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Adult-born neurons modify excitatory synaptic transmission to existing neurons

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Cite as: eLife 2017;6:e19886 doi: 10.7554/eLife.19886

Abstract

Adult-born neurons are continually produced in the dentate gyrus but it is unclear whether synaptic integration of new neurons affects the pre-existing circuit. Here we investigated how manipulating neurogenesis in adult mice alters excitatory synaptic transmission to mature dentate neurons. Enhancing neurogenesis by conditional deletion of the pro-apoptotic gene Bax in stem cells reduced excitatory postsynaptic currents (EPSCs) and spine density in mature neurons, whereas genetic ablation of neurogenesis increased EPSCs in mature neurons. Unexpectedly, we found that Bax deletion in developing and mature dentate neurons increased EPSCs and prevented neurogenesis-induced synaptic suppression. Together these results show that neurogenesis modifies synaptic transmission to mature neurons in a manner consistent with a redistribution of pre-existing synapses to newly integrating neurons and that a non-apoptotic function of the Bax signaling pathway contributes to ongoing synaptic refinement within the dentate circuit.

https://doi.org/10.7554/eLife.19886.001

eLife digest

Neurogenesis, the creation of new brain cells called neurons, occurs primarily before birth. However, a region of the brain called the dentate gyrus, which is involved in memory, continues to produce new neurons throughout life. Recent studies suggest that adding neurons to the dentate gyrus helps the brain to distinguish between similar sights, sounds and smells. This in turn makes it easier to encode similar experiences as distinct memories.

The brain’s outer layer, called the cortex, processes information from our senses and sends it, along with information about our location in space, to the dentate gyrus. By combining this sensory and spatial information, the dentate gyrus is able to generate a unique memory of an experience. But how does neurogenesis affect this process? As the dentate gyrus accumulates more neurons, the number of neurons in the cortex remains unchanged. Do some cortical neurons transfer their connections – called synapses – to the new neurons? Or does the brain generate additional synapses to accommodate the newborn cells?

Adlaf et al. set out to answer this question by genetically modifying mice to alter the number of new neurons that could form in the dentate gyrus. Increasing the number of newborn neurons reduced the number of synapses between the cortex and the mature neurons in the dentate gyrus. Conversely, killing off newborn neurons had the opposite effect, increasing the strength of the synaptic connections to older cells. This suggests that new synapses are not formed to accommodate new neurons, but rather that there is a redistribution of synapses between old and new neurons in the dentate gyrus.

Further work is required to determine how this redistribution of synapses contributes to how the dentate gyrus works. Does redistributing synapses disrupt existing memories? And how do these findings relate to the effects of exercise – does this natural way of increasing neurogenesis increase the overall number of synapses in the system, potentially creating enough connections for both new and old neurons?

https://doi.org/10.7554/eLife.19886.002

Introduction

Continual neurogenesis in the adult dentate gyrus (DG) produces new granule cells (GCs) that integrate into the hippocampal circuit by establishing synapses with existing neurons (Espósito et al., 2005; Ge et al., 2006; Toni et al., 2008; Dieni et al., 2013). During a transient period of maturation, new GCs exhibit intrinsic and synaptic properties distinct from mature GCs, potentially underlying the contribution of neurogenesis to memory encoding (Schmidt-Hieber et al., 2004; Ge et al., 2007; Aimone et al., 2011; Marín-Burgin et al., 2012; Dieni et al., 2013; Brunner et al., 2014; Dieni et al., 2016). Yet computational models also suggest that remodeling of pre-existing circuits by continual neurogenesis can degrade established memories (Weisz and Argibay, 2012; Chambers et al., 2004), a possibility that has recently gained experimental support from the observation that neurogenesis facilitates ‘forgetting’ (Akers et al., 2014; Epp et al., 2016). Circuit remodeling could occur by synaptic redistribution, wherein existing terminals that synapse onto mature GCs are appropriated by newly integrating GCs. This possibility is supported by anatomical evidence that immature dendritic spines transiently receive a high proportion of synapses from multiple-synapse boutons (Toni et al., 2007; Toni and Sultan, 2011). Furthermore, dramatically increasing the number of new neurons does not alter the density of spines and synapses in the molecular layer, suggesting a readjustment of synaptic connections (Kim et al., 2009). Yet whether synaptic integration of new GCs is accompanied by changes in synaptic function and structure of mature GCs is not known.

The number of integrating new GCs can be selectively altered by genetic manipulations targeted to adult stem cells that regulate the survival of progeny (Enikolopov et al., 2015). Adult-born neurons undergo a period of massive cell death during the first weeks after cell birth that is rescued by deletion of the pro-apoptotic protein Bax (Sun et al., 2004; Kim et al., 2009), and conditional Bax deletion in Nestin- expressing progenitors enhances the number of adult-born neurons without affecting other cell populations (Sahay et al., 2011; Ikrar et al., 2013). Similarly, inducible expression of the diphtheria toxin receptor in Nestin-expressing stem cells allows selective ablation of adult-born neurons (Arruda-Carvalho et al., 2011). These approaches have been used to identify contributions of adult born neurons in hippocampal-based behaviors, with the understanding that behavioral outcomes could either reflect unique functions of adult-born neurons themselves or homeostatic adaptions within the network (Singer et al., 2011). Physiological stimuli like exercise and environmental enrichment also enhance dentate neurogenesis, yet it is unclear whether genetically targeted manipulations of neurogenesis mimic the circuit function in the same manner as physiological stimuli.

To identify network adaptions resulting from synaptic integration of new GCs, here we tested how manipulating the number of adult-born GCs affects perforant path-evoked excitatory synaptic currents (EPSCs) in mature GCs. We measured synaptic transmission to pre-existing mature GCs in response to selective genetic manipulations of Nestin-expressing stem cells, using inducible Bax deletion to enhance, or diphtheria toxin-induced ablation to reduce, the number of new neurons. We also tested synaptic transmission to immature GCs and mature GCs with Bax deletion to investigate potential non-apoptotic functions of the Bax signaling pathway in synaptic function (Jiao and Li, 2011; Ertürk et al., 2014). Finally, we tested whether enhancing neurogenesis by a physiological stimulus likewise alters excitatory transmission to mature neurons. Our results show that selectively manipulating the number of immature GCs modifies synaptic function of mature GCs in a manner consistent with synaptic redistribution, with an inverse relationship between the number of new neurons and perforant-path evoked EPSCs. In contrast, enhancing neurogenesis via the non-selective paradigm of environmental enrichment generates a net increase in functional connectivity of mature neurons. Together these results demonstrate the capacity of mature GCs to alter synaptic function in response to genetic and experiential circuit manipulations.

Results

Enhancing immature neurons decreases EPSCs and spine density of mature neurons

We sought to test synaptic transmission to mature GCs after selectively enhancing the number of integrating new GCs by manipulating cell survival, given that most proliferating DG progenitors and newborn neurons undergo apoptosis (Sierra et al., 2010). Cell death of progenitors and new GCs requires the pro-apoptotic protein Bax, a member of the BCL-2 family of proteins in the intrinsic apoptotic pathway (Sun et al., 2004). Both germ line and conditional Bax deletion block cell death of adult-generated GCs without altering proliferation or the gross structural integrity of the DG (Sun et al., 2004; Kim et al., 2009; Sahay et al., 2011). As previously described (Sahay et al., 2011; Ikrar et al., 2013), we increased the population of adult-born GCs by crossing inducible Nestin-CreERT2 mice with a Bax conditional knockout mouse line to selectively block apoptotic cell death in proliferating cells and their progeny (Materials and methods; Figure 1—figure supplement 1A). Four-to-six weeks after tamoxifen-induced recombination at two months of age, we compared the number of new GCs and synaptic responses from pre-existing mature GCs in hippocampal slices from BaxKOimmature mice (referred to as BaxKOim) and controls (Figure 1A). We crossed some BaxKOim mice with a transgenic reporter line that labels early postmitotic GCs (Overstreet et al., 2004) to reveal a ~40% increase in the number of newborn GCs and overtly normal dentate structure (Figure 1B,C).

Figure 1 with 2 supplements see all
Increasing neurogenesis reduces EPSCs in mature GCs.

(A) The experimental timeline showing recording 4–6 weeks after tamoxifen (TMX)-induced Bax deletion in Nestin-expressing progenitors. (B) Confocal images of newborn neurons expressing eGFP in fixed sections (50 μm) from control and BaxKOim mice. (C) Stereological cell counts of eGFP+ newborn cells revealed neurogenesis was enhanced by 41% (control 16,881 ± 1422 cells, n = 4; BaxKOim 23,756 ± 2166 cells, n = 4; unpaired t-test p=0.038). (D) Schematic showing experimental paradigm, with simultaneous fEPSPs and whole-cell recordings of EPSCs from mature GCs. All experiments were performed in the presence of picrotoxin to block GABAA receptor-mediated currents. (E) Examples of fEPSPs (top) with the fiber volley (FV, top inserts) and EPSCs (bottom) in slices from control and BaxKOim mice. Synaptic responses were evoked by increasing intensity stimulation by a patch pipette placed in the middle molecular layer. fEPSPs and EPSCs were binned by FV amplitude. Stimulus artifacts are blanked for clarity. (F) The fEPSP versus FV plot illustrates the effectiveness of FV normalization, with fEPSP increasing linearly with axonal recruitment. There was no difference in fEPSPs in slices from BaxKOim and control mice (two-way ANOVA, 0.076). FVs are binned by 100 μV and each symbol denotes the mean and SEM of 10–38 responses from 15 control and 14 BaxKOim slices (with four responses in the largest 300–400 μV FV control bin). (G) Left, a decrease in synaptic strength to mature GCs was revealed by the EPSC plotted against FV amplitude (two-way ANOVA, Fgenotype (1,167)=54.41 p<0.0001; p<0.05 for all bins with Bonferroni post-tests). Right, the overall EPSC/FV ratio was reduced in BaxKOim slices (unpaired t-test, n = 86, 95).

https://doi.org/10.7554/eLife.19886.003

To assess excitatory transmission from entorhinal cortex across the population of GCs and onto individual mature GCs, we stimulated the medial perforant path while simultaneously recording field excitatory postsynaptic potentials (fEPSPs) and excitatory postsynaptic currents (EPSCs) from mature GCs (Figure 1D,E). All experiments were performed in the GABAA receptor antagonist picrotoxin to isolate glutamatergic synaptic responses. There was no difference in fiber volleys (FVs; a measure of axonal activation) or fEPSPs between slices from BaxKOim and control mice (Figure 1—figure supplement 2A) (Sahay et al., 2011), as well as no difference in fEPSPs when responses were binned by the FV to account for differences in the number of stimulated axons across slices (Figure 1F). We targeted mature GCs located near the mid or outer edge of the granule cell layer and confirmed their maturity by morphology and intrinsic membrane properties (Figure 1—figure supplement 2B,C). Interestingly, we found that mature GCs in BaxKOim mice exhibited smaller EPSCs than mature GCs in controls across all FV amplitudes (Figure 1G, left), and an overall lower EPSC/FV ratio (Figure 1G, right). There was no difference in the EPSC/FV ratio between mature GCs in Cre+ and Cre- controls, and the difference in EPSCs persisted when only Baxfl/fl genotypes were analyzed (Figure 1—figure supplement 1B,C). Thus mature GCs in BaxKOim slices had reduced excitatory transmission.

To assess the pre- or postsynaptic locus of reduced EPSCs in mature GCs from BaxKOim mice, we first compared the paired-pulse ratio (PPR), a measure of presynaptic release probability. There was no difference in the PPR of evoked EPSCs at an interstimulus interval of 100 ms (Figure 2A), implying that adult-born neurons do not regulate transmission to mature GCs by secreting a factor that alters the release probability. However, mature GCs in BaxKOim mice displayed a lower frequency of spontaneous EPSCs (sEPSCs) with no change in amplitude (Figure 2B), suggesting a reduction in the number of active synapses with no change in postsynaptic responsiveness. Furthermore, using Sr2+ to desynchronize evoked release in order to detect single site EPSCs (Bekkers and Clements, 1999; Rudolph et al., 2011; Williams et al., 2015), we found a reduction in the frequency but not the amplitude of desynchronized events (Figure 2C). Thus, enhanced numbers of newly generated neurons were associated with reduced excitatory synaptic transmission to mature GCs that appeared to be mediated by fewer functional synapses.

Fewer functional synapses on mature GCs in BaxKOim mice.

(A) The paired-pulse ratio of evoked EPSCs (100 ms ISI) was similar in BaxKOim and control mature GCs (unpaired t-test p=0.90; n = 15 controls,14 BaxKOim). (B) Spontaneous EPSCs in mature GCs from BaxKOim mice had lower frequency and similar amplitudes as sEPSCs in mature GCs from control mice (unpaired t-test p=0.027 and 0.79, respectively; n = 11 controls, 9 BaxKOim). (C) Asynchronous EPSCs were generated by desynchronizing synaptic release with 1 mM Ca2+ and 4 mM Sr2+. Uniquantal aEPSCs were detected following the synchronous EPSC. Left, 40 traces overlaid with examples of averaged aEPSCs. Middle, there was no difference in the average amplitude across genotypes but a reduction in the frequency of aEPSCs (1743 events in 8 GCs from controls, 1015 events in 9 GCs from BaxKOim; unpaired t-test p=0.51 and 0.007 respectively). There was no difference in the average rise time or decay of aEPSCs (not shown).

https://doi.org/10.7554/eLife.19886.006

To further examine the locus of change, we assessed the PPR of EPSCs in mature GCs across a range of interstimulus intervals (20–1000 ms). In this protocol, mature GCs in BaxKOim and control mice again exhibited similar passive and active properties (Figure 3—figure supplement 1). The PPR was mildly depressing (Petersen et al., 2013), with no difference in ratios between genotypes (Figure 3A), as previously reported using fEPSPs (Sahay et al., 2011). During the recordings, we filled GCs with biocytin for posthoc spine analysis, focusing on dendrite segments in the middle molecular layer where medial perforant path synapses are located (Figure 3B). Consistent with reduced evoked and sEPSCs, there was a robust reduction in the density of spines in mature GCs from BaxKOim mice compared to controls (Figure 3C). We classified spines by shape (mushroom, thin, stubby) to determine the percentage of each spine type in control and BaxKOim mice. There was a slight increase in the percentage of stubby spines in BaxKOim mice (Figure 3D), with no significant difference in the percentage of thin and mushroom spines. Together, these results support the functional data showing that increasing the number of newborn GCs decreases synaptic transmission to mature GCs by reducing the number of synapses.

Figure 3 with 1 supplement see all
Mature GCs in BaxKOim mice exhibit low spine density.

(A) There was no difference in the paired-pulse ratio of EPSCs in mature GCs from BaxKOim and control mice across a range of interstimulus intervals (2-way ANOVA, p=0.31, n = 8,12 mature GCs). (B) Examples of reconstructed mature GCs from the recordings in (A). Red boxes indicate regions used for spine analysis. (C) Left, example images of dendritic spines from mature GCs. Scale bar, 10 μm. Middle, the density of dendritic spines was lower in BaxKOim mice (14 ± 0.8 spines/10 μm, 936 total spines counted on 15 dendritic segments in two control mice; 10 ± 0.6 spines/10 μm, 676 total spines on 12 dendritic segments from 3 BaxKOim mice; p=0.0007 unpaired t-test). (D) Classifying spines as stubby, thin and mushroom revealed a significant increase in the percentage of stubby spines in mature GCs from BaxKOim mice (p=0.04 unpaired t-test) with no change in the percentage of thin spines (p=0.07 unpaired t-test) or mushroom spines (p=0.45 unpaired t-test).

https://doi.org/10.7554/eLife.19886.007

Ablation of immature neurons increases synaptic transmission to mature neurons

We next tested whether genetically ablating adult-generated neurons alters excitatory transmission to mature GCs. We crossed Nestin-CreERtm4 mice (Kuo et al., 2006a) to Cre-inducible diphtheria toxin receptor (iDTR) mice (Buch et al., 2005; Arruda-Carvalho et al., 2011). Six weeks after tamoxifen-induced recombination, DT injections were given to ablate immature adult-born GCs in Nestin-Cre+/iDTR+ offspring (termed Ablatedim mice; Figure 4A) with Cre- littermates used as controls. Ten days after injections, there was a 27% reduction in the number of Dcx-expressing immature cells in the dentate of Ablatedim mice (Figure 4B; 5601 ± 262, n = 2, compared to 7648 ± 332, n = 4, p=0.016), noting that re-population of Dcx-expressing cells in the period after DT injection can lead to an underestimation of ablation efficiency (Vukovic et al., 2013; Yun et al., 2016). Performing simultaneous field and whole-cell recordings from mature GCs in Ablatedim mice and controls at 1–2 weeks after DT injections suggested no change in total synapses, assayed by the FV and fEPSP slopes (Figure 4C,D, Figure 4—figure supplement 1A). We also assayed synaptic terminals by immunodetection of the vesicular glutamate transporter (vGlut1) in the molecular layer, and found no differences between controls and either Ablatedim or BaxKOim slices (Figure 4—figure supplement 1B). Furthermore, there was no change in the fEPSP normalized to the FV (Figure 4D). However, there was enhanced synaptic transmission to individual mature GCs, shown by larger EPSC amplitudes across FVs (Figure 4E) and an overall larger EPSC/FV ratio (2.2 ± 0.1 in control compared to 3.7 ± 0.4 in Ablatedim mice; n = 42, 47 respectively, p=0.001 unpaired t-test). The change in synaptic strength was not associated with any changes in the intrinsic properties of mature GCs (Figure 4—figure supplement 2A). These results suggest that reducing the number of immature GCs increases the strength of synaptic transmission to mature GCs, an effect that cannot be explained by altered inhibition as GABAA receptors were blocked in these experiments (Singer et al., 2011; Temprana et al., 2015; Drew et al., 2016). There was no difference in PPR, suggesting that release probability was unchanged (Figure 4—figure supplement 2B). We were unable to detect differences in the average frequency or amplitude of sEPSCs in mature GCs from Ablatedim mice (Figure 4—figure supplement 2C), making it unclear whether reduced EPSCs resulted from pre- or postsynaptic mechanisms. Since the frequency of spontaneous activity in GCs is low, the threshold for detecting differences in synaptic function using spontaneous activity may be higher than for evoked transmission with FV normalization, and it appears that neurogenesis was altered by a greater degree in BaxKOim mice compared to Ablatedim mice (~40% versus 25% change in new neuron number). However, we also cannot rule out the possibility that separate pools of synaptic vesicles contribute to differences between results obtained with evoked and spontaneous assays (reviewed in Kavalali, 2015).

Figure 4 with 2 supplements see all
Ablating neurogenesis increases synaptic transmission to mature GCs.

(A) Experimental timeline showing ablation of immature GCs that are <6 weeks of age. Recordings from mature GCs were done 1–2 weeks after ablation. (B) Confocal images of Dcx-expressing immature neurons in control and Ablatedim mice. (C) Example of fEPSPs (top) with fiber volleys (FV, top insets) and simultaneously recorded EPSCs from mature GCs (bottom) in control and Ablatedim mice. (D) There was no difference in the fEPSP slope versus FV between Ablatedim and control mice (two-way ANOVA p=0.879, each symbol represents 8–22 responses from 7 control and 7 Ablatedim mice; FVs were binned by 75 μV). (E) The EPSC amplitude plotted against FV was larger in mature GCs from Ablatedim mice compared to controls (two-way ANOVA, Fgenotype (1,91)=30.31 p<0.0001; ***p<0.001 Bonferonni post-test). There was an increase in the overall EPSC/FV ratio in mature GCs from Ablatedim mice (unpaired t-test, p=0.0008, n = 42, 47).

https://doi.org/10.7554/eLife.19886.009

In summary, manipulating the number of immature GCs was inversely associated with excitatory synaptic strength of mature GCs. These manipulations did not affect global measures of axonal activation, synaptic strength or presynaptic terminals, suggesting that changing the number of newly generated neurons did not alter the total number of afferent axons or synapses. The idea that global measures of basal synaptic transmission and release probability are independent of the number of dentate GCs is in agreement with prior results in the conditional BaxKO (Sahay et al., 2011) as well as the observation that perforant path synapse density is unaltered in germline BaxKO mice which exhibit dramatically enhanced numbers of dentate GCs (Kim et al., 2009). Together these results support the idea that synaptic integration of newborn GCs involves a redistribution of existing synapses from old to new cells (Tashiro et al., 2006; Toni et al., 2007; McAvoy et al., 2016).

Bax deletion enhances synaptic strength of immature neurons

One assumption inherent to this idea, however, is that synaptic integration of newborn neurons is unaffected by manipulating their number, such that the increase in new cell number is paralleled by an increase in the total number of new synapses. We therefore sought to confirm synaptic integration of BaxKO immature GCs by crossing BaxKOim and control mice with a tdTomato reporter line (Ai14) to target BaxKO and BaxWT immature GCs for recordings (Figure 5A). The input resistance is a measure of cell maturity (Overstreet-Wadiche and Westbrook, 2006; Dieni et al., 2013) and as expected, labeled immature GCs (six weeks post-tamoxifen) had higher input resistance than mature GCs, with no difference between genotypes (Figure 5B). This confirms that the immature GCs were at a similar stage of maturation and is consistent with the similar dendrite development reported in this model (Sahay et al., 2011). FVs and fEPSP slopes were the same between genotypes, replicating the results of Figure 1 and further suggesting a similar level of axonal activation and number of total synapses after conditional Bax deletion (Figure 5—figure supplement 1). Consistent with the low excitatory connectivity of immature GCs (Dieni et al., 2016), in control mice the EPSC/FV ratio of immature GCs (1.24 ± 0.07 n = 80) was lower than the EPSC/FV ratio in mature GCs (2.44 ± 0.16 n = 86, p<0.0001 unpaired t-test). But unexpectedly, simultaneously recorded fEPSPs and EPSCs revealed that EPSCs in BaxKO immature GCs were significantly larger than EPSCs in BaxWT immature GCs across FV bins, and the overall EPSC/FV ratio was greater (Figure 5C). Thus BaxKO immature GCs showed enhanced synaptic transmission compared to WT immature GCs. The PPR of EPSCs in immature GCs was similar between genotypes and there was not a significant difference in the frequency or amplitude of sEPSCs (Figure 5—figure supplement 2), again noting that the low frequency of spontaneous activity in immature GCs (Mongiat et al., 2009; Dieni et al., 2016) makes it difficult to interpret the lack of change in sEPSCs. These results confirm that BaxKO immature GCs acquired synapses during integration and, in fact, suggest Bax deletion promotes the synaptic integration of new GCs.

Figure 5 with 4 supplements see all
Bax deletion enhances EPSCs in adult born neurons.

(A) Whole cell recordings were made from immature GCs in control and BaxKOim slices at six weeks post-tamoxifen injection, using picrotoxin to isolate glutamatergic EPSCs. Simultaneous fEPSPs were recorded in the molecular layer as in Figure 1. (B) Immature GCs in control and BaxKOim tdT mice had a similar input resistance that was higher than mature GCs (n = 12, 12, 16, respectively; one-way ANOVA p=0.0004, *p<0.05,***p<0.0001 Bonferroni post hoc test). (C) Left, examples of fEPSPs (top) and EPSCs (bottom) recorded in immature GCs. Middle, an increase in synaptic transmission to immature BaxKO GCs was revealed by the EPSC plotted against fiber volley (two-way ANOVA, Fgenotype (1,143)=18.55 p<0.0001, n = 12 control tdT, 12 BaxKOim tdT; *p<0.05 with Bonferroni post-tests). Right, the EPSC/FV ratio for all stimulus intensities (control 1.24 ± 0.07, n = 80; BaxKOim1.59 ± 0.09, n = 75; unpaired t-test p=0.0029). (D) Schematic showing simultaneous recordings from adjacent tdT- (BaxWT) and tdT+. (BaxKO) GCs in slices from BaxKOim tdT mice at 16 weeks after tamoxifen. (E) Adult-generated BaxKO GCs had larger EPSCs than simultaneously recorded unlabeled mature GCs. EPSCs were normalized to the maximum amplitude of the unlabeled (BaxWT) GC in each slice (two-way ANOVA, Fgenotype (1,94)=11.59 p=0.001, n = 6 pairs), scale bars: 10 ms, 100 pA. Comparing raw EPSCs between pairs of unlabeled and tdT+ GCs across all stimulus intensities confirmed EPSCs were larger in tdT+ GCs (not shown, paired t-test, p<0.0013).

https://doi.org/10.7554/eLife.19886.012

To further test the role of Bax in excitatory transmission to postmitotic GCs, we compared synaptic activity of adult-born BaxKO and unlabeled GCs at 16 weeks after tamoxifen-induced recombination, well after excitatory synaptic integration is complete (Mongiat et al., 2009). We directly compared EPSCs using simultaneous recordings from neighboring BaxWT (tdT-) and BaxKO (tdT+ GCs; Figure 5D). In this paradigm, FV normalization is unnecessary because the number of stimulated axons is the same for both recorded cells. To compare across cell pairs with different numbers of stimulated fibers in each slice, we normalized EPSCs to each BaxWT GC. Consistent with a role of Bax suppressing synaptic depression, EPSCs in BaxKO GCs were larger than EPSCs in BaxWT GCs (Figure 5E). There was no difference in the mature intrinsic properties of BaxWT and BaxKO GCs, again showing that Bax deletion does not alter intrinsic cell properties (Figure 5—figure supplement 3). Thus, enhanced synaptic transmission in Bax deficient GCs persists when adult-born neurons are fully mature.

Bax deletion in mature neurons increases EPSCs and spine density

Our results show that Bax deletion increases excitatory synaptic integration of adult born GCs, consistent with growing evidence that the Bax/caspase signaling cascade has non-apoptotic functions in synaptic plasticity (Unsain and Barker, 2015). Prior work suggests that Bax activation is an intermediary step between NMDAR-Ca2+ influx and local activation of caspase-3, which in turn is necessary and sufficient for LTD and subsequent spine pruning (Li et al., 2010; Jiao and Li, 2011; Ertürk et al., 2014; Sheng and Ertürk, 2014). The high level of Bax mRNA throughout the adult dentate gyrus (Lein et al., 2007) raises the possibility that this pathway contributes to activity-dependent synaptic remodeling of mature GCs in addition to controlling the number of integrating new GCs via apoptosis. Given that synaptic strength may depend on Bax expression, we tested whether overall Bax levels are altered in BaxKOim mice. Western blot analysis revealed no difference in Bax protein levels in hippocampal lysates from BaxKOim and control mice, showing that deletion of Bax from a small percentage of GCs does not lead to widespread changes in Bax protein (Figure 5—figure supplement 4).

To further probe the synaptic function of Bax, we next tested whether enhanced synaptic strength persists in mature neurons when Bax is deleted from postmitotic GCs throughout development. We generated conditional BaxKO in postmitotic GCs (termed BaxKOmature) using POMC-Cre to direct recombination in dentate GCs throughout development (Gao et al., 2007; Figure 6—figure supplement 1). Expression of tdTomato (tdT)reporter revealed that most, but not all, NeuN-expressing GCs in the granule cell layer expressed Cre and that NeuN-lacking proliferating progenitors in the subgranular zone were Cre negative (Figure 6A), consistent with transient activity of the POMC promoter in early postmitotic GCs (Overstreet-Wadiche et al., 2006; Overstreet et al., 2004). We compared EPSCs in simultaneous recordings from neighboring tdT+ (BaxKO) and tdT- (BaxWT) mature GCs (Figure 6B), again normalizing EPSCs to each WT cell to compare EPSCs across cell pairs. EPSCs in BaxKO GCs were larger than EPSCs in BaxWT GCs across a range of stimulus intensities (Figure 6C). To confirm that the increase in EPSC amplitude resulted from Bax deletion, we repeated the experiment in POMC-Cre/BaxWT/tdT mice (Figure 6D,E). EPSCs were the same in neighboring tdT+ and tdT- mature GCs (Figure 6F), indicating that the difference shown in Figure 6C requires the Baxfl/fl genotype. Thus, Bax deletion from immature GCs decreases EPSCs in mature GCs via a non-cell autonomous mechanism (Figures 13), whereas here we show a cell-autonomous effect of Bax deletion that increases EPSCs in mature GCs (Figure 6). These counterintuitive results could occur if Bax deletion generates presynaptic actions that are most evident when Bax is deleted from a large population of GCs. We addressed potential presynaptic alterations in BaxKOmat mice by testing the Ca2+-dependence of synaptic transmission. However, we found no difference in presynaptic function as assessed by comparing EPSC amplitudes and PPRs across a range of extracellular Ca2+ concentrations (Figure 6—figure supplement 2).

Figure 6 with 2 supplements see all
Bax deletion increases EPSCs and spine density of mature GCs.

(A) Confocal image of fixed tissue from a BaxKOmat /tdTomato mouse showing tdT (red) and NeuN (blue). Note the larger fraction of tdT+ GCs compared to (D), consistent with enhanced survival of GCs that lack Bax (confirmed in Figure 7A). (B) Adjacent tdT- (BaxWT) and tdT+ (BaxKO) mature GCs were recorded simultaneously. (C) Examples of EPSCs in tdT- and tdT+ mature GCs to the same stimuli. EPSCs were normalized to the maximum EPSC of the unlabeled cell in each slice. EPSCs were larger in tdT+ GCs (two-way ANOVA, Fgenotype (1,198)=21.14 p<0.0001, n = 12 cell pairs). (D) Confocal image of fixed tissue from a BaxWT/POMC-Cre+/tdTomato mouse, in which both tdT+ and unlabeled GCs are BaxWT(red tdT, blue NeuN). (E) Adjacent tdT+ and unlabeled mature GCs were recorded simultaneously. (F) There was no difference in EPSCs between BaxWT tdT+ and unlabeled cells (two-way ANOVA p=1.0, n = 8 cell pairs), confirming the difference in panel C requires the Bax-/- genotype. (G) Posthoc dendrite reconstructions (top) revealed higher spine density in BaxKO GCs from (A) (10.5 ± 0.53 spines/10 μm in BaxWT compared with 17.60 ± 1.3 BaxKO, unpaired t-test p<0.0001) with no change in spine head diameter (unpaired t-test, p=0.7, n = 21 segments from 5 BaxWT, 18 segments from 9 BaxKO). Lower images illustrate spine analysis.

https://doi.org/10.7554/eLife.19886.017

Since Bax activation is necessary and sufficient to activate caspase-3, which acts as a mediator of activity-dependent hippocampal LTD and synaptic pruning (Li et al., 2010; Jiao and Li, 2011; Ertürk et al., 2014; Lo et al., 2015), we wondered whether enhanced synaptic transmission to BaxKO GCs resulted from a deficit in synaptic pruning. We analyzed dendritic spines in BaxKO and BaxWT GCs by filling cells with biocytin during recordings. Posthoc analysis revealed a significant increase in the density of spines in BaxKO mature GCs, with no change in head diameter (Figure 6G). Together these results show that loss of Bax in GCs generates a persistent enhancement of synaptic transmission consistent with a deficit in synaptic pruning.

Neurogenesis-induced loss of synaptic strength requires intact Bax signaling

Based on the above results, we predicted that neurogenesis-induced loss of synapses from mature GCs might require intact Bax signaling to allow synaptic pruning. We thus assayed neurogenesis-induced synapse loss from mature GCs in BaxKOmat mice, where most mature GCs lack Bax. First, we confirmed that BaxKO in newly postmitotic GCs increases the number of integrating new neurons by assessing neurogenesis using POMC-eGFP expression. Consistent with the later period of cell death that occurs in newly postmitotic GCs (Sierra et al., 2010), we found that the number of newborn integrating neurons was enhanced to a similar degree as observed in BaxKOim mice (Figure 7A). However, neurogenesis-induced suppression of synaptic transmission to mature GCs was absent, since the evoked EPSC was similar to controls across all stimulus intensities and the average EPSC/FV ratio was unchanged (Figure 7B). Similar to Ablatedim and BaxKOim mice, there was no difference in axonal activation or total synapse number, measured by the FV amplitude and fEPSP slope versus FV, respectively (Figure 7—figure supplement 1A). Intrinsic properties of mature GCs were the same in BaxKOmat and control mice, showing that Bax deletion does not affect these measures of cellular excitability (Figure 7—figure supplement 1B). There was also no difference in the PPR, sEPSC frequency or sEPSC amplitude between mature GCs in control and BaxKOmat mice (Figure 7—figure supplement 2). However, there was considerable variability in EPSC/FV ratios and sEPSC frequencies in mature GCs from BaxKOmat mice, potentially indicative of the heterogeneous population of BaxWT and BaxKO GCs (as in Figure 6A) with mixed susceptibility to neurogenesis-induced synapse impairment. Together, these results suggest that neurogenesis-induced loss of synaptic strength to mature GCs requires intact Bax signaling.

Figure 7 with 2 supplements see all
Neurogenesis-induced loss of synaptic transmission requires Bax in mature GCs.

(A) Confocal images of newborn neurons expressing eGFP in BaxKOmat mice. Stereological cell counts revealed neurogenesis was enhanced by ~48% (control 17,910 ± 900 cells, n = 7; BaxKOmat26,508 ± 2728 cells, n = 6; unpaired t-test), similar to enhanced neurogenesis in BaxKOim mice. (B) Left, examples of fEPSPs (top) and EPSCs in mature GCs (bottom) from control and BaxKOmat mice. Middle, there was no difference in EPSCs across FVs (two-way ANOVA p=0.990, n = 12 control, 19 BaxKOmat GCs) or in the EPSC/FV ratio (unpaired t-test p=0.387, n = 88 control, 129 BaxKOmat), although there was greater variability in the BaxKOmat group (CV = 52% vs. 43%) consistent with a mixed population of mature Bax-/- and Bax+/+ GCs (as shown in Figure 6).

https://doi.org/10.7554/eLife.19886.020

Environmental enrichment increases synaptic strength of mature neurons

Our experiments revealed that selective manipulations of adult-born neurons are sufficient to alter functional synaptic transmission to mature neurons, raising the question of whether enhancing neurogenesis by physiological stimuli likewise affects synaptic function of mature neurons. One long-established strategy to enhance neurogenesis is housing rodents with environmental enrichment (EE) that includes exploration of novel objects, social interactions, and running wheels. EE enhances both the number of newborn GCs and their synaptic integration (van Praag et al., 1999; Tashiro et al., 2007; Ambrogini et al., 2010; Chancey et al., 2013; Bergami et al., 2015), as well as altering structural plasticity in the dentate and other brain regions (Green and Greenough, 1986; Foster et al., 1996; Eadie et al., 2005; Foster and Dumas, 2001; Stranahan et al., 2007).

We enhanced neurogenesis by housing WT mice with EE (Figure 8A), a treatment reported to generate a 1.5–2-fold increase in the number of integrating new GCs (van Praag et al., 1999; Brown et al., 2003; Olson et al., 2006). We previously found that housing mice with running wheels alone for four weeks increases the number of POMC-eGFP labeled GCs to 146% of age-matched controls (Overstreet et al., 2004), suggesting that EE enhances neurogenesis to a similar or greater extent as observed in BaxKOim mice (Figure 1B,C). To assess the strength of excitatory transmission from entorhinal cortex across the population of GCs and onto individual mature GCs, we again stimulated the medial perforant path while simultaneously recording fEPSPs and EPSCs from mature GCs in GABAA receptor antagonists (Figure 8B). As previously reported (Green and Greenough, 1986; Foster et al., 1996), the fEPSP slope was enhanced in slices from EE mice with no difference in the FV, suggesting an increase in total synaptic strength with no change axonal excitability (Figure 8—figure supplement 1A,B). Indeed, normalizing the fEPSP slope to the FV to account for differences in the number of stimulated axons across slices revealed a significant increase in the fEPSP (Figure 8C). We targeted mature GCs located near the outer edge of the granule cell layer and confirmed their maturity by intrinsic membrane properties (Figure 8—figure supplement 1C). Consistent with the enhanced fEPSPs, EPSCs in mature GCs were larger in slices from mice housed in EE (Figure 8D), such that the overall EPSC/FV ratio was 2.6 ± 0.16 in EE compared to 1.6 ± 0.07 in control (n = 88, 58 respectively, p<0.0001 unpaired t-test). Enhanced synaptic strength after EE could result either from increased release probability, increased number of synapses or increase in the number of receptors per synapse. We found no difference in the PPR of evoked EPSCs, suggesting that release probability is unchanged (Figure 8E), as previously reported (Foster et al., 1996). However, the frequency of sEPSCs was increased with no change in sEPSC amplitude (Figure 8F), similar to the recently reported increase in miniature EPSCs in mature GCs after EE (Kajimoto et al., 2016). Together these results suggest that enhanced evoked EPSCs in mature GCs result from greater number of functional synapses, consistent with increased spine density in Golgi-stained (presumably mature) dentate GCs (Eadie et al., 2005; Stranahan et al., 2007). These results show that mature GCs exhibit experience-dependent synaptic enhancement that argues against the recently described restricted period for experience-dependent plasticity of dentate GCs (Bergami et al., 2015). However, these results cannot resolve whether integration of EE-induced newborn GCs affects synaptic function of mature neurons. Increased connectivity of mature neurons is likely a parallel phenomenon independent of neurogenesis, since similar increases in synaptic transmission and spine density occur in non-neurogenic regions (Rampon et al., 2000; Malik and Chattarji, 2012; Jung and Herms, 2014). Thus, the magnitude of increased connectivity of mature GCs could be reduced by neurogenesis-induced synaptic redistribution. Altogether, these results highlight the capacity of mature GCs to undergo changes in synaptic connectivity in response to both genetic and experiential circuit manipulations.

Figure 8 with 1 supplement see all
Environmental enrichment increases synaptic transmission to mature GCs.

(A) The experimental timeline showing recordings performed 4–6 weeks after EE. (B) Left, simultaneous fEPSPs and whole-cell recordings from mature GCs, as shown in Figure 1. Examples of fEPSPs (top) with FV (insert) and EPSCs in mature GCs (bottom) in slices from control and EE mice. (C) Slices from EE mice exhibited an increase in the fEPSP slope plotted against FV amplitude (two-way ANOVA, Fmanipulation (1,116)=9.59, p=0.0025, n = 11 control, 9 EE). FVs were binned by 75 μV. (D) Left, an increase in synaptic transmission to mature GCs was revealed by the EPSC plotted against FV (two-way ANOVA, Fmanipulation (1,150)=52.88, p<0.0001, n = 11 control, 9 EE). *p<0.01with Bonferroni post-tests. Right, the overall EPSC/FV ratio was enhanced by EE (unpaired t-test, p<0.0001). (E) The paired-pulse ratio of EPSCs (100 ms ISI) was similar in EE and control mice (p=0.181 unpaired t-test, n = 9 controls, 9 EE). (F) Spontaneous EPSCs in mature GCs from EE mice had higher frequency (p=0.0081 unpaired t-test) but similar amplitude as sEPSCs in mature GCs from control mice (p=0.46, n = 9 controls,10 EE).

https://doi.org/10.7554/eLife.19886.023

Quantitative estimate of synapse transfer between mature and immature neurons

Immature GCs make up a small percent of total GCs, and yet when neurogenesis was selectively manipulated the change in synaptic strength to mature GCs was unexpectedly robust. To determine whether the magnitude of altered transmission to mature GCs could be explained by a redistribution of existing synapses to integrating new GCs, we made a quantitative estimate of the proportion of mature synapses that would be transferred to new GCs over the time course of our experiments. We simulated the BaxKOim condition, since in this condition we quantified excitatory input to mature GCs and immature GCs, as well as the increase in new cells induced by Bax deletion. Other parameters were based on reported rates of neurogenesis (Chancey et al., 2013; Gil-Mohapel et al., 2013), cell death (Sierra et al., 2010) and excitatory synaptic integration (Dieni et al., 2013, 2016). The simulation is based on a static number of synapses that re-distribute to immature GCs according to their number and time-dependent synaptic integration (Figure 9—figure supplement 1). The simulation showed a steep increase in the proportion of synapses occupied by immature GCs in BaxKOim mice starting at the time point when immature GCs start to integrate into the network (Figure 9A, red line). The robust transfer of synapses resulted not only from the increased number of immature GCs, but also from the increased acquisition of immature synapses resulting from Bax deletion. The predicted reduction in mature synapse number (expressed as a %) at days 36–43 in the simulation was similar to the % change in mature EPSCs measured experimentally (Figure 9B). Despite the small proportion of immature GCs within the network (initially set at 5%), the continuous increase in cell number along with enhanced synaptic integration was compounded over time to attenuate synapses on pre-existing neurons to a degree that could account for the magnitude of reduced synaptic strength observed in the BaxKOim experiments.

Figure 9 with 1 supplement see all
Simulation of neurogenesis-induced synaptic redistribution.

(A) Distribution of synapses occupied by mature and immature GCs using quantitative synaptic transfer simulation (see Materials and methods). Lines indicate the percentage of synapses on mature and immature GCs across the duration of the BaxKOim experiment, with the total number of synapses held constant. (B) Experimentally measured %change in EPSCs in mature GCs (left axis) compared to the %change in mature synapse number predicted by the simulation at time points t = 36 through t = 43 (right axis). Experimental data is the mean mature GC EPSC amplitude in BaxKOim mice normalized to control from each FV bin shown in Figure 1G. (C) Graphic depiction of synaptic integration of adult born neurons showing that new GCs (green) gain EC synapses (orange) through two possible sources: (C1) New EC terminals may form to innervate new GCs. In this case, increasing neurogenesis would increase the total number of synapses over time but the synapses per individual mature GC would remain constant. (C2). Alternatively, new GCs may take over existing EC synapses from surrounding mature GCs. In this case, the total number of synapses would remain constant over time and the number of synapses per mature GC would decrease. The reduced synaptic input to mature GCs in BaxKOim mice coupled with the apparent lack of change in total synapses (Figures 13) supports the synaptic redistribution model.

https://doi.org/10.7554/eLife.19886.025

Discussion

Here we tested how manipulating the number of adult-born GCs affects excitatory synaptic transmission to mature GCs. We found that selectively manipulating adult-born neurons inversely correlated with synaptic strength of mature neurons with no detectable changes in global measures of synaptic transmission. We reasoned that there are two ways that integrating newborn GCs can acquire synapses; new GCs can form new synaptic connections with existing afferent axons or new GCs can take pre-existing synapses from neighboring mature GCs. If synaptic integration of developing GCs triggers formation of new presynaptic terminals, then neurogenesis will increase the total number of synapses within the DG network but will not affect the number of synapses per mature GC (Figure 9C1). In contrast, appropriation of existing synaptic terminals would cause mature GCs to lose synapses while the total number of synapses in the network remains constant (Figure 9C2). By comparing measures of total synapses (fEPSPs) and synapses per mature GC (EPSCs) after selectively altering neurogenesis, our results support the latter model wherein newborn GCs appropriate existing synapses and consequently modify synaptic input to mature GCs.

Enhancing neurogenesis reduces mature neuron synaptic transmission and spine density

Our results showing that increasing neurogenesis decreased synaptic transmission and spine density of mature GCs is consistent with the idea that immature neuron synaptic integration is a competitive process (Tashiro et al., 2006; Toni and Sultan, 2011; McAvoy et al., 2016). Anatomical analysis has suggested that multisynaptic boutons (MSBs) represent an intermediary structure in the transfer of functional synapses from mature to immature GCs (Toni et al., 2007; Toni and Sultan, 2011). Although we did not find evidence for alterations in the total number of functional synapses reflecting the presence of MSBs when neurogenesis was manipulated, shared transmission from MSBs may be functionally silent due to lack of AMPA receptors on new neurons (Wu et al., 1996; Chancey et al., 2013), or may be below the detection limits of field potential recordings. Furthermore, recent work suggests MSBs are a common feature of mature GCs and the complexity of MSB innervation increases with GC maturation (Bosch et al., 2015), so it is unclear how our functional results relate to prior anatomical studies. Nevertheless, our results unambiguously demonstrate that neurogenesis modifies synaptic transmission to existing mature GCs through a mechanism that involves reduced number of functional synapses. Unlike prior reports of alterations in DG excitability following selective manipulations of neurogenesis, we isolated excitatory synaptic transmission using GABAA receptor antagonists, thus our results cannot be attributed to differential recruitment of local inhibitory circuits by immature GCs (Singer et al., 2011; Massa et al., 2011; Ikrar et al., 2013; Temprana et al., 2015). In addition to such feedback inhibition, regulation of the density of mature GC excitatory synapses could potentially contribute to the counter-intuitive finding that the number of immature GCs is inversely related to the excitability of the mature network (Ikrar et al., 2013; Drew et al., 2016).

Our interpretation that integrating new GCs acquire synapses from mature GCs relies on the assumption that modulation of EPSCs reflects changes in synapse number. Several pieces of evidence support this assumption. First, to account for differences in the number of stimulated axons across slices, we normalized EPSCs in mature GCs to the simultaneously recorded fiber volley, a common approach used in synaptic plasticity studies. Thus, differences in EPSCs cannot result from systematic differences in the number of stimulated axons. Second, reduced evoked EPSCs were accompanied by reduced frequency of sEPSCs with no change in amplitude and no change in the PPR. These characteristics are widely accepted indicators of changes in synapse number. Third, strontium-evoked asynchronous EPSCs likewise supported the idea that small EPSCs in mature GCs from BaxKOim mice resulted from fewer active synapses rather than a postsynaptic change in sensitivity. We also found no difference in the Ca2+ sensitivity of EPSCs between BaxKOmat and control mice. This suggests that Bax deletion from the majority of GCs did not affect Ca2+ dependence of release processes, making it unlikely that a secreted factor acts presynaptically to alter release following Bax manipulation. Finally, we found that the density of mature GC spines was reduced after selective enhancement of neurogenesis. Our results are consistent with a model wherein newly generated GCs usurp pre-existing synapses from mature GCs, perhaps through an activity-dependent competitive process (Tashiro et al., 2006), yet we cannot rule out other non-competitive mechanisms by which newly generated cells affect the number of synapses on mature GCs. The recent observation that conditional suppression of spines on mature GCs enhances the integration of newborn GCs further supports the interactions between new and existing neurons (McAvoy et al., 2016).

A synaptic re-distribution model predicts that the addition of new neurons does not alter the total number of synapses within the circuit (Figure 9C). We used fEPSPs as a primary measure of total synapses, and presumably the fEPSP does not change despite the loss of EPSCs in mature GCs due to the additional contribution of synapses on immature neurons. Although we did not detect differences in fEPSPs (or vGluT expression), it is important to note that fEPSPs may not be particularly sensitive to synaptic density and will also be affected by intrinsic excitability. We did not detect any differences in the intrinsic excitability of mature GCs in our genetic models, but it is expected that the higher intrinsic excitability of immature neurons would enable a greater contribution to fEPSPs compared to mature GCs (for a given number of active AMPAR-containing synapses). However, newborn GCs have a high fraction of silent synapses that may limit their contribution to fEPSPs (Chancey et al., 2013). Most importantly, our interpretation of synaptic redistribution is not affected if the immature GC contribution to the fEPSP does not fully compensate for the loss of transmission to mature neurons (that is, if the fEPSP was reduced in BaxKOim mice). Only an increase in the fEPSP in BaxKOim mice would lend support a synaptic addition model. Even so, changes in fEPSPs are somewhat tangential to our novel finding that EPSCs in mature GCs are altered by selective manipulations of newborn GCs.

Non-apoptotic role of the Bax signaling pathway in synaptic function

Our results indicate that Bax is required in mature GCs for neurogenesis-induced loss of transmission, suggesting that a change in the Bax signaling pathway is involved in spine loss from mature GCs. The contribution of Bax in our experiments is thus complex. We show that mature GCs exhibit a non-cell autonomous effect of Bax deletion from adult-born GCs (Figures 13, decreased EPSCs) that is opposite to the cell-autonomous effect of Bax deletion in both cell types (Figures 56, increased EPSCs). Remarkably, the cell autonomous function is required for the non-cell autonomous effect (Figure 7). This complexity, however, makes sense when we consider the role of Bax signaling in both cell death and synapse pruning. We propose that the non-cell autonomous effect results from enhanced neurogenesis (supported by the observation that ablation of neurogenesis produced the opposite outcome, Figure 4), whereas the cell autonomous effect results from a contribution of the Bax pathway in synaptic depression and spine pruning.

Although the Bax signaling pathway is best known in the context of programmed cell death, it also has a non-apoptotic role in synaptic plasticity that is mediated by downstream caspases, the same family of cysteine proteases that initiate cell apoptosis (Sheng and Ertürk, 2014). Caspases mediate dendritic remodeling during neural development (Kuo et al., 2006b; Williams et al., 2006; Riccomagno and Kolodkin, 2015), and more recent work shows that caspase-3 activation is necessary and sufficient for NMDAR-mediated AMPA receptor internalization and LTD at hippocampal synapses (Li et al., 2010Jiao and Li, 2011). LTD is associated with spine shrinkage and is typically considered a herald of synapse pruning (Oh et al., 2013; Wiegert and Oertner, 2013), thus it appears that pathways mediating cellular destruction also contribute to synaptic destruction (Sheng and Ertürk, 2014). Indeed, local induction of caspase-3 activity in dendrites triggers spine elimination whereas caspase-3 KO mice exhibit increased GC spine density (Ertürk et al., 2014; Lo et al., 2015), similar to our results of increased spine density in BaxKO GCs. Our findings that Bax deletion enhanced synaptic strength and spine density while blocking neurogenesis-induced loss of mature GC synaptic strength are consistent with the idea that on-going synaptic refinement controls the strength of excitatory transmission and that continual neurogenesis promotes a competitive environment for redistribution of synapses (McAvoy et al., 2016).

Implications for dentate function

These results have potential implications for understanding the role of neurogenesis and plasticity in DG function. First, both enhancing neurogenesis and blocking output from mature GCs improves performance on the same context discrimination task (Sahay et al., 2011; Nakashiba et al., 2012), suggesting that neurogenesis could contribute to DG function by modifying mature GC activity. Synaptic depression and subsequent pruning are activity-dependent processes that typically require NMDA receptor activation (Shipton and Paulsen, 2014). Hence, re-distribution of active terminals away from mature GCs could transiently sparsify population activity, if new GCs initially have insufficient excitatory connectivity to allow recruitment (Dieni et al., 2016). Second, since eliminating Bax in progenitors leads to greater innervation as well as greater survival of neural progeny, enhancing neurogenesis by blocking the apoptotic pathway likely promotes competition to a greater extent than other methods of increasing neurogenesis. This could have implications for understanding the potential role of enhanced neurogenesis using Bax deletion in behavioral outcomes assessing pattern separation, stress resilience and forgetting (Sahay et al., 2011; Akers et al., 2014; Hill et al., 2015). Finally, our results showing that deletion of Bax signaling in postmitotic GCs enhances synaptic transmission is consistent with increased activation of DG neurons observed in caspase-3-/- mice, which also show behavioral deficits in attending to relevant stimuli (Lo et al., 2015). Together, we speculate that synaptic redistribution between immature and mature GCs may contribute to activity-dependent synaptic remodeling that allows salient stimuli to receive precedence in DG encoding and may also contribute to circuit remodeling that degrades established memories (Weisz and Argibay, 2012; Chambers et al., 2004Akers et al., 2014; Epp et al., 2016).

Materials and methods

Transgenic mice

All animal procedures followed the Guide for the Care and Use of Laboratory Animals, U.S. Public Health Service, and were approved by the University of Alabama at Birmingham Institutional Animal Care and Use Committee (protocol# 8674 and 10134). Mice of either gender were maintained on a 12 hr light/dark cycle with ad libitum access to food and water.

BaxKOimmature mice were generated by crossing heterozygous loxP-flanked Bax mice (Jackson #006329, the Bak1 null allele was bred out) with Nestin-CreERt2 mice (Jackson #016261). The offspring were crossed with each other to produce Nestin-Cre+ or -/Baxfl/fl, Baxfl/+, or Bax+/+ animals (see Figure 1—figure supplement 1). Eight week-old mice were injected with tamoxifen (TMX, from a 20 mg/ml stock dissolved in sunflower seed oil, 75 mg/kg for three consecutive days) to induce recombination and experiments were done 4–6 weeks post-injection. Control Nestin-Cre- or Bax+/+ genotypes received TMX injections with the same protocol. For knockdown of neurogenesis, homozygous iDTR mice (Jackson #007900) were crossed with male Nestin-CreERtm4 mice provided by Chay Kuo (Kuo et al., 2006a) to obtain offspring that were iDTR+ and either Nestin-Cre+ (Ablatedimmature) or Nestin-Cre- (control group). All mice were given TMX injections between 6–8 weeks of age, followed by diphtheria toxin injections six weeks later (DT, 16 µg/kg in sterile saline for three consecutive days). To conditionally delete Bax from postmitotic GCs, we crossed POMC-Cre mice (Jackson #005965) with Baxfl/fl mice (see Figure 6—figure supplement 1). Conditional knockouts were maintained on a mixed 129 and C57BL/6J background using sibling controls. For counting newborn GCs, mice were crossed with POMC-eGFP transgenic mice (Jackson #009593). In some experiments, we visualized Cre-expressing cells by crossing conditional lines with Ai14 reporter mice (Jackson #007914). Tissue from homozygous germ line BaxKO mice (Jackson #002994) was used to validate Bax antibodies in western blots, with Bax+/- mice crossed with each other to generate both Bax-/- and control Bax+/+ genotypes. All experiments were performed in adult P60-P120 mice.

Electrophysiology

Mice were anesthetized and perfused intracardially with cold cutting solution containing (in mM): 110 choline chloride, 25 D-glucose, 2.5 MgCl2, 2.5 KCl, 1.25 Na2PO4, 0.5 CaCl2, 1.3 Na-ascorbate, 3 Na-pyruvate, and 25 NaHCO3. The brain was removed and 300 μm horizontal slices were taken on a Vibratome 3000EP or Leica VT1200S (Leica Biosystems, Wetzlar, Germany). After recovery in artificial CSF (ACSF) containing (in mM): 125 NaCl, 2.5 KCl, 1.25 NaH2PO4, 2 CaCl2, 1 MgCl2, 25 NaHCO3, and 25 glucose, recordings were performed at 30°C in ACSF +100 μm picrotoxin (PTX) to block GABAA receptors. Patch pipettes were filled with the following (in mM): 115 K-gluconate, 20 KCl, 4 MgCl2, 10 HEPES, 4 Mg-ATP, 0.3 Na-GTP, 7 phosphocreatine, 0.1 EGTA, pH 7.2 and 290 mOsm (2–4 MΩ). In some cases, a 0.2% biocytin was included in the patch pipette. Field pipettes were placed in the middle molecular layer and filled with ACSF (1–2 MΩ). A patch pipette filled with 1M NaCl (1 MΩ) was used to stimulate the middle molecular layer using an isolated stimulator (Digitimer, Letchworth Garden City, UK). The minimum stimulation intensity that evoked an EPSC was first established and the stimulus intensity was increased at multiples of the threshold intensity until response saturation was evident. In some experiments we tested a pre-set range of stimulus intensities, again ceasing stimulation after responses saturated. Both methods used the same range of intensities (0 to 100 V) with each approach generating fewer independent observations at progressively higher stimulus intensities due to saturation of axonal recruitment. The response of 10 stimuli at each intensity was averaged. Averaged field EPSPs (fEPSPs) and EPSCs were binned by their corresponding fiber volley (FV) amplitude. This normalizes for differences in stimulus intensities across experiments and removes the parameter ‘stimulus intensity’ from data sets.

Immunohistochemistry

Anesthetized mice were perfused intracardially with 0.9% NaCl or 0.1 M PBS and chilled 4% PFA before brains were removed and post-fixed overnight in PFA. Free-floating horizontal slices were taken on a Vibratome 1000 (50 μm). To enhance endogenous GFP expression, slices were blocked in TBS block buffer (0.1M TBS, glycine, 3% bovine serum albumin, 0.4% Triton X-100 and 10% normal goat serum) and incubated overnight with anti-GFP conjugated Alexa 488 (1:1000, Invitrogen, Carlsbad, CA). For NeuN and Dcx, slices were washed in TBST (50 mM Tris, 0.9% NaCl and 0.5% Triton X-100) and treated with antigen retrieval solution (10 mM sodium citrate, 0.5% tween 20) and 0.3% hydrogen peroxide before block with TBST +10% normal goat serum, followed by 48 hr incubation in rabbit anti-NeuN antibody (1:1000, Millipore, Billerica, MA) or rabbit anti-Dcx antibody (1:500, Abcam, Cambridge, UK), respectively. For NeuN, this was followed by incubation of 4 hr with goat anti-rabbit Alexa 647 (Invitrogen). For Dcx, a 3 hr incubation with biotinylated goat anti-rabbit (1:800, Southern Biotech, Homewood, AL) was followed by a 30 min incubation with streptavidin conjugated to Alexa Fluor 647 (1:200, Invitrogen). Slices were mounted with Prolong Gold or VectaShield mounting medium (Invitrogen). To visualize spines, acute brain slices containing biocytin-filled cells were post-fixed in 4% PFA for at least 24 hr then stained with streptavidin conjugated to Alexa Fluor 647 (1:1000, Invitrogen).

Stereology

EGFP+ cells and doublecortin (Dcx+) cells were counted using the optical fractionator method from every sixth slice through the entire left dentate gyrus using StereoInvestigator software (MBF Bioscience, Williston, VT). Counting frame and SRS grid sizes were set to give a Gunderson coefficient of error of <0.1 by an investigator blinded to genotype.

Spine counting

For Figure 3, mature GCs from control and BaxKOim mice were patched using an internal solution that included 0.2% biocytin. After fixation, GC dendrites and spines were imaged on an Olympus Fluoview 300 confocal microscope with a 60X objective and a 3X digital zoom using a z-step of 0.1 μm. Dendritic segments that were relatively horizontal to the plane of the slice were selected for spine analysis by an investigator blinded to genotype (avg segment length = 46 ± 7 µm in control and 56 ± 4 µm in BaxKOim mice, p=0.3). Analysis of spine density and type was performed by an investigator blinded to genotype using NeuronStudio software (Rodriguez et al., 2008).

For Figure 6, TdTomato+ (Bax-/-) or tdTomato- (Bax+/+) cells were patched in alternating slices from BaxKOmature mice and processed as described above. Spine density, length and head width were analyzed using Imaris software (Bitplane, Belfast, Northern Ireland)(Swanger et al., 2011).

Bax protein analysis

Hippocampal lysates were prepared by homogenizing flash frozen subdissected hippocampi using RIPA buffer (150 mM NaCl, 50 mM Tris, pH 7.5, 1% Triton-X 100, 0.5% sodium deoxycholate, 1% sodium dodecyl sulfate) containing protease inhibitors (Fisher Scientific, Hampton, NH). Following BCA assay (Pierce), 20 μg of lysate was separated through 12% polyacrylamide gels and transferred to low-fluorescent PVDF (Biorad, Hercules, CA). Membranes were blocked with casein blocking buffer (Sigma-Aldrich, St. Louis, MO) in Tris buffered saline with 0.1% Tween 20 (TBST) and incubated with primary antibody (in 0.3% BSA in TBST) at 4°C overnight using antibodies to detect Bax (Fisher Scientific) or beta-tubulin (Developmental Studies Hybridoma Bank). Secondary antibodies conjugated to Alexa-680 (Fisher Scientific) allowed detection and quantification by scanning with an Odyssey Imaging System (Licor Biosciences, Lincoln, NE).

Statistics

Data are expressed as mean ± SEM. We set the alpha level at 0.05 and accepted significant results with p<0.05 for all statistical tests. When determining the effect of genotype between two samples, data sets that satisfied normality criteria were analyzed with two-tailed paired or unpaired t tests, while non-normal data sets were analyzed with Mann-Whitney or Wilcoxon tests. For comparing two genotypes across multiple stimulus intensities, a two-way ANOVA was used. When EPSCs or fEPSPs were binned by FV amplitude, the number of data points varied between samples requiring an unweighted means analysis. Statistics were performed using Graphpad Prism.

Quantitative estimate of synapse redistribution

The purpose of the calculation is to predict the proportion of mature GC synapses that will be appropriated by immature cells over a 6-week time period in a control or BaxKOim DG. Time (t) is expressed in days, where t = 0 represents the starting point when 8-week-old animals are injected with TMX. New GCs are continually added to an existing network comprised of mature and immature GCs. Each new GC gains synaptic strength beginning two weeks after cell birth (Ge et al., 2006; Mongiat et al., 2009; Dieni et al., 2013), acquiring innervation from a finite pool of synapses with synaptic strength defined as the number of synapses per cell. The total number of GCs was initially set at 200,000 (unilateral cell count in the adult mouse DG (Pugh et al., 2011). The number of mature GCs (>8 weeks cell age) was set at 95% of the total (190,000), while the initial number of immature cells (2–8 weeks cell age) was set at 5% of the total (10,000) (Imayoshi et al., 2008). The baseline number of mature GC synapses at t = 0 was set at 100%, defined as 100 per cell, giving initial mature synapse number, SM:

SM=100(190,000×0.95)

We approximated the increase in synaptic strength, Y(t), of developing GCs by fitting the amplitude of evoked EPSCs in immature GCs at progressive ages (Dieni et al., 2013) by the equation:

Y(t)=71.1ln(14+t)187.7

For example, a 2-week-old control GC receives ~5% as many excitatory synapses as a mature GC, a 5-week-old GC contains ~65% as many excitatory synapses, and an 8-week-old GC achieves ‘mature’ levels of 100% synaptic strength. To determine the initial number of immature synapses, SI(0), we divided the number of initial immature GCs by 43 (the number of days of maturation and thus the number of different synaptic strengths) and multiplied this quantity by the sum of all synaptic strengths:

SI(0)=10,000/43×t=143Y(t)

This result plus the initial number of mature synapses gives the total synapses in the system:

SM + SI(0)

which remains static throughout the simulation (~19.6 million).

To calculate the number of synapses appropriated by immature GCs each day, we considered cell proliferation P(t), the rate of cell survival, and synaptic strength Y(t). The rate of decrease in progenitor proliferation was defined by a best-fit equation (Gil-Mohapel et al., 2013), adjusted to give ~8000 progenitor cells at t = 14, (stereological ki67 counts from 8-week-old mouse) (Chancey et al., 2013), giving the available progenitor cell number, P(t):

P(t)=4×106(42+t)1.5

The survival rate for new WT cells is 20% (Sierra et al., 2010). In the BaxKOim group, new GCs incorporating into the network at t = 14 (2 weeks after TMX-induced recombination) have a survival rate of 70% (assuming partial efficiency of Cre expression)(Lagace et al., 2007). The number of immature GCs added to the system per day, I(t), is:

I(t)=P(t)×survival rate

All immature GCs will gain synaptic strength daily. The immature synapses appropriated each day, SI(t), is the cumulative sum of the surviving GCs times their respective synaptic strengths:

SI(t)=(I(t)×Y(1))+(I(t1)×Y(2))+(I(t2)×Y(3))

Importantly, BaxKO GCs possess ~35% more synapses than control due to lack of Bax-dependent synapse pruning (Figure 3E, EPSC increase at highest FV bin).

In both groups, the cumulative number of immature synapses divided by the total synapses (multiplied by 100) equals the percent synapses appropriated by the immature population:

%im=SI(t)SM + SI(0)×100

Since there is a static number of total synapses defined at the start of the simulation, the percent mature synapses remaining is:

%mat=100%im

The %synapses occupied by all cell groups across time is plotted in Figure 7. Since the experiment is less than eight weeks in duration, immature GCs never convert into mature GCs, and we did not account for the conversion of pre-existing WT immature GCs because that population would not differ between control and BaxKOim conditions. To calculate the predicted difference in mature synapse number in BaxKOim vs. control conditions, we took the ratio of %mat in BaxKOim to %mat in control at each time point from t = 36 through t = 43 (multiplied by 100).

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Decision letter

  1. John Huguenard
    Reviewing Editor; Stanford University School of Medicine, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your work entitled "Adult Born Neurons Modify Excitatory Synaptic Strength of Existing Neurons" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

Notably, each of the reviewers appreciated the potential impact of the paper, as little is known regarding the incorporation of newly born neurons into existing circuitry, and in particular, the potential compensatory adaptations of existing neurons that occur when new neurons are integrated. And the approach to increase or decrease neurogenesis and examine effects on excitability is sound. However each of the reviewers had distinct concerns, regarding validation of the neurogenic modifying approaches or electrophysiological analysis and results. While each of these could be addressed with additional experimental results it would not be possible to complete these within a reasonable time frame. As you are probably aware, eLife has a policy to not ask authors to do more than 2 months of work to return a revision, as we believe that asking authors to do considerable new experiments has slowed down the progress of science. Therefore, we are rejecting this manuscript now so that you are free to move on to another journal with the work in its present state should you wish. Of course, if you agree with the reviewers, you may wish to do the work suggested, in any case.

Reviewer #1:

The paper from Overstreet-Wadiche's lab seeks to understand the synaptic-level interrelationship between newborn GCs and mature GCs in the hippocampal dentate circuit. One view of in regards to the functional contribution of newborn GCs is that they possess unique intrinsic properties in the network, but an emerging view is that there are homeostatic adaptations within the network (e.g., feedback inhibition from adult-born GCs to mature GCs) that contributes to the circuit. This paper provides additional experimental support for the latter view, which is potentially significant in the field.

The main thesis of this manuscript is to understand how mature granule cells in the dentate gyrus respond to integrating adult-born GCs. They use two primary methods to manipulate endogenous neurogenesis; BaxKO in Nestin progenitors to boost neurogenesis and iDTR to ablate. They use a series of electrophysiological recordings in hippocampal slices to measure synaptic function. By stimulating afferent axons, they measured total synaptic function by means of fEPSPs and synaptic responses per GC by whole cell EPSCs under both conditions.

The two key findings are:

1) Bax deletion in newborn neurons enhance neurogenesis and results in a decreased synaptic transmission of mature GCs. This is demonstrated by decreased frequency of sEPSCs, but not amplitude, recorded from individual mature granule cells. In addition, using Sr+ to disrupt synchronous vesicular release, the data suggest a postsynaptic mechanism.

2) Conversely when NG is ablated synaptic transmission onto mGCs is increased. Similarly this is demonstrated by an increase in EPSC from whole cell recordings of mature granule cells.

They interpret these results with a model that suggest that as immature neurons start to integrate, the mature neurons reduce their synaptic strength in favor of the immature neurons, a so-called redistribution of pre-existing functional synapses.

To further support this provocative interpretation, they provide evidence that BaxKO new neurons have increased EPSC amplitudes compared to aged matched WT new neurons. And BaxKO in mature granule cells also showed larger amplitude EPSCs than neighboring BaxWT mature GCs. While this demonstrates that Bax plays a role in regulation synaptic function, the data is not yet convincing to support their major conclusion that the observed effects is due to a redistribution of pre-existing synapses. Also, there are some issues with their WT controls and validation of the Bax KO and iDTR genetic tools, which the authors could address with their existing resources.

Major Comments:

1) The generation of control for BaxKO appears to be littermates that lack NestinCreERT2 or floxed Bax gene. When the data is presented, it appears that both controls are combined, or at least not distinguished. It would be informative to separate out genotype controls to determine if the results are consistent with both controls to ensure there is not an effect specifically due to Cre expression.

2) The authors should show the number of newborn GCs reduced with their inducible DT-R model. Although this model has been previously published, there may be different efficiencies of newborn neuron ablation based on the number of cells that express DT-R after TAM administration and the length of DT treatment.

3) To compliment the functional data and bolster support for the redistribution hypothesis include immunohistochemistry (e.g. for AMPA receptors) of mGCs in BaxKO vs WT to anatomically quantify synaptic appropriation.

4) For Figure 5 and associated Supplementals, the data is comparing TdTom+ vs TdTom- or eGFP+ vs eGFP- and assuming the reporter is a direct indication of cre recombination in that cell. However, because it is independently expressed it is possible for the reporter to falsely identify recombination efficiency. How do you know TdTom+ cell has Bax deleted? Is there an antibody for Bax that could be counterstained with TdTom to show they do not colocalize?

Reviewer #2:

The paper by Adlaf and colleagues investigates how the incorporation of newly generated granule cells of the adult hippocampus alters excitatory connections onto preexisting granule cells. This is the other side of the coin that has not been looked at previously, focusing on how networks are remodeled to accommodate development and integration of new neurons. The authors used conditional Bax deletion to prevent apoptotic death of new neurons to increase neurogenesis, and found reduced excitation onto mature preexisting neurons due to a reduced number of synapses. When neurogenesis was reduced using diphtheria toxin, synaptic transmission onto mature neurons was increased, suggesting a competition for presynaptic terminals between new and old neurons. The authors also found that Bax deletion increases excitation in a cell-autonomous manner both in developing and in mature granule cells, and that synaptic competition requires Bax signaling. This is a very interesting work that approaches circuit remodeling by from an original perspective, necessary to understand in depth the consequences of neuronal addition in functional networks.

There are some experimental caveats that need to be addressed to strengthen the main conclusions in the manuscript.

1) In the experiments shown in Figure 1, the experimental design is such that Bax is specifically deleted in adult-born neurons, and the strength of synaptic transmission is decreased in old preexisting granule cells. Given that synaptic strength depends on the levels of Bax expression, it would be important to check whether Bax levels in old preexisting neurons are altered under these conditions that increase neurogenesis.

2) In Figure 2, the authors could look at spine density and see the structural correlates to functional changes. This is a simple experiment within the scope of the manuscript that would reinforce that changes in the number of synaptic connections are occurring.

3) In Figure 3, it is important to quantify the extent to which neurogenesis has been reduced by DT.

4) In the data shown in the Suppl data to Figure 3. no changes were found in neurotransmitter release probability, or freq of spont events or postsynaptic amplitude associated with the reduction in EPSC amplitude in iDTR mice. Thus, there are no parameters to explain the reduced strength. The authors state that it is unclear whether the reduction is pre or postsynaptic. However, this inconsistency between data obtained from evoked vs spontaneous release needs to be clarified experimentally or adequately explained (describing possible underlying causes).

5) Expt in Figure 6 show that only increasing neurogenesis is not sufficient to promote reduction of synaptic strength in preexisting neurons; Bax is required for this sort of competition (there is more neurogenesis, but in the absence of Bax in all neurons there is no difference in synaptic strength). Bax is required for what seems to be a neuron/neuron competition. Going back to the experiments shown in Figure 1, in this context it would be important to test whether increasing neurogenesis in a manner that is independent of Bax renders a similar reduction in EPSC strength. For instance, housing mice in enriched environment, that increases survival of adult-born neurons, should induce similar effects in old neurons. This is a very simple experiment.

6) Figure 4 Suppl also show no changes in parameters measured from spontaneous synaptic currents. Similarly to what was discussed in point 4, it would be expected that BaxKO neurons with more synapses show higher frequency of spontaneous synaptic currents. This discrepancy also needs clarification.

Reviewer #3:

The key finding of this study is that selective manipulation of adult born neurons is inversely correlated with the numbers of excitatory synapses on mature neurons, with no detectable changes in so-called global measures of synaptic transmission. The conclusion is that it is a re-appropriation of existing synaptic terminals that causes mature GCs to lose synapses while the total synapse number in the network remains constant. This is a potentially interesting finding with significant implications. However, there are several major concerns.

1) There is uncertainty concerning the basic finding in Figure 1. The fiber volley vs stimulus intensity plot in Figure 1E (left panel) shows clear saturation beyond 10x threshold, with the FVs reaching a plateau at around 100 microV even with a doubling of the stimulus intensity (compare 10x with 20x). Thus, stimulation beyond 10x clearly does not recruit any more afferent fibers. The saturation effect is also obvious in the fEPSP slope vs stimulation intensity plot (Figure 1E, middle panel), where again the response reaches a plateau beyond 10x threshold. Therefore, it appears meaningless to use stimulation intensities beyond 10x threshold, or the corresponding FV value of 200 microV. But if we accept the latter premise, the argument that there is a reduced synaptic transmission to mature neurons without a change in fEPSP falls apart. To wit, in the fEPSP slope vs FV plot (Figure 1E, right panel), the two BaxKOim data points at 100 and 200 microV are clearly (and probably significantly) smaller than their control counterparts. At these smaller stimulation intensities, the fEPSP vs FV and the EPSC vs FV plots actually look very similar. Note also that the largest difference in the EPSC amplitude vs FV plot in Figure 1F appears at the strongest stimulation intensity. Thus, the basic finding about the unchanged fEPSP in the face of altered single cell EPSC does not appear to be valid when responses to physiologically meaningful stimuli are considered.

2) Another fundamental methodological issue concerns the alleged lack of changes in PPR. The authors apparently used 100 ms ISI only, which invariably yields PPR values around 1 (Figure 2A), meaning that at this ISI there was no short-term plasticity. Thus, the negative findings concerning PPR are not really interpretable from these data.

3) The authors only examined responses to stimulation of the middle molecular layer. Given that the study is concerned with global measures of synapse numbers in the dentate gyrus, other major excitatory inputs should be also examined and taken into account.

4) Why is it that the sEPSP frequency did not change in the immature neuron ablation experiment, whereas it did so when the number of immature neurons was enhanced?

5) A central conclusion of the study is that the induction of neurogenesis does not change the total number of synapses, and this argument is based on the (allegedly) unchanged fEPSP amplitude (but see above). Can one really use a relatively gross measure such as the fEPSP amplitude to argue for unchanged synapse numbers in a circuit, without direct anatomical assessment of synaptic density? This question is especially pertinent in a situation where EPSPs in different GC populations are expected to contribute to the field EPSP unevenly, since young and old GCs have different intrinsic properties.

6) The term synaptic strength is used in an ambiguous way in the text. Synaptic strength is usually used to define the efficacy of a given synapse, while here the proposed mechanism is a redistribution of functional synapses in the network. Receiving less or more inputs is not usually taken to mean that the synaptic strength changes.

7) What is the proposed mechanism of the hypothesized process of the transfer of synapses between immature and mature GCs?

https://doi.org/10.7554/eLife.19886.030

Author response

Notably, each of the reviewers appreciated the potential impact of the paper, as little is known regarding the incorporation of newly born neurons into existing circuitry, and in particular, the potential compensatory adaptations of existing neurons that occur when new neurons are integrated. And the approach to increase or decrease neurogenesis and examine effects on excitability is sound. However each of the reviewers had distinct concerns, regarding validation of the neurogenic modifying approaches or electrophysiological analysis and results. While each of these could be addressed with additional experimental results it would not be possible to complete these within a reasonable time frame. As you are probably aware, eLife has a policy to not ask authors to do more than 2 months of work to return a revision, as we believe that asking authors to do considerable new experiments has slowed down the progress of science. Therefore, we are rejecting this manuscript now so that you are free to move on to another journal with the work in its present state should you wish. Of course, if you agree with the reviewers, you may wish to do the work suggested, in any case.

We thank the reviewers for constructive comments. We have added substantial new data, analysis and text revisions to address the concerns. In particular, we think the new data, summarized below, has strengthened our conclusions:

1) Analysis of FV/EPSC ratio in control groups (reviewer 1; new panels in Figure 1—figure supplement 1)

2) Quantification of iDTR ablation (reviewer 1 and 2; new panel in Figure 4)

3) Immunohistochemical detection of synapses (reviewer 1; new panels in Figure 4—figure supplement 1)

4) Spine density/type in mature neurons from BaxKOim and control mice (reviewers 1& 2; new Figure 3)

5) Assay of Bax expression in mature neurons (reviewer 2; new Figure 5—figure supplement 4)

6)Additional interstimulus intervals for the paired-pulse ratio (reviewer 3; new Figure 3)

7) Using environmental enrichment to enhance neurogenesis (reviewer 2; new Figure 8)

8)Data showing unchanged Ca2+ dependence of EPSCs (reviewer 3; new Figure 6—figure supplement 2)

We’d like to highlight a recently published article that likewise supports the idea of synaptic redistribution. McAvoy et al., Neuron (2016), show that synaptic integration of new neurons increases following conditional reduction of mature GC spines. We now show that selectively increasing neurogenesis decreases mature spine density, providing anatomical support for our main conclusion that adult born neurons modify synaptic connectivity of mature neurons.

Reviewer #1:

[…]

Major Comments:

1) The generation of control for BaxKO appears to be littermates that lack NestinCreERT2 or floxed Bax gene. When the data is presented, it appears that both controls are combined, or at least not distinguished. It would be informative to separate out genotype controls to determine if the results are consistent with both controls to ensure there is not an effect specifically due to Cre expression.

This is a good point. Since we recorded EPSCs primarily from mature neurons that do not ever express Cre, it seemed unlikely that Cre expression in newborn neurons affects our results. Nevertheless, to explicitly control for this possibility, we have now added new panels to Figure 1—figure Supplement 1. Panel B shows there is no difference in the EPSC/FV ratio across control groups. We also included an additional analysis showing the EPSC/FV ratio difference persists using only mice with the Baxfl/fl genotype, since Baxfl/fl mice were originally obtained on a mixed B6;129 genetic background (panel C). We referenced these results in the text (Results section paragraph two). Finally, we did not detect differences in the synaptic properties of mature dentate GCs that underwent Cre-mediated recombination in POMC-Cre/Baxwt/wt mice (Figure 6F), further suggesting that transient Cre expression does not affect our results.

2) The authors should show the number of newborn GCs reduced with their inducible DT-R model. Although this model has been previously published, there may be different efficiencies of newborn neuron ablation based on the number of cells that express DT-R after TAM administration and the length of DT treatment.

We have now added stereological cell counts of Dcx-expressing immature neurons showing ~25% reduction in Dcx+ cells along with representative images (Figure 4B). We included this data in the text (subsection “Ablation of immature neurons increases synaptic transmission to mature neurons”) along with the explanation that re-population of Dcx-expressing cells during the 1-2 weeks after DT-mediated ablation underestimates the ablation efficiency, as described in similar models (Yun et al., 2016).

3) To compliment the functional data and bolster support for the redistribution hypothesis include immunohistochemistry (e.g. for AMPA receptors) of mGCs in BaxKO vs WT to anatomically quantify synaptic appropriation.

We have now included data showing that expression of the presynaptic protein vGluT1 is unaltered in the molecular layer of BaxKOimmature and Ablatedimmature slices (Figure 4—figure supplement 1, subsection “Ablation of immature neurons increases synaptic transmission to mature neurons”). However, this approach to label synapses by pre- or postsynaptic markers only assays total synapses, since it is not feasible to distinguish between mature or immature GC synapses.

Unfortunately, even using tdTom expression in immature neurons would not be sufficient to distinguish between mature and immature dendrites/spines because tdTom does not label the entire population of immature neurons.

To further address the request to anatomically quantify synaptic appropriation, we have now performed spine counts from mature GCs in BaxKOim and control mice after whole cell recordings. We find that spine density of mature GCs is reduced, supporting the interpretation of our functional results. This data is included in new Figure 3 (with text in subsection “Enhancing immature neurons decreases EPSCs and spine density of mature neurons”).

4) For Figure 5 and associated Supplementals, the data is comparing TdTom+ vs TdTom- or eGFP+ vs eGFP- and assuming the reporter is a direct indication of cre recombination in that cell. However, because it is independently expressed it is possible for the reporter to falsely identify recombination efficiency. How do you know TdTom+ cell has Bax deleted? Is there an antibody for Bax that could be counterstained with TdTom to show they do not colocalize?

We agree that TdTom expression may overestimate recombination efficiency, and thus the difference in EPSCs shown in original Figure 5 (now Figure 6) underestimates the effect of Bax deletion. At the time we did the experiment, we were aware that Bax antibodies are notorious for poor labeling in brain slices and we subsequently confirmed this (see below). Thus we tested whether the Baxfl/fl allele was necessary for the difference in EPSCs in TdTom+ vs TdTom- cells (new Figure 6C) by performing the functional control experiment shown in Figure 6F. This experiment shows that the Baxfl/fl allele is necessary for the difference in EPSCs because repeating the experiment in TdTom+ Baxwt/wt cells revealed no difference. We think this is a very strong control experiment, which also supports a lack of effect of transient Cre expression on synaptic function (see point 1, above).

To our knowledge, the only validated immunohistochemical identification of Bax in hippocampal slices shows weak diffuse labeling excluded from cell bodies using a non-fluorescent method (Figure 2 of Sun et al., J Neurosci 2004). Unfortunately we are unable to reliably detect Bax using this and other antibodies with fluorescence protocols, with comparison to tissue from germline BaxKO mice (Author response image 1). This may be the reason why previous studies using conditional Bax deletion have not confirmed deletion with immunohistochemistry (Sahay et al., 2011; Ikrar et al., 2013; Hill et al., 2015). However, the increase in newborn neurons in BaxKOimmature and BaxKOmature mouse lines identified by POMC-GFP expression independently verifies that Bax has been deleted from a significant fraction of newborn cells. In addition to the results shown in Figure 6C, we found increased evoked EPSCs in 6 week-old and 16 week-old adult born conditional Bax-deleted GCs. Thus our data shows that EPSCs are enhanced in three separate experimental paradigms with Cre-mediated recombination in Baxfl/fl cells, making it highly unlikely to be an experimental artifact.

Author response image 1
Bax immunostaining from WT slices (top) and germline BaxKO slices (bottom), using both standard (left) and antigen-retrieval methods (right).

We are unaware of an antibody that reliably identifies Bax in hippocampal slices.

https://doi.org/10.7554/eLife.19886.027

Reviewer #2:

[…]

There are some experimental caveats that need to be addressed to strengthen the main conclusions in the manuscript.

1) In the experiments shown in Figure 1, the experimental design is such that Bax is specifically deleted in adult-born neurons, and the strength of synaptic transmission is decreased in old preexisting granule cells. Given that synaptic strength depends on the levels of Bax expression, it would be important to check whether Bax levels in old preexisting neurons are altered under these conditions that increase neurogenesis.

We did not intend to imply that synaptic strength depends of the level of Bax expression. Previous work shows that Bax is required for LTD because Bax activation is an intermediary step between NMDAR-Ca2+ influx and activation of caspase-3 that is necessary and sufficient for LTD (Li et al., Cell, 2010; Li and Jiao, Neuron 2011). Thus while Bax deletion prevents LTD (which leads to spine

shrinkage and eventual spine elimination (Nägerl et al., Neuron 2004; Zhou et al., Neuron 2004)), we are not aware of evidence that the level of Bax protein correlates with Bax activation or synaptic function/spine density. We have clarified the point that Bax activation is a key requirement in NMDAR- dependent hippocampal LTD in the text (subsection “Bax deletion in mature neurons increases EPSCs and spine density”).

Our results indicate that Bax is required in mature GCs for neurogenesis-induced loss of transmission (Figure 7), suggesting that a change in the Bax-caspase 3 signaling pathway is indeed involved in spine loss from mature GCs. Thus the contribution of Bax to our results is complex. We show that mature GCs exhibit a non-cell autonomous effect of Bax deletion from adult-born GCs (Figures 13, decreased EPSCs) that is opposite to the cell-autonomous effect of Bax deletion in both cell types (Figures 56, increased EPSCs). Remarkably, the cell autonomous function is required for the non-cell autonomous effect (Figure 7). This complexity makes sense when we consider the role of Bax-caspase 3 signaling in both cell death and synapse pruning (these points are now clarified in the Discussion).

We speculate that synapse loss from mature neurons involves enhanced Bax-caspase 3 activation rather than changes in Bax expression, as is reported for NMDAR-dependent LTD (Li and Jiao, 2011). We have now included western blot analysis showing that overall Bax protein levels are unchanged in BaxKOim mice, confirming that deletion of Bax from a small percentage of cells does not change global levels of Bax protein (new Figure 5 —figure supplement 4). In the absence of a reliable antibody for cell-type specific quantification in slices, this question cannot be more directly addressed with our currently available approaches. We are very interested in understanding this signaling pathway and its consequences for GC synaptic connectivity, but addressing this in more detail is likely beyond the scope of the current study.

2) In Figure 2, the authors could look at spine density and see the structural correlates to functional changes. This is a simple experiment within the scope of the manuscript that would reinforce that changes in the number of synaptic connections are occurring.

We thank the reviewer for this suggestion. We have now included spine analysis that shows reduced spine density in mature GCs from BaxKOim mice (new Figure 3, text in subsection “Enhancing immature neurons decreases EPSCs and spine density of mature neurons”). This data confirms the interpretation of our functional analysis (Figures 12) of reduced number of synapses onto mature GCs in BaxKOim mice.

3) In Figure 3, it is important to quantify the extent to which neurogenesis has been reduced by DT.

We have now added stereological cell counts of Dcx-expressing immature neurons showing >25% reduction in Dcx+ cells along with representative images (revised Figure 4). This data is included in the text (subsection “Ablation of immature neurons increases synaptic transmission to mature neurons”) along with the explanation that re-population of Dcx-expressing cells during the 1-2 weeks after DPT-mediated ablation underestimates the ablation efficiency (Yun et al., 2016).

4) In the data shown in the Suppl data to Figure 3. no changes were found in neurotransmitter release probability, or freq of spont events or postsynaptic amplitude associated with the reduction in EPSC amplitude in iDTR mice. Thus, there are no parameters to explain the reduced strength. The authors state that it is unclear whether the reduction is pre or postsynaptic. However, this inconsistency between data obtained from evoked vs spontaneous release needs to be clarified experimentally or adequately explained (describing possible underlying causes).

This is a good point that we now explain in the text. The frequency of spontaneous activity in mature dentate GCs is low (~1Hz), and primarily mediated by single site events since the amplitude is not affected by TTX (i.e. Dieni et al., J Neuro 2013, Parent et al., E J Neurosci 2016). Mature granule cells are estimated to have up to -5,000 synapses, such that spontaneous events represent the activity of only a small fraction of synapses, and there is high variability in the frequency of activity between individual GCs. Due to dendritic filtering and imperfect space clamp, it is also likely that spontaneous synaptic activity arising at proximal locations (i.e. synapses in the IML) are over represented, potentially contributing to “noise’ within and between experiments that does not occur in the evoked assay. While it is informative to see changes in spontaneous activity, it may be difficult to detect relatively small differences using this “lower resolution” assay, whereas evoked transmission repeatedly assays a higher fraction of synapses and there is less variability between cells after we use the FV to normalize the number of stimulated axons across slices. It appears that the change in neurogenesis in BaxKOim mice is greater than in iDTR mice (~40% vs ~25% change), and thus the “lower resolution” assay is unable to detect a difference in the iDTR model whereas the “higher resolution” assay can detect both. Finally, there is also ongoing debate about whether evoked and spontaneous synaptic release events actually arise from the same pool of vesicles, potentially underlying many reported differences in experimental results obtained by evoked and spontaneous activity (reviewed by Kavalali, Nat Rev Neurosci 2015). We have added this explanation in subsection “Ablation of immature neurons increases synaptic transmission to mature neurons”.

5) Expt in Figure 6 show that only increasing neurogenesis is not sufficient to promote reduction of synaptic strength in preexisting neurons; Bax is required for this sort of competition (there is more neurogenesis, but in the absence of Bax in all neurons there is no difference in synaptic strength). Bax is required for what seems to be a neuron/neuron competition. Going back to the experiments shown in Figure 1, in this context it would be important to test whether increasing neurogenesis in a manner that is independent of Bax renders a similar reduction in EPSC strength. For instance, housing mice in enriched environment, that increases survival of adult-born neurons, should induce similar effects in old neurons. This is a very simple experiment.

We have now included the requested experiment of increasing neurogenesis with EE/running (new Figure 8). We did not include this data in the original submission because this simple experiment requires a complex interpretation. Studies over the last 30 years have shown that EE increases synaptic connectivity in the DG, seen as increases in fEPSPs without changes in fiber volleys or paired-pulse ratios (Green and Greenough, JNeurophys 1986; Foster, Brain Res 1996) that can be explained by the increased spine density of mature GCs (Eadie et al., J Comp Neurol 2005; Stranahan et al., Hippocampus 2007, Glasper et al., Hippocampus 2010). We replicated the longstanding findings on fEPSPs and FVs (new Figure 8—figure supplement 1), and now directly show enhanced synaptic transmission to mature neurons with both evoked and spontaneous EPSC assays (new Figure 8). Our results are thus consistent with the prevailing evidence that EE increases synaptic connectivity of pre-existing mature GCs, in contrast to recent evidence that only adult born neurons in a critical period of development exhibit increased connectivity after EE (Bergami et al., Neuron 2015). Several differences in the cell populations tested or timing of EE could contribute to these differences. But altogether our results demonstrate the capacity of mature GCs to exhibit synaptic plasticity (in the form of altered connectivity) in response to both genetic and experiential circuit manipulations, a point that we now make clear in the Introduction.

Because EE increases the connectivity of mature neurons as well as the number of new neurons, this paradigm cannot resolve whether there is a redistribution of synapses. Increased synaptic connectivity (based on both anatomical and functional measures) induced by EE occurs in CA1 and other brain regions, indicating that it is a parallel mechanism independent of neurogenesis (Rampoon et al., Nat Neuro 2000; Stranahan et al., 2007; Glasper et al., 2010, De Bartolo et al., Brain Struct Funct, 2015; Bechard et al., Behav Brain Res 2016, etc). Thus EE could induce both outcomes (an increase in mature connectivity and an increase in new neuron synaptic redistribution), such that the “net effect” of EE on mature GC EPSCs shown in Figure 8 may be reduced by synaptic re-distribution to EE-induced new neurons. Extensive and additional experiments beyond the scope of this manuscript would be required to sort out this question.

We wholly agree that it would be useful to replicate our main result using an approach that is truly selective for enhancing the number of new neurons without affecting their integration or synaptic function, but we are unaware of an approach that has been fully validated in this regard. It seems that many proteins have dual roles in cell survival/cell cycle and synaptic function. Supporting this point, both proteins that were used to alter spine density in mature neurons in McAvoy et al., (2016; Klf9 and Rac1) may also be involved in apoptotic pathways (Stankiewicz and Linseman, Front Cell Neurosci 2014; Lebrun et al., Mol Cell Neurosci 2013) and our impression is that most genetic approaches that alter the number of new GCs also affect their maturation/dendrite structure.

While we cannot identify and validate a new approach to selectively increase neurogenesis without affecting integration, we point out that we used an alternative method to selectively reduce neurogenesis (Figure 4). In our view, demonstrating opposite manipulations of neurogenesis that lead to opposite changes in mature neuron EPSCs provides compelling evidence for our main conclusion that the number of new neurons affects synaptic connectivity of mature neurons.

6) Figure 4 Suppl also show no changes in parameters measured from spontaneous synaptic currents. Similarly to what was discussed in point 4, it would be expected that BaxKO neurons with more synapses show higher frequency of spontaneous synaptic currents. This discrepancy also needs clarification.

We reiterate that the low frequency of spontaneous activity makes it a less sensitive assay for changes in synaptic function than evoked activity. There are also many examples in the literature of discrepancies between evoked and spontaneous activity, which has led to ongoing debate about whether evoked and spontaneous synaptic release events arise from the same pool of vesicles (reviewed by Kavalali, Nat Rev Neurosci 2015). We have mentioned these points in the text. See also point 4 above.

Reviewer #3:

The key finding of this study is that selective manipulation of adult born neurons is inversely correlated with the numbers of excitatory synapses on mature neurons, with no detectable changes in so-called global measures of synaptic transmission. The conclusion is that it is a re-appropriation of existing synaptic terminals that causes mature GCs to lose synapses while the total synapse number in the network remains constant. This is a potentially interesting finding with significant implications. However, there are several major concerns.

1) There is uncertainty concerning the basic finding in Figure 1. The fiber volley vs stimulus intensity plot in Figure 1E (left panel) shows clear saturation beyond 10x threshold, with the FVs reaching a plateau at around 100 microV even with a doubling of the stimulus intensity (compare 10x with 20x). Thus, stimulation beyond 10x clearly does not recruit any more afferent fibers. The saturation effect is also obvious in the fEPSP slope vs stimulation intensity plot (Figure 1E, middle panel), where again the response reaches a plateau beyond 10x threshold. Therefore, it appears meaningless to use stimulation intensities beyond 10x threshold, or the corresponding FV value of 200 microV.

We respectfully disagree that there is uncertainty in the result in Figure 1. This concern appears to be a misunderstanding of our experimental protocol and analysis, which likely resulted because we did not provide sufficient detail in the methods. We will explain the misunderstanding in detail, and describe how we have altered the figure and text to avoid this misunderstanding for other readers.

Author response image 2
Examples from six randomly selected experiments showing FV versus stimulus intensity relationships.

Although the average graph (bottom right) suggests saturation of FVs around 200 µV (dotted line), the individual experiments reveal high variability in maximum FVs and saturation points (bottom left). We monitored saturation during each experiment and stopped stimulating when saturation was apparent (i.e. slice A,D,E).

https://doi.org/10.7554/eLife.19886.028

Reviewer 3 is correct that the averaged FV and fEPSP data shown in Author response image 3E gives the appearance saturation. The averaged data, however, is not a good indicator of saturation in individual experiments because the stimulus-response relationships are highly variable between slices. We have illustrated this variability in a few examples of raw data and their average (Author response image 2). The raw data shows variability both in the maximum FV amplitude (ordinate) as well as the stimulus intensity at which the maximum FV amplitude is achieved (the point of saturation on the abscissa). Hence the “linear range” of the stimulus response in each experiment occurs across a variable range of stimulus intensities. The goal of our stimulating protocol was to stimulate at multiple points within the linear range to acquire useful data across that range, but not above it. When saturation appeared (i.e. Slice A, D, E), we did not acquire responses at higher stimulus intensities. This results in variability in the stimulus intensities and intervals used (compare slice B versus E), and fewer data points in the averages for the highest range of stimulus intensities. Each individual response includes a range of FV amplitudes with a minimal number of saturated FVs. We have added the following description to the Methods to clarify how the experiments were performed:

“A patch pipette filled with 1M NaCl (1 MΩ) was used to stimulate the middle molecular layer using an isolated stimulator (Digitimer). The minimum stimulation intensity that evoked an EPSC was first established and the stimulus intensity was increased at multiples of the threshold intensity until response saturation was evident. In later experiments we tested a pre-set range of stimulus intensities, again ceasing stimulation after responses saturated. Both methods used the same range of intensities (0 to 100 V) with each approach generating fewer independent observations at progressively higher stimulus intensities due to saturation of axonal recruitment. The average response of 10 stimuli at each intensity was used for analysis. Field EPSPs (fEPSPs) and EPSCs were binned by their corresponding fiber volley (FV) amplitude. This normalizes for differences in stimulus intensities across experiments and removes the parameter “stimulus intensity” from all data sets.”

Thus the reviewer’s conclusion that responses are saturated at 10x threshold or at FV values above 100-200 µV are due to an over-interpretation of the averaged data plots of stimulus intensity and our failure to explicitly explain these plots. Whether the number of recruited axons is limiting (or saturated) is best assessed by the right panel of Author response image 3E (circled) that plots the fEPSP slope versus FV. This linear relationship indicates that the postsynaptic response (fEPSP) does not saturate well above FV values beyond 100-200 µV.

We included all three plots in Author response image 3E as an attempt to illustrate that the linear fEPSP vs FV plot (shown in most of our figures) results from the transformation of the two “messy” stimulus intensity plots (FV vs Stim and fEPEP vs Stim), revealing the linear (non-saturated) fEPSP-FV relationship once the “Stim Intensity” parameter is removed. We apologize if this presentation caused confusion rather than clarifying how the circled graph was generated.

But if we accept the latter premise, the argument that there is a reduced synaptic transmission to mature neurons without a change in fEPSP falls apart.

As shown in Author response image 3 and described above, we did not include saturated responses in our analysis. The fEPSP versus FV relationship is linear.

To wit, in the fEPSP slope vs FV plot (Figure 1E, right panel), the two BaxKOim data points at 100 and 200 microV are clearly (and probably significantly) smaller than their control counterparts.

The fEPSP slope vs FV plot of slices from control and Baxim is not statistically significant (2way ANOVA). This outcome was replicated when we repeated the experiment (Figure 5—figure supplement 1; note this is the same experiment as shown in Figure 1, except that the mice include the Cre reporter TdTom and whole cell recording targets TdTom+ immature rather than unlabeled mature GCs).

At these smaller stimulation intensities, the fEPSP vs FV and the EPSC vs FV plots actually look very similar. Note also that the largest difference in the EPSC amplitude vs FV plot in Figure 1F appears at the strongest stimulation intensity.

The normalization procedure removes the parameter “stim intensity” from our analysis, leaving only the FV parameter that is a more robust measure of the number of axons recruited. Each FV bin contains data from a range of stimulus intensities (shown in lower left panel of Author response image 2; it does not make sense to refer to stimulus intensities in binned FV data). Furthermore, most of our fEPSPs and EPSCs are in the smallest FV bins (now clarified in the figure legends) and all FV bins in Figure 1F are significantly different by Bonferonni post-tests (now added to legend).

Thus, the basic finding about the unchanged fEPSP in the face of altered single cell EPSC does not appear to be valid when responses to physiologically meaningful stimuli are considered.

Importantly, our interpretation about synaptic redistribution does not require that the fEPSP is unchanged, only that fEPSP does not increase (as one might initially predict from enhancing neurogenesis). Based on smaller EPSCs in mature GCs (Figures 13), one might indeed expect smaller fEPSPs since mature GCs make up the vast majority of GCs. Presumably the fEPSP does not change because of the additional contribution of immature synapses. Yet, our interpretation of synaptic redistribution does not require that the immature neuron contribution to the fEPSP fully compensates for the loss of transmission to mature neurons (that is, if we had actually found a reduction in the fEPSP, or if the sensitivity of extracellular recording is too low to detect such a difference). Only an increase in the fEPSP in Figure 1 would lend support a synaptic addition model (but even so, the decrease in mature EPSC and spine density still does not fit with an addition model). These are interesting and relevant points that we have now expanded on in the discussion, but the main point is that a smaller fEPSP does not affect our conclusions.

We reiterate that our central finding, that EPSCs in mature GCs are reduced in BaxKOim mice, is a robust result; the difference is apparent in the raw traces, it is accompanied by changes in frequency of spontaneous EPSCs and Sr2+ evoked EPSCs, and we now show that it is associated with reduced spine density of mature GCs (new Figure 3).

To avoid these misunderstanding by other readers, we have reformatted Figure 1 to make the presentation consistent with the other figures. We moved the averaged FV and fEPSP plots to Figure 1—figure supplement 2 and we provided more detail in the legends. We added a new panel to Figure 1 that shows the robustness of the difference in EPSC/FV ratios (independent of the FV amplitude) and we revised the methods to clarify that saturation was assessed in each individual experiment.

2) Another fundamental methodological issue concerns the alleged lack of changes in PPR. The authors apparently used 100 ms ISI only, which invariably yields PPR values around 1 (Figure 2A), meaning that at this ISI there was no short-term plasticity. Thus, the negative findings concerning PPR are not really interpretable from these data.

We have now included additional ISIs for the paired pulse ratio experiment in new Figure 3. There was no difference in PPR at any ISI, as reported previously using fEPSPs in a similar model (S Figure 9C from Sahay et al., Nature 2011). We now cite Petersen et al., Neuroscience (2013) that shows MPP stimulation generates mildly depressing PPRs that depend, in part, on the stimulus intensity because it is difficult to isolate MPP from LPP stimulation (which displays PPF). We did not attempt to isolate the MPP rather we simply placed the stimulating electrode in the middle molecular layer, making it likely that our responses have some contamination from the LPP.

We are surprised by the assertion that a PPR near 1 is insensitive to changes in release probability, since manipulating release probability usually changes the PPR at short ISIs regardless of the initial PPR (i.e. Figure 8B,D of Debanne et al., 1996; Chancey et al., J Neuro 2014). To further address this concern, we now include data showing that the Ca2+ dependence of release is unaltered by Bax deletion from the majority of GCs (new Figure 6—figure supplement 2). This figure confirms that PPRs at 100 ms ISI are sensitive to changing the release probability and further rule out presynaptic changes following Bax deletion.

3) The authors only examined responses to stimulation of the middle molecular layer. Given that the study is concerned with global measures of synapse numbers in the dentate gyrus, other major excitatory inputs should be also examined and taken into account.

We reported changes in sEPSCs in Figure 1 that represent a global measure of synapse number rather than strictly medial perforant path synapses. While the results from additional examination of the A/C pathway or the lateral perforant path would be interesting and may perhaps help to define the extent of the change in mature neuron synaptic function, it is not clear that results from such experiments could alter our main finding since this request for additional data does not address a specific concern about our current data set.

4) Why is it that the sEPSP frequency did not change in the immature neuron ablation experiment, whereas it did so when the number of immature neurons was enhanced?

This is a good point that we now clarify in the text. The frequency of spontaneous EPSCs in mature dentate GCs is low (~1Hz), and primarily mediated by single site events since the amplitude is not affected by TTX (i.e. Dieni et al., J Neuro 2013, Parent et al., E J Neurosci 2016). Mature granule cells are estimated to have up to -5,000 synapses, such that spontaneous events represent the activity of only a small fraction of synapses, and there is high variability in the frequency of activity between individual GCs. Due to dendritic filtering and imperfect space clamp, it is also likely that spontaneous synaptic activity arising at proximal locations (i.e. synapses in the IML) are over represented, potentially contributing to “noise’ within and between experiments that does not occur in the evoked assay. While it is informative to see changes in spontaneous activity, it may be difficult to detect relatively small differences using this “lower resolution” assay, whereas evoked transmission repeatedly assays a higher fraction of synapses and there is less variability between cells after we use the FV to normalize the number of stimulated axons across slices. It appears that the change in neurogenesis in BaxKOim mice is greater than in iDTR mice (~40% vs ~25% change), and thus the “lower resolution” assay is unable to detect a difference in the iDTR model whereas the “higher resolution” assay can detect both. Finally, there is also ongoing debate about whether evoked and spontaneous synaptic release events actually arise from the same pool of vesicles, potentially underlying many reported differences in experimental results obtained by evoked and spontaneous activity (reviewed by Kavalali, Nat Rev Neurosci 2015). We have added this explanation in subsection “Ablation of immature neurons increases synaptic transmission to mature neurons”.

5) A central conclusion of the study is that the induction of neurogenesis does not change the total number of synapses, and this argument is based on the (allegedly) unchanged fEPSP amplitude (but see above).

Our central conclusion is indicated by the title (Adult-Born Neurons Modify Excitatory Synaptic Transmission to Existing Neurons) and in the last sentence of the Abstract:

“Together these results show that neurogenesis modifies the synaptic function of mature neurons in a manner consistent with a redistribution of pre-existing synapses to newly integrating neurons and that a non-apoptotic function of the Bax signaling pathway contributes to ongoing synaptic refinement within the DG circuit.”

As described in Point 1 above, we have a much softer interpretation of total synapse number than indicated by Reviewer 3 and smaller fEPSPs would not alter our conclusions.

Can one really use a relatively gross measure such as the fEPSP amplitude to argue for unchanged synapse numbers in a circuit, without direct anatomical assessment of synaptic density? This question is especially pertinent in a situation where EPSPs in different GC populations are expected to contribute to the field EPSP unevenly, since young and old GCs have different intrinsic properties.

We agree that the fEPSP may not be particularly sensitive to synaptic density and will be affected by intrinsic excitability (this is why we measured intrinsic excitability of mature GCs in all our protocols). We also agree that high intrinsic excitability will potentially allow immature GCs to have a greater contribution to fEPSPs (per AMPAR-containing synapse), potentially leading to similar fEPSPs despite smaller EPSCs in mature GCs.

Our new EE data includes a “positive control” that we can detect larger fEPSPs in a condition that is well known to increase the number of spines in the DG (replicating prior results of Green and Greenough, 1986 and Foster et al., 1996). We also cited prior work that found no change in anatomical assessment of synaptic density (measured with EM) despite a large increase in newborn cells in germline Bax knockout mice, and we have now also included an immunohistochemical assay suggesting presynaptic terminal density does not change (Figure 4—figure supplement 1). Even so, whether the fEPSP is smaller is tangential to our novel finding that EPSCs in mature GCs are altered by selective manipulations of newborn GCs.

6) The term synaptic strength is used in an ambiguous way in the text. Synaptic strength is usually used to define the efficacy of a given synapse, while here the proposed mechanism is a redistribution of functional synapses in the network. Receiving less or more inputs is not usually taken to mean that the synaptic strength changes.

We purposefully used the somewhat ambiguous term synaptic strength because it refers to a functional measure of synaptic transmission without assigning pre- or postsynaptic mechanisms to define it. Now that we have anatomical data that confirms the reduction in EPSCs in Figures 1 and 2 are associated with fewer anatomically defined synapses, we have edited the text, including the title, to use more precise terminology.

7) What is the proposed mechanism of the hypothesized process of the transfer of synapses between immature and mature GCs?

Our data shows that deletion of Bax increases synaptic strength and spine density (new Figure 5 & 6) and that deletion of Bax in mature GCs blocks neurogenesis-induced loss of synaptic transmission (Figure 7). Since the Bax-/caspase-3 pathway is necessary and sufficient for AMPA receptor trafficking, NMDAR-dependent synaptic depression (LTD) and synapse pruning (references cited in the text), we speculate that the transfer of synapses involves synaptic depression (LTD) and subsequent pruning of mature GC synapses. We have clarified our discussion of this in the Results section and Discussion.

https://doi.org/10.7554/eLife.19886.031

Article and author information

Author details

  1. Elena W Adlaf

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    EWA, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  2. Ryan J Vaden

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    RJV, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  3. Anastasia J Niver

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    AJN, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  4. Allison F Manuel

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    AFM, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  5. Vincent C Onyilo

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    VCO, Conception and design, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  6. Matheus T Araujo

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    MTA, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  7. Cristina V Dieni

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    CVD, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  8. Hai T Vo

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    HTV, Acquisition of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  9. Gwendalyn D King

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    GDK, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon 0000-0002-3659-9241
  10. Jacques I Wadiche

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    JIW, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    1. jwadiche@uab.edu
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon 0000-0001-8180-2061
  11. Linda Overstreet-Wadiche

    1. Department of Neurobiology, University of Alabama at Birmingham, Birmingham, United States
    Contribution
    LO-W, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    1. lwadiche@uab.edu
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon 0000-0001-7367-5998

Funding

Civitan International (Emerging Scholars Award)

  • Elena W Adlaf
  • Hai T Vo

National Institutes of Health (F31NS098553)

  • Ryan J Vaden

National Institutes of Health (P30 NS047466)

  • Gwendalyn D King
  • Jacques I Wadiche
  • Linda Overstreet-Wadiche

National Institutes of Health (NS065920)

  • Jacques I Wadiche

National Institutes of Health (NS064025)

  • Linda Overstreet-Wadiche

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank members of the Wadiche labs for helpful discussions throughout this project, Mary Seelig for technical assistance and Nancy Gallus for help with immunohistochemistry. This work was supported by Civitan International Emerging Scholars awards (EWA and HTV), F31NS098553 (RJV), NIH NS064025 (LOW), NIH NS065920 (JIW) and NIH P30 NS047466.

Ethics

Animal experimentation: All animal procedures followed the Guide for the Care and Use of Laboratory Animals, U.S. Public Health Service, and were approved by the University of Alabama at Birmingham Institutional Animal Care and Use Committee (protocol# 8674 and 10134).

Reviewing Editor

  1. John Huguenard, Reviewing Editor, Stanford University School of Medicine, United States

Publication history

  1. Received: July 21, 2016
  2. Accepted: January 5, 2017
  3. Version of Record published: January 30, 2017 (version 1)

Copyright

© 2017, Adlaf et al

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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