Led light

Dual‐stimulation experiments were performed by first stimulated with ATP and then with intracellular IP3 by uncaging a previously loaded caged IP3 in MCF10A cells that stably express genetically encoded calcium activity reporter GCaMP5 (Akerboom et al, 2012) fused to mCherry (Su et al, 2013). The mCherry was used to normalize for expression level by dividing the GCaMP signal for each cell by the average mCherry signal over the same segmentation mask. Prior to experiments, cells were first incubated in 1:2,000 dilution of 10 mg/ml Hoechst and then incubated in 1.5 μM membrane permeable caged IP3 for 1 h (Enzo Life Sciences, Yan et al, 2015). During the experiment, the cells were first stimulated with 10 μM ATP and imaged for the subsequent 15 min. After the 15‐min period, cells were allowed 35 min to recover from the ATP stimulated while still imaged to enable cell tracking. After the recovery period, cells were exposed to 365‐nm LED light (Thorlabs) for 5 s to uncage IP3. The subsequent calcium response from the cells was imaged for 5 min.

For light-evoked terminal glutamate release impinged onto SPNs, ChR2-containing corticostriatal axons in the pDMS were stimulated by pulsing an LED light (473 nm, ThorLabs, New Jersey, LED light source attached to microscope) onto the slice under 40× water-immersion objective. Continuous trains (at 0.5 Hz) of light pulses (1 ms width, interpulse interval 50 ms and 20 pulses per train) were delivered onto the slice at a maximum intensity of 3.9 mW to evoke action potentials arising from excitatory postsynaptic potentials. The stimulation protocol was chosen to mimic in vivo recordings in SPNs. For halorhodopsin experiment, EYFP positive and negative SPNs were injected repeatedly with +25, +50, or +100 pA current (duration 200 ms pulse, triggered at 0.5 Hz) to evoke action potentials. After initial 10 s baseline, LED 625 nm (1 mW) was shone onto the slice pulsing at 40 Hz for 150 s, to mimic our in vivo experimental protocol.