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Primer on Cellular and Tissue Temperature Control

Temperature Control Primer

Temperature plays a crucial role in the up and down regulation of metabolic activity.  Thus the temperature of a preparation under study is a very important consideration when doing any physiology or imaging experiment. Doing work at physiologic temperature simulates real world conditions and working below that can sometimes help slow down processes that otherwise would be too fast to study. In any event, whether temperature is the dependent or independent variable, it is important to have robust systems to adequately provide for all temperature control needs. 

Thermal needs in experiments are typically divided into two categories, above room temperature and below room temperature. Above, will require heating devices and below will require cooling devices. Heating systems typically use resistive elements to generate thermal energy.  Cooling devices are mostly derived from thermoelectric elements that take advantage of the Peltier effect whereby when an electric current is passed through a special alloy of metals known as a Peltier device, heat moves in one direction across the element so a cold heat sink can be created to provide a cooling element and remove heat. 

Heating a Chamber

Typically when cells or tissue are under study, they are confined to a small space in a Petri dish or chamber.  Usually these chambers hold a small amount of liquid in which the cells or tissue is submerged. Keeping the tissue warm means heating the chamber that the sample is in.  By default enough power must be present to heat the chamber and then also the liquid.  Heating a chamber is a static load in that loss will be limited to radiational cooling and maybe some evaporation.  Typically these loses are linear and in proportion to the temperature above ambient that the devices is heated to.  The warmer you want to keep the device, the more energy you must push into it, and the more heat loss.  Sufficient wattage must be present in the heating device and its power supply to meet the demand or the device will not get nor stay warm. 

Having sufficient power is just the first part in the heating process, the second, critical part, is feedback control. When sufficient power is present to heat a device to a temperature significantly above room temperature, there must be some sort of temperature feedback so that the device will not be heated out of range with the risk of damaging or destroying the tissue or sample. A simple thermostatic feedback that merely turns on the power to the heater when the temperature drops below set point, and then turns it off when the temperature rises above is very crude and not useful for precise scientific experiments. To this end a PID (Proportional Integral Derivative) loop is utilized so that feedback from the sensor can be analyzed and then a proportional amount of energy applied in relation to the set point required, the temperature at any given moment, and the ongoing rate of temperature change.  PID control allows the temperature of a chamber to be set at a specific level and then to rise to that level and come to a precise stop at the desired point much the way an elevator stops evenly at a particular floor.  This kind of control can only happen in a “closed loop” where the temperature, at all times, is fed back to the control algorithm so appropriate changes in power are created to insure the desired temperature is maintained.  Sensors must respond quickly, their selection and location within heated devices is critical so that they give back fast and accurate temperature information.

Sensor feedback is critical as we have stated above.  The location of a sensor within a device can have a profound effect on the accuracy and speed of its feedback.  Positioning feedback sensors within metal parts of the device and in close proximity to fluids that are to be heated is very helpful. These types of locations where the senor is in close attachment to parts that are directly influenced by the derived energy of the device serve to insure that the feedback loop will not be lost.  The above, not withstanding, at times it is preferable to have the feedback sensor located in the cell bath directly.  In such a location the feedback sensor is subject to influence by contact with the atmosphere or the interface between the cell bath and the air—a naturally cooling location due to evaporation.  Such influences may create a false reading compared to what the temperature really is where the cells or tissue are. These types of sensors are very useful for telling the temperature right at the sample, and using them as a control point is at times critical in order to get the sample to the precise experimental temperature.  However, a sensor in this position is vulnerable to false readings if it emerges from the bath due to movement or evaporation of the fluids etc. Therefore when controlling from the cell bath, precautions must be taken to keep the senor submerged and it is helpful to have a control system that can respond to a situation where the sensor looses contact to prevent over-heating and damage to the experiment or the devices in use. 

Heating Flowing Liquids

Much of the same principles listed above are relevant to heating a flowing liquid, but two additional challenges are present:  The first is that flowing liquid can absorb a lot of thermal energy, so power capacity and thermal transfer can be a challenge and the other is that the flow rate can vary which can make temperature control difficult since the energy demand will vary with the flow rate.  If perfusion heater is not made well, the liquid flowing through it may be able to pass through without absorbing the necessary energy to bring it to the desired temperature. Not only does the amount of power available need to be sufficient, but the flow path and materials in the heating device need to be able to transfer that power in the form of heat to the flowing liquid efficiently.

When a liquid is heated often its density goes down.  You may notice that when you start warming a flowing liquid, its flow rate decreases. This can affect an experiment in two ways, one, the amount of thermal energy carried by the liquid to the preparation will decrease as the mass of liquid moving to the prep is reduced, and second gasses dissolved in the liquid can come out of solution and begin to fill spaces within the device reducing the flow paths, reducing the surface area that the liquid can contact and disrupting the flow as bubbles emerge from the device. Systems must be robust and designed to handle these anomalies. 

Furthermore the location of feedback sensors within the device can have a large impact on accurate temperature control.  If a sensor is located in the beginning of the flow path, the cool liquid entering the system will pass over the sensor making the device “think” that it is colder than it actually is and the unit may actually over-heat the liquid as it tries to compensate.  It is always best if the sensor is located in the flow path at the exit of the device—but not where it can be affected by the ambient temperature.

The temperature can be measured at the output of the device as the liquid flows into a chamber.  After all, that is where it will meet the prep, so really that is the temperature that counts the most.  The only problem here is that the sensor can be influenced by the temperature of the liquid already in the chamber. The flow must be maintained and the position of the sensor in the flow stream is critical. 

Cooling a Chamber

On Earth, cooling an object is the harder task vs. heating it up.  Even though entropy would suggest that cooling is easier, our methods to remove heat from an object or area require large amounts of energy—think of an air conditioner—one of the largest users of electricity in the average home. In the laboratory when we are trying to cool a perfusion liquid or small chamber the usual method is to involve thermoelectric elements known as Peltier devices.  Peltiers use electric current to move heat in one direction across an alloy of metals.  A Peltier device has two surfaces usually made of a ceramic.  The two surfaces are in close proximity; 3-5mm apart.  As one side gets cool, the other side gets hot.  The hot side receives all the heat energy removed from the side that is getting cold, plus the heat of the current needed to do that work.  Thus, good heat-sinking and cooling is necessary for the side of the Peltier that gets warm, otherwise the cold side will not stay cold at all. Peltiers are not efficient.  They consume a lot of power and it has to be DC since they are polarized. If you reverse the polarity the side that was cold gets hot and vise versa. This is a very useful property since you can do both heating and cooling with a Peltier.  However, the close proximity of the cold to the hot surface ads to the inefficiency.   

A Peltier device and have a ΔT, the temperature difference between the hot side and cold side of as much as 80°C.  Such a large delta separated by a few millimeters is inherently inefficient.  The advantage is all that power to move heat in a small package.  In the laboratory, confined in the experimental setup, space is at a premium, but power is readily available. A standard wall outlet has more than enough power to move heat around in the small confines of a cell preparation.

Typically cells or living tissue is/are kept in a chamber that has a clear optical path and capacity to hold one to several milliliters or more of fluid. The chamber must have good surface area contact with the part of the device that will get cool. Since cooling is less efficient than heating, the surface volume is crucial.  Also, heat loss, or rather cold loss, through un-insulated surfaces is very high and much of the cooling power is lost this way.  In humid environments, the cold surfaces may sweat as moisture from the air condenses on them.  One should consider that moisture from the air can even condense in the cell bath and may dilute it—if it is not a circulating system.  In addition to robbing precious cooling calories, the moisture, as it beads up, creates a multiplier of additional surface area that robs even more cold from the system.  We cannot emphasize enough how important good contact between the chamber and the cooling device is.  Cooling is so in-efficient that care must be taken at every junction point to be sure that it is as efficient as possible.  Good fits and thermal grease are a must in many systems.

One Achilles heel of cooling with thermoelectric modules (Peltier devices) is that care must be taken to remove heat from the device.  It will not function properly unless the “hot” side is cooled to approx. room temperature or a little less.  Failure to cool it can result in permanent damage to the Peltier, but first it will go into a feed-forward state where the whole element will just get hotter and hotter until either failure or power is cut off. While flowing air can be used, water with more than three thousand times the volumetric heat carrying capacity as air, is much more efficient at removing heat and allows smaller foot-print heat exchangers to be used.  The challenge is to provide a system that circulates water and is free from leaking even a drop.  Fortunately CPU coolers made for cooling computer CPUs are well suited in terms of cooling capacity to be used for cooling temperature controlled devices in the laboratory. In some cases even a small flow from a sink can be used offering a very low cost cooing solution.  (We should mention here that devices that employ thermoelectrics can be used for heating as well, but a flow of cooling water should be maintained so that a feed-forward state does not result.)

Cooling Flowing Liquids

Cooling a flowing liquid in any environment is challenging.  As we stated above the cooling capacity of water is very high, one of the highest of any fluid on Earth and so removing heat from it requires a lot of power.  The faster it flows and the more volume the more energy.  There is no question that given enough space and power, devices for cooling in and around the microscope stage can cool things off way below the freezing point of water if necessary.  However, the limits on space and power mean that there will be limits on how much cooling we can provide in the region around the prep. 

Sometimes cooling flowing liquids will be done with a chiller bath since they have a lot of cooling capacity.  Ultimately their final temperature will be limited to several degrees above freezing since they cannot cool the water in the chiller below freezing.  Mostly, cooling liquids around the microscope stage is done using thermoelectric devices.  In this case they are attached to heat exchangers on both sides of the thermoelectric device, one side for cooling and removing heat, the other side for cooling the liquid that is headed to the prep. 

In order to remove heat from the thermoelectric devices, a steady flow of cooling water is advisable.  Air based coolers can be used, but they require so much air volume that the movement of air can cause vibrations, not to mention acoustic noise, that can even interfere with sensitive electronic recordings. A flow of tap water can be used, or better, CPU coolers developed for gaming computers can be utilized.  They are quite and efficient.  

As with heating devices, cooling devices for flowing liquids work best when the temperature sensor for feedback control is embedded within the device where it has close contact to the fluid being cooled. Cold liquids flowing through a tube seem to warm up even faster than liquids of the same volume and flow rate cool off after being heated.  Cooling is really very perishable.  Care has to be taken to keep flow paths short and insulated if possible. 

Heating and Cooling Controllers

The back end of any heating and cooling system is critical.  Good electronics with adequate power and easy controls take the chore out of temperature controlling the prep and associated fluidics. Since most heating and cooling devices for cellular physiology run on DC power, a controller that has 2.5 to 3 Amps of deliverable power is usually essential, since at 12 V that will give available wattage in the mid 20’s.  Think of that amount as a powerful soldering iron and you can picture how that heat can be used to warm up a small cell bath or a few ml/min of perfusion liquid. 

The control part of the device cannot be overlooked.  The control panel should be easy to read and intuitive to use, without too many button presses and menus etc.  An on/off switch that is easy to access should not be overlooked as this can be very important if something goes wrong and you need to kill power.  Preventing something bad from happening is also the part of a good controller.  Proper feedback on the temperature of anything that is being actively heated or cooled is critical as well. It is this information that will drive the controllers output, so the input needs to be accurate and flawless.  But alas things can go wrong; sensors de-couple from the prep or an errant voltage setting occurs, so it is important to have a controller with fail safe systems.  Some artificial intelligence that looks at a temperature situation and can determine if things seem okay, or if something is out of line and likely to overheat or freeze.  If that should happen, an alarm should alert the user to the situation, and the controller should automatically go in to a safe mode to protect the prep and the equipment from a damaging overheat or freeze situation.

Additional important features are PID control that prevents overshoot when you need a fast temperature jump.  Inputs to drive the temperature changes you need via remote device like a data acquisition board.  Also outputs that allow the temperature that is being monitored to be recorded or tracked by a data acquisition system. 

Finally, devices used in electrophysiology need to be low noise in terms of EMI and electrical noise.  Since so much power is being switched on and off to the temperature control devices that are close to sensitive electronics like patch clamp amplifiers, precautions must be taken on the part of the electronics designers to smooth out sharp electrical power jumps and fast signals in the electronics that can radiate out and be detected by sensitive amplifiers, adding noise to the recorded signal.

Primer on Perfusion Control

Perfusion Control Primer

In order to do accurate experiments on living tissue, viability must be maintained.  Cells must remain happy and healthy to yield accurate data.  The problem is that we must create an artificial environment that mimics as closely as possible the natural one that the cells came from.  Metabolic activity requires nutrients and oxygen, and the removal of wastes.  In most organisms, blood flow takes care of this requirement, when we put cells in an artificial environment, we must provide this for them.

Typically, mimicking the natural environment is done with two factors, maintaining physiological temperature and providing liquid perfusion of the cell bath.  Cells living within an organism are used to constant blood flow so placing them in an environment that differs from the constant supply of nutrients and removal of waste can be stressful.  Temperature is also a factor, mammalian cells like to be warm.  Systems that provide perfusion must also be capable of addressing the temperature control issues.

In most living organisms a muscular organ takes care of providing the blood flow.  In the lab, we typically replace that with a pump or a gravity fed source of liquid.  Such a system will run at room temperature unless we artificially heat or cool it.  In the paragraphs below, we will address the issues of liquid flow and temperature control.

Gravity Flow

The easiest flow to obtain on earth is gravity flow.  All liquids flow down hill, so all you have to do is suspend a reservoir of liquid above the place where you want it to wind up and it can easily be made to get there with no mechanical intervention.  Gravity flow is very nice because it is smooth and the lack of pumps means no electrical noise or vibration.  One disadvantage is that as the reservoir empties the head pressure decreases.  This is because the total height of the liquid is what sets the pressure in gravity systems.  There are ways to keep the head height level by using an inverted bottle that drains into the primary reservoir as the fluid height goes down and air can flow into the mouth of the bottle, releasing more liquid into the primary reservoir. 

If the reservoir in a gravity system is set high enough, then as the fluid level decreases, the percentage of the overall height of the reservoir is less, so this small change in height will not have a big effect on flow.  When reservoirs are low to the table, then a small change in fluid height can have a big impact on flow rate.  In some case these differences in speed can be compensated for by a flow restriction valve.  The valve can act as an integrator to help keep the flow rate the same as the fluid height changes.  Generally gravity systems do not have enough head pressure to make these kinds of valves work very well at maintaining flow as the fluid height changes.  The orifice needs to be quite small to really integrate flow over a wide range.  If the orifice gets too small flow slows down and can only be overcome by adding pressure to the system to push the fluid through.

Pumped Flow

There are a variety of pumps that can be used in typical experiments.  Gear pumps, vain pumps with impellers and diaphragm pumps are not typically used.  They can damage proteins in fluids by the harsh action of their pump actuators.  In the lab, the most typical pump is the peristaltic pump followed by the syringe pump.  The peristaltic pump works by employing a series of rollers that impinge on a tube to cause a bolus of liquid to be trapped in the soft tubing typically used in the pump.  As the pump rotates, bolus after bolus of liquid are moved along.  Peristaltic pumps can be large or very small, using tubing with only a 1mm bore.  The tubing must be soft and flexible. One problem with these pumps is that the flow always has a small pulsatile rate to it.  As each bolus is released by the pump the flow rate changes slightly and observing the flow on a flow meter will show a small oscillation of flow.  While some tissue may be very tolerant of this pulsation, being used to it in the host organism, the pulsation can be problematic for recording electrical signals as capacitance of the bath can change as the volume shifts with each pulse.  Generally peristaltic pumps can provide almost any flow rate desired.  Each pump usually has a pretty good range of flow rates with any particular tubing size, and then a change of tubing can bring a whole new range of flows.

Syringe pumps are also very reliable for moving liquids.  A mechanical armature powered by a motor with a lead screw, is used to either compress or extend the plunger of a typical injection syringe.  Syringe pumps can be very accurate and a typical feature is the ability to set the flow rate in mL/sec or minute.  Often using a small syringe gives more accurate flow, but the limitation of the pump is that when the syringe is empty, the experiment must be paused to replenish the syringe.  Often times, especially with larger syringes, there is a tendency for the plunger to stick or flex a little which can result in uneven flow rates.  Syringes with large volumes of liquid inside may have a tendency to out gas forming a bubble in the syringe that displaces extra solution out of the syringe above the desired flow rate.

Metering Pumps

Metering pumps are designed to pump a set volume of liquid.  Typically there is a plunger in a fixed size cavity.  Each time the plunger oscillates in the cavity, a fixed amount of liquid moves through the pump, keep track of the cycles and you know how much volume has been moved.  They can be very accurate, but are better at volumes larger than what is typically used on the bench for a small cell chamber.  The reciprocating motion of the pump can make it un-friendly for electrophysiology as the pulsations can be large..  But in any application where continuous flow of a known volume is required, metering pumps rule.

Regulatory Devices for Flow Control

Typically a valve is used to regulate flow, and in this case it is either a valve that completely blocks flow when closed, or a restrictive valve that can limit flow.  Valves that completely stop flow are used most often.  Sometimes these valves make a pulse of fluid when they open or close.  This pulse can be disruptive to an experiment if it causes a mechanical disturbance to a recording electrode or a cell membrane that is sensitive to vibration or movement.  Precautions should be taken to lessen the pressure pulse by dovetailing the closing of one valve directly with the opening of another, or reducing the tubing size to squelch the pulse (this can reduce flow rate) or opening and closing the valve at a reduced speed.

Variable flow valves can be used to help regulate flow, but their cost usually makes them prohibitive.  Also, controlling them electronically can be difficult since it often requires an analog voltage that needs to be varied in order to operate the valve.  Flow control valves are not fast, but they can be the most gentle way to start and stop a flow.

Another way to control flow is with a restricted orifice.  A restricted orifice forces the liquid to pass through a certain size hole that will automatically reduce the flow rate.  Usually the orifice has a very precise ID and thus the flow rates of various liquids passing through the orifice can be calculated if the density and pressure on the liquid are known.  One very nice way to implement accurate flow control is to incorporate a device that restricts flow and then use pressure to push the liquid through.  This gives a system where the flow rate will be directly proportional to the pressure being applied.

Two Electrode Voltage Clamp of Oocytes

The voltage clamp technique is a method that allows ion flow across the cell membrane to be measured as an electric current as the transmembrane potential is held under constant experimental control with a feedback amplifier. Ion channels expressed in Xenopus oocytes can be studied using the two-microelectrode voltage clamp. The membrane of the oocyte is penetrated by two microelectrodes, one for voltage sensing and one for current injection. The membrane potential as measured by the voltage-sensing electrode and a high input impedance amplifier is compared with a command voltage, and the difference is brought to zero by a high gain feedback amplifier. The injected current is monitored via a current-to voltage converter to provide a measure of the total membrane current.

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Macropatch Recording

The technique for macropatching is similar to tight seal single-channel recording. The electrodes are pulled to larger tip diameters than would be used for single-channel recording; however, with single-channel recording, following initial contact with the membrane, suctions is applied to form an electrically and mechanically tight seal in gigohm range. Recordings can be obtained in the cell-attached configuration or from inside-out or outside-out membrane patches.

The objective, for many laboratories using this technique, is to record from specific areas of the membrane relative to known specializations, for example, the distribution of the ion channels in the pre- or postsynaptic membrane, dendritic vs somatic membranes. Macropatch electrodes have resistances in the range 0.5-3MOhms and are in the order of 2-8um diam. Our range of Axon and NPI patch clamp amplifiers are particularly suited for such recordings.

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Single-Channel Recording

Single-channel recording is achieved by pressing a fire-polished glass pipette, which has been filled with a suitable electrolyte solution, against the surface of a cell and applying light suction. Under such conditions, the glass pipette and the cell membrane will be less than 1 nm apart. The tighter the seal will have two advantages, 1) better electrical isolation of the membrane patch and 2) a high seal resistance reduces the current noise of the recording, permitting good time resolution of single channel currents, currents whose amplitude is in the order of 1 pA. Classically, three different configurations of the patched membrane can be used for single-channel recording: cell-attached, outside-out and inside-out patches. Cell-attached configuration contacts the cell membrane forming a gigaohm seal. Long-term stable recordings with low background noise can be performed in this configuration with minimal disruption to the intracellular milieu. For the outside-out configuration, the external surface of the patch is exposed to the external recording media. Offering the opportunity to repetitively expose the channels to different drugs and at various concentrations. In the inside-out patch configuration, it is the internal face of the membrane that is exposed to the external solution. This provides access to intracellular receptor binding sites and also enables studies of intracellular signaling pathways.

It is now possible to record single-channel current activity from many cell types, that is, from mammalian species, insects, invertebrates and also plants. The recording of single-channel currents enables detailed kinetic analyses of native and recombinant ion channels, including those that have been subject to natural or intended mutations to their structure.

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Whole-Cell Recording

Whole cell recording is the most commonly used configuration of the patch clamp technique. It is achieved by rupturing the patch of membrane isolated by the patch pipette bringing the cell interior into contact with the pipette interior. Using the whole cell patch clamp design of experiment one can then record the electrical activity of the entire cell via several modes. Voltage clamp, where the potential difference across the cell membrane is controlled and current measured, or current clamp, controlling the current and measuring the voltage across the membrane are the two main modes of whole cell recording. These recording configurations are very powerful techniques in the study of ion channel activity, aspects of neuronal behaviour and synaptic transmission. Our range of Axon and NPI patch clamp amplifiers are perfect for carrying out whole cell patch clamp recordings.

One major problem in whole recording is series resistance. Employing the ‘Discontinuous single electrode voltage clamp’ (dSEVC) technique is a very useful procedure to overcome the series resistance. The dSEVC separates the current injection from potential measurement in time, by rapid switching between a current injection mode and potential measuring mode. This ensures that no current passes through the resistor created at the pipette/cell interface during potential recording and completely eliminates series resistance artifacts. Provided the switching frequency between the current injection- and voltage measuring-mode is high enough, the plasma membrane can be clamped to a steady membrane potential. Our NPI SEC range of amplifiers are designed specifically for dSEVC being the fastest and most accurate single-electrode current – and voltage – clamp systems available.

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Cardiac Rhythmicity

The spontaneous depolarization and repolarization events that occurs in a repetitive and stable manner within the cardiac muscle is often abnormal or lost in cases of cardiac dysfunction or cardiac failure. The underlying mechanisms of this rhythmicity are based on the myriad of voltage dependent ion channels found in cardiac myocytes. These ion channels can then be studied at the single channel, single cell or tissue level using various techniques and equipment supplied by ALA Scientific Instruments.

The use of the patch clamp technique is very powerful at the single channel and single cell (whole cell) level. Such amplifiers as the manufactured by Axon and NPI are ideal. In addition, ALA Scientific Instruments range of perfusion systems make cardiac experiments very easy. At the tissue level, multielectrode arrays are capable of recording from distinct regions of cardiac slices simultaneously.

In Vitro Cell and Tissue Preparations

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In Vivo Multielectrode Recording

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Systems Neuroscience

The function of neuronal circuits and systems can only be studied with preparations and experimental models where neurons are connected together to form neural networks. In vitro preparations such as acute brain slices and organotypic slices are ideal models from which to investigate neural circuits. Acute slice and organotypic preparations are used in conjuction with multielectrode arrays to record from many neurons, or populations of neurons, simultaneously. Experimenters can utilize these systems for diverse projects including, long term potentiation in hippocampal slices, micro electroretinograms from retinal explants or spontaneous rhythmic activity from cardiomyocyte cultures.

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Gap Channel Conductance

Gap junctions are formed between the opposing membranes of neighbouring cells. Hemichannels (connexons)on each side dock to one another to form conductive channels between the two cells. The main role of gap junctions is in the electrical synaptic transmission of the nervous system allowing direct and rapid communication between neurons. The measurement of the electrophysiological properties of gap junctions is carried out using the dual whole cell voltage clamp technique. The dual voltage clamp method is the common method to assay the electrical properties of gap junctions, such as junctional conductance and transjunctional voltage dependence.

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Exocytosis

Exocytosis is the process in which the contents of secretory vesicles are released by their fusion to the plasma membrane and plays a vital role in signaling of the nervous and endocrine system. Exocytosis can be detected by several physical and chemical means. By chemically oxidizing the released secretory products at a fixed electrode potential, carbon fiber amperometry provides excellent temporal and spatial resolution in detecting exocytosis. Among the most intensively studied products include, norepinephrine, epinephrine, dopamine, and serotonin. ALA can provide full systems suitable for recording this process. The VA-10 amplifier is a sensitive (picoampere range) current amplifier that is specifically designed for voltammetric and amperometric measurements with carbon-fiber microelectrodes in biological systems. It was designed at the Max-Planck-Institute for Experimental Medicine in Göttingen as an economically priced alternative to do-it-yourself systems and expensive commercial systems.

It can be used for either DC amperometry using the built-in voltage source, or it can be operated with user-supplied external voltage waveforms (e.g. for cyclic voltammetry). The VA-10X is ideally suited for measurements from single cells plated onto glass cover slips and with carbon-fiber disk electrodes having diameters of 10 µM or less. Carbon fibre electrodes, CFE-1, are hand made at ALA Scientific Instruments and individually tested to ensure viability.

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Ion Channel Pharmacology

Ion channels are involved in many cellular processes, drugs acting on ion channels have long been used for the treatment of many diseases, especially those affecting electrically excitable tissues. Combined with advancements in high throughput screening of ion channels in drug discovery, elucidating ion channel pharmacology has grown into an enormous global task. Because of the high information content, voltage clamp is the best way to study ion-channel function which is superbly provided by the Axon and NPI systems.

Ion channel pharmacology is ultimately dependent upon the application of the particular drug in question. Such drug application is provided depending on your needs. Computer controlled local perfusion is provided by ALA Octaflow and VC3 systems. Your drug of choice can also be applied to the internal of cells during whole cell recording using the ALA 2PK+ system. A wide variety of manifolds, perfusion chambers and other accessories allows the ion channel pharmacologist to easily design a system to suit their requirements.

Classical Compound Application

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Pipette Internal Solution Exchange

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Brain/Spinal Cord Slice Recording

Techniques developed in 1989 by two groups (Edwards et al & Blanton et al) allowed direct patch clamping from neurons in acute slices of brain and spinal cord. This technical achievement overcame the problems of using neurons in culture which had been treated with enzymes and displayed altered gene expression and synaptic functions. The use of brain and spinal cord slices have since been extensively used to answer some fundamental questions in neurobiology such as the fundamental mechanism of inhibitory and excitatory neurotransmission as well as identifying single channel characteristics of ion channels found natively in the brain.

Many ALA products are suited for brain slice electrophysiology. Constant perfusion is a key requirement for the viability of slices and the ALA VC3 perfusion systems are ideal for controlling the change in bathing solutions during experiments. Extracellular field recordings are ideally carried out using NPI EXT-02F extracellular amplifier and ISO-Stim Isolated Stimulator combination.

Whole cell recordings are routinely carried out in slices using Axon MuiltiClamp 700A or Axopatch 200B amplifiers or the NPI ELC or SEC amplifiers. Slices are also the most common preparation used with multielectrode array experiments.

Brain Slice Recording Signals

Multi Electrode Array Recording 

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Extracellular Using Micropipettes

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Retinal Physiology

Classical Extracellular Recording

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Multielectrode Array Recording

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Invertebrate Preparations

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