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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Neuron. Author manuscript; available in PMC Sep 9, 2011.
Published in final edited form as:
PMCID: PMC2939042
NIHMSID: NIHMS228900

CDK5 serves as a major control point in neurotransmitter release

Abstract

CDK5 is an important kinase in nervous system function, controlling neural development and postsynaptic signal integration. Here we show that CDK5 plays a major role in controlling neurotransmitter release. Inhibition of CDK5 activity, by either acute or genetic means leads to profound potentiation of presynaptic function, including unmasking of previously “silent” synapses. Removal of CDK5 activity additionally unlocks access to the resting synaptic vesicle pool, which normally remains recalcitrant to exocytosis and recycling even following prolonged action potential stimuli. Presynaptic CDK5 levels are additionally severely depleted by chronic neuronal silencing, a treatment that is functionally similar to CDK5 KD with regard to presynaptic potentiation. Thus CDK5 appears to be an integral element in presynaptic homeostatic scaling and the resting vesicle pool appears to provide a potent functional presynaptic homeostatic control parameter. These studies thus pinpoint CDK5 as a major control point for modulation of neurotransmitter release in mammalian neurons.

Cyclin-dependent kinase 5 (CDK5), a proline directed serine/threonine kinase, has emerged in the last decade as an important enzyme critical for neural development (Kwon and Tsai, 2000; Ou et al., 2010), neural signaling pathways and neurogenesis (Lagace et al., 2008). In the basal ganglia, CDK5 serves as regulator of DARP32, a central control point in dopamine signaling (Chergui et al., 2004). CDK5 has been implicated in certain forms of synaptic plasticity, such as hippocampal-dependent spatial learning (Hawasli et al., 2007) and learned fear extinction (Sananbenesi et al., 2007). Additionally CDK5 has become a potential therapeutic target (Wei and Tomizawa, 2007), as it has been implicated in a number of pathological conditions, including Alzheimer's disease (Cruz and Tsai, 2004). A number of presynaptic proteins have been identified as CDK5 substrates (Cousin and Robinson, 2001; Samuels et al., 2007; Taniguchi et al., 2007), and in nerve terminals this enzyme has been largely attributed in playing a role in synaptic vesicle endocytosis (Anggono et al., 2006; Tan et al., 2003; Tomizawa et al., 2003). Although acute application of a small molecule inhibitor of CDK5 was previously shown to significantly enhance synaptic transmission in hippocampus (Tomizawa et al., 2002), it has been suggested that this results from off-target effects of this drug via interaction with calcium channels (Yan et al., 2002). This conclusion relied on the fact that small molecule inhibitors potentiated calcium currents equally well in the absence of p35, a major cyclin for CDK5 in the brain, however residual activity driven by p39, a synaptically-localized CDK5 cyclin (Humbert et al., 2000), had not been ruled out.

In order to investigate the potential role of CDK5 in neurotransmitter release we made use optical assays of exocytosis that unambiguously report measures of presynaptic function without hindrances arising from potential postsynaptic effects. We took advantage of both acute pharmacological inhibition of CDK5 as well as shRNA-meditated ablation of CDK5 activity in combination with these optical assays. Our experiments revealed that CDK5 normally serves to potently inhibit neurotransmitter release which, at the extreme, results in complete silencing of nerve terminals. Acute inhibition of CDK5 unmasks these previously inactive presynaptic sites. In active terminals we found that CDK5 controls access to the “resting” synaptic vesicle pool. At nerve terminals synaptic vesicles recycle to sustain neurotransmitter release during repetitive action potential (AP) firing. The size of the pool of vesicles that recycles and the kinetics of each of the recycling steps all play important roles in determining the efficacy of synaptic function during repetitive stimulation (Fernandez-Alfonso and Ryan, 2006). We and others have previously shown that in addition to vesicles that undergo exocytosis and recycling in response to stimulation (the readily-releasable and reserve pools), a large fraction of vesicles fail to participate in recycling, and the size of this fraction varies from synapse to synapse (Fernandez-Alfonso and Ryan, 2008; Fredj and Burrone, 2009; Harata et al., 2001; Li et al., 2005). Our experiments show that the size of this resting vesicle pool is determined by the balance of calcineurin and CDK5 activity, and that resting vesicles can be converted to recycling vesicles through CDK5 inhibition. Furthermore the size of the resting vesicle pool can be increased by removal of calcineurin. Finally we show that the endogenous balance of CDK5 and calcineurin activities is determined by the history of chronic activity, as long term neuronal silencing by TTX phenocopies both acute and shRNA-mediated removal of CDK5 activity with respect to changes in presynaptic function. We show that that neuronal silencing leads to a loss of presynaptic CDK5 indicating that this enzyme is a substrate in homeostatic plasticity and that the resting vesicle pool provides an important presynaptic control parameter in homeostatic scaling. These studies thus identify CDK5 as an important control point in determining neurotransmitter release properties.

Results

In order to measure exocytosis with high fidelity, we made use of vGlut1-pHluorin (vG-pH) or vGlut1-pHluorin-mCherry (vG-pH-mCh) expressed in dissociated hippocampal neurons. pHluorin, a modified form of GFP whose fluorescence is quenched upon protonation (pKa ~7.1) (Sankaranarayanan et al., 2000), can be targeted to the lumen of synaptic vesicles by attachment to one of several different synaptic vesicle proteins. Fusion of pHluorin into one of the intralumenal loops of the vesicular glutamate transporter (vGlut1) (Voglmaier et al., 2006) results in very low surface expression and provides high sensitivity for measurement of exocytosis (Balaji and Ryan, 2007). We additionally fused a copy of mCherry, a pH-insensitive and red-shifted protein, to the cytosolic C terminus of vG-pH. When targeted to the synaptic vesicle lumen, pHluorin fluorescence is quenched and upon AP stimulation and exocytosis it is relieved (Figure 1A). Following exocytosis, the signal decays due to endocytic capture of vG-pH and vesicle reacidification. Although during the stimulus train the signal in principle reflects the difference between exocytosis and ongoing endocytosis and reacidification, we previously showed that during the initial 10 s of stimulation the signal represents almost purely exocytosis, as the combined kinetics of endocytosis and reacidification are too slow to impact the measurement (Kim and Ryan, 2009b). In order to examine the impact of CDK5 inhibition on presynaptic function, we made use of Roscovitine, a small molecule inhibitor, as well as shRNA-mediated knockdown of CDK5. Double transfection of vG-pH and shRNA targeting CDK5 resulted in a ~92% reduction in CDK5 levels (Figure S1A and C). We monitored vG-pH responses to 100 AP stimuli at 10 Hz across ~20–40 boutons of an individual neuron simultaneously. Signals were normalized at each bouton to the fluorescence value obtained by rapid alkalization of the entire labeled vesicle pool using NH4Cl, thus correcting the signals for possible variation in expression levels. Application of Roscovitine resulted on average in a ~2.5-fold potentiation of exocytosis of the ensemble response (Figure 1A and 1B), while control cells treated only with vehicle remained unchanged (Figure S2). Comparison of the response amplitude distribution across several hundred boutons from many cells showed that shRNA-mediated removal of CDK5 activity resulted in very similar responses to wild type (WT) neurons treated with Roscovitine (Figure 1C and 1D). Furthermore treatment of shRNA-treated neurons with Roscovitine resulted in only a very small further increase in amplitude to a 100 AP stimulus (Figure 1A and B), and transfection of KD neurons with an shRNA-insensitive cDNA encoding CDK5 fully restored the 100 AP response to WT levels (Figure S5A and S5B). These data demonstrate that CDK5 normally provides a strong inhibitory influence on presynaptic function and rule out any significant CDK5-independent impact of Roscovitine in these measurements.

Figure 1
The Exocytosis elicited by brief action potential stimuli (100AP) is potentiated following removal of CDK5 activity

Unmasking presynaptically silent synapses by inhibition of CDK5

We noted in our control experiments that a small percentage of synapses showed no measurable signal during a 100 AP stimulus (Figure 1C). These synapses were identified by the signal obtained following NH4Cl treatment or by the mCherry channel. Following Roscovitine treatment however these previously silent nerve terminals showed very robust responses to 100 AP stimuli (Figure 2A and 2B). We quantified the magnitude of the unmasking by specifically measuring the impact of Roscovitine treatment on those synapses whose signal in response to first 100 AP stimulus was within 1 standard deviation of the baseline noise in the recording, which represented ~ 11 % of the entire distribution of synapses in the histograms in Figure 1C. Following CDK5 inhibition, a large fraction of these specific synapses showed 100 AP responses that were now well above the noise floor (Figure 2). The average response of these synapses grew from an exocytic response that corresponded to 0.5% of the entire vesicle pool (<1 vesicle, assuming an SV pool < 200 vesicles) to 8% of the entire pool following Roscovitine treatment. Following Roscovitine treatment ~5% of synapses remained unresponsive to AP-driven activity, almost identical to the fraction of synapses that were unresponsive in CDK5 knockdown neurons (6.1%). Of the synapses whose responses were initially within the baseline noise, but subsequently gave responses above the baseline noise (the ~5% of the population that became”unmasked”), the mean response was ~14%, close to the central mode of the entire distribution of potentiated synapses (Figure 1C).

Figure 2
Acute inhibition of CDK5 unmasks silent synaptic boutons

CDK5 inhibition unlocks access to the resting synaptic vesicle pool

Under control conditions we previously showed that on average no more than ~50% of the entire vesicle pool typically undergoes exocytosis even after several hundred AP stimuli. Following Roscovitine treatment we noticed that for individual boutons the 100 AP stimulus resulted in signals as high as 80–90% of the entire vesicle pool as judged by the fluorescence obtained by direct alkalinization with NH4Cl treatment (Figure 1E). The fact that a mere 100 AP was now driving individual synapses to exocytose virtually all of their SV pool suggests that CDK5 inhibition might be increasing the size of the available SV pool, presumably through conversion of non-recycling vesicles into recycling vesicles. We tested this hypothesis in two ways. Following exocytosis synaptic vesicle components are recaptured by endocytosis and the regenerated synaptic vesicle is reacidified by a V-type ATPase. During prolonged stimulation, vG-pH signals represent the balance of exocytosis, endocytosis and reacidification, however in the presence of a V-ATPase inhibitor, the signal represents the cumulative amount of exocytosis (Sankaranarayanan and Ryan, 2001). An example of this is illustrated in Figure 3A. After several hundred AP stimuli the fluorescence reaches a plateau, which arises from the fact that all vesicles that can fuse with the plasma membrane have done so, and have become trapped in the alkaline state, as even though endocytosis persists, no reacidification occurs. Thus the entire recycling pool of synaptic vesicles becomes alkaline at this point. Subsequent application of NH4Cl allows one to determine if any vesicles remained in the acidic state following this protocol. Under control conditions these measurements indicate that on average only ~51% of the vesicle pool labeled with vG-pH can participate in recycling (Figure 3A and 3C) in agreement with previous published results (Fernandez-Alfonso and Ryan, 2008; Li et al., 2005). When we repeated this protocol on neurons following acute application of Roscovitine, the size of the recycling pool grew substantially to ~80% (Figure 3B and 3C). This trend persisted across all neurons examined, where the average recycling pool size following CDK5 inhibition was 81%. Single bouton analysis derived from many neurons also supported this view (Figure 3D and 3E) and demonstrated that following acute inhibition of CDK5, recycling pools at individual synapses spanned a much narrower range of values than in untreated neurons.

Figure 3
Acute Inhibition of CDK5 increases the recycling pool and decreases the resting pool in nerve terminals

We next used a different strategy, based on uptake of an exogenous tracer (FM 4–64) into recycling vesicles. FM dyes are a family of amphipathic molecules that are impermeant to the plasma membrane, fluoresce when bound to membranes and can be loaded during vesicle cycling into synaptic vesicles. Following a 600 AP stimulus at 10 Hz in the presence of FM 4–64, nerve terminals become maximally loaded, as exposure to the dye during a second round of stimulation leads to only a small increase in total fluorescence (Figure 4B and 4C, control) in agreement with previous results (Ryan and Smith, 1995). In contrast, if prior to the attempt to load additional vesicles in a second round of stimulation the nerve terminals are incubated in the presence of Roscovitine, a much larger fluorescence load is achieved (Figure 4B and 4C, Roscovitine). In the continued presence of Roscovatine these newly loaded vesicles were fully usable again, as additional stimulation in the absence of extracellular FM 4–64 led to destaining down to levels that were similar to controls. On average the loading increased by 86% in the presence of Roscovitine compared to only 11% in controls (Figure 4C and 4D). These data combined with the results using vG-pH demonstrate that acute inhibition of CDK5 leads to a very large increase in the size of the recycling vesicle pool by converting resting vesicles into functional recycling vesicles.

Figure 4
Acute CDK5 inhibition increases the size of the recycling vesicle pool as determined by FM 4–64 uptake

A balance of CDK5 and Calcineurin activity determines the size of the recycling and resting vesicle pools

In order to further verify that the increase in pool size was due to inhibition of CDK5 and not an-off-target effect we used the vG-pH-based protocol (Figure 3) in CDK5 knockdown neurons as well as in neurons transfected with an shRNA targeting the regulatory subunit of calcineurin (calcineurin B, CNB), one of the main proline-directed serine/threonine phosphatases known to counteract CDK5 in nerve terminals (Cousin and Robinson, 2001). The regulatory and catalytic subunits of calcineurin form an obligate heterodimer, and loss of the regulatory subunit leads to complete loss of calcineurin function (Zeng et al., 2001). Transfection of shRNA-targeting CNB resulted in a ~94% loss of CNB (see Figure S1B and S1C). We measured the balance of recycling and resting vesicles in both CDK5 KD and CNB KD neurons and found that in the absence of CNB the recycling fraction was 20% (Figure 5A) similar to observations made under purely pharmacological blockade (Kumashiro et al., 2005), and much smaller than in WT, while in the absence of CDK5 it was 79%, almost identical to the value obtained following acute inhibition. Both CDK5 and CNB KD phenotypes could be fully rescued by expression of shRNA-resistant plasmids (Figure S5). Furthermore acute treatment of CNB KD neurons with Roscovitine also counterbalanced the loss of CNB, restoring vesicle pool sizes to near WT levels (Figure S4). The fact that removal of a phosphatase leads to changes that are opposite to that of removal of the kinase by either acute or genetic means implies that it is CDK5's enzymatic activity that is needed for control of the recycling and resting synaptic vesicle pool sizes. Single bouton analysis of the recycling fraction also showed that, like with Roscovitine treatment, the range of values exhibited by synapses in either CDK5 KD or CNB KD was much smaller than in WT, although with CNB removal the values spanned the range of 0 to ~50%.

Figure 5
CDK5 and Calcineurin control the recycling pool size

Chronic silencing of neuronal activity depletes presynaptic CDK5 activity

In WT neurons the values of the recycling synaptic vesicle pool fraction showed a large degree of variation from synapse to synapse (Figure 3D), but following acute inhibition of CDK5 this range was reduced by roughly half. These data suggest that the local balance of calcineurin and CDK5 is determined in part at the individual synapse level and varies from synapse to synapse. We hypothesized that one potentially important variable across synapses that might determine this local balance of calcineurin and CDK5 activity was the chronic local history of activity. We tested this hypothesis by measuring the recycling synaptic vesicle pool fraction in neurons that had been silenced for 72 hrs using TTX. Figure 6 shows an example of the ensemble response of presynaptic terminals from an individual neuron that had been silenced in such a fashion prior to the experiment. The size of the recycling vesicle pool fraction was significantly larger than in wild-type (Figure 6A and 6C), similar to CDK5 KD neurons. Furthermore the size of the recycling pool was identical in CDK5 KD neurons that had also been silenced with TTX (Figure 6B and 6C). Analysis of the recycling pool values from individual boutons also showed that, as is in the case where CDK5 activity is removed, the range of recycling pool sizes was restricted to a narrower range (Inset, Figure 6A and 6B). The remarkable similarity of chronic neuronal silencing and explicit ablation of CDK5 activity upon the size of the recycling vesicle pool strongly suggests that this silencing directly impacts either presynaptic CDK5 activity levels or the availability of a relevant CDK5 substrate. In order to examine this issue we directly measured protein expression levels with two different approaches in control compared to chronically silenced neurons. Western blot analysis of primary neuronal cell cultures indicated that CDK5 expression decreased to ~50% in TTX-treated compared to control neuronal cultures, while other presynaptic proteins, including several known CDK5 substrates, showed no significant change (Figure 6D and 6E). Given that CDK5 appears to be localized throughout the neuron and is also expressed in glial cells (Figure S1) we surmised that the partial global loss of CDK5 might not accurately reflect presynaptic CDK5 levels. We examined the impact of chronic silencing on presynaptic CDK5 levels by direct co-transfection of neurons with GFP-synapsin I and an epitope-tagged CDK5 (HA-CDK5) that would permit robust immunocytochemical localization and quantification. These neurons were then treated with or without TTX for several days after allowing 4 days for initial protein expression following transfection at DIV 8. Immunocytochemical analysis of GFP-synapsin I and HA-CKD5 revealed that chronic neuronal silencing leads to a dramatic suppression of presynaptic CDK5 levels down to ~13% of control neurons (Figure 6F and 6G). These data indicate that CDK5 is a substrate of presynaptic homeostatic plasticity and that changes in recycling versus resting pools driven by silencing can be accounted for by changes in presynaptic CDK5 levels. As a further test of the hypothesis that TTX treatment impacts the functionality of CDK5 we examined the impact of TTX treatment on neurons in which CNB activity had previously been ablated. These experiments revealed that in the absence of the counterbalancing phosphatase, the recycling pool sizes returned to WT levels in TTX silenced neurons (Figure 7A). Finally we also examined the proportion of presynaptically silent synapses in TTX treated neurons. This analysis showed that the presynaptically silent pool was reduced to 5.6% of synapses examined, very similar to that of CDK5 KD (5%) and Roscovitine treated neurons (6.1%, Fig 2).

Figure 6
Chronic suppression by tetrodotoxin phenocopies the loss of CDK5 activity
Figure 7
The balance of CDK5 and Calcineurin activities determine key properties of neurotransmitter release

The balance of CDK5 and calcineurin activities control the kinetics of exocytosis

In addition to the large increase in the size of the pool of synaptic vesicles that can engage in exocytosis and recycling, we also found that the kinetics of exocytosis of that pool depends critically on the balance of CDK5 and calcineurin activities. The kinetics of exocytosis can be determined by examining the time course of the exponential rise in fluorescence of vG-pH during stimulation in the presence of bafilomycin (Figures 3A and 3B, 5A and 5B, 6A and 6B). Data from all the different experimental conditions tested are illustrated in Figure 7B which shows that all conditions that resulted in elimination of CDK5 activity as well as the chronic neuronal silencing led to exocytosis kinetics that were ~ 2-fold faster than in WT, while in the absence of calcineurin, exocytosis kinetics slow ~3-fold compared to WT (Figure 7C). Similar results were obtained by examining the impact of acute CDK5 inhibition on the kinetics of FM 4–64 destaining (Figure S6). Our data thus show that exocytosis kinetics, as probed with repetitive stimulation is strongly potentiated by loss of CDK5. Under such stimulus conditions the rate of exocytosis might be controlled at a variety of different biochemical steps, including the efficiency of exocytosis and vesicle pool refilling. Examination of responses to single or paired action potential stimuli revealed that removal of CDK5 had little impact compared to controls (Figure S7). These data suggest that the CDK5 is likely acting on a step upstream of exocytosis in controlling sustained exocytosis rates and underscore the central claim in this work, that CDK5 normally provides a strong suppression of presynaptic function both influencing the number of vesicles that can participate as well as the ability to efficiently use those vesicles in driving neurotransmitter release. The experiments also demonstrate that suppression by CDK5 is counterbalanced by the activity of calcineurin in the control of exocytosis kinetics.

Discussion

The results we report here demonstrate 1) that the kinase CDK5 normally acts as a strong suppressor of neurotransmitter release; 2) that is counterbalanced by the activity of calcineurin; 3) that the balance of these activities acts on both the kinetics of exocytosis as well as in regulating access to the resting synaptic vesicle pool; 4) that the functional consequence of the balance of these two activities in turn is influenced by chronic activity levels indicating that these activities are likely potent presynaptic substrates of homeostatic scaling; 5) that presynaptic CDK5 levels are highly suppressed by chronic neuronal silencing, indicating that CDK5 itself is a critical substrate in homeostatic feedback. Previously, suppression of neurotransmitter release by the CDK5 inhibitor Roscovitine has been attributed to direct interaction of the drug with Ca2+ channels, independent of CDK5 itself. These previous experiments however had not examined the impact of these inhibitors in the absence of CDK5, only, in one case, in the absence of one of CDK5's activators, the cyclin p35 (Yan et al., 2002). Our results demonstrate that with respect to the experiments performed here, the action of Roscovitine is via CDK5, as the removal of CDK5 by shRNA phenocopies the impact of Roscovitine in each of the assays performed, the impact of Roscovitine is largely occluded in the absence of CDK5 and removal of a counterbalancing phosphatase activity has opposite effects to CDK5 inhibition.

Our experiments revealed that a proportion of nerve terminals (~10%) have very little activity in terms of exocytosis, releasing less than 1 vesicle in a 100 AP train at 10 Hz, but ~half of these nerve terminals could be shifted to the active state by acute Roscovitine treatment. Those that did shift showed very robust responses, well within the central mode of the distribution of all Roscovitine-potentiated synapses. In the absence of CDK5 or in chronic suppression by TTX, the fraction of inactive synapses was only ~5%, similar to the remaining inactive population in WT after Roscovitine treatment, while in the absence of calcineurin this population grows significantly (~42%). These data imply that in principle shifting the balance of CDK5 and calcineurin can be used to control the population of presynaptically silent or presynaptically active synapses and that a portion of synapses are normally effectively clamped to an inactive state by CDK5.

The observation that synaptic terminals contain a fraction of vesicles in the “resting” state that do not normally recycle was first made by carrying out photoconversion of FM dye loaded terminals followed by electron microscopy. These studies revealed that in spite of maximal dye loading, fewer than 1/3rd of vesicles appeared to have loaded the dye and therefore undergone recycling (Harata et al., 2001). Subsequent experiments using pHluorin-tagged SV proteins came to similar conclusions (Fernandez-Alfonso and Ryan, 2008; Li et al., 2005). We and others hypothesized that such a reservoir of vesicles could provide an important dynamic variable by which to modify synaptic performance, however no known mechanisms for tapping into this pool had been uncovered. For example, previous work demonstrated that increasing calcium entry (for example by using higher stimulus frequencies) does not change the size of the accessible vesicle pool (Fernandez-Alfonso and Ryan, 2008). Recent work suggested that this resting pool provides the source of vesicles for spontaneous fusion events and not AP evoked neurotransmitter release (Fredj and Burrone, 2009; Sara et al., 2005), however this idea has been directly challenged (Groemer and Klingauf, 2007). The studies reported here demonstrate that access to this resting pool with respect to AP-driven exocytosis is determined by the balance of CDK5 and calcineurin activities, and that the consequence of this balance in turn is set by long term neuronal activity through control of CDK5 levels. These studies suggest that dynamic control of this balance could provide a potent control system for tuning synaptic performance.

Previously, the major role attributed to CDK5 within the nerve terminal has been to phosphorylate a number of endocytic proteins (the dephosphins). One study that examined the impact of CDK5 inhibition on endocytosis concluded that inhibition of CDK5 led to an acceleration of endocytosis (Tomizawa et al., 2003). Our data indicates that the apparent acceleration can be attributed to the fact that a much larger pool of vesicles becomes loaded following CDK5 inhibition. The work we present here has not examined the impact of CDK5 inhibition on endocytosis kinetics, and will be the subject of future studies.

A number of studies in the last decade have pointed to the fact that neuronal circuits appear to be governed by homeostatic scaling principles, whereby both pre and postsynaptic efficacy can be auto-tuned to compensate for long term changes in levels of synaptic transmission (Davis, 2006). Significant evidence has accumulated indicating that postsynaptic scaling is manifest as a change in glutamate receptor number or composition (Turrigiano, 2008). In peripheral synapses, a mechanism involving Eph receptors, Cdc42 and voltage gated Ca2+ channels has been shown to operate during homeostatic regulation in nerve terminals (Frank et al., 2009). In hippocampal neurons, chronic silencing of networks in cell culture have been shown to increase synapse size and presynaptic efficiency (Murthy et al., 2001; Thiagarajan et al., 2005) as well as changes in the ratio of two isoforms of CaMKII (Thiagarajan et al., 2002). The fact that chronic silencing of neuronal activity decreases expression of presynaptic CDK5 indicates that this enzyme is an important element of homeostatic feedback regulation that results in upregulation of presynaptic function.

Two of the most important questions for future studies relating to this work are 1) to identify the substrate(s) of CDK5 that are controlling access to the resting pool and are modulating the kinetics of exocytosis and 2) to determine how CDK5 is normally regulated at nerve terminals. At present it is unknown if the acceleration of vesicle pool turnover is a direct consequence of unlocking the resting vesicle pool or if it results from CDK5/Calcineurin-based modulation of proteins that control a rate-limiting step in vesicle pool turnover. In addition to the originally identified set of endocytic proteins (dynamin, amphiphysin, and synaptojanin) collectively referred to as dephosphins, several other proteins that are known to be found in nerve terminals have also been identified as CDK5 substrates. These include Synapsin (Jovanovic et al., 1996; Matsubara et al., 1996), Septin 5 (Amin et al., 2008), Munc18 (Fletcher et al., 1999; Shuang et al., 1998), CASK (Samuels et al., 2007), CRIMP (Cole et al., 2008) and the CDK5-like kinase Pctaire (Cheng et al., 2002; Liu et al., 2006). Although Roscovitine treatment has been shown to prolong the open channel lifetime of both P/Q and N-type Ca2+ channels, these measurements were not made in the context of nerve terminals. Furthermore the impact on Ca2+ channels had been attributed to direct interaction with Roscovitine, although these experiments had not been carried out in the absence of CDK5. Our studies suggest that at least in the context of the synapse, Ca2+ channels may be modified in a CDK5 dependent fashion to control neurotransmitter release properties. Previous work indicated that for hippocampal neurons the effects of Roscovitine appear to be mediated through P-type Ca2+ channels, and one study showed that the synprint loop of P/Q-type Ca2+ channels can be phosphorylated in vitro by CDK5(Tomizawa et al., 2002). Thus it is plausible that the acceleration of exocytosis kinetics caused by removal of CDK5 may result from enhancement of Ca2+ channel function. Relatively little is currently known about how CDK5 is regulated, other than the identity of the two known cyclins, p35 and p39. It has been suggested that CDK5, like calcineurin, can be modulated by elevations in intracellular calcium, as CDK5 has been shown to interact with calmodulin in a calcium-dependent manner (Dhavan et al., 2002).

The close functional correspondence between neurons whose activity has been chronically silenced with those whose CDK5 activity has been ablated and the demonstration that presynaptic CDK5 levels are suppressed during silencing indicates a convergence at the level of molecular pathways for these experiments An important direction for future studies will be to decipher how silencing mediates loss of presynaptic CDK5.

The studies we describe here report previously unappreciated and potent control system for neurotransmitter release demonstrating that CDK5 normally suppresses presynaptic function in at least two ways. This work should open up new avenues in understanding how synaptic transmission is controlled in both normal and diseased states of brain function.

Experimental procedures

Cell culture and optical setup

Hippocampal CA3-CA1 regions were dissected from 2 day old Sprague Dawley rats, dissociated, and plated onto poly-ornithine-coated glass for 14–21 days as previously described (Ryan, 1999). All constructs were transfected 8 days after plating. Experiments were performed 14 – 21 days after plating (6 – 13 days after transfection), and the coverslips were mounted in a rapid-switching, laminar-flow perfusion and stimulation chamber (volume ~75 μl) on the stage of a custom-built laser illuminated epifluorescence microscope. Live cell images were acquired with an Andor iXon+ (Model # DU-897E-BV) back illuminated EMCCD camera in epifluorescence. A solid state diode pumped 488 nm or 532 nm laser (for mCherry excitation) that weres shuttered using Acousto-Optic Tunable Filters (AOTF) in all not data acquiring periods served as a common light source for setup. Fluorescence excitation and collection was done through a 40× 1.3 NA Fluar Zeiss objective using 515–560 nm emission and a 510 nm dichroic filters (for pHluorin) and a 572–647 nm emission filter (for mCherry). Action potentials were evoked by passing 1 ms current pulses, yielding fields of ~10 V/cm via platinum-iridium electrodes. Cells were continuously perfused (0.2 ml/min) in a saline solution containing (in mM) 119 NaCl, 2.5 KCl, 2 CaCl2, 2 MgCl2, 25 HEPES (buffered to pH 7.4), 30 glucose, 10 μM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), and 50 μM D,L-2-amino-5-phosphonovaleric acid (AP5). Temperature was clamped at 30°C to decrease effect from temperature fluctuation. Unless otherwise noted, all chemicals were obtained from Sigma (St. Louis, MO). NH4Cl applications were done with 50 mM NH4Cl in substitution of 50 mM of NaCl (buffered to pH 7.4). Advasep was obtained from CyDex (Lenexa, KS) and used at 1 mM where indicated. Bafilomycin (Calbiochem) was used at 1 μM where indicated. Roscovitine (Calbiochem) was used at 100μM where indicated. Tetrodotoxin (Alomone labs, Israel) was used at 1μM.

Immunofluorescence and quantification

To quantify the efficiency of shRNA-mediated knockdown, following live cell imaging, neurons were fixed with 4% paraformaldehyde and permeabilized with 0.2% Triton X-100, blocked with 5% BSA, and subsequently incubated with appropriated primary antibodies [anti-GFP(invitrogen, CA), anti-CDK5(invitrogen.CA), anti-calcineurin B (Millipore,MA), anti-HA (Covance)]. Alexa 488 or Alexa 546 conjugated secondary antibodies were applied in primary antibody incubated samples with different color combinations as needed. Immunofluorescence images of fixed cells were acquired using epi-fluorescence microscope with EM CCD camera. Expression levels of CDK5 and calcineurin were measured at cell bodies corrected for background in surrounding regions to avoid possible spatial overlap with other cells. This was compared to the fluorescence intensity in non transfected (GFP-negative) cell bodies. To quantify presynaptic CDK5 levels neurons were co-tranfectected with GFP-Synapsin I and HA-CDK5. Four days after transfection (DIV 12) neurons were treated with or without TTX (1μM) for 3 ~5 days prior to immunostaining. Cells were incubated with anti-HA (Covance, CA) and anti-GFP antibodies, subsequently incubated with Alexa 546 and Alexa 488 conjugated secondary antibodies HA-CDK5 levels were determined by centering ROIs on the synapsin channel and determining the ratio of the HA-CDK5 intensity to that of GFP-synapsin across many boutons. GFP-synapsin levels were not significantly affected by TTX treatment (not shown).

Western blot analysis

Hippocampal CA3-CA1 regions were dissected from 2~3 day old Spraque Dawley rats, dissociated, and plated onto poly-ornithine-coated 6-well dishes for 15~17 Days. Neurons in 3 wells were treated with TTX (1μM) at 12 days in vitro (DIV) for 3~5 days and neurons in the other 3 wells were not treated with TTX as controls. All experiments were performed as parallel with same condition. Cells were lysed with l buffer containing 10 mM Tris (pH 7.4), 1 % SDS, 10 mM NaF, 1mM PMSF, supplemented with proteases inhibitor mixture (Complete mini;Roche, Germany). Subsequently, concentration of lysates was quantified with BCA (Bicinchoninic acid) assay (Thermo,IL). Similar amounts of lysate were subjected to SDS-PAGE and subsequently transferred to PVDF membranes. Membranes were incubated with 5% non-fat dry milk for blocking, and subsequently incubated with appropriated antibodies [anti-dynamin I (BD transduction, CA), anti-Amphiphysin (gift from Dr. De Camilli), anti-synapsin I (synaptic system, Germany), anti-VAMP2 (calbiochem, CA), anti-Munc18 (sigma,MO), anti-Tubulin (Santacruz, CA)). Band intensities were quantified and measured ratio of [TTX+]/ [TTX−] after background subtraction.

Plasmids

Synthetic oligonucleotides containing the rat CDK5 cDNA target sequences (CCTCCGGGAGATCTACTCAAA) of CDK5 and rat calcineurin B subunit cDNA target sequences (CTATGTGTGACATCTTGTG) of calcineurin B regulatory subunit for cloning in pSUPER vector were synthesized (Invitrogen,CA), annealed and ligated into pSUPER with BglII and HindIII enzyme site according to manufacturer's instruction. vGlut1-pHluorin-mCherry (vG-pH-mCh) was subcloned that mCherry was connected in frame to C-terminus of vG-pH exposed to cytoplasmic region. The HA-CDK5 construct resistant to shRNA CDK5 was generated using Quick Change Site-Directed mutagenesis (Stratagene, La Jolla, CA). The sequence for shRNA CDK5 corresponding to nucleotide positions, 144–168, was mutated to CCTGCGCGAAATATGTCTACTCAAA leaving the amino acid sequence unchanged. For rescue of CN KD, human cDNA of calcineurin B was utilized, which has one nucleotide mismatch.

Image and Data Analysis

Images were analyzed in ImageJ (http://rsb.info.nih.gov/ij) using a custom written plugin (http://rsb.info.nih.gov/ij/plugins/time-series.html). All visible varicosities were selected for analysis by testing their responsiveness to test application of NH4Cl. Fluorescence time course traces were analyzed using Origin Pro (ver 7.5). The peak of amplitude at 100 AP was selected at the end of stimulation at 10Hz 10s. BAF and NH4Cl were analyzed by mean value of plateau regions. Silent boutons were defined as those where the response of 100APs was smaller than the standard deviation (σ) of the baseline before stimulation (ΔF100 − σ ≤ 0). In order to measure the rate of exocytosis, the traces in the presence of BAF were fitted with single exponential as previously described (Kim and Ryan, 2009a).

Supplementary Material

01

Acknowledgements

The authors would like to thank Jeremy Dittman and members of the Ryan lab for valuable discussions. We thank Ricky Kwan for excellent technical assistance. vG-pH was a gift from Rob Edwards and Susan Voglamaier (UCSF). This work was supported in part by NIH grants to TAR (NS036942, MH085783) and a sponsored research agreement with Galenea Corp (Cambridge MA).

Footnotes

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