Phasic and delayed glutamate release occurs at axon-OPC synapses upon axonal stimulation with trains of 20 pulses at 100 Hz

Our first step on the way to understanding the response of OPCs to different patterns of neuronal activity was to study functional changes at axon-OPC synapses upon repetitive axonal stimulation in situ. We performed whole-cell patch-clamp recordings of callosal OPCs in brain slices prepared from postnatal day 19–22 (P19-P22) NG2-DsRed mice and stimulated callosal axons electrically with trains of stimuli. The recorded cells had membrane resistance of 767 ± 157 MOhm (n = 36) and showed voltage-gated outward K⁺ currents upon depolarization (n = 36), identifying them as OPCs [20–22]. We initially used the paradigm of 20 pulses at 100 Hz. The response of OPCs to train stimulation of axons consisted of two kinetically distinct components: phasic excitatory postsynaptic currents (EPSCs) during the train and delayed EPSCs, which occurred after the final stimulus in the train and continued for several hundreds of milliseconds (Fig 1A). To exclude the possibility that delayed axon-glia EPSCs are triggered by polysynaptic activation of callosally projecting neurons or result from dissimilar conduction velocities in individual fibers, we performed all subsequent experiments in slices where the corpus callosum has been isolated from the cortex (see Materials and methods) and used the minimal stimulation paradigm (Fig 1A and 1B), designed to stimulate a single axon contacting the recorded cell [10, 23]. Under these conditions, we were still able to record phasic and delayed axon-glial EPSCs, although we observed more failures of release during the train and fewer delayed events after the train.

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Fig 1. Phasic and delayed currents occur in callosal oligodendrocyte precursor cells (OPCs) upon train stimulation of callosal axons with 20 pulses at 100 Hz.

(A) Representative example traces showing phasic and delayed currents in OPCs (holding potential [V_h] = −80 mV) upon nonminimal (top) and minimal (bottom) stimulation of callosal axons. (B–J) Unitary, delayed, and miniature excitatory postsynaptic currents (EPSCs) in OPCs. (B, E, H) Original example events. (C, F, I) Average current waveforms generated by averaging of (C) 34 unitary events without failures, (F) 170 delayed events, and (I) 106 miniature events. Red lines indicate monoexponential fits to the decaying phase of the current. (D, G, J) Amplitude distribution histograms fitted with lognormal function (red line). Total of (D) 265 unitary events from 20 OPCs, (G) 2,089 delayed events from 14 OPCs, and (J) 370 miniature events from 4 OPCs were pooled for each histogram. Blue empty histogram in (D) represents amplitude distribution histogram of 894 failures pooled from the same 20 OPCs that were used for the histogram of responses. Black empty histograms in (G, J) indicate noise distribution. (K–M) Average current amplitude including failures, response probability, and response potency during minimal stimulation train. Each point represents the mean value from 14 cells for a given stimulus. To ease visualization of facilitation, a dashed red line is drawn at the level corresponding to the value at the first stimulus. One-way ANOVA with post hoc Dunnett’s test was used to compare the values for each stimulus to the value measured at the first stimulus (S1 Table). Each point represents mean ± SEM. (N) Average rate of delayed events after the minimal stimulation train. 1,702 events from 14 cells were pooled for this histogram. Red line indicates monoexponential fit to the events rate. (O–P) Rise time and decay time constants of unitary, delayed, and miniature EPSCs (mEPSCs) in OPCs. The values were measured on the same events used for the amplitude distribution histograms in (D, G, J). One-way ANOVA (S1 Data). *p < 0.05, **0.001 < p < 0.01, ***p < 0.001. Box and whisker plots: the bottom and top of each box represent 25th and 75th percentiles of the data, respectively, while whiskers represent 10th and 90th percentiles. The midline represents the median. The numerical data used in K–M and O–P are included in S2 Data.

https://doi.org/10.1371/journal.pbio.2001993.g001

The amplitude distribution histogram of unitary axon-glia EPSCs (without failures) could be fitted with lognormal function and showed one peak at 2.8 pA (n = 20; Fig 1B–1D), which was comparable to the histogram peak of miniature EPSCs (mEPSCs) at 2.2 pA (n = 4; Fig 1H–1J). Unitary EPSC amplitude distribution was skewed towards larger values (Kolmogorov-Smirnov test, p < 0.001), indicating that more than one vesicle can be released at a single axon-OPC connection [10]. The initial average amplitude of phasic axon-glia EPSCs (including failures) was 1.58 ± 0.33 pA, and the failure rate estimated from minimal stimulation was 73 ± 5% (n = 14 cells, Fig 1K and 1L), which is similar to the values reported previously [10]. The EPSC amplitude showed progressive facilitation at the beginning of the train, doubled by the third pulse, and returned to baseline by the end of the train (n = 14, Fig 1K). The release probability displayed comparable dynamic, while the response potency remained mostly unaltered (n = 14, Fig 1L and 1M), pointing to the fact that activity-dependent synaptic enhancement at axon-OPC synapses during the train is of presynaptic origin.

In addition to facilitation of phasic release, repetitive axonal stimulation increased the likelihood of spontaneous axon-glia EPSCs after the train. The rate of these delayed EPSCs decayed monoexponentially with a time constant of 98.7 ms (Fig 1N). The amplitude distribution histogram of delayed events was similar to the histograms of unitary EPSCs and mEPSCs: it could be fitted with lognormal function and showed one peak at 3.3 pA (n = 14 cells; Fig 1E–1G). The 10%–90% rise time and decay time constant of unitary, delayed, and mEPSCs were also comparable (Fig 1O and 1P). Hence, each delayed axon-glia EPSC likely represents a quantal response, as it is assumed for neuronal synapses [23].

Phasic and delayed axon-glia EPSCs were blocked by tetrodotoxin (TTX) (0.5 μM, n = 3, not shown) or by 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) (10 μM, n = 3, not shown), indicating that they depend on action potential propagation and are mediated by alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA)/kainate receptors.

Phasic and delayed release at axon-OPC synapses depends on P/Q- and N-type voltage-gated Ca²⁺ channels

As activity-dependent Ca²⁺ entry into callosal axons plays a crucial role for vesicle fusion and neurotransmitter release at axon-OPC synapses [10], our next goal was to find out whether phasic and delayed release at these synapses depends on the same types of voltage-gated Ca²⁺ channels (VGCCs). We recorded callosal OPCs, stimulated axons electrically, and bath-applied a selective blocker of N-type VGCCs ω-conotoxin-GVIa (Ctx) (1 μM) or a selective blocker of P/Q-type VGCCs ω-agatoxin-IVA (Atx) (0.5 μM). Ctx reduced the initial mean amplitude of axon-glia EPSCs by 95% (control 1.93 ± 0.69 pA, Ctx 0.09 ± 0.30 pA, Fig 2A and 2D), while Atx reduced it by 88% (control 1.78 ± 0.44 pA, Atx 0.2 ± 0.05 pA, Fig 2B and 2E). Each toxin also caused a decrease of the average axon-glia current amplitude during the train, which was especially pronounced during the first 5–10 stimuli (Fig 2D and 2E). Changes in the probability of axon-glia responses upon application of each toxin followed a similar time course during the train as changes in the current amplitude (Fig 2G and 2H), while the response potency was largely unaffected (Fig 2J and 2K). Peak rate of delayed axon-glia EPSCs was also significantly reduced upon application of Ctx (6.3 ± 1.1 Hz before versus 3.3 ± 0.8 Hz after toxin application, Fig 2M), or Atx (13.2 ± 5.2 Hz before versus 4.5 ± 1.2 Hz after toxin application, Fig 2N). Simultaneous application of the two blockers strongly reduced but did not completely abolish phasic or delayed axon-glia EPSCs (S1 Fig).

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Fig 2. Reducing presynaptic Ca²⁺ level impairs short-term facilitation and delayed glutamate release at axon–oligodendrocyte precursor cell (OPC) synapses.

(A–C) Example traces showing average currents (including failures) in three OPCs in response to the 1st, 3rd, 5th, 10th, and 20th stimuli in the train: under control conditions (before drug application, black), after 15 min perfusion of (A) ω-Conotoxin-GVIA (Ctx, 1 μM), (B) ω-Agatoxin-IVA (Atx, 0.5 μM), (C) after 10 min perfusion of ethylene glycol tetraacetic acid acetoxymethyl ester (EGTA-AM) (20 μM) via the bath. (D–L) Average current amplitude including failures, response probability, and response potency upon each stimulus of the train during control conditions and after perfusion of Ctx (n = 5 cells), Atx (n = 5 cells), and EGTA-AM (n = 6 cells). Each point represents mean ± SEM. Paired t test. (S2 Table, S3 Table and S4 Table). *p < 0.05, **0.001 < p < 0.01. (M–O) Peak rate of delayed currents in OPCs during control conditions and after perfusion of (M) Ctx, (N) Atx, (O) EGTA-AM. Each grey point represents an average peak rate in an individual experiment. Each point in color represents the mean peak rate within the experimental group, i.e., control, Ctx, Atx, and EGTA-AM. Paired t test (S3 Data). Throughout the figure: concentration of a given drug is the same. The shown traces during drug application are taken from the time period when the drug effect on the amplitude of the evoked currents or rate of delayed currents was on the steady state. The numerical data used in D–O are included in S4 Data.

https://doi.org/10.1371/journal.pbio.2001993.g002

Taken together, these findings indicate that both phasic and delayed glutamate release at axon-OPC synapses in the corpus callosum is mainly mediated by N- and P/Q-types of VGCCs. However, a small component of release may be mediated by other subtypes of axonal Ca²⁺ channels or may be Ca²⁺ independent.

Accumulation of free Ca²⁺ in callosal axons underlies activity-dependent facilitation at axon-OPC synapses during the train and delayed release after the train

Accumulation of presynaptic Ca²⁺ plays an important role in triggering activity-dependent enhancement of synaptic efficacy at neuronal synapses [24]. Hence, after determining the source of Ca²⁺ mediating glutamate release at axon-OPC synapses, we tested whether accumulation of residual presynaptic Ca²⁺ underlies the facilitation of release during and after the train. We loaded callosal axons with ethylene glycol tetraacetic acid acetoxymethyl ester (EGTA-AM) (20 μM), a membrane-permeant high-affinity Ca²⁺ buffer with a slow Ca²⁺ binding kinetic [25], and studied the effects of EGTA-AM on phasic and delayed release. EGTA-AM did not significantly change either the mean amplitude of the first axon-glia EPSC in the train or its probability (Fig 2C and 2I), indicating that the basal transmission at axon-glia synapses relies on Ca²⁺ microdomains rather than on free residual axonal Ca²⁺ and that the distance between Ca²⁺ entry site and Ca²⁺ sensor is small [10]. In contrast, short-term synaptic enhancement during the train was completely abolished in the presence of EGTA-AM (Fig 2F). This effect was of presynaptic origin because EGTA-AM caused significant increase in the number of failures after each stimulus in the train, while the response potency remained unaffected (Fig 2I and 2L). EGTA-AM also caused significant reduction of the peak rate of delayed axon-glia EPSCs (4.8 ± 1.3 Hz before versus 0.9 ± 0.2 Hz after EGTA-AM application, Fig 2O).

Thus, repetitive stimulation of callosal axons triggers the rise of residual intra-axonal Ca²⁺level, which mediates facilitation of glutamate release at axon-OPC synapses during the train as well as delayed release after the train.

Activity-dependent changes at axon-OPC synapses during the train differ depending on the stimulation paradigm

Having demonstrated that OPCs detect phasic and delayed glutamate release at synapses with neurons, we next investigated how these two components of release depend on the stimulation paradigm and whether OPCs distinguish differences in release. We used 6 stimulation paradigms: 2 pulses at 25 Hz; 5 pulses at 5, 25, and 100 Hz; and 20 pulses at 25 and 100 Hz and first focused on release during the train. Upon stimulation of callosal axons with 5 pulses at 5, 25, or 100 Hz, the average amplitude of phasic axon-glia EPSCs showed progressive facilitation, which reached its maximum at third–fifth pulse, but the facilitation rate did not depend on the stimulation paradigm: 2.88 ± 0.8 for 5 pulses at 5 Hz (n = 5), 3.75 ± 0.9 for 5 pulses at 25 Hz (n = 11), and 2.67 ± 0.7 for 5 pulses at 100 Hz (n = 19) (Fig 3A). For all three paradigms, the facilitation was of presynaptic origin (Fig 3B and 3C). If longer trains (20 pulses) were applied, the time course and the amount of synaptic facilitation were determined by the stimulation paradigm: facilitation was larger and more sustained upon stimulation at 25 Hz (n = 6) versus 100 Hz (n = 14) (Fig 3D–3F). As the response potency remained constant during the trains (Fig 3H), differences in synaptic enhancement between the two stimulation paradigms can be explained by the distinct changes in the probability of glutamate release (Fig 3G).

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Fig 3. Short-term facilitation and desynchronization of glutamate release at axon-glia synapses during the train depend on the stimulation paradigm.

(A–B) Normalized average current amplitude including failures and response probability and (C) absolute response potency during minimal stimulation train of 5 pulses at 5 Hz (n = 5 cells), 25 Hz (n = 11 cells), or 100 Hz (n = 19 cells). Each point represents mean ± SEM. The values for each stimulus were compared between the different paradigms with one-way ANOVA. No statistically significant differences were found (S5 Data). (D, E) Example traces showing average currents (including failures) in two oligodendrocyte precursor cells (OPCs) upon minimal stimulation with 20 pulses at (D) 100 Hz and (E) 25 Hz. Fifty sweeps were averaged for each paradigm. Stimulation artifacts are truncated for clarity. Dotted lines are drawn at the level corresponding to the peak amplitude of the first response within the train. (F, G) Normalized average current amplitude including failures and response probability and (H) absolute response potency during minimal stimulation train of 20 pulses at 25 Hz (n = 6 cells) or at 100 Hz (n = 14 cells). Independent sample t test (S5 Table). (I, J) Average variance of onset latency of the phasic responses during the stimulation train of 20 pulses at (I) 100 Hz (n = 14 cells) or (J) 25 Hz (n = 6 cells). Paired t test was used to compare the value for each stimulus to the value at the first stimulus (S6 Table). (K) Schematic drawing explaining two approaches for generating average current waveforms used to study desynchronization of release during the train. The results of this analysis are shown in panels (L–O). Bottom row (“aligned”): events are aligned on the time of their onset (red dot) and then averaged. Top row (“not aligned”): average event is generated from the raw traces, which are by default aligned on the stimulation time (green dot) but not on their onset. If desynchronization of release occurs during the train while properties of individual events do not change, it is expected that (1) kinetics of the average nonaligned event is slower at the end of the train than at the beginning, and (2) kinetics of the average aligned event does not change during the train. (L, N) Average rise time and (M, O) average decay time constant of aligned and nonaligned responses (excluding failures) after each stimulus within the train of 20 pulses at (L, M) 100 Hz or (N, O) 25 Hz. Average aligned and average nonaligned events for each stimulus were generated as explained in (K). The same cells were used as in (F–J). Independent sample t test was used to compare the values from nonaligned and aligned averages for each stimulus (S7 Table). *p < 0.05; **0.001 < p < 0.01. The numerical data used in A–C, F–J, and L–O are included in S6 Data.

https://doi.org/10.1371/journal.pbio.2001993.g003

Repetitive axonal stimulation leads to buildup of presynaptic Ca²⁺ at axon-OPC synapses (Fig 2), which, in analogy to neuronal synapses, may contribute not only to short-term plasticity but also to activity-dependent desynchronization of phasic release [26–28]. We therefore investigated whether activity-dependent desynchronization of glutamate release occurs at axon-OPC synapses and how it differs depending on the stimulation paradigm. We found that the variance of the onset latency of phasic axon-glia EPSCs increased by 80% over the time course of the 100 Hz but not 25 Hz stimulation train (Fig 3I and 3J), pointing to desynchronization of release. To substantiate this finding, for each stimulus of the train we took the same set of phasic axon-glia EPSCs and averaged them either as they have been recorded (“nonaligned”) or after aligning them on the event onset (“aligned”) (Fig 3K). In case of 100 Hz stimulation, the kinetics of the average EPSC generated from the nonaligned events was slower compared to the aligned events, and this difference was more pronounced at the end than at the beginning of the train (n = 14, Fig 3L and 3M), designating broadening of the release timing during the train. A similar tendency was observed for 25 Hz stimulation, but the differences were not statistically significant (n = 6, Fig 3N and 3O).

Thus, activity-dependent synaptic enhancement and desynchronization of release at axon-OPC synapses differ depending on the stimulation paradigm, and OPCs discriminate the differences.

Delayed glutamate release at axon-OPC synapses depends on the stimulation paradigm

We next investigated how different stimulation paradigms affect delayed release at axon-OPC synapses. If we stimulated the axons at 25 Hz, the peak rate of delayed axon-glia EPSCs was higher when longer trains were applied: 0.8 ± 0.2, 1.8 ± 0.4, and 6.1 ± 1.9 Hz after 2, 5, and 20 pulses, respectively (Fig 4A). A similar result was obtained when we stimulated the axons at 100 Hz and compared 5 versus 20 pulses (Fig 4D & 4G). If we kept the number of pulses in the train constant and varied their frequency, the peak rate of delayed events was larger for higher stimulation frequencies (Fig 4D and 4G), although there was no difference between the stimulation at 25 versus 100 Hz. The rate of delayed events decayed monoexponentially to the basal rate of spontaneous axon-glia EPSCs, which was 0.31 ± 0.05 Hz (Fig 4B, 4E and 4H). For the same stimulation frequency, the decay time constant was larger for longer than for shorter trains (Fig 4B and 4C). In contrast, when the number of pulses was the constant parameter, the dependence of the decay time constant on the stimulation frequency was less pronounced (Fig 4E, 4F, 4H and 4I). The mean amplitude, rise, and decay time of the delayed axon-glia EPSCs were comparable in all cells, indicating that these parameters are not influenced by the type of prior axonal activity (not shown).

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Fig 4. The rate and the time course of delayed glutamate release at axon-oligodendrocyte precursor cell (OPC) synapses depend on the stimulation paradigm.

(A) Average peak rate of the delayed events after the stimulation with 2 pulses (n = 6 cells), 5 pulses (n = 6 cells), or 20 pulses (n = 8 cells) at 25 Hz. The box “Spont.” shows the frequency of the spontaneous events recorded before each stimulation train. One-way ANOVA (S7 Data). (B) Average rate of delayed events after the stimulation with 2, 5, or 25 pulses at 25 Hz. Solid lines indicate monoexponential fits to the events rate. The same cells used as in (A). (C) Average decay time constants of the delayed events rate after the stimulation with 2, 5, or 20 pulses at 25 Hz. The same cells used as in (A). One-way ANOVA (S7 Data). (D) Data and statistical comparisons are as in (A) but for the stimulation paradigms of 5 pulses at 5 Hz (n = 7 cells), 25 Hz (n = 7 cells), and 100 Hz (n = 6 cells). One-way ANOVA (S7 Data). (E) Data as in (B) but for the stimulation paradigms of 5 pulses at 5, 25, and 100 Hz. The same cells used as in (D). (F) Data and statistical comparisons are as in (C) but for the stimulation paradigms of 5 pulses at 5, 25, and 100 Hz. The same cells used as in (D). One-way ANOVA (S7 Data). (G) Data and statistical comparisons are as in (A) but for the stimulation paradigms of 20 pulses at 25 Hz (n = 8 cells) and 100 Hz (n = 13 cells). One-way ANOVA (S7 Data). (H) Data as in (B) but for the stimulation paradigms of 20 pulses at 25 and 100 Hz. The same cells used as in (G). (I) Data and statistical comparisons are as in (C) but for the stimulation paradigms of 20 pulses at 25 and 100 Hz. The same cells used as in (G). One-way ANOVA (S7 Data). Box and whisker plots: the bottom and top of each box represent 25th and 75th percentiles of the data, respectively, while whiskers represent 10th and 90th percentiles. The midline represents the median. The numerical data used in A, C, D, F, G, and I are included in S8 Data.

https://doi.org/10.1371/journal.pbio.2001993.g004

Thus, the delayed glutamate release differs depending on the number of stimuli in the train and the stimulation frequency, and OPCs can distinguish these differences.

Temporal profile of synaptic charge transfer at axon-OPC synapses differs depending on the stimulation paradigm

When studying facilitation of transmitter release at axon-OPC synapses, we have so far considered only changes in the amount of facilitation from stimulus to stimulus in the train (Fig 3A and 3F) but not in real time. To fill this gap, we analyzed synaptic charge transfer through AMPA/kainate receptors in OPCs upon axonal stimulation (during and after the train) and plotted it versus real time. We found that the temporal profile of charge transfer, i.e., its distribution over time, was determined by the stimulation paradigm (Fig 5A) and was entirely different even for those paradigms in which the amount of facilitation and the total synaptic charge transfer were similar (Figs 3A & 5A). For instance, although the same amount of charge was transferred during stimulation with 5 pulses at 5 Hz and at 25 Hz (Fig 5B) and the amount of facilitation was comparable (Fig 3A), in the former case, the transfer of synaptic charge oscillated at 5 Hz and was completed after 810 ms (Fig 5A, blue trace), while in the latter case, the transfer of charge oscillated at 25 Hz and was completed already after 170 ms (Fig 5A, red trace). Similar differences were also observed when we compared stimulation with 20 pulses at 25 Hz (Fig 5A, second red trace) versus 100 Hz (Fig 5A, black trace).

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Fig 5. The time course and the amount of synaptic charge transfer through alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors at axon–oligodendrocyte precursor cell (OPC) synapses depend on the stimulation paradigm.

(A) Average synaptic charge transfer upon stimulation of callosal axons with trains of four different frequencies and durations plotted versus real time. Each color (blue, red, dark red, or black) represents mean charge transfer from n = 5 cells (5 pulses at 5 Hz), n = 5 cells (5 pulses at 25 Hz), n = 5 cells (20 pulses at 25 Hz), and n = 10 cells (20 pulses at 100 Hz). Charge transfer is shown in 5-ms bins. Note that plotting charge transfer versus real time allows observing not only the amount of synaptic facilitation but also its distribution over time, and it differs dramatically depending on the stimulation paradigm (Fig 3A & 3D). (B) Total average synaptic charge transfer during the stimulation trains of different frequencies and durations. Both phasic and asynchronous charge transfer during the trains are considered for these bar graphs. The same cells used as in (A). One-way ANOVA (S9 Data). (C) Total synaptic charge transferred by the delayed currents occurring after the stimulation trains of different frequencies and durations. The same cells used as in (A). One-way ANOVA (S9 Data). (D) Percentage contribution of synaptic charge transferred during (phasic + asynchronous charge) and after (delayed charge) the stimulation trains of different frequencies and durations. White numbers on the bars indicate the proportion of charge transferred during the train. Box and whisker plots: the bottom and top of each box represent 25th and 75th percentiles of the data, respectively, while whiskers represent 10th and 90th percentiles. The midline represents the median. The numerical data used in B–C are included in S10 Data.

https://doi.org/10.1371/journal.pbio.2001993.g005

When comparing the relative contribution of charge transfer during and after the stimulation train, we found that, independently of the stimulation paradigm, 85%–98% of charge transfer occurred during the train (Fig 5D, Fig 5B & 5C). At four tested paradigms, the delayed glutamate release contributed only 2%–3% of total charge transfer. The largest contribution of 14% was taking place upon stimulation of axons with 20 pulses at 100 Hz (Fig 5D). This is similar to the situation at some neuronal synapses but not at the others [29].

In adult animals, phasic and delayed glutamate release at axon-OPC synapses also depends on the stimulation paradigm

Repetitive axonal stimulation (20 pulses at 25, 100, or 300 Hz) triggered phasic and delayed glutamate release at axon-OPC synapses also in adult (P50–53) mice (Fig 6). In contrast to juvenile mice, we observed little/no difference in presynaptic facilitation of phasic axon-glia EPSCs between 25 and 100 Hz stimulation paradigms; however, the amount of facilitation was lower for 300 Hz stimulation (Fig 6A–6D). (Note that the temporal profile of charge transfer is expected to differ between all three paradigms [Fig 5]). The incidence of delayed EPSCs was transiently elevated after cessation of stimulation, similar as it occurred in the juvenile mice. The delayed events occurred over a slightly longer time window and their peak rate was higher for 25 and 100 Hz stimulation versus 300 Hz trains (Fig 6E–6H). Hence, changes in glutamate release at axon-OPC synapses are determined by the stimulation paradigm also in adult mice, but the effect of a given paradigm may differ from that in the juvenile animals.

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Fig 6. Activity-dependent changes at axon-oligodendrocyte precursor cell (OPC) synapses are determined by the stimulation paradigm also in the adult (P50–53) mice.

(A–C) Normalized average (A) current amplitude including failures, (B) probability of responses, and (C) absolute potency of responses upon each stimulus of the minimal stimulation train of 20 pulses at 25 Hz (n = 4 cells), 100 Hz (n = 6 cells), or 300 Hz (n = 5 cells). One-way ANOVA (S11 Data). 25 Hz versus 300 Hz stimulation: *p < 0.05, **0.001 < p < 0.01. 100 Hz versus 300 Hz stimulation: #p < 0.05. 25 Hz versus 100 Hz stimulation: ^p < 0.05 (D) Example traces showing average currents (including failures) in 3 OPCs in response to the 1st, 5th, 10th, and 20th stimulus of the minimal stimulation train of 20 pulses at 25 Hz, 100 Hz, and 300 Hz. At least 15 trials were averaged to generate each example trace. (E) Original example traces showing delayed currents in callosal OPC (holding potential [V_h] = −80 mV) occurring after the stimulation train of 20 pulses at 25 Hz. (F) Average rate of delayed currents occurring after the stimulation train of 20 pulses at 25 Hz, 100 Hz, or 300 Hz. Solid lines indicate monoexponential fits to the events rate. (G) Average decay time constants of the delayed event rate after the stimulation train with 20 pulses at 25 Hz, 100 Hz, or 300 Hz. One-way ANOVA (S11 Data). (H) Average peak rate of the delayed events after the stimulation with 20 pulses at 25 Hz, 100 Hz, or 300 Hz. The box “Spont.” shows the frequency of the spontaneous events recorded before each stimulation train. One-way ANOVA (S11 Data). Box and whisker plots: the bottom and top of each box represent 25th and 75th percentiles of the data, respectively, while whiskers represent 10th and 90th percentiles. The midline represents the median. The numerical data used in A–C and G–H are included in S12 Data.

https://doi.org/10.1371/journal.pbio.2001993.g006

Differentiation and proliferation of OPCs in vivo depend on the pattern of transient changes in axonal activity

Having demonstrated that response of OPCs to repetitive axonal stimulation in brain slices differs depending on the stimulation pattern, we were wondering whether also in vivo OPCs respond to different stimulation paradigms in a distinct way. We implanted an electrode array into the corpus callosum of adult mice and transiently changed the activity of callosal axons in freely behaving animals by stimulating the corpus callosum at 5 Hz, 25 Hz, or 300 Hz (Fig 7A and 7B). Importantly, none of the stimulation paradigms triggered an unusual behavior or seizures in any of the animals. We then assessed changes in proliferation and differentiation of OPCs 7 days after the stimulation (Fig 7C–7F).

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Fig 7. Transient stimulation of callosal axons in vivo at 5 Hz but not at 25 Hz or 300 Hz promotes differentiation of oligodendrocyte precursor cells (OPCs) into oligodendrocytes (OLs).

(A) Scheme describing experimental design for studying effects of callosal stimulation in vivo on proliferation and differentiation of OPCs. (B) Scheme showing sagittal section of mouse brain and the position of an electrode array used for electrical stimulation of the corpus callosum. (C) Schematic drawings of the investigated cell types. For cell counting, OPCs were identified as NG2⁺CC1^- cells, premyelinating OLs (pre-OLs) as NG2⁺CC1⁺ cells, and myelinating OLs as NG2^- cells expressing CC1 in their soma. Within the OL lineage, 5-ethynyl-2´-deoxyuridine (EdU) only labels proliferating OPCs. However, the progeny of an EdU⁺ OPC will be EdU⁺. (D) Coronal sections of the corpus callosum. Maximum intensity projection (from 14 successive confocal planes) showing triple channel immunofluorescent labelling with DAPI (top left, blue), NG2 (top right, green), CC1 (bottom left, red), and the overlay of 3 channels (bottom right). White dashed line denotes the middle region of the corpus callosum used for cell counting. White arrow indicates midline of the brain. Scale bars: 100 μm. (E) As in (D), but higher magnification example of one NG2⁺CC1^- OPC (arrow) and three NG2^-CC1⁺ OLs (arrowheads). Maximum intensity projection was generated from a z-stack of 3 successive confocal planes. Note that some processes of an NG2⁺ OPC are clearly visible. Scale bars: 10 μm. (F) As in (E), but an example of one NG2⁺CC1⁺ pre-OL (arrow). Note that the expression level of NG2 is weaker than in OPCs, and the expression level of CC1 is weaker than in OLs (Fig 7E). Scale bars: 10 μm. (G–I) Average density of (G) OPCs, (H) pre-OLs, and (I) OLs in corpus callosum upon electrical stimulation of callosal axons at 5 Hz (n = 5 mice, total 13 slices), 25 Hz (n = 5 mice, total 16 slices), or 300 Hz (n = 5 mice, total 17 slices) versus sham-stimulated controls (n = 7 mice, total 25 slices). Note that differentiation rate was significantly increased by 5 Hz but not by 25 Hz or 300 Hz stimulation (H). Nested ANOVA and post hoc Tukey were used for statistical analysis (S13 Data). Box and whisker plots: the bottom and top of each box represent 25th and 75th percentiles of the data, respectively, while whiskers represent 10th and 90th percentiles. The midline represents the median. The numerical data used in G–I are included in S14 Data.

https://doi.org/10.1371/journal.pbio.2001993.g007

Stimulation of axons at 5 Hz and to a lesser extent at 25 Hz, but not at 300 Hz, resulted in substantial increase in the density of premyelinating OLs (pre-OLs): 1,551 ± 380 cells/mm³ in sham-treated animals versus 3,625 ± 911 cells/mm³ upon 5 Hz stimulation versus 2,488 ± 421 cells/mm³ upon 25 Hz stimulation versus 1,230 ± 387 cells/mm³ upon 300 Hz stimulation (Fig 7H). At the same time, none of the 3 stimulation paradigms significantly altered the density of OPCs or mature OLs (Fig 7G and 7I). To explore whether corresponding alterations might have occurred at earlier time points after the stimulation, we have also counted cells 3 days after the stimulation. We found, however, that the changes took the same path at that time point: stimulation triggered differentiation of OPCs, and 25 Hz stimulation was more effective with this respect than 300 Hz stimulation (S2 Fig). Hence, at this earlier time point, we restricted our analysis to only two stimulation paradigms. The density of OPCs 3 days after the stimulation was reduced compared to sham-treated animals (S2 Fig); this was most likely the consequence of their differentiation rather than cell death, as we have not observed caspase 3 (Casp-3)–positive OPCs in the corpus callosum (see below).

We next studied the effect of three different stimulation paradigms on proliferation of OPCs (Fig 8A). Stimulation of callosal axons at 25 Hz or 300 Hz triggered a statistically significant increase in the density of proliferating cells in the corpus callosum, while 5 Hz stimulation appeared less effective (Fig 8B). Furthermore, a clear trend was observed for 25 Hz stimulation being more efficient with this respect than 300 Hz: we counted 3,260 ± 839 cells/mm³ in the sham-treated group, 7,651 ± 930 cells/mm³ (130% increase) upon 25 Hz stimulation, and 5,652 ± 854 cells/mm³ (70% increase) upon 300 Hz stimulation. The ratio of 5-ethynyl-2´-deoxyuridine (EdU⁺) oligodendroglial cells within the total population of cycling cells in the corpus callosum was also enhanced after the stimulation (Fig 8C). The major cause of this enhancement was the rise in the number of EdU⁺ myelinating OLs (Fig 8D). Interestingly, upon stimulation of axons at 5 Hz, we observed a clear trend to enhanced numbers of pre-OLs versus sham-treated animals, versus the group of animals stimulated at 25 Hz and also at 300 Hz (Fig 8D). Hence, pre-OLs may also contribute to the higher ratio of EdU⁺ oligodendroglial cells within the total population of cycling cells after electrical stimulation.

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Fig 8. Callosal stimulation in vivo enhances proliferation rate of oligodendrocyte precursor cells (OPCs) and promotes differentiation of newly born OPCs into oligodendrocytes (OLs).

(A) Coronal sections of corpus callosum. Maximum intensity projection image (from 14 successive confocal planes) showing fluorescent 5-ethynyl-2´-deoxyuridine (EdU) labelling in corpus callosum. White dashed line marks the region of interest used for cell counting. Top panel: overview. Scale bar: 100 μm. Bottom panels: higher magnification of the area marked by the white square at the top panel; left, EdU (false color look-up table (LUT) “Green Fire Blue”); right, overlay of EdU and DAPI (blue) channels. Arrowheads indicate EdU⁺ cells. Scale bar: 10 μm. (B) Density of EdU⁺ cells in corpus callosum. (C) Proportion of OL lineage cells (OPCs + pre-OLs + OLs) labelled with EdU, within the total population of EdU⁺ cells. (D) Proportion of EdU⁺ OPCs (green), premyelinating OLs (pre-OLs) (red-green) and OLs (red) within the total population of EdU⁺ cells. (E) Single layer confocal image showing quadruple channel fluorescent labelling with DAPI (blue), NG2 (green), CC1 (red), and EdU (false color LUT “Green Fire Blue”), and the overlay of four channels (top right). The arrowhead indicates an EdU⁺ OPC. Scale bars: 10 μm. The box plot shows proportion of EdU⁺ OPCs within the total OPCs population. (F) Single layer confocal image showing quadruple-channel fluorescent labelling with DAPI (blue), NG2 (green), CC1 (red), and EdU (false color LUT “Green Fire Blue”), and the overlay of four channels (top right). The arrowhead points to an EdU⁺ OL. Scale bar: 10 μm. The box plot shows proportion of EdU⁺ OLs within the total OLs population. Throughout this figure, the same mice and slices were analyzed as in Fig 7. Nested ANOVA and post hoc Tukey test were used for statistical analysis (S15 Data). Box and whisker plots: the bottom and top of each box represent 25th and 75th percentiles of the data, respectively, while whiskers represent 10th and 90th percentiles. The midline represents the median. The numerical data used in B–F are included in S14 Data.

https://doi.org/10.1371/journal.pbio.2001993.g008

We next evaluated the fraction of EdU⁺ cells within the population of OPCs and within the population of OLs, separately. Stimulation of axons at 25 Hz or 300 Hz caused a statistically significant increase in the proportion of proliferating (EdU⁺) OPCs within the OPC population, while 5 Hz stimulation was less effective (Fig 8E). Furthermore, all stimulation paradigms promoted differentiation of the newly born (EdU⁺) OPCs into OLs. This was reflected in the higher numbers of the EdU⁺ OLs within the total population of OLs in stimulated versus sham-treated mice (Fig 8F). The 5 Hz (248% increase) and the 25 Hz stimulation paradigms (230% increase) were slightly more effective with this respect than 300 Hz stimulation (160% increase).

Our findings so far suggest that, depending on the paradigm, electrical stimulation promotes proliferation and/or differentiation of OPCs in the corpus callosum. These effects may be mediated by direct axon-OPC contacts and/or by diffusible substances, e.g., neurotransmitters, growth factors, etc. Alternatively, however, the primary effect of callosal stimulation may be the death of OPCs and/or OLs; proliferation and differentiation may then occur as a consequence of this effect. To test for this, we performed staining of brain slices for a cell death marker cleaved Casp-3, as it has been done previously (Gibson et al. 2014) [5]. We chose to stain for Casp-3 three days after the stimulation because in mouse models of diseases, Casp-3 is detectable 24 hours after the onset and stays for at least 7 days [30]. Similar to the study of Gibson et al., we have not observed any Casp-3–expressing cells in the middle part of the corpus callosum (which we used for counting oligodendroglial cells) in any of the stimulated animals (independently of the stimulation paradigm) and also not in sham-treated mice: n = 3 animals for 25 Hz (not shown); n = 3 animals for 300 Hz (S3 Fig); n = 2 sham-treated animals (S3 Fig). At the site of electrode implantation in the cortex, some Casp-3–positive cells were present (S3 Fig). Hence, our stimulation paradigms are most likely too mild to trigger cell death in the corpus callosum, and the observed effects of stimulation on proliferation and/or differentiation of OPCs are unlikely to be triggered by cell death.

Taken together, increased density of pre-OLs upon 5 Hz stimulation and increased ratio of EdU⁺ OPCs upon 25 Hz and 300 Hz stimulation suggest that some frequencies (5 Hz in our case) are more efficient at triggering differentiation, while other frequencies (25 Hz and 300 Hz in our case) are more efficient at triggering proliferation.

Microglia was not activated in the stimulated corpus callosum

To check if callosal stimulation triggered activation of microglia, we sacrificed the animals 3 days after the stimulation and performed cluster of differentiation 68 (CD68) staining. The time point of 3 days was chosen because several independent studies on brain trauma have demonstrated that activated microglia are observed 12 hours after trauma and stay in the brain tissue for up to 2–4 weeks [31–33]. We could not find CD68-positive cells in the middle part of the corpus callosum (which we used for counting oligodendroglial cells) in either stimulated or sham-treated animals (S4 Fig; n numbers are as for Casp-3 above). At the same time, we observed microglia activation at the site of electrodes implantation (close to the cortical surface) and along the track of the electrodes reaching down to the external capsule/corpus callosum (S4 Fig). Importantly, the distance between the microglia activation site and the nearest area used for cell counting was at least 500–600 μm. Hence, even if microglia release factors which could influence oligodendroglial behavior, the concentration of these factors at the cell-counting site is expected to be significantly diminished by diffusion. Therefore, in our study, microglia activation most likely had no/little effects on the proliferation and differentiation of OPCs.

Ethics statement

All experiments were performed in accordance with the guidelines of the Animal Care and Use Committee at the University of Tübingen. All experimental protocols were approved by the Regierungspräsidium Tübingen. All efforts were made to minimize the suffering of the animals. For surgery, the animals were anesthetized with a mixture of isoflurane and oxygen (3%–5% v/v), and metacam (1 mg/kg body weight) was injected subcutaneously in order to prevent pain suffering of the animals. Before decapitation, the animals were anesthetized with a mixture of isoflurane and oxygen (3% v/v).

Animals

NG2DsRedBAC transgenic mice [11] were used in all experiments. Breeding pairs were originally obtained from The Jackson Laboratory (stock 008241) and bred in house. Mice were kept in 12–12 hours of light-dark cycle; food and water were available ad libitum.

Slice preparation

Coronal brain slices containing corpus callosum were prepared from 19–22- or 50–53-day-old mice of both sexes. Mice were anesthetized with a mixture of isoflurane and oxygen (3% v/v) and decapitated. The brain was dissected in ice-cold N-methyl-D-glucamine (NMDG)-based solution containing (in mM): 135 NMDG, 1 KCl, 1.2 KH₂PO₄, 20 choline bicarbonate, 10 glucose, 1.5 MgCl₂, and 0.5 CaCl₂ (pH 7.4, 310 mOsm), gassed with carbogen (95% O₂, 5% CO₂). 270-μm-thick coronal brain slices were cut in the same solution using Leica VT 1200S vibratome. The slices were transferred to the 32°C Haas-type interface incubation chamber and perfused with Ringer solution containing (in mM): 124 NaCl, 3 KCl, 1.25 NaH₂PO₄*H₂O, 2 MgCl₂, 2 CaCl₂, 26 NaHCO₃, 10 glucose; 300 mOsm/kg; 7.4 pH; gassed with carbogen. The chamber was gradually cooled down to room temperature.

Electrophysiology

At least 1 hour after the preparation, individual slices were transferred to the submerged recording chamber mounted on a stage of an upright microscope (FN-1, Nikon, Japan) equipped with infrared differential interference contrast (IR-DIC) filters and a fluorescence light source. The slices were kept at room temperature and superfused continuously (about 2 ml/min) with gassed Ringer solution. OPCs were selected for recordings based on their red fluorescence and were distinguished from pericytes based on their morphology. Patch pipettes were pulled from borosilicate glass capillaries (Science Products, Germany) on a vertical puller (Model PC10, Narishige, Japan). Pipettes had resistance of 5–7 MOhms when filled with internal solution containing (in mM): 125 K-gluconate, 2 Na₂ATP, 2 MgCl₂, 0.5 EGTA, 10 HEPES, 20 KCl, 3 NaCl; 280–290 mOsm/kg, titrated to pH 7.3 with KOH. Cells were voltage clamped at the holding potential V_h = −80 mV with an EPC-8 amplifier (HEKA, Germany) and V_h was corrected for a −13 mV liquid junction potential before seal formation. Liquid junction potential was calculated using the software JPCalc for Windows (Peter H. Barry, Sydney, Australia). Series resistance was not compensated. After establishing the whole-cell configuration, 10 depolarizing voltage steps (increment +10 mV) were applied to each cell from V_h = −80 mV, and corresponding current responses were recorded in order to verify that the selected cell was an OPC [21]. Evoked synaptic currents were elicited with isolated pulse stimulator (A-M Systems, Model 2100, Science Products, Germany) using a monopolar glass electrode (resistance 5–6 MΩ) filled with Ringer solution and placed at 50–250 μm from the recorded cell. Single or paired (40 ms interpulse interval) biphasic rectangular pulses of 100–250 μs duration were applied every 15–30 s. Trains of stimuli were applied each 30 s. Minimal stimulation was performed as described by us previously [10] in brain slices where the corpus callosum was isolated from the cortex by 4 cuts: 2 parallel and 2 perpendicular to the orientation of callosal axons. The rationale for doing this was to exclude the possibility that delayed axon-glia currents are triggered by polysynaptic activation of callosally projecting neurons or result from dissimilar conduction velocities in individual callosal fibers. When recording stimulated and spontaneous synaptic currents, we applied a voltage step of −10 mV at the beginning of each sweep in order to monitor the dynamic of the series resistance. Whole-cell currents in response to voltage steps were low-pass filtered at 10 kHz and digitized with a sampling frequency of 20 kHz (ITC-18, HEKA Instruments Inc, USA). All synaptic currents were low-pass filtered at 1 kHz and digitized with a sampling frequency of 10 kHz. Data acquisition was performed using Recording Artist (written by Rick Gerkin, Arizona State University, USA) running under Igor Pro 6.3 (WaveMetrics, Lake Oswego, USA). All recordings of evoked or spontaneous synaptic currents were performed in the presence of NMDA-receptor antagonist (RS)-3-(2-Carboxypiperazin-4-yl)-propyl-1-phosphonic acid (CPP, 10 μM, Tocris) and GABA_A-receptor antagonist (RS)-3-(2-Carboxypiperazin-4-yl)-propyl-1-phosphonic acid (gabazine, 5 μM, Sigma). Miniature synaptic currents were recorded in the presence of 10 μM CPP, 5 μM gabazine, and 0.5–1 μM tetrodotoxin citrate (TTX, Abcam). To verify that synaptic currents are mediated by ionotropic glutamate receptors, we used 6-Cyano-7-nitroquinoxaline-2,3-dione (CNQX, 10 μM, Abcam). To test the role of calcium and VGCCs for synaptic currents in OPCs, we used ethylene glycol tetraacetic acid acetoxymethyl ester (EGTA-AM, 20 μM, Invitrogen), N-type calcium channels blocker Ctx (1 μM, Alomone Labs), or P/Q-type calcium channels blocker Atx (0.5 μM, Alomone Labs). Before applying Ctx or Atx, Albumin Fraction V (BSA, 0.1%, Carl Roth) was perfused via the bath for at least 25 minutes. All drugs were dissolved in Ringer solution and applied via the bath. All patch-clamp recordings were performed at room temperature using Ringer solution unless indicated differently.

Analysis of electrophysiology data

Only those recordings were considered for the analysis in which the offset drift by the end of the experiment was smaller than ±5 mV and the change of the series resistance was <30% of the original value. The series resistance was between 20 and 40 MΩ.

To study stimulated and spontaneous synaptic currents in OPCs, 30–100 sweeps of 5 s length were analyzed per cell. No additional digital filtering was applied to the recorded sweeps before the analysis. Each sweep was cut into 2 pieces: one containing the events during the train and the second containing the delayed events after the train. The detection and the subsequent analysis of the events during and after the train were performed separately.

For the pharmacology experiments, at least 50 sweeps were recorded before the drug perfusion. EGTA-AM effect was analyzed after at least 10 minutes of drug perfusion. Ctx or Atx effects were analyzed after at least 15 minutes of drug perfusion.

Surgery, in vivo stimulation, and EdU application

Adult male mice (6–8 weeks old) from the NG2DsRedBAC transgenic mouse line, but not expressing the transgene, were anesthetized with a mixture of isoflurane and oxygen (3%–5% v/v) and fixed in the stereotaxic frame (Stoelting, USA). The depth of the anaesthesia was verified by testing the reaction of the mouse to a toe pinch. Bepanthen (Bayer) was applied to the eyes in order to protect them from dehydration. Metacam (1 mg/kg body weight, Boehringer Ingelheim) was injected subcutaneously in order to prevent pain suffering of the animals. The body temperature was monitored using a rectal thermometer and kept at 36°C using the heating pad. The skin above the skull was disinfected, a small cut was made, and Xylocaine (2%, Astra Zeneca) was applied locally. The surface of the skull was cleaned with bone scraper and hydrogen peroxide (3%). Two windows were opened in the skull using a dental drill (NSK Mio drill): on the right side, a small longitudinal window was opened for the implantation of an electrode array used for stimulation, and on the left side, a larger window was opened for the recording of field potentials. An array of 5 electrodes was implanted into the right side of the corpus callosum using the following coordinates: posterior −1.06, lateral 0.8, and ventral 1.3 mm from Bregma and was fixed with dental cement (IvoclarVivadent). The electrode array was self-made: glass-coated platinum wires (Thomas Recording, Germany) were sharpened (outer diameter: 80 μm, platinum diameter: 25 μm, impedance was about 500 kΩ), and 5 of them were arranged parallel to each other. The distance between the tips was 250 μm. To verify that action potentials propagate through the middle part of the corpus callosum used for subsequent counting of cells, we exposed the surface of the cortex contralateral to the implanted electrode array and attached the recoding ball electrode to it. We then applied low frequency stimulation to the array and tested whether we were able to record field potentials at the contralateral side. This was possible in each mouse. In some animals, we applied TTX (1 μM) or CNQX (10 μM) in saline onto the surface of the cortex to verify that the field potential was dependent on the action potentials and ionotropic glutamate receptor activation. Each drug inhibited the field potential. These animals were not used for further experiments.

After the surgery, mice recovered from anesthesia fast and were returned to their home cages. One week (“7 days group”) or 11 days (“3 days group”) after surgery, mice were briefly anesthetized to connect the stimulator. This way, all mice were sacrificed 14 days after the surgery, and therefore any possible long-term impact of the surgical procedure was expected to be similar in all animals. Each mouse was stimulated in its home cage, where it was allowed to move freely. Each mouse underwent one of the following types of stimulation: (a) sham, i.e., the mouse was connected to the stimulator but the system was switched off, (b) 20 pulses at 5 Hz were applied every 5 minutes 36 times, (c) 20 pulses at 25 Hz were applied every 5 minutes 36 times, or (d) 20 pulses at 300 Hz were applied every 5 minutes 36 times. Every mouse underwent only 1 stimulation session, which lasted 3 hours, i.e., 36 stimulations at 5-min intervals. Video monitoring was performed for the majority of animals during the whole stimulation session. If, in rare cases, the electrodes got disconnected from the mouse, the mouse was excluded from the subsequent analysis. After the end of the stimulation session, each mouse was taken back to its original housing room. From that moment on and during the following 48 hours, the drinking water of each mouse contained 0.2 mg/ml 5-ethynyl-2’-deoxyuridine (EdU, Thermo Fisher) [59].

Immunohistochemistry

One week or 3 days after the stimulation session, mice were sacrificed, the brain was removed, and 300-μm-thick coronal slices were cut using the Leica VT 1200S vibratome. The cutting was performed in a solution of the following composition (in mM): 87 NaCl, 2.5 KCl, 1.25 NaH₂PO4*H2O, 7 MgCl₂, 0.5 CaCl₂, 25 NaHCO₃, 25 glucose, and 75 sucrose. The slices were fixed overnight at 4°C in 4% paraformaldehyde which was dissolved in 10 mM phosphate-saline buffer (PBS). Then, the 300-μm-thick slices were washed, embedded into agar, and resectioned in PBS to 30-μm-thick slices using Microtome (HM 650V, Thermo Fischer Scientific). All stainings were performed on 30-μm free-floating slices placed into multiwell plates. For EdU visualization, we followed the protocol recommended by Thermo Fisher Scientific. For antigen retrieval, we incubated the slices in 10 mM citric acid (pH = 6.0) at 37°C. After washing, we applied blocking solution containing 0.1 M Tris-buffer saline (TBS), 3% Albumin Fraction V, and 0.2% Triton-X for 1 hour at 37°C. Primary antibody was incubated overnight in 0.1 M TBS (pH = 7.6) containing 0.2% Triton-X. The following primary antibody was used: rabbit or guinea pig anti-NG2 (1:500, gift from Bill Stallcup, Burnham Institute, La Jolla, USA), mouse anti-APC (1:50, Ab-7, CC-Calbiochem), rabbit anti-Cleaved-Casp-3 (1:500, Cell Signalling Technology), rat anti-CD68 (1:100, Serotec). Detection was performed using the following secondary antibody: goat anti-rabbit Alexa Fluor 488 (1:1,000, Invitrogen), goat anti–guinea pig Alexa Fluor 633 (1:1,000, Invitrogen), goat anti-mouse Alexa Fluor 555 (1:500, Invitrogen), goat anti-rat Alexa Fluor 488 (1:1,000, Invitrogen). Secondary antibody was applied for 3 hours at 37°C. For counterstaining of the nuclei, we used Diamidino-2-phenylindole dihydrochloride (DAPI, 0.2 μg, Sigma).

Image acquisition

Laser scanning microscope (LSM) 710 system (Zeiss, Germany) was used for image acquisition. Images containing corpus callosum were acquired and saved as z-stacks with a 16-bit pixel depth. Each z-stack was 11–21 μm thick and consisted of 12–22 z-slices, and the z-step was 1 μm. Each layer of a z-stack was acquired as a multiple tile scan (vertical x horizontal: 2 x 5 images) in which each tile was 512 x 512 pixels in size. Pixel size was 0.42 x 0.42 μm. Each tile scan represented a triple-channel fluorescence image, in which channels were acquired sequentially in ZEN software using 40x oil-immersion objective (NA = 1.3). The following excitation laser lines and emission-detection ranges were used: for DAPI excitation 405 nm, emission 414–490 nm; for Alexa-488 excitation 488 nm, emission 497–556 nm; for Alexa-555 excitation 561 nm, emission 569–633 nm; for Alexa-633 excitation 633 nm, emission 650–740 nm. The beam splitters for each dye matched the excitation laser lines. The pinhole was set to 1 airy unit for each channel. Laser power, detector gain, and offset were adjusted such that in the final scan (averages from 2), we had good signal-to-background-noise ratio. All images presented in the figures are representative examples and have not been additionally processed with any software after acquisition. For presentation purposes, the color of the far red channel used for visualizing EdU signals has been changed in the figures to “Green Fire Blue” LUT in ImageJ in order to distinguish it from the red channel.

Cell counting

We counted cells in unprocessed z-stack images using ImageJ (NIH, USA). The corpus callosum was identified based on the maximum intensity projection of the CC1 and DAPI staining. To define the lateral borders of the region of interest used for counting, in each coronal slice we determined the midline and outlined the area of the corpus callosum approximately 500 μm to each side from the midline, i.e., the region of interest was approximately 1 mm broad. In dorso-ventral direction, we considered the full thickness of the corpus callosum, which was easily identifiable based on the visualization of the cortex and the hippocampus. Anterior and posterior borders for counting were approximately from −0.5 mm to 1.5 mm from Bregma.

For the cells located close to the predefined border of the region of interest, only those cells were included into the analysis whose nucleus touched the border. OPCs were identified as NG2⁺CC1^- cells, pre-OLs as NG2⁺CC1⁺ cells, and myelinating OLs as NG2^- cells expressing CC1 in their soma. The following groups of animals were used: (1) sham-treated mice (n = 7), (2) mice stimulated at 25 Hz and sacrificed 3 days after the stimulation (n = 3), (3) mice stimulated at 300 Hz and sacrificed 3 days after the stimulation (n = 3), (4) mice stimulated at 5 Hz and sacrificed 7 days after the stimulation (n = 5), (5) mice stimulated at 25 Hz and sacrificed 7 days after the stimulation (n = 5), and (6) mice stimulated at 300 Hz and sacrificed 7 days after the stimulation (n = 5). Cells were counted in 2–4 slices from each mouse: in total, 25 slices from sham-treated; 11 slices from 25 Hz, 3 days; 10 slices from 300 Hz, 3 days; 13 slices from 5 Hz, 7 days; 16 slices from 25 Hz, 7 days; and 17 slices from 300 Hz, 7 days groups. After counting, the data was normalized to the volume in which the counting was performed.

Statistics

All data acquisition was randomized (animals for in vivo stimulation, cells during patch-clamp experiments). Throughout the study, we made all efforts to avoid pseudoreplications, both when performing experiments in slices and in vivo. For patch-clamp recordings, we used in total 32 mice of the age P19–22 (Figs 1–5) and 13 mice of the age P50–53 (Fig 6). If several OPCs were recorded on the same day from the same animal, different slices were used and different stimulation paradigms were applied. In some cases, multiple stimulation paradigms were applied while recording from the same cell. During the data analysis, we verified again that all recordings for a given stimulation paradigm always stem from a different mouse. In this sense, the “n” numbers given in the text reflect simultaneously the number of cells and the number of animals. For in vivo experiments, cells were counted in 2–4 slices per animal; the counts within one animal were averaged and treated as one data point for the ANOVA.

Statistical analysis was performed using SPSS, including tests for homoscedasticity and normal distribution. Where 2 groups were compared, independent t tests were used. Pharmacology data was tested with paired t test (control versus drug conditions). When more than 2 data sets were compared, we used one-way ANOVA with post hoc Bonferroni or Dunnett's test. For evaluating the in vivo experiments, we used nested-ANOVA design (stimulation group: fixed factor; mice: random factor; cell density per slice: dependent variable, “nested” in mice) with post hoc Tukey test.

In all graphs, statistically significant differences are indicated as follows: *p < 0.05, **0.001 < p < 0.01, ***p < 0.001, or the p values are written on the graph.

Box and whisker plots: the bottom and top of each box represent 25th and 75th percentiles of the data, respectively, while whiskers represent 10th and 90th percentiles. The midline represents the median. For graphs other than box plots, each point represents the mean ± SEM unless indicated differently in the corresponding figure legend. Data in text is also represented as mean ± SEM.