Whole‐Cell Recording In Vivo

Michael R. DeWeese1

1 Cold Spring Harbor Laboratory, Cold Spring Harbor, New York
Publication Name:  Current Protocols in Neuroscience
Unit Number:  Unit 6.22
DOI:  10.1002/0471142301.ns0622s38
Online Posting Date:  January, 2007
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In vivo whole‐cell patch‐clamp recording provides a means for measuring membrane currents and potentials from individual cells in the intact animal. Patch‐clamp methods have largely been developed in vitro. This body of work has contributed enormously to the understanding of many important phenomena in excitable cells—including synaptic plasticity in the mammalian central nervous system, and the behavior of individual protein channels. In recent years, an increasing number of groups have applied whole‐cell recording techniques in the intact animal. Such in vivo studies offer the tantalizing possibility of uncovering the underlying principles and mechanisms of neural interactions within the natural context of fully intact biological networks. This unit focuses on strategies for overcoming the specific technical challenges posed by in vivo whole‐cell recording. A straightforward procedure is described for obtaining whole‐cell records from the cortex of the anesthetized rat; this procedure has also been applied successfully to awake animals and other rodent species with minor modifications.

Keywords: whole‐cell; patch clamp; in vivo; intracellular recording; cortex; neuron; rat

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Table of Contents

  • Strategic Planning
  • Basic Protocol 1: In Vivo Whole‐Cell Patch‐Clamp Recording
  • Reagents and Solutions
  • Commentary
  • Literature Cited
  • Figures
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Basic Protocol 1: In Vivo Whole‐Cell Patch‐Clamp Recording

  • Physiological buffer (see recipe)
  • Normal saline: 0.9% (w/v) NaCl
  • Rats (male or female, post‐natal day 17 to 30; Sprague‐Dawley)
  • General anesthetic (e.g., 60 mg ketamine/0.5 mg medetomedine per kg)
  • Internal solution (potassium‐ or cesium‐based, see reciperecipes)
  • Agarose solution (1% to 2% agarose by weight in physiological buffer; Type III‐A, A9793, Sigma), melted and kept up to 4 hr at ∼40°C
  • Electrode puller (e.g., Narishige 2‐stage vertical puller)
  • Electrode glass (e.g., filamented, fire‐polished, thin‐walled, borosilicate electrode glass 3 in. (75 mm) length, 1.5 mm o.d., World Precision Instruments)
  • Dissecting microscope
  • Patch pipet storage container with cover
  • Disposable 1‐ml and 30‐ml syringes (for anesthesia and pipet pressure control, respectively)
  • Disposable syringe needles: 25‐G (for rats) or 27‐G (for mice)
  • Temperature controller with heating pad and rectal thermometer
  • Stereotaxic frame (for rats) that allows access to desired cortical region
  • Cotton swabs
  • High speed pneumatic dental drill
  • Gel foam sponges
  • Dural hook
  • Recording chamber, electrically shielded
  • Computer‐based data acquisition/analysis system, including A/D board and software (see units 6.1& 6.6)
  • Silver ground wire coated with AgCl at tip (e.g., model E201Ag‐AgCl pellet; Axon Instruments)
  • Amplifier with headstage (e.g., Axopatch 200B from Axon Instruments)
  • Micromanipulater for headstage (e.g., MP‐285 model, Sutter Instruments)
  • Small plastic alligator clip
  • Pipet holder with silver electrode wire coated with AgCl at tip
  • Tubing for pressure control (made of hard plastic, ∼3 mm o.d.)
  • Three‐way valve
  • Pressure gauge (e.g., DPM‐1B model, Bio Tek Instruments)
  • Additional reagents and equipment for injection of rodents ( appendix 4F) and patch‐clamp techniques (units 6.1, 6.3, 6.6, 6.7, 6.10, & 6.16)
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Literature Cited

Literature Cited
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