect photocycle kinetics.
Consistent with our predictions, kinetics of V48I and G80R differentiate ZTL and FKF1 type chemistry. Specifically, variants extend the ZTL lifetime from 1.4 hr (WT) to 6.6 (G80R) or 10.7 (V48I) hours (Figure 1—figure supplement 3 and Table 1). Double variants G46S:G80R and V48I:G80R function cooperatively to extend the ZTL photocycle to 21 hr or >65 hr respectively, very similar to WT FKF1 (62.5 hr) (Pudasaini and Zoltowski, 2013) (Figure 1—figure supplement 3 and Table 1). In addition, the difference in values for k3 in these variants are sufficient to elicit theoretical differences in PRR5/TOC1 degradation rates (Figure 1B). To verify that rate-altering variants do not also perturb allosteric regulation of ZTL function it is essential to obtain crystal structures of ZTL variants and to map an allosteric trajectory that couples C4a adduct formation to regulation of ZTL function. Towards these aims we solved 2.5 Å, 2.6 Å, and 2.1 Å crystal structures of dark-state WT ZTL, G80R and V48I:G80R (residues 29–165) respectively (see Table 2). In addition, the long-lived V48I:G80R variant enabled direct crystallization (2.3 Å) of the light-state adduct allowing for interrogation of signal transduction pathways.
All ZTL variants form solution dimers, crystallize in the same space group (P3121) and demonstrate topologically equivalent structures consistent with PAS/LOV proteins (Figure 2, Figure 2—figure supplements 1 and 2). These structures reveal functional differences from all known LOV structures that differentiate the effects of V48I and G80R on ZTL signaling mechanisms. Namely, G80R structures are analogous to WT in all manners. The primary difference is the formation of a π-cation interaction that stabilizes the Dα/Eα linkage and C82 that is necessary for C4a adduct formation (Figure 2B). Increased rigidity of C82 imposed by the π-cation interaction is consistent with having an effect only on photocycle kinetics. In contrast, examination of V48I containing structures reveals distinct differences that identify allosteric signal transduction mechanisms to the N/Ccap. These mechanisms identify V48I as a residue that disrupts allosteric regulation of ZTL function. Below, we provide detailed analysis of ZTL in dark and light states to highlight these functional differences. We focus on two reported aspects of ZTL group function, LOV:LOV mediated homodimerization and allosteric regulation of Ncap and Ccap elements.
Previous solution studies of ZTL group proteins suggest they function as obligate dimers (Ito et al., 2012; Nakasako et al., 2005), however ZTL group dimers have not been observed in vivo and their function is unknown (Han et al., 2004). Consistent with solution studies, the crystallographic lattice is defined by two sets of anti-parallel dimers formed by extensive contacts along the β-scaffold. We term these interfaces the ‘compact’ and ‘elongated’ dimers based on a 2 Å translation that is coupled to the degree of order present within the Ncap (Figure 2). In the compact dimer, clear density for Ncap residues (31-43) is observed in one molecule that contacts the helical interface of the adjacent monomer (Figure 2C,D). In contrast, the elongated dimer contains no density for Ncap residues (Figure 2E,F). A stabilizing element in both dimers is a hydrophobic core composed of a tetrad of Ile residues (l151 and I153).
Additional contacts along the helical interface define a possible secondary dimer. The helical dimer is parallel in orientation and involves contacts between the E-F helices and associated loop. Specifically, R95 forms a salt bridge with the phosphate moiety of FMN in the neighboring molecule (Figure 2—figure supplement 2). However, based on two lines of evidence, we conclude that the β-scaffold interface represents the solution ZTL and FKF1 homodimers. First, FKF1 lacking the entire E-F loop and ZTL variants that disrupt R95 contacts remain dimeric (Figure 2—figure supplement 2) (Nakasako et al., 2005). Second, I151R (and I160R in FKF1) variants abolish dimer formation in vitro (Figure 2A and Figure 2—figure supplement 1). Thus, ZTL and FKF1 solution dimers are formed by equivalent anti-parallel contacts along the β-scaffold, similar to other PAS/LOV proteins (Card et al., 2005).
Given the unknown role of ZTL dimers in vivo and the known role of Ncap and Ccap elements in LOV allostery, we turned our attention to the structural differences between the different dimers. Close examination of residues linking the Ncap and Ccap to the active site FMN identifies distinguishing interactions that may be involved in signal transduction. The loss of Ncap density in the elongated dimer directly follows a CGF motif (C45-G46-F47-(V48)) that links the Ncap, FMN binding pocket and the V48 position (Figure 3). An analogous hinge involving a Cys residue is known to mediate signal transduction in fungal LOV proteins, where the hinge directs both a conformational change and integration of oxidative and osmotic stress (Lokhandwala et al., 2015; Zoltowski et al., 2007, 2009; Lokhandwala et al., 2016; Lamb et al., 2009; Zoltowski and Crane, 2008). Further, the CGF motif differentiates ZTL and FKF1, where FKF1 contains the G46S and V48I (equivalent) mutations, thereby highlighting the region as a possible factor regulating the divergent functions of these closely related proteins.
In ZTL, signal transduction diverges from known LOV signaling mechanisms (Figures 3 and 4). In contrast to other LOV proteins, where a conserved Gln (Q154) acts as a molecular bridge linking the FMN to a residue in Aβ, the dark-state structure of ZTL contains a heterogeneous orientation of the active site Q154 (Figure 3 and Figure 3—figure supplement 1) (Zoltowski et al., 2007; Halavaty and Moffat, 2007; Möglich and Moffat, 2007). The altered orientation is coupled to contacts in the Ncap, Ccap and helical interface. Given that all three sites have been implicated in PAS/LOV signaling (Harper et al., 2004; Card et al., 2005; Zoltowski and Crane, 2008; Partch et al., 2009), we examine each interaction below. We define them as the Aβ-Ncap hinge, Iβ-Ccap hinge and Cα (helical interface) to specify structural elements that may be involved in signal transduction.
In ZTL, Q154 does not adopt a specific orientation as observed in all other LOV structures, rather it varies in all four molecules in the asymmetric unit of both WT-ZTL and G80R (Figure 3A and Figure 3—figure supplement 1). The altered conformations contact the Aβ-Ncap hinge through G46 in the CGF motif (Ncap; Figure 3B). Here, Q154 abuts (3.4 Å) the Cα carbon of G46 (Ser in FKF1). The lack of a side chain at G46 permits favorable interactions for Q154 at the flavin O4 position and allows insertion of F156 into the flavin active site. In this manner an unusual orientation of Q154 links the Iβ-Ccap hinge and helical interface through a QFF motif (Q154-F155-F156). In most LOV proteins, the equivalent residue to F156 is hydrophilic and adopts a solvent exposed position (Figure 3). The resulting cavity occupied by F156 is filled by a Met or Leu residue (V69 in ZTL) from Cα (Figure 3B,D). In ZTL, the altered orientation of Q154 clashes with typical orientations of Cα. Steric clashes lead to movement of conserved F66 and rotation of V69 away from Q154 and the FMN binding pocket.
The combined interactions stemming from Q154 positions it ideally to have concerted conformational changes within Ncap and Ccap elements. Although, the Gln locus has been cited as the primary source of signaling in LOV proteins (Zoltowski et al., 2007; Halavaty and Moffat, 2007; Freddolino et al., 2013), the lack of interactions near N5 and Aβ is atypical. Thus, it is likely that ZTL does not involve an H-bonding switch in signal transduction and that canonical mechanisms of LOV signaling may not be universally conserved. Rather, we propose that steric interactions involving the flavin O4 position and G46 shift a dynamic equilibrium in the Q154 position to stabilize the ZTL protein in a light-state configuration. We posit that the heterogeneous orientation of Q154 destabilizes ZTL in the absence of covalently attached FMN, consistent with in vitro studies demonstrating a tendency to lose flavin rapidly. Upon light activation, ZTL is stabilized by covalent attachment of FMN (through C82) and results in movement of Q154 that necessitates rearrangement of the CGF, QFF and Cα sites to elicit signal transduction.
Dark and light state structures of V48I:G80R confirm concerted movement of Q154 that is linked to conformational changes in V/I48. The Q154 conformation is not heterogeneous in V48I:G80R, rather, the Ile side chain sterically directs Q154 to an exposed conformation near O4 that has not been observed in other LOV proteins. Comparing the dark-state to V48I:G80R grown in the light, confirms density for the C4A adduct and confirms direct crystallization of the light-state (Figure 4). Difference density maps of the light-state molecule indicate that adduct formation is coupled to rotation of Q154 to a buried position (Figure 4C). Unexpectedly, rotation of Q154 requires concerted movement of I48 that reorients the Ncap. Thus, the presence of G46 selects for a heterogeneous conformation of Q154 and V48I biases the heterogeneous Q154 towards the exposed conformation in the dark. In the light, V48I disrupts rotation of Q154 to the buried conformation, thereby leading to a predominantly exposed conformation regardless of lighting conditions. Thus, unexpectedly, V48I is both an allosteric variant and alters photocycle kinetics. In particular, V48I retains an ability to form the covalent C4a adduct, which is more stable than WT (V48I k3 = 0.094 hr−1; WT k3 = 0.71 hr−1). However, in V48I allosteric regulation of the Ncap through Q154 is disrupted leading to V48I selecting for a distinct exposed Q154 conformation despite adduct formation (Figure 4).
Based on our structural results we can refine our models on how ZTL rate altering variants will perturb ZTL function. G80R does not impact ZTL structure or allosteric regulation of Ncap or Ccap elements. In this manner, G80R acts as a photocycle variant only and allows direct testing of LOV photocycle kinetics on targeted degradation of PRR5 and TOC1 (see Equation 1). In contrast, despite being photochemically active and stabilizing the C4a adduct, V48I blocks allosteric regulation of the Ncap through incomplete rotation of Q154 and selection of a distinct exposed conformation of Q154. Thus, we propose that V48I is an allosteric variant that mimics the degradation-active dark state. Combining our proposed mechanism with Equation 1 above and existing literature on light-dark formation of ZTL:GI complexes we make the following testable hypotheses: (1) V48I should demonstrate weaker interactions with GI. (2) V48I should show constitutive activity regardless of lighting conditions in targeting PRR5 and TOC1 for degradation. Thus, PRR5 and TOC1 levels should be constitutively low in variants containing the V48I mutation.
We examined ZTL variants for their ability to form light and dark regulated complexes with GI and ARABIDOPSIS SKP1-LIKE 2 (ASK2) of the SCF complex. CoIP results confirm that G80R retains light regulated complex formation with GI comparable to WT. In contrast, Ncap variants G46S and V48I both alter light driven complex formation with GI (Figure 4—figure supplement 1). Whereas, V48I leads to dampened light-driven formation of the ZTL:GI complex, G46S enhances GI complex formation in both the dark- and light-states (Figure 4—figure supplement 1A). These results support our allosteric model of ZTL regulation and demonstrate that Ncap variants decouple allosteric regulation of signal transduction from photochemical formation of the C4a adduct. Further, the data implicates light-driven conformational changes near the Ncap in dictating GI interactions. Specifically, where V48I mimics the weak GI interacting dark-state and G46S mimics the strong GI interacting light state. We note that the G46S results reported here diverge from the effects of the G46E mutation reported by Kim et al. (2007), where G46E abolishes GI interactions due to apparent misfolding of the LOV domain. These results are informative on the nature of mutations at the G46 site, namely the long side chain present in a G46E variant leads to steric clashes and likely would force E46 into a hydrophobic pocket (see Figure 4—figure supplement 2). We contend such clashes leads to the destabilization of the LOV domain and abolition of GI interactions as reported previously for G46E (Kim et al., 2007). In contrast, the shorter sidechain in G46S, can be easily accommodated by a subtle rotation of the active site Q154 towards the proposed light-state buried conformation (modeled in Figure 4—figure supplement 2). Such results are consistent with enhanced GI interactions in G46S. Combined, the results implicate the G46/V48 locus in ZTL allostery and regulation.
In contrast to GI, where G46/V48 mutations affect function in a light/dark manner, all ZTL variants complex with ASK2 in a light-independent manner comparable to WT (Figure 4—figure supplement 1B). The protein:protein interaction data confirms G80R behaves as WT in known biochemical functions of ZTL and only differs in photocycle kinetics. In contrast, V48I acts as an allosteric variant mimicking the dark-state conformation. Based on these results and previously published in vivo studies showing enhanced degradation activity in the dark (Más et al., 2003a), we have tools to test the effect of photocycle kinetics (G80R) and Ncap allostery (G46S/V48I). In this manner, G80R variants isolate effects of photocycle kinetics allowing testing of predictions based on equation 1.
The decrease in GI affinity in the V48I variant could complicate in vivo phenotypes for these variants. Literature indicates ZTL stability is dictated by GI interactions (Kim et al., 2007). Thus, we could expect low ZTL levels in V48I variants. These low ZTL levels would act in opposition to any increased targeted degradation of PRR5/TOC1 and could mask allosteric phenotypes. Based on our experimental conditions, we do not expect any complications to result. Prior studies indicate that ZTL and GI reciprocally stabilize each other (Kim et al., 2013, 2007), but increased ZTL stability occurs in a circadian phase dependent manner (Kim et al., 2003). Light does enhance ZTL:GI affinity and reciprocal stability by three fold, but ZTL retains some stabilization during the subjective circadian day regardless of lighting conditions (Kim et al., 2007, 2003). These previous findings suggest that in consideration of the ZTL:GI equilibrium, GI is limiting except during the subjective circadian day. During the day, GI levels rise sufficiently to shift the equilibrium to saturate the ZTL:GI complex regardless of lighting conditions. Given that our V48I variant retains light-state affinity comparable to WT dark-state protein, GI expression during the subjective day under LL conditions should rescue the decrease in affinity. Concomitant with ZTL-ox, reciprocal stabilization of ZTL/GI should lead to high ZTL levels and ZTL should no longer oscillate. Indeed, under our experimental conditions cycling of ZTL protein is lost and V48I variants show enhanced stability in vivo (Figure 5—figure supplement 1), thereby the role of GI in ZTL stability is masked under our conditions and our results likely reflect the effect of ZTL photochemistry (G80R) and allosteric activation of PRR5/TOC1 degradation (V48I) independent of the effects on GI binding.
Based on the data above we make the following predictions. In comparing G80R-ox to WT-ox under LD conditions, the fast decay of WT-ox will lead to higher populations of the active-dark state early in the evening. In contrast, slower adduct decay in G80R will lead to prolonged population of the less active light-state and delays in ZTL degradation activity. As a result, we should observe an enhanced delay in PRR5/TOC1 degradation in G80R variants as shown in Figure 1 and Figure 5—figure supplement 2. For V48I and V48I:G80R, the allosteric effects should enhance degradation of PRR5 and TOC1 leading to constitutively low PRR5/TOC1 levels.
To test our predictions, we constructed Arabidopsis transgenic ZTL overexpression lines (ZTL-ox) containing WT, V48I, G80R and V48I:G80R. G46S variants were excluded due to poor yields from E. coli that render solution or structural information intractable for the isolated G46S variant (see Materials and methods). These constructs were then examined for their effect on PRR5 and TOC1 degradation as well as circadian period and amplitude. All transgenic lines were selected based on similar ZTL transcript levels and ZTL protein levels were measured (Figure 5—figure supplement 1).
Consistent with our predictions, G80R-ox (#22) leads to delayed degradation of PRR5 and TOC1 under LL and LD (Figure 5). Specifically, despite being overexpressed (Figure 5—figure supplement 1A), G80R-ox variants demonstrate peak amplitudes of TOC1 consistent with WT (Figure 5A,B). Further, comparison of apparent rate constants for PRR5 degradation confirms a direct effect of ZTL photocycle kinetics on PRR5 degradation, where G80R #22 (0.34 hr−1 LD, 0.14 hr−1 LL) exhibits smaller rate constants compared to WT-ox (0.5 hr−1 LD, 0.13 hr−1 LL) under LD conditions where k3 is most relevant (Table 3). These results are consistent with a more active dark-state ZTL. Thus, ZTL photocycle kinetics regulate degradation of TOC1/PRR5.
Examining V48I and V48I:G80R variants confirms a role of V48I in altering light-dark regulation of ZTL activity. Strains harboring V48I and V48I:G80R show constitutively low levels of PRR5 and TOC1, consistent with high degradation activity regardless of lighting conditions (Figure 5). Both variants show degradation rate constants for PRR5 of ~0.8–1 hr−1 under LD conditions, similar to maximum rate constants (0.8 hr−1) predicted from computation models of clock function (De Caluwé et al., 2016; Pokhilko et al., 2013) (see Materials and methods and Table 3). V48I and V48I:G80R also exhibit high activity under LL conditions, demonstrating a partial loss of light-dark regulation (Figure 5 and Table 3). Combined, G80R confirms a direct role of LOV photocycle kinetics on ZTL activity and that V48I acts in an allosteric switch to enable light-state V48I (adduct formation) to mimic the more degradation active and less GI-binding competent dark-state.
Given the effect of V48I, G80R and V48I:G80R on PRR5 and TOC1 degradation, these variants should have predictable effects on circadian period. Previous studies of ZTL-ox variants demonstrate a dose dependent effect of ZTL on circadian period, where high protein levels lead to short-period phenotypes progressing to arrhythmicity and ZTL-nulls having a long-period phenotype (Somers et al., 2004). Similarly, TOC1 overexpression strains have a long-period phenotype and TOC1-null strains have short periods (Más et al., 2003b; Gendron et al., 2012). Thus, we predict that despite overexpression the defect in degradation of TOC1 by G80R-ox should lead to WT periods. In contrast, low TOC1 levels in V48I and V48I:G80R should lead to arrhythmic phenotypes under expression levels comparable to WT-ox strains demonstrating normal periods.
Indeed examination of circadian periods in WT and mutant-ox strains confirms predicted effects of photocycle kinetics (G80R) and the signaling defect (V48I) on circadian period. All strains containing V48I lead to an arrhythmic phenotype consistent with heightened degradation activity in these variants (Figure 6 and Table 3). In contrast, G80R-ox strains harboring 3-times more ZTL protein than WT has a circadian period indistinguishable from WT and WT-ox (see Table 3; 35S:HA-ZTL (G80R) #22; 24 hr, compared to WT; 24 hr and 35S:HA-ZTL #17; 24 hr). Only when protein levels exceed 10-fold higher than WT-ox is the period shortened to 19 hr (35S:HA-ZTL (G80R) #23) (Figure 6 and Table 3). These results confirm that ZTL photocycle kinetics are coupled to selection of circadian period through PRR5/TOC1 degradation.
In Arabidopsis thaliana, ZTL and FKF1 diverge in their diurnal pattern of transcription and function. Whereas, FKF1 function and transcription is specific to the day, ZTL demonstrates constitutive transcription and distinct functions during the day and night (Song et al., 2014). These distinct functions allow ZTL to impact photoperiodic flowering through morning specific and GI-dependent destabilization of CO, and circadian function through night specific degradation of TOC1 and PRR5 (Más et al., 2003a; Song et al., 2014; Kiba et al., 2007). These results suggest that both light (CO destabilization/GI interaction) and dark (TOC1/PRR5 degradation) activate ZTL for function. Such observations seem paradoxical; however, examination of our structural and kinetic results indicates ZTL may use a divergent allosteric mechanism to enable dual light/dark functions.
The ability of LOV domains to function as photoswitchable proteins is predicated on LOV proteins specifying a distinct light- and dark-state configuration. In all dark and light-state LOV structures currently in the protein data bank, the active site Gln (Q154 in ZTL) adopts a distinct buried conformation in the dark, with the NH moiety of the Gln side chain near flavin-N5 (Halavaty and Moffat, 2007; Zoltowski et al., 2007). Existing light-state structures indicate that C4a adduct formation and N5 protonation induces rotation of the Gln side chain to favor an H-bond between flavin-HN5 and the O moiety of the Gln side chain (Vaidya et al., 2011). In those structures, the Gln residue maintains the buried conformation and is only distinguished by the nature of H-bonds. In our ZTL structures this is not the case, rather dark-state structures have a heterogeneous orientation of Q154 that samples orientation near both the buried conformation and an exposed conformation (Figure 3—figure supplement 1). A heterogeneous conformation of the key signaling switch would suggest poor regulation of function under dark-state conditions. This is exactly what is observed in ZTL. ZTL retains fairly robust activity for GI interactions and TOC1/PRR5 degradation in both the light and dark, with light enhancing GI interactions and repressing TOC1/PRR5 degradation by 3–5 fold (Kim et al., 2007) and (Table 3). As such, GI-dependent function in CO destabilization is likely not a light-regulated event, but rather is a combined result of constitutive transcription of ZTL, day-specific expression of GI and poor signal amplification following light-dark interconversion due to the heterogeneous orientation of Q154. Thus, the unusual Gln orientations appear to be evolutionarily selected to permit light- and dark-state functions of ZTL and to differentiate ZTL and FKF1.
Combining our dark- and light-state structures with in vivo data allows construction of a putative signaling mechanism differentiating ZTL from other LOV proteins. Examination of the ZTL structures identifies two key residues involved in regulating ZTL allostery. The heterogeneous conformation of Q154 is permitted by the lack of a sidechain in G46 (model in Figure 4—figure supplement 2). Introduction of V48I, directs Q154 to a single exposed conformation in the dark. Biasing the Q154 to the exposed conformation results in disrupted interactions with GI and high TOC1/PRR5 degradation, in vivo, consistent with selecting for a distinct dark-state conformation. Further, the light-state V48I:G80R molecule indicates partial Q154 rotation to a buried conformation that is impeded by V48I. This impediment does not allow robust sampling of the buried conformation under any lighting conditions and coincides with constitutively high TOC1/PRR5 degradation in V48I containing variants. Combined we propose a putative model of ZTL signaling (Figure 4). ZTL retains functionality in the light and dark due to a heterogeneous orientation of Q154 permitted by G46 and V48. Differences in functionality between dark/light result from subtle biases in the orientation of Q154 between buried and exposed conformations. Based on V48I:G80R structures and activity, we propose that biasing towards the exposed conformation as the dark-state configuration (enhanced PRR5/TOC1 degradation) and biasing towards the buried conformation as the light-state configuration (enhanced GI binding).
Based on our proposed mechanism distinguishing ZTL signal transduction from other LOV proteins, one would predict evolutionary selection of G46 and V48 in ZTL proteins to permit the heterogeneous orientation of Q154. Phylogeny of LOV sequences in planta demonstrates evolutionary selection of residues G46, V48 and F156 to differentiate ZTL-like, FKF1-like and phototropin-like (LOV2) proteins and putative signal transduction pathways (Figure 7 and Figure 7—figure supplement 1). Specifically, in monocots and dicots ZTL-like proteins conserve the G46 that is necessary for selection of the exposed conformation of Q154. In FKF1 (monocots: A46 and dicots: S46), phototropins (N46) and all other structurally characterized LOV proteins, an H-bond or sterically directing residue occupies this position.
Residue selection at position 46 is coupled to V48. V48 is conserved in ZTL, but substitutions are permitted in FKF1, phototropins and fungal LOVs (Figure 7—figure supplement 1) (Lokhandwala et al., 2015; Zoltowski et al., 2007). The altered signaling mechanism in ZTL is supported by examination of LOV proteins in Brassica rapa. ZTL-like proteins in Brassica and AtLKP2 contain a Q154L substitution, resulting in an LXF consensus sequence at that locus. The presence of a Q154L substitution is unexpected, since these substitutions abrogate blue light signaling in other LOV proteins due to an inability to undergo the Gln-flip mechanism (Zoltowski et al., 2007; Nash et al., 2008; Nozaki et al., 2004). In the proposed ZTL mechanism a Gln-flip is not necessitated and Q→L substitutions are permitted. In this manner, our understanding of a canonical model of LOV signal transduction is incomplete. We propose that evolutionary selections at G46 and V48 tune allostery to the Ncap through the Q154 orientation to differentiate modes of LOV signal transduction. Given the evolutionary selection of A/S residues and permission of V48 substitutions in FKF1, we propose that FKF1 functions in a manner analogous to other LOV proteins, where light activates the biological function (CO stabilization/CDF degradation) of the primarily nuclear FKF1 protein. In this manner, evolutionary selection at position 46 may dictate day functional (FKF1) and day/night functional (ZTL) differentiation, thereby implicating LOV photocycle kinetics as being imperative for proper signal transduction.
Currently, delays in TOC1 and PRR5 degradation are explained using competition between GI and TOC1/PRR5 for the available ZTL pool (Más et al., 2003a; Kim et al., 2007; Kiba et al., 2007; Fujiwara et al., 2008). In these models, GI expression during the day both stabilizes and sequesters ZTL. In the evening, GI pools decline leading to active free ZTL. In light of our data, the competition model must be incomplete. In the competition model there should be no differences in the LL/LD characteristics in WT and G80R strains. In the competition model GI expression profiles should dictate delays and these would be unaffected by rate-altering ZTL variants. WT and G80R interact with GI to comparable levels, yet G80R demonstrates enhanced delays in PRR5 and TOC1 degradation (Figure 5D). These results are inconsistent with a competition model alone. In contrast, modeling PRR5 degradation using the time dependent conversion of ZTL-light to ZTL-dark can predict with reasonable certainty the extended delay in G80R and predicts distinct differences under LL conditions between the two proteins (Figure 5—figure supplement 2). In this manner, it is clear that ZTL photocycle kinetics are instrumental in dictating delays in PRR5/TOC1 turnover and circadian period.
Based on all these elements we propose that circadian period is regulated in a complex manner involving a ZTL:TOC1:GI circuit, whereby competitive inhibition and ZTL photocycle kinetics act in concert to dictate ZTL protein levels and ZTL activity in a circadian phase dependent manner. Adduct decay in ZTL then enhances degradation of PRR5 and TOC1, impacting circadian period through two methods: (1) Degradation of PRR5, helps diminish TOC1 levels through increased cytosolic retention and accessibility to ZTL (Wang et al., 2010) and (2) Degradation of cytosolic TOC1. Both factors require proper ZTL photocycle kinetics. Thus, LOV photocycle kinetics are instrumental in evolutionary selection for a 24 hr period.
Examination of recent mathematical models of plant circadian clocks reveals a common method of incorporating light-dark dependent degradation of PRR5 and TOC1 by ZTL. In Pokhilko et al., PRR5 and TOC1 levels are treated as follows (Pokhilko et al., 2013):
Where Cx represent protein concentrations, Cxm, mRNA levels, px, nx and mx are parameters fit to data sets. ZG represents the ZTL:GI complex. D represents darkness, where D = 1 at night and D = 0 during the day. Both equations are constructed of similar elements, a protein synthesis term defined by mRNA levels and a degradation term defined by PRR5/TOC1 protein levels, and in the case of TOC1 the total ZTL protein pool. We note, that ZTL protein levels are not incorporated into existing models of PRR5 degradation (De Caluwé et al., 2016; Pokhilko et al., 2013) suggesting that overexpression has a weak enough effect on the overall rate of PRR5 degradation that models lacking ZTL concentration can accurately predict degradation kinetics. Although biologically such an analysis is incomplete, for modeling purposes the accuracy of these prior models suggests this assumption is reasonably valid. Therefore, they normalize the ZTL concentration to one and it does not appear in the PRR5 degradation term. Further, in the fit parameters, p5 = 1, which allows simplification as shown in Equation S4, where cZTL now equals the total pool of ZTL protein regardless of whether free or in complex with GI.
For our purposes, ZTL rate altering variants should not perturb the transcription rates, therefore differences in PRR5/TOC1 degradation should be limited to the degradation terms. For PRR5 and TOC1 the degradation terms are as follows
The above equations do not incorporate ZTL photochemistry, rather, the term m*D to enhance the degradation rate constant under dark state conditions. In the dark, D = 1 and the degradation rate constant (k2) becomes (m17 + m24) or (m6 + m7) for PRR5 and TOC1 respectively. In the light, D = 0 and the rate constants (k1) reduce to m17 and m6. To add ZTL photochemistry to these equations we rewrite these degradation equations in terms of light and dark-state ZTL. We also note additional complications in the TOC1 equation. The term m8*cTOC1 is a non-ZTL dependent degradation term, presumably accounting for nuclear degradation of TOC1. Because of the additional complexities in TOC1 expression and non-ZTL dependent degradation we do not model TOC1 degradation in vivo, we do however show below that the rate constant for LOV adduct decay, k3, will impact TOC1 in a manner analogous to PRR5.
Where k1, k2 are the rate constant for degradation in the light and dark respectively. Similarly, cZTL-L and cZTL-D represent the light and dark-state levels of ZTL. We also normalize the total ZTL concentration to 1. Doing so allows calculation of cZTL-L and cZTL-D as a function of time during the dark-phase of LD cycles, by incorporating the rate constant for adduct decay, k3.
It follows that:
We note, that a compact model of clock function by De Caluwe et al. demonstrates that existing data can be adequately fit for both PRR5 and TOC1 levels using an analogous equation for both PRR5 and TOC1 (only the degradation term has been extracted) (De Caluwé et al., 2016).
Where, kD, kL are the light and dark degradation rate constants respectively, and L and D reference the lighting conditions, where L = 1, D = 0 during the day and L = 0, D = 1 during the evening. CPT represents either the PRR5 or TOC1 level. Applying our method above to the De Caluwe et al. model (Equation S13), results in an analogous expression as Equation S10.
Thus, in terms of ZTL chemistry, PRR5 and TOC1 degradation should be dictated by k3. Using mutants that affect only k3, see Table 1, one should be able to accurately model PRR5 decay rates in vivo. Again we note that the expression pattern of TOC1 and complex regulation of TOC1 mRNA complicates TOC1 levels during the circadian cycle. For these reasons, we use the simpler PRR5 degradation data to mathematically test our model above and use qualitative differences in degradation patterns to examine effects of k3 on TOC1.
To predict how k3 may affect delays in PRR5 degradation we required estimates of k1 and k2. Examining Pokhilko et al. and De Caluwe et al. provides reasonable bounds for parameter estimation. First, PRR5 models suggest that PRR5 can be accurately modeled excluding differences in ZTL expression levels (De Caluwé et al., 2016; Pokhilko et al., 2013), this is consistent with our data showing that WT, WT-ox and G80R-ox all have the same decay rate under LL conditions despite differences in protein levels. Second, in De Caluwe et al. the maximum degradation rate constant for PRR5 degradation was fit as 0.78 hr−1. Similarly, in Pokhilko et al. a max value of 0.5 hr−1 was obtained for PRR5 and a kmax of 0.8 hr−1 for TOC1 by combining all degradation terms. Given the similarity of these maximum values we chose a kmax for the dark-state of 0.8 hr−1. The lowest fit rate constant in mathematical models is m6 (0.2 hr−1) for TOC1 degradation under light-state conditions. Thus, we used 0.2 hr−1 as an estimate of PRR5/TOC1 degradation in the light.
k1 = 0.2 hr−1
k2 = 0.8 hr−1
k3 = 0.7 hr−1 (WT), 0.15 hr−1 (G80R), 0.09 hr−1 (V48I), 0.05 hr−1 (G46S:G80R), 0.02 hr−1 (V48I:G80R)
Values for k3 are derived from experimental values for the adduct decay time constant (τ) present in Table 1, where k3 = 1/τ.
Our structural data indicates that V48I mimics the dark-state regardless of lighting conditions, thus, V48I degradation kinetics under LD serves as a reasonable estimate for k2, the dark-state rate constant. We note, that our experimental value for V48I (0.8 hr−1) is identical, to the kmax values present in both Pokhilko et al. and De Caluwe et al, which should reference the more active dark-state value.
Under LL conditions we observe a rate constant for PRR5 degradation of 0.13–0.14 hr−1 for WT, WT-ox and G80R. Under our lighting conditions (broad spectrum; 40–50 µmol m−2s−1), the light-state should be near saturated, and thus in the absence of allosteric effects should report the light-state degradation rate constant. We use 0.14 hr−1 then as an estimate for k1 the light-state rate constant.
We note the similarity of values between the three strains despite different expression levels. These results suggest that the models by Pokhilko et al. and De Caluwe et al. demonstrating a weak dependence of PRR5 degradation on ZTL expression levels is reasonably valid under our conditions. Further, it provides a reasonable estimate for k1 the light-state degradation term.
Thus, to test the accuracy of our model we numerically solved Equation S9 in matlab using:
k1 = 0.14 hr−1
k2 = 0.8 hr−1
k3 = 0.7 hr−1 (WT), 0.15 hr−1 (G80R)
Initial PRR5 levels were taken from Figure 5D at t = 12 hr (initial dark).
Values for k3 are derived from experimental values for the adduct decay time constant (τ) present in Table 1, where k3 = 1/τ.
Results were plotted in Figure 5—figure supplement 2.
ZEITLUPE construct ZTL-S composed of residues 29–165 were cloned into both p-His and pGST parallel vectors using NcoI and XhoI restriction sites. Proteins were purified as reported previously (Pudasaini and Zoltowski, 2013). These DNA sequences were verified by GENEWIZ sequencing service. The plasmids were then isolated and tested for protein expression. All constructs were transformed into E. coli (JM109 or JM109DE3) cells and grown overnight at 37°C as starter culture till O.D600 ~0.6–0.7. The rich culture was then transferred into 1.0 L of LB media for large-scale expression. These cultures were grown for 2–3 hr at 37°C till O.D600 ~0.5–0.6 then the temperature was lowered to 18°C. The culture was then induced with 200 µM Isopropyl-β-D-thio-galactopyranoside (IPTG) after 1 hr incubation at 18°C. After induction, the cells were grown at 18°C for about 18–20 hr. Finally, the cells were centrifuged at 4000 rpm to collect and store the cells in stabilizing buffer (50 mM Tris pH 7or 50 mM Hepes pH 8 with 100 mM and 10% glycerol). The harvested cell-pellets were stored for later use at −80°C.
We note that ZTL is difficult to express and purify from E. coli as most WT proteins are confined to inclusion bodies. Typical experiments required cell pellets from 18 L of cells. Such difficulties make studies of variants such as G46S difficult in the absence of G80R. The G80R variant, likely due to the stabilizing effects of the R80-F84 π-cation interaction (Figure 2B), enhances protein yields in E. coli substantially. This allows access to G46S:G80R, despite intractable yields of the isolated G46S variant.
ZTL-S was purified using affinity chromatography followed by size exclusion chromatography. Prior to purification the cells were lysed by sonication at 4°C. After sonication the protein solution was centrifuged at 18 k rpm at 4°C for 60 min. The supernatant was then purified using Ni-NTA or GST affinity columns. 6xHis and GST tags were cleaved using 6His-TEV-protease followed by an additional round of Ni-NTA affinity chromatography to remove the 6His tag and 6His-TEV-protease. The final eluted protein was subjected to Fast Protein Liquid Chromatography (FPLC) using a Hiload Superdex 200 16/60 gel filtration column equilibrated with stabilizing buffer.
Solution characterization of purified proteins was done using a Superdex 200 10/300 analytical column (GE Lifesciences). Protein concentrations were determined using the absorbance at 450 nm (ext. coefficient 12,500). Apparent molecular weights were calculated by comparing the elution volume of known standards (sweet potato β-amylase (200 kDa)−12.4 ml; yeast alcohol dehydrogenase (150 kDa)−13.31 ml; bovine albumin (66 kDa)−14.61 ml; carbonic anhydrase (29 kDa)−17.03 ml; horse heart cyctochrome c (12.4 kDa)−18.32 ml)) (Sigma Aldrich). Absolute molecular weights of ZTL 16–165, ZTL 29–165 and FKF1 28–174 were determined by subjecting samples to refractive index and light-scattering detectors on a Wyatt Minidawn light-scattering instrument following a Superdex 200 10/300 analytical column. MW’s were determined using ASTRA software from Wyatt Technologies (Santa Barbara, California). All SEC and multi-angle light scattering experiments were conducted in stabilizing buffer (see above).
Site specific protein variants of ZTL-S constructs were obtained using the quickchange protocol. Following PCR amplification samples were treated with 1 µL of DpnI and incubated at 37°C for 2.5–3 hr to cleave the methylated template DNA. A single colony of DH5α E. coli was grown at 37°C overnight and plasmid DNA was isolated and verified by DNA sequencing (Genewiz). Rate altering variants of ZTL were expressed, purified and characterized using the method described above.
ZTL-S and its variants were initially screened with Hampton Screens (HR2-110 and HR2-112) via the hanging drop methods using 1.5 µL well solution with 1.5 µL of ZTL-S at various concentration range of 5–10 mg/ml. Optimum crystallization conditions for WT (0.1 M Tris pH 8.5, 0.1 M Magnesium Chloride hexahydrate, 28% w/v PEG 4k), G80R (0.1 M Tris pH 8.5, 0.2 M Magnesium Chloride hexahydrate, 30% w/v PEG 4k), V48I:G80R (0.1 M Tris ph 8.5, 0.2 M Sodium Acetate trihydrate, 30% w/v PEG 4K). Protein for crystallographic studies was purified in the same stabilizing buffer.
Light state crystals of V48I:G80R were obtained as follows. Prior to setting screens V48I:G80R protein was exposed to a broad spectrum white flood light (150 W), while on ice for two minutes. Saturation of the light-state was confirmed by UV-vis spectra analysis, by verifying disappearance of the 450 and 478 nm absorption bands characteristic of the dark-state protein. The light-state protein was then crystallized directly using the hanging drop method outlined above. Crystals appeared within 24 hr, and trays were exposed to white light once a day to maintain population of the light-state species.
Diffraction data was collected at the F1 beamline at the Cornell High-Energy Synchrotron Source (CHESS). Data for WT and all variants was collected at 100 K. The following cryoprotectants were added: V48I:G80R dark (25% Glycerol v/v), V48I:G80R light (25% Glycerol v/v), G80R (25% ethylene glycol v/v), and WT (25% ethylene glycol v/v). Data was scaled and reduced in HKL2000 (Otwinowski and Minor, 1997) (see Equation S2 for refinement statistics). The initial WT structure was solved using molecular replacement in PHASER (McCoy et al., 2007) and PHENIX (Adams et al., 2010) with the LOV1 domain of Arabidopsis phototropin 2 (PDBID 2Z6D) as a search model. Structures of ZTL variants were solved using the same method with WT ZTL as the search model. Rebuild cycles were completed in COOT (Emsley and Cowtan, 2004) and refinement with REFMAC5 (Murshudov et al., 1997) and PHENIX (Adams et al., 2010). All ZTL structures contain four molecules per asymmetric unit that is composed of two anti-parallel LOV dimers. Residues 29–31 are not visible in the electron density in any molecule. In several molecules residues 29–43 and 164–165 are unable to be resolved and have not been built. For light-state structures clear density is observed for the adduct state in two of four molecules. Although electron density suggests an adduct for the remaining two molecules, we do not model them with an adduct and the reduced density likely reflects reduction by x-rays during data collection as has been observed previously (Zoltowski et al., 2007).
Purified protein fractions were concentrated to 30–60 µM for UV-Vis absorption spectroscopy measurements and kinetics. Samples were exposed to a broad spectrum white flood light (150 W), while on ice for two minutes. An Agilent UV3600 spectrophotometer was used to characterize the absorption spectra of all constructs in both light and dark states. The light state peak of 378 nm (ext. coefficient 8500) and dark state peak 450 nm (ext. coefficient 12,500) were used to estimate the protein concentrations for experiments.
Photocycle recovery kinetics were analyzed by measuring the absorption at 450 and 478 nm as a function of time. Spectra were collected at intervals ranging from 100 seconds-2 hours to ensure minimal repopulation of the light-state by the probe source. Time intervals were chosen to maintain approximately 10–20 measurements per half life. Kinetic traces at 450 and 478 nm were then fit with a monoexponential decay of the form y = y0 + A*e-k*x. The rate constant k and time constant (1/k) were abstracted. Results are presented in Table 1 as the average of three independent measurements.
The Columbia-0 (Col-0) plant that possesses CCA:LUC reporter was previously described (Pruneda-Paz et al., 2009). To generate overexpressors of HA-ZTL, HA-ZTL (V48I), HA-ZTL(G80R) and HA-ZTL (V48IG80R), the nucleotide sequences encoding HA tag was incorporated into the ZTL forward primer (5’-CACCATGTACCCATACGATGTTCCAGATTACGCTGAGTGGGACAGTGGTTC-3’, the underline indicates the sequences that encodes HA tag). Amino acid substitutions on ZTL coding region were generated by using megaprimer-based PCR amplification method (Burke and Barik, 2003). The primers used for generating the mutated ZTL coding sequence are followings; ZTL (V48I)_R (5’- AACGGCATCAGTAACAATGAATCCACAAGGCGC-3’), ZTL (G80R)_R (5’- CAAGAAGCGGCAATTTCGTCCGAGAACTTCCTC-3’), ZTL_R (5’- TTACGTGAGATAGCTCGCTA-3’). Amplified PCR fragments were cloned into pENTR/D-TOPO vector (Invitrogen). After verifying sequences, HA-ZTL and mutated HA-ZTL coding regions were transferred into pB7WG2 or pH7WG2 binary vector (Karimi et al., 2002) using LR Clonase II enzyme mix (Invitrogen) to generate 35S:HA-ZTL, 35S:HA-ZTL (V48I), 35S:HA-ZTL (G80R) and 35S:HA-ZTL (V48I:G80R). The binary vectors were introduced into the CCA1:LUC plants by conventional Agrobacterium-mediated transformation method. The T3 generations of transgenic lines in which the expression levels of ZTL variant mRNAs were similar were chosen for the circadian analysis. The plants were grown on soil or Linsmaier and Skoog (LS) media (Caisson) in plant incubator (Percival Scientific; Perry, Iowa) set at 22°C under full-spectrum white fluorescent light (70–90 µmol m−2s−1: F017/950/24’, Octron Osram Sylvania) in long days (16 hr light/8 hr dark).
Bioluminescence Imaging and analysis were performed as previously described with minor modifications (Fenske et al., 2015). Seedlings were grown on LS media in the plant incubator (Percival Scientific) in 12 hr light/12 hr dark photoperiod for 10 days before being transferred to continuous light (40–50 µmol m−2s−1) conditions. 9-day-old seedlings were pretreated with 5 mM D-luciferin (Biosynth) in 0.01% Triton X-100 solution, and incubated one day before imaging. The bioluminescence generated from the CCA1:LUC reporter was imaged for 15 min at every 2 hr using NightOwl system (Berthold; Germany) and analyzed using IndiGO software (Berthold). Period length estimation was performed using fast Fourier Transform-Nonlinear Least Squares (FFT-NLLS) analysis in the Biological Rhythms Analysis Software System (BRASS) (http://millar. bio.ed.ac.uk/PEBrown/BRASS/BrassPage.htm).
For gene expression analyses, 14-day-old seedlings grown on LS agar plates were harvested at ZT16 and used for RNA extraction using illustra RNAspin Mini kit (GE Healthcare; Chicago, Illinois). 2 µg of RNA was reverse-transcribed using iScript cDNA synthesis kit (Bio-Rad; Hercules, California). The cDNA was diluted by adding 40 µL of water, and 2 µL of cDNA was used for quantitative polymerase chain reaction (q-PCR) using MyiQ real-time thermal cycler (Bio-Rad). IPP2 expression was used as an internal control to normalize cDNA amount. Primers and PCR conditions used for IPP2, PRR5, TOC1 and ZTL amplification were previously described (Baudry et al., 2010). Expression of ZTL, PRR5 and TOC1 was calculated from three biological replicates.
To analyze the expression levels of HA fused ZTL protein, endogenous PRR5, and TOC1 proteins, seedlings were grown in 12 hr light/12 hr dark conditions for 14 days. Total protein was extracted using extraction buffer [50 mM Na-P pH7.5, 150 mM KCl, 1 mM DTT, 1 mM EDTA, 0.05% Sodium deoxycholate, 0.1% SDS, 50 µM MG-132, Protease inhibitor cocktail (Pierce)}. Protein was separated in 9% SDS-PAGE gels and transferred to Nitrocellulose membrane (Bio-Rad). HA-ZTL and Actin were detected using anti-HA (3F10, Roche) and anti-actin (C4, Millipore) antibodies, respectively. Western procedure for detecting TOC1 and PRR5 proteins was previously described (Baudry et al., 2010). For protein quantification, western blot images were analyzed using Image J (Schneider et al., 2012).
For Figure 4—figure supplement 1A, Agrobacteria containing both GI and ZTL variants were coinfiltrated into 4-week-old N. benthamiana leaves. The infiltrated plants were incubated under LD for two days and transferred to continuous light or dark with additional 24 hr incubation. Co-IP was performed according to Fujiwara et al. (2008). The immuno-complexes were resuspended in SDS sample buffer and heated briefly. GFP antibody (Invitrogen, A11120) was used for immunoprecipitation of GI-GFP protein. ZTL variants were detected by western blotting with HA antibody. For Figure 4—figure supplement 1B, Agrobacterium harboring each overexpression construct was mixed according to combinations indicated and was infiltrated into 3-week-old N. benthamiana leaves and incubated in LD and either light or darkness as described above. Sample preparation, the IP buffer condition, the IP method and immunoblot procedure were described previously (Song et al., 2012).
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- Brian D Zoltowski
- Brian D Zoltowski
- Young Hun Song
- Takato Imaizumi
- Brian D Zoltowski
- Takato Imaizumi
- David E Somers
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
This work is based on research conducted at the Cornell High Energy Synchrotron Source (CHESS). CHESS is supported by the NSF and NIH/NIGMS via NSF award DMN-0936384, and by NIGMS award GM-103485. We thank the Herman Frasch Foundation (739-HF12 to BDZ) the NIH (R15GM109282 to BDZ, R01GM079712 to TI and R01GM093285 to DES), Next-Generation BioGreen 21 Program (SSAC, PJ01117501 to YHS and PJ01117502 to TI, Rural Development Administration, Republic of Korea) and NSF (MCB-1613643 to BDZ) for funding. We thank Elaine Tobin for generously providing TOC1 antibody and Robert Green for experimental assistance.
- Richard Amasino, University of Wisconsin, United States
© 2017, Pudasaini et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.