Chronic low-dose Δ9-tetrahydrocannabinol (THC) treatment stabilizes dendritic spines in 18-month-old mice

THC treatment long-lastingly increased spine density in 18-month-old, but not in 3-month-old mice

To test this idea, we repeatedly imaged dendritic segments of layer V pyramidal neurons of the somatosensory cortex in 3- and 18-month-old GFP-M male mice through a chronic cranial window. The imaging started 1 week before THC treatment, lasted throughout the 28-day THC treatment and ended 28 days after THC treatment had terminated (Fig. 1a, Extended Fig. 1). Consistent with previous studies^10,11, old mice showed increased spine density, higher spine turnover and decreased spine stability compared to young mice during baseline conditions (Extended Fig. 2). Notably, THC treatment for 28 days had no effect on spine density in young mice (Fig. 1b), but spine density increased in old mice in comparison to the vehicle-treated mice (Fig. 1c). This effect became significant at day 25 of the treatment and manifested throughout the entire post-treatment observation period (Fig. 1c). Correspondingly, THC significantly increased spine density at days 35, 42, 49 and 56 of the treatment in comparison to baseline day -1. Whereas, spine density decreased in the vehicle treated group at day 56 (post-hoc results not shown in the graph).

THC treatment altered spine dynamics differently in 3- and 18-month-old mice

Although spine density was not altered in young mice during and after THC treatment, spine dynamics showed a transient alteration. Between day 7 and 13 of the treatment, the spine turnover ratio was increased (Fig. 2a), with a rise in spine formation at day 7 and and a subsequent rapid loss of most of the recently gained spines (Fig. 2 b and c). This consequently led to an increased rate of transient, unstable spines (Fig. 2d), also reflected by a significant reduction of the probability of spine survival (Fig. 2e). In contrast, in old mice, no change in spine gain (Fig. 2g), but a reduction in spine loss (Fig. 2h) led to a lowered turnover ratio (Fig. 2f) and decreased the number of transient spines (Fig. 2i). Together, this resulted in an elevated probability of spine survival underlying the increasing spine density (Fig. 2j). Moreover, opposite to the young mice, in old mice changes were smaller in magnitude, but long-lasting, starting from around day 13–16 into the treatment. Thus, cumulatively resulting in observed elevated spine density.

Cranial window implantation

Mice were anaesthetized with an intraperitoneal (i.p.) injection of medetomidin 0.5 mg/kg, midazolam 5 mg/kg, fentanyl 0.05 mg/kg body weight (BW). 5 mg/kg BW carprofen was injected s.c. for perioperative analgesia. Mouse breathing was checked throughout the surgery, body temperature was kept constant using a heating pad. Eye ointment was used to protect from eye dryness (Bepanthen). The mouse head was epilated and fixed in a stereotaxic frame. A circular trepanation 3–4 mm in diameter was carefully done over the left somatosensory cortex exposing the dura. The brain was covered with a 5 mm glass plate (Menzel) and sealed with cyanoacrylate (UHU) and dental cement (Paladur). A small Delrin bar with threaded holes was glued on the right hemisphere to fix the animal under the microscope. After surgery the anesthesia was reversed with atipamezol 2.5 mg/kg, flumazenil 0.5 mg/kg, naloxone 1.2 mg/kg BW. The mouse was placed under a warming lamp and monitored until fully awake. Every 12 h for 3 days, 0.1 mg/kg BW buprenorphine was applied s.c. for post-surgery analgesia. After the cranial window surgery, mice were single-housed. Animals had 3–4 weeks to recover before the chronic imaging was started.

Drug treatment

Twenty eight-day releasing osmotic pumps (alzet; model 1004) were prepared according to the manufacturer’s protocol to attain a 3 mg/kg/day dose with either THC or vehicle (ethanol:cremophor:saline, 1:1:18) solutions. Osmotic pumps were implanted subcutaneously on the back of the mice under anesthesia. Mice were distributed between the treatment group and the control group to ensure equal numbers and quality of dendritic segments to analyze. The experimenter was blinded for the treatment during all imaging sessions and spine analysis.

2-photon imaging

For imaging mice were sedated with an i.p. injection of medetomidin 0.5 mg/kg, midazolam 5 mg/kg BW and the head was fixed under the microscope. The body temperature was kept constant and the eyes were protected from dehydration with eye ointment (Bepanthen). We used a custom built 2-photon microscope driven with a Ti:sapphire Laser (Chameleon Vision-S, Coherent) running at 910 nm for GFP excitation. The setup was controlled by ScanImage (Vidrio Technologies) software³⁸, running on MATLAB (Mathworks). 3–5 regions of interest (ROIs) were obtained from each mouse using a 40 × objective (NA 0.8, LumPlanFl, Olympus). Each ROI included multiple apical dendritic segments from layer V pyramidal neurons. To confirm that the dendritic segments belonged to layer II/III or V neurons, we followed dendritic segment towards the soma. Only ROIs in which spines were clearly visible were included into the analysis. To repeatedly find the same dendritic segments, blood vessels and the shape of neighboring dendrites were used as landmarks. In each time point from each imaging region one z-stack image was acquired of 1024 × 1024 pixels (0.065 μm/pixel) resolution and 16-bit depth using a 0.43 Hz frame rate resulting in a pixel dwell time of 2,000 ns. In the young group a step size of 0.51 µm was used, but in old animals it was changed to 0.8 µm to decrease the imaging time. Each frame was acquired 4 times and averaged to increase signal to noise ratio. Overview pictures were done during every imaging session to observe the stability of the window. After imaging, mice were woken up by an i.p. injection of atipamezol 2.5 mg/kg, flumazenil 0.5 mg/kg BW. Mice were placed under a warming lamp and monitored until fully awake.

Dendritic spine analysis

Spine analysis was performed using a MATLAB script “Spine Analysis”¹⁴ provided with r3.8 ScanImage³⁸. In general, 2–3 ROIs were analyzed per animal, in some cases we could only analyze 1 ROI (5 out of 26 mice) or even 4 ROIs (2 out of 26 mice). Only images that allowed for a visual identification of spines were used in the spine analysis. The number of ROIs and mice analyzed in each time point can be found in Supplementary Table ¹. Image analysis consisted of two phases: annotation and correlation. First, each spine was manually annotated as a single line from the dendritic shaft to the tip of the protrusion. Only protrusions longer than 0.4 µm were considered as spines¹⁴. Next, annotated protrusions were correlated between time points. Protrusions could fall into one of the following categories: stable, gained, lost, and transient depending on their consecutive presence or absence (Fig. 1b). Spines were defined as stable: when the element was present on both consecutive time points; gained: when the element was not there on one time point but appeared on the consecutive one; lost: when the element was gone on the consecutive time point; transient: when gained on one time point and lost in the next one. Turnover ratio was calculated as the number of lost and gained spines between two time points divided by total number of spines in both time points. Within each ROI, we have measured the total length of each dendritic segment and quantified the number of present spines within the measured dendritic segment. To obtain the spine density we divided the sum of all annotated spines by the total length of all measured dendrites within a ROI. The reported spine dynamics were calculated using a custom written Python script (https://github.com/danielmk/spine_annotations_processing).

We would like to thank Dorit Glass and Isabelle Blameuser for excellent technical support and the workshop at the Institute of Physiology II for custom-made imaging setups. We would like to thank Dr. Daniel Müller-Komorowska for the help with writing the script for spine analysis.