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Patch clamping is the gold standard measurement technique for cell-type characterization in vivo, but it has low throughput, is difficult to scale, and requires highly skilled operation. We developed an autonomous robot that can acquire multiple consecutive patch-clamp recordings in vivo. In practice, 40 pipettes loaded into a carousel are sequentially filled and inserted into the brain, localized to a cell, used for patch clamping, and disposed. Automated visual stimulation and electrophysiology software enables functional cell-type classification of whole cell-patched cells, as we show for 37 cells in the anesthetized mouse in visual cortex (V1) layer 5. We achieved 9% yield, with 5.3 min per attempt over hundreds of trials. The highly variable and low-yield nature of in vivo patch-clamp recordings will benefit from such a standardized, automated, quantitative approach, allowing development of optimal algorithms and enabling scaling required for large-scale studies and integration with complementary techniques. NEW & NOTEWORTHY In vivo patch-clamp is the gold standard for intracellular recordings, but it is a very manual and highly skilled technique. The robot in this work demonstrates the most automated in vivo patch-clamp experiment to date, by enabling production of multiple, serial intracellular recordings without human intervention. The robot automates pipette filling, wire threading, pipette positioning, neuron hunting, break-in, delivering sensory stimulus, and recording quality control, enabling in vivo cell-type characterization.

The activities of groups of neurons in a circuit or brain region are important for neuronal computations that contribute to behaviors and disease states. Traditional extracellular recordings have been powerful and scalable, but much less is known about the intracellular processes that lead to spiking activity. We present a robotic system, the multipatcher, capable of automatically obtaining blind whole-cell patch clamp recordings from multiple neurons simultaneously. The multipatcher significantly extends automated patch clamping, or 'autopatching', to guide four interacting electrodes in a coordinated fashion, avoiding mechanical coupling in the brain. We demonstrate its performance in the cortex of anesthetized and awake mice. A multipatcher with four electrodes took an average of 10 min to obtain dual or triple recordings in 29% of trials in anesthetized mice, and in 18% of the trials in awake mice, thus illustrating practical yield and throughput to obtain multiple, simultaneous whole-cell recordings in vivo., Competing Interests: SK, FF, GH, AS, XH, EB, EB, CF No competing interests declared, (© 2018, Kodandaramaiah et al.)

In vivo studies of neurophysiology using the whole cell patch-clamp technique enable exquisite access to both intracellular dynamics and cytosol of cells in the living brain but are underrepresented in deep subcortical nuclei because of fouling of the sensitive electrode tip. We have developed an autonomous method to navigate electrodes around obstacles such as blood vessels after identifying them as a source of contamination during regional pipette localization (RPL) in vivo. In mice, robotic navigation prevented fouling of the electrode tip, increasing RPL success probability 3 mm below the pial surface to 82% ( n = 72/88) over traditional, linear localization (25%, n = 24/95), and resulted in high-quality thalamic whole cell recordings with average access resistance (32.0 MΩ) and resting membrane potential (-62.9 mV) similar to cortical recordings in isoflurane-anesthetized mice. Whole cell yield improved from 1% ( n = 1/95) to 10% ( n = 9/88) when robotic navigation was used during RPL. This method opens the door to whole cell studies in deep subcortical nuclei, including multimodal cell typing and studies of long-range circuits. NEW & NOTEWORTHY This work represents an automated method for accessing subcortical neural tissue for intracellular electrophysiology in vivo. We have implemented a novel algorithm to detect obstructions during regional pipette localization and move around them while minimizing lateral displacement within brain tissue. This approach leverages computer control of pressure, manipulator position, and impedance measurements to create a closed-loop platform for pipette navigation in vivo. This technique enables whole cell patching studies to be performed throughout the living brain., (Copyright © 2017 the American Physiological Society.)

Many experimental biological techniques utilize hollow glass needles called micropipettes to perform fluid extraction, cell manipulation, and electrophysiological recordings For electrophysiological recordings, micropipettes are typically fabricated immediately before use using a "pipette puller", which uses open-loop control to heat a hollow glass capillary while applying a tensile load. Variability between manufactured micropipettes requires a highly trained operator to qualitatively inspect each micropipette; typically this is achieved by viewing the pipette under 40-100x magnification in order to ensure that the tip has the correct shape (e.g., outer diameter, cone angle, taper length). Since laboratories may use hundreds of micropipettes per week, significant time demands are associated with micropipette inspection. Here, we have automated the measurement of micropipette tip outer diameter and cone angle using optical microscopy. The process features repeatable constraint of the micropipette, quickly and automatically moving the micropipette to bring its tip into the field of view, focusing on the tip, and computing tip outer diameter and cone angle measurements from the acquired images by applying a series of image processing algorithms. As implemented on a custom automated microscope, these methods achieved, with 95% confidence, ±0.38 µm repeatability in outer diameter measurement and ±5.45° repeatability in cone angle measurement, comparable to a trained human operator. Accuracy was evaluated by comparing optical pipette measurements with measurements obtained using scanning electron microscopy (SEM); optical outer diameter measurements differed from SEM by 0.35 ± 0.36 µm and optical cone angle measurements differed from SEM by -0.23 ± 2.32°. The algorithms we developed are adaptable to most commercial automated microscopes and provide a skill-free route to rapid, quantitative measurement of pipette tip geometry with high resolution, accuracy, and repeatability. Further, these methods are an important step toward a closed-loop, fully-automated micropipette fabrication system.

Whole-cell patch clamping in vivo is an important neuroscience technique that uniquely provides access to both suprathreshold spiking and subthreshold synaptic events of single neurons in the brain. This article describes how to set up and use the autopatcher, which is a robot for automatically obtaining high-yield and high-quality whole-cell patch clamp recordings in vivo. By following this protocol, a functional experimental rig for automated whole-cell patch clamping can be set up in 1 week. High-quality surgical preparation of mice takes ∼1 h, and each autopatching experiment can be carried out over periods lasting several hours. Autopatching should enable in vivo intracellular investigations to be accessible by a substantial number of neuroscience laboratories, and it enables labs that are already doing in vivo patch clamping to scale up their efforts by reducing training time for new lab members and increasing experimental durations by handling mentally intensive tasks automatically.

Amplification of multiple unique genetic targets using the polymerase chain reaction (PCR) is commonly required in molecular biology laboratories. Such reactions are typically performed either serially or by multiplex PCR. Serial reactions are time consuming, and multiplex PCR, while powerful and widely used, can be prone to amplification bias, PCR drift, and primer-primer interactions. We present a new thermocycling method, termed thermal multiplexing, in which a single heat source is uniformly distributed and selectively modulated for independent temperature control of an array of PCR reactions. Thermal multiplexing allows amplification of multiple targets simultaneously-each reaction segregated and performed at optimal conditions. We demonstrate the method using a microfluidic system consisting of an infrared laser thermocycler, a polymer microchip featuring 1 μl, oil-encapsulated reactions, and closed-loop pulse-width modulation control. Heat transfer modeling is used to characterize thermal performance limitations of the system. We validate the model and perform two reactions simultaneously with widely varying annealing temperatures (48 °C and 68 °C), demonstrating excellent amplification. In addition, to demonstrate microfluidic infrared PCR using clinical specimens, we successfully amplified and detected both influenza A and B from human nasopharyngeal swabs. Thermal multiplexing is scalable and applicable to challenges such as pathogen detection where patients presenting non-specific symptoms need to be efficiently screened across a viral or bacterial panel.

Quantitative PCR (qPCR) techniques have become invaluable, high-throughput tools to study gene expression. However, the need to measure gene expression patterns quickly and affordably, useful for applications such as stem cell biomanufacturing requiring real-time observation and control, has not been adequately met by rapid qPCR instrumentation to date. We report a reverse transcription, microfluidic qPCR system and its application to DNA and RNA amplification measurement. In the system, an environmental control fixture provides mechanical and thermal repeatability for an infrared laser to achieve both accurate and precise open-loop temperature control of 1 μl reaction volumes in a low-cost polymer microfluidic chip with concurrent fluorescence imaging. We have used this system to amplify serial dilutions of λ-phage DNA (10(5)-10(7) starting copies) and RNA transcripts from the GAPDH housekeeping gene (5.45 ng total mouse embryonic stem cell RNA) and measured associated standard curves, efficiency (57%), repeatability (~1 cycle threshold), melting curves, and specificity. This microfluidic qRT-PCR system offers a practical approach to rapid analysis (~1 h), combining the cost benefits of small reagent volumes with the simplicity of disposable polymer microchips and easy setup., (Copyright © 2013 Elsevier B.V. All rights reserved.)