Mice and rats were maintained in a 12 h light/dark cycle with food and water ad libitum. Every effort to minimize the distress and pain of animals was made. All experimental procedures were performed according to the German animal welfare legislation, approved by the appropriate animal welfare committee and the animal welfare officer of the University of Freiburg.

Mice of the strain C57BL/6J and rats of the strain Wistar (Crl:WI) of both sexes were used in this study.

Organotypic tissue cultures were prepared from mice and rats of both sexes at postnatal day 3–5 and allowed to mature for 3 weeks in a humidified atmosphere prior to experimental assessment. Cultures were stimulated in a standard 35 mm petri dish with rMS (10 Hz, 900 pulses); sham-stimulated cultures underwent the same handling but without stimulation. Whole cell voltage-clamp and current-clamp recordings were conducted 2–4 h after stimulation. High density microelectrode array (HD-MEA) recordings and current-clamp recordings were also performed on non-stimulated slice cultures to determine species-specific differences. CA1 pyramidal neurons from non-stimulated tissue cultures were post hoc stained and reconstructed using Neurolucida 360 (ver. 2019.1.3; MBF Bioscience). These reconstructions were utilized for multiscale single-cell modeling.

300 μm thick organotypic tissue cultures containing the hippocampus and the entorhinal cortex were prepared at postnatal day 3–5 from mice and rats of either sex as described previously (Vlachos et al., 2012; Galanis et al., 2021). The tissue cultures were maintained in an incubator at 35°C with 5% CO2 for at least 18 days before any experimental assessment. Tissue culture medium was changed 3 times per week and consisted of 50% (v/v) MEM, 25% (v/v) basal medium eagle (BME), 25% (v/v) heat-inactivated normal horse serum, 25 mm HEPES, 0.15% (w/v) NaHCO3, 0.65% (w/v) glucose, 0.1 mg/mL streptomycin, 100 U/mL penicillin, and 2 mm Glutamax (pH 7.3 with HCl or NaOH).

Tissue cultures were transferred in a standard 35 mm petri dish filled with standard extracellular solution (129 mM NaCl, 4 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 4.2 mM glucose, 10 mM HEPES, 0.1 mg/mL streptomycin, 100 U/mL penicillin, pH 7.4, preheated to 35°C; 365 mOsm with sucrose). A 70 mm figure-of-eight coil (D70 Air Film Coil, Magstim) connected to a Magstim Super Rapid2 Plus1 (Magstim) was placed 1 mm above the lid of the petri dish and the cultures were stimulated with a protocol consisting of 900 pulses at 10 Hz. Tissue cultures were orientated in a way that the induced electric field within the tissue was approximately parallel to the dendritic tree of CA1 pyramidal neurons. Species- and time-matched cultures were not stimulated, but otherwise identically treated served as the controls.

Whole-cell voltage-clamp recordings of CA1 pyramidal cells were conducted as previously described (Vlachos et al., 2012; Lenz et al., 2016; Galanis et al., 2021). Recordings were conducted at 35°C. The bath solution contained 126 mM NaCl, 2.5 mM KCl, 26 mM NaHCO3, 1.25 mM NaH2PO4, 2 mM CaCl2, 2 mM MgCl2, and 10 mM glucose and was saturated with 95% O2/5% CO2. Miniature α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor-mediated excitatory postsynaptic currents (mEPSCs) were recorded in the presence of 10 μM D-APV and 0.5 μM TTX in the bath solution while the patch pipettes contained 126 mM K-gluconate, 4 mM KCl, 4 mM ATP-Mg, 0.3 mM GTP-Na2, 10 mM PO-creatine, 10 mM HEPES, and 0.1% (w/v) biocytin (pH 7.25 with KOH, 290 mOsm with sucrose). Miniature inhibitory postsynaptic currents (mIPSCs) were recorded in the presence of 0.5 μM TTX, 10 μM D-APV, and 10 μM CNQX in the bath solution while the patch pipettes contained 125 mM CsCl, 5 mM NaCl, 2 mM MgCl2, 2 mM Mg-ATP, 0.5 mM Na2-GTP, 0.1 mM EGTA and 10 mM HEPES (pH = 7.33 with CsOH; 275 mOsm with sucrose). Neurons were recorded at a holding potential of −70 mV. Series resistance was monitored in 2–4 min intervals and recordings were discarded if the series resistance reached ≥30 MΩ and the leak current changed significantly.

Whole-cell current-clamp recordings of CA1 pyramidal cells were conducted at 35°C. The bath solution contained 126 mM NaCl, 2.5 mM KCl, 26 mM NaHCO3, 1.25 mM NaH2PO4, 2 mM CaCl2, 2 mM MgCl2, 10 mM glucose, 10 μM D-APV, 10 μM CNQX, and 10 μM bicuculline methiodide and was saturated with 95% O2/5% CO2. Patch pipettes contained 126 mM K-gluconate, 4 mM KCl, 4 mM ATP-Mg, 0.3 mM GTP-Na2, 10 mM PO-creatine, 10 mM HEPES, and 0.1% (w/v) biocytin (pH 7.25 with KOH, 290 mOsm with sucrose). Neurons were hyperpolarized with −100 pA and then depolarized up to +400 pA with 1-s-long 10 pA current injection steps. Recordings were discarded of the series resistance reached ≥15 MΩ.

HD-MEA recordings of mouse and rat tissue cultures were conducted at 35°C. The bath solution was similar to the one used for voltage-clamp recordings without the addition of any drugs. Cultures were placed on an Accura HD-MEA chip (3Brain, Switzerland) and acclimatized for 2 min before recording. Each tissue culture was recorded for 10 min with a BioCAM DupleX (3Brain, Switzerland).

CA1 pyramidal neurons were patched with pipettes containing 126 mM K-gluconate, 4 mM KCl, 4 mM ATP-Mg, 0.3 mM GTP-Na2, 10 mM PO-creatine, 10 mM HEPES, and 1% (w/v) biocytin (pH 7.25 with KOH, 290 mOsm with sucrose). The neurons were kept in the whole-cell configuration for at least 10 min during which they were depolarized with 100 ms current injections of 200 pA at 5 Hz. Tissue cultures were fixed in a solution of 4% (w/v) PFA and 4% (w/v) sucrose in 0.01 M PBS for 1 h and further processed and images as previously described (Galanis et al., 2021).

CA1 pyramidal cells were reconstructed using Neurolucida 360 (ver. 2019.1.3; MBF Bioscience) as described previously (Shirinpour et al., 2021). Somata were reconstructed in 2D using the contour method in order to avoid overestimation of soma volume due to the somatic whole cell recording method used to fill the cells and the high signal intensity of these compartments.

Finite element method was used to create a three-dimensional mesh model consisting of two compartments, representing the bath solution and tissue cultures. The physical dimensions of the mesh model were based on the physical parameters of the in vitro settings, with a coil-to-Petri dish distance of 1 mm and the coil positioned above the culture. Electrical conductivities of 1.654 S/m and 0.275 S/m were assigned to the bath solution and culture, respectively. The rate of change of the coil current was set to 1.4 A/ms at 1% MSO and scaled up to higher stimulation intensities. Simulations of macroscopic electric fields were performed using SimNIBS (3.2.6) and MATLAB (2023a). A validated 70 mm MagStim figure-of-eight coil was utilized in all simulations (Thielscher and Kammer, 2004). The 99th percentile of the E-field, which represents the robust maximum value, was extracted from the volume compartment of the tissue culture.

Reconstructions were imported into the NeMo-TMS pipeline and endowed with a Jarsky model (Jarsky et al., 2005). When axons are “swapped,” the original axon is removed from the cell at the point of intersection with the soma or dendrite, and replaced with the axon of another cell that has been severed at the same point. Each cell is oriented with the apical dendrite pointing in the positive y direction, and axon orientations relative to this are preserved in the swapping process. For single-cell simulations, TMS is simulated as a uniform electric field of varying intensity, with the threshold defined as the smallest TMS amplitude that elicits a somatic action potential.

Analyses were performed with the person analyzing the data blind to the experimental condition. For this project, we used one or two tissue cultures from each animal. Electrophysiological data were analyzed using pClamp 11.2 software suite (Molecular Devices), the Easy Electrophysiology 2.5.0.2 (Easy Electrophysiology Ltd.) and BrainWave (3Brain) software. Statistical comparisons were made using Mann–Whitney test (to compare two groups) two-way ANOVA and Kruskal-Wallis test as indicated in the figure captions and text (GraphPad Prism 7). p values of <0.05 were considered a significant difference. All values represent mean ± SEM.

Confocal image stacks were exported as 2D projections and stored as TIFF files. Figures were prepared using Photoshop graphics software (Adobe). Image brightness and contrast were adjusted.

A 10 Hz stimulation protocol consisting of 900 pulses at 50% MSO was used to assess the effects of rMS on synaptic plasticity in brain tissue cultures prepared from mice of either sex (Figures 1A–C). Individual CA1 pyramidal neurons were patched and AMPA receptor-mediated mEPSCs were recorded 2–4 h after stimulation. In line with our previous work (c.f., Vlachos et al., 2012; Lenz et al., 2015, 2020; Eichler et al., 2023) a significant increase in mean mEPSC amplitude was observed as compared to age−/time-matched control cultures that were treated in the exact same way except for 10 Hz rMS (control; Figures 1D,E).

In a different set of cultures, we assessed 10 Hz rMS-induced changes in GABA receptor mediated mIPSCs onto CA1 pyramidal neurons using the experimental approach described above. A reduction in mean mIPSC amplitude was observed in these experiments as reported in our previous study (Figures 1F,G; c.f., Lenz et al., 2016). These results confirm the robust effects of 10 Hz rMS on mEPSC and mIPSC amplitudes of CA1 pyramidal neurons in mouse entorhino-hippocampal tissue cultures, which are consistent with a potentiation of excitatory synapses and a depression of inhibitory synapses.

The same 10 Hz protocol (10 Hz, 900 pulses, 50% MSO) was applied to tissue cultures prepared from rat brains (Figure 2), aiming to assess the rMS effects across two similar yet distinct neuronal networks. Age-matched rat entorhino-hippocampal cultures displayed a larger cross-section than mouse tissue cultures (Figure 2A), without any apparent morphological differences in CA1 pyramidal neurons (Figure 2B). Recordings of AMPA receptor-mediated mEPSCs from CA1 pyramidal neurons showed no statistically significant differences between control and 10 Hz rMS-stimulated preparations (Figures 2C,D). Inhibitory synaptic strength was also unaffected, as no significant differences in mean mIPSC amplitude and frequency were detected 2–4 h after stimulation (Figures 2E,F).

The electric field (E-field) strength induced in the mouse and rat slice cultures was described using computational modeling (Saturnino et al., 2019). Three-dimensional mesh models were created with two compartments (i.e., bath solution and slice cultures) using the finite element method (Figure 3A). The physical dimensions of the mesh models were adapted from data obtained in mouse and rat brain issue cultures (Figure 3B). Macroscopic modeling of the E-field revealed that stimulation at 50% MSO induces a stronger electric field in the mouse (20.4 V/m) when compared to the rat tissue culture (19.3 V/m). Based on the modeling we determined that 53% MSO stimulation of rat tissue cultures would result in an E-field that is comparable to what we estimated in the mouse tissue cultures stimulated with 50% MSO (Figure 3C). Accordingly, another set of rat tissue cultures was stimulated with 53% MSO (10 Hz, 900 pulses) and AMPA receptor-mediated mEPSCs were recorded from CA1 pyramidal neurons 2–4 h after stimulation. No significant differences in mean mEPSC amplitude and frequency were observed in these experiments (Figure 3D). We conclude that simulation-based standardization of electric fields may not suffice to achieve comparable biological effects in mouse and rat CA1 pyramidal neurons, i.e., in neurons embedded in networks with comparable architectures and properties.

To test for differences in spontaneous network activity between mouse and rat entorhino-hippocampal slice cultures basal firing rates and field potential rates were recorded in a different set of 3-week-old mouse and rat tissue cultures using HD-MEA recordings (Figures 4A,B). No significant differences between mouse and rat tissue cultures were observed in firing and field potential (FP) rates in these experiments (Figures 4C–F). We conclude that baseline network activity is not responsible for the inability of rMS to induce plasticity in rat CA1 pyramidal neurons.

To investigate whether differences in CA1 pyramidal neuron size and complexity could explain the variation in rMS outcome, we reconstructed biocytin-filled and streptavidin-A488 stained CA1 pyramidal neurons from both rat and mouse hippocampal tissue cultures and analyzed their dendrites and axons (Figure 5). This was motivated by the observation that the brain sizes of mice and rats, as well as their tissue cultures, differ.

No significant differences were observed between the two groups in apical and basal dendritic length (Figures 5C,D). Sholl and diameter/volume analyses (Figures 5E–G) did not show any statistical significance between CA1 dendrites and their complexity of rat and mouse CA1 pyramidal neurons in entorhino-hippocampal tissue cultures. Similarly, no significant differences were observed when CA1 axons were reconstructed and compared in mouse and rat tissue cultures (Figures 5I–L). We conclude, that structural properties of CA1 pyramidal neurons are not statistically different and cannot explain why the rat tissue cultures do not respond to 10 Hz rMS even when the E-field is closely matched based on E-field simulations.

We assessed the impact of rMS on CA1 pyramidal neurons through a multiscale computational model that connects the physical input parameters of rMS to dendritic and axonal morphologies (Figure 6). This approach was necessary because our morphological analysis might not have encompassed distinctions pertinent to the neuronal activation induced by rMS.

When examining the dendritic architecture of CA1 neurons in mice and rats, and employing a standardized artificial axon across all cells (c.f., Aberra et al., 2018; Shirinpour et al., 2021; Eichler et al., 2023), our simulations revealed no significant difference in the depolarization threshold elicited by rMS (Figures 6A,B). Subsequently, we investigated whether axonal morphologies might underlie the observed variability in our experimental outcomes. An additional series of simulations was conducted, this time integrating the authentic axonal morphologies of these neurons. Again, no significant differences in the depolarization thresholds were observed between the two groups (Figure 6C).

A noteworthy insight emerged from these simulations, confirming previous observations (e.g., Aberra et al., 2018, 2020): the axon’s influence is pivotal in establishing the rMS-induced depolarization threshold (Table 1). We followed up on this observation, by establishing connections between the axons responsible for the lowest and highest rMS depolarization thresholds across all mouse and rat cells. Indeed, an almost 2-fold difference in the depolarization thresholds was observed in these simulations across all reconstructed neurons (Figure 6D). Yet, despite these simulation results, the dissimilarity in rMS-triggered plasticity between mouse and rat tissue cultures remained unresolved, eluding a complete explanation based solely on the interactions of dendritic and axonal morphologies.

Next, active and passive membrane properties were recorded from CA1 pyramidal neurons and analyzed. Indeed, this set of experiments identified significant differences in the passive and active properties between mouse and rat CA1 pyramidal neurons (Figure 7).

While no significant differences in membrane capacitances were noted between the two species (mouse: 90.7 ± 5.863 pF and rat: 101.7 ± 6.204 pF; Mann–Whitney test; p = 0.37; data not shown), the input resistance of mouse CA1 pyramidal neurons was significantly higher as compared to rat CA1 pyramidal neurons (mouse: 156.8 ± 11.65 MOhm and rat: 67.25 ± 4.909 MOhm; Mann–Whitney test; p < 0.001; U = 279), while the cells of both mice and rats were resting at comparable membrane potentials (Figures 7A–C). Consistently, the current–voltage (I/V) curves demonstrated that depolarizing mouse CA1 pyramidal neurons required less current compared to those in rat slice cultures.

Looking at the active membrane properties (Figures 7D–F) a similar trend was observed with the most striking differences being in the action potential induction threshold (mouse: −31.81 ± 0.877 mV; rat: −28.47 ± 0.744 mV; Mann–Whitney test; p = 0.0021; U = 794) and the first spike latency (mouse: 419.8 ± 56.03 ms; rat: 715 ± 77.36 ms; Mann–Whitney test; p = 0.0074; U = 15; data not shown). Figure 7F, shows that current injections produced stronger responses in mouse CA1 pyramidal neurons than in rat neurons, i.e., higher action potential frequencies at a lower current injection. These results indicated that mouse CA1 pyramidal neurons are more excitable than rat neurons, suggesting that higher stimulation intensities may be needed to induce rMS-induced plasticity in rat tissue cultures.

Subsequently, we tested whether a 10 Hz stimulation protocol applied at a higher intensity would induce plasticity in rat CA1 pyramidal neurons. Indeed, when rat tissue cultures were stimulated with 10 Hz rMS at 60% MSO a robust increase in the mean mEPSC amplitude was detected (Figure 8A), similar to what we observe in the mouse cultures stimulated at 50% MSO (cf., Figures 1E). In addition, a significant reduction in mean mIPSC amplitude was evident 2–4 h after rMS stimulation at 60% MSO in a different set of rat tissue cultures (Figure 8B; c.f., Figure 1G). These results demonstrate that rat CA1 pyramidal neurons do express rMS-induced plasticity, but require a higher stimulation intensity for rMS-induced potentiation of excitatory synapses and depression of inhibition to occur.