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When you would have to move your patch-clamp lab in a new-to-be-built lab, how would you take action to minimize noise levels later on during measurements?
So for example, how can you make sure the PCR machine or freezer in the lab next door does not cause noise in your setup?
I guess an independent fuse box is the minimum. But what else? Hoping for some answers without losing me in too technical jargon :-)
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There are those who take fancy precautions, like enclosing their experiment rooms in Faraday cages, or planting dedicated copper stakes for grounding in their basements. In my opinion, that isn't really necessary. I think it is most practical to just set the rig up, and de-noise using standard methods. The basic idea is to tie every piece of equipment to a common ground, avoiding ground loops. For example, if the air table serves as the common ground, then all devices ground to that, not to each other. We do use Faraday cages to shield our in vivo and in vitro rigs. They help a lot. It takes fiddling. Best of luck.
-Matthew
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I need training in rodent's brain electrophysiology.
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Then probably your best options are either to attend a school/workshop that also teaches ephys (but you would gain only limited experience), or to look for a postdoc in which you can combine both ephys and also the techniques in which you are already trained. Good luck! :)
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I have doubts about which is the best and most suitable amplifier. We own the AxoClamp 2B and the Axopatch 200B. Thanks a lot for the help.
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Depends on your budget. If that's not a problem, get yourself an Axopatch 700B. You can do more than just field potentials. You can, when needed, do whole-cell experiments (say, when buffering internal Calcium with EGTA or Bapta, blocking a channel from the cytosolic side...etc.,) and unitary activity channel recordings (all configurations: C-A, I-O or O-O). Furthermore, you can perform , ion-selective electrode recording, amperometry/voltammetry and bilayer recordings.
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I am planning to perform in vitro electrophysiology experiments on rats model. Is it possible to store whole rat brain for invitro electrophysiology experiments or one have to perform the experiment on fresh tissue. How long such samples can be stored and at what temperature before performing experiment and what kind of reagents/solutions are required for storage.
Kindly guide.
Thank you
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I agree with Max Anstötz in general. Mammalian brain tissue requires continuous temperature control, oxygenation, nutrient supply, and waste removal if you want to study electrical activity that resembles physiological. On the other hand, there are ways to keep brain slices in culture for up to several months, through the tissue "flattens out," and the neuronal phenotypes may drift over time as some synaptic connections are lost and others change. You may want to look at the work by the Gähwiler laboratory in Switzerland.
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Hello everyone,
I am trying to record single channel events on my HEK293 cells stably expressing Cav2.2 channels, using the cell-attached patch clamp technique.
I used the following solutions:
Extracellular: 150 KCl, 10 HEPES, pH 7.3 with KOH
Pipette solution: 110 BaCl2, 10 HEPES, pH 7.4 with KOH
Unfortunatelly, I haven't been able to find the channels so far.
- I want to step the cell to 0, 10, 20 and +30 mV from a holding potential of -100 mV. I understand that to hold the membrane at -100 mV I have to set the V-command in my protocol to +100 mV (and steps to 0, -10, -20, and -30 mV). Is this correct?
- In that case, the currents I expect are patch inward currents (barium ions flowing from the pipette into the cell). I understand these currents should appear as outward currents (positive) in my recordings, is that right?
- Should I invert the polarity of both voltage and currents for analysis and representation?
- Finally, I am trying to use the Patch mode in my Axopatch 200B amplifier, but every time I try this a regular spike noise will appers. Do you know why this might be happening?
Thank you very much for your attention!
Diego
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I am also trying to get some single channel recordings of the Nav.
The solutions look fine in my opinion, but the protocol in my opinion is not correct.
The holding that you have in figure 2 is 0mV and not +100mV. If I am not mistaken in the Clampex software when you create a new protocol you decide the holding potential under the option outputs and there you put the desired holding.
Afterwards in the option waveform you can decide the first step and the delta level.
And yes the current you expect to see in the raw recording should be "positive".
About the regular spikes in the Patch mode, I was also wondering and decided to ask the Axon support and this was their reply "Instead of resistor, a feedback capacitor is used in the Patch mode in Axopatch 200B amplifier. The voltage across the feedback capacitor cannot ramp in one direction forever. At some point the capacitor voltage will approach the supply limits and the integrator must be reset to start again near zero volts. Thus the current record must be interrupted for 50 us while the integrator and differentiator reset. The frequency of resets depends on the current passing through the headstage, with the larger current requiring more frequency resets."
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I think both are used mainly for blocking GABAA receptors. Since gabazine is more selelctive for GABAA receptors, is it better to use for studying EPSC? Is it enough to block IPSC by only blocking GABAA receptors? Thanks.
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Hello Lingdi!
Of these two drugs, gabazine is more convenient to use, as it is water soluble and requires a smaller concentration to block the evoked IPSC (in rat cortical slices we use 5-10 uM of gabazine or 50-100 uM of PTX). However, gabazine is more expensive than PTX.
Bicuculline methiodide also blocks the GABAaR-mediated IPSC (it is water soluble and is cheaper than gabazine). Note that it affects the SK-channels.
If the IPSC in your preparation is mediated by both GABAaRs and GABAbRs you should also apply CGP-55845 or use the pipette solution with cesium salts.
Good luck with your experiments!
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Hi everyone,
I have been trying to get a gigaseal from primary cell-culture cells from rat’s DRGs, but without success. I don’t know exactly what the problem is.. I have tried after 2 hours form getting the cells, after 24 hours, and 48 hours, I have changed the glass used to fabricate the pipettes, (I tried both thin walled glass and thick walled glass), I have tried with different pipettes resistances (from 2MΩ to 10 MΩ) but without success.
A PDF that I prepared is associated to explain what I have done, containing screenshots of what I get in the “SutterPatch” Program, and picture of the pipettes I used.
Do you have any recommendations or a solution to help forming gigaseal ?
Thank you in advance!
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Thanks very much, we will try this and see what we will get
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I'm finding it really hard to get a gigaohm seal when recording from CHO cells. Are they very finicky? Is there anything I should know about them? I have no problem patching from cultured neurons using the same solutions (Ringers external and KMeSulfonate internal) or pipettes, so guessing that it is the cells that are the issue. Thanks!
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I did patch CHO cells when I was in S. Korn Lab (UCONN). I didn’t stay long, but his students: P. Andalib, J. Trapani and J. Consiglio were using them routinely with HEK293 too without any particular problem. I suggest you google Trapani’s name along with CHO and look at some of their work regarding Kv channels. Good luck!
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Can anybody recomend me any solution or way how to clean Ussing chambers? The problem is we've got a signal even from chambers without dextran (which has been used to measure dextran permeability previously).
Thank you    
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We wash properly with hydrochloric acid following rinsing with distilled water
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Dear All,
I am wondering, could anyone help me out with a manual or datasheet with specs for this good ol` beast the RK 300 from Bio-logic? It would be very helpful and very much appreciated!
Thanks in advance for taking the time!
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We did have photocopy manual for biologic 400.
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Dear fellow electrophysiologists,
We currently have a Sutter p-1000 micropipette puller in our lab that we use to pull pipettes for patching cultured neurons (R 2.5 - 4.5 mOhm and short taper). However, we are having a lot of stability issues with this puller, most likely due to daily heavy use.
So now we are considering buying a p-2000 laser-based puller, but unfortunately we cannot get it for a trial period.
Therefore, I was wondering if anyone who is using a p-2000 (and perhaps also has experience with other puller systems) could share some experiences, or give some thoughts on the following questions; What is your experience with the P-2000? Is it stable over time? Would you recommend it given its considerably higher price?  Is there a limit to the number of pulls per day?
Thank you very much!
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Dear Maaike,
Consistency of the pull results, or rather lack thereof, is the most common issue in puller tech support. It can have a variety of reasons, either mechanical issues, or inappropriate choice of glass or the pull parameters. For some of the mechanical issues, I made a diagnostic video a few years ago: https://www.youtube.com/watch?v=75y3dfGwP58 You also want to make sure your drierite still has some blue left: https://www.youtube.com/watch?v=BLvYLLJmcnY
As for the pull parameters, be sure to follow the Pipette Cookbook. The latest revision can be downloaded from the Sutter web site. In a nutshell, you start with the program recommendation in the Cookbook, find the midpoint velocity (p. 30), then write the program out into individual lines (using line repeats on the P-1000) and fine tune your tip size on the last line. If that does not get you to consistent pull results, fill out the 15 questions worksheet on the support page (https://www.sutter.com/PDFs/15questions.pdf) and send it to adair@sutter.com. She will help you get to better pipettes and can also answer questions regarding the P-2000 Puller.
In general, heavy use in itself will not make the pull results inconsistent. Only if those many users constantly bump the filament and then don't realign it and run a ramp test, you will obviously see variability. Wear parts like the filament and drierite will also age faster with frequent use. You should not need to come to the same conclusion as Paula! Sorry to hear about your unsatisfactory experience, Paula. Did you contact Sutter about your issues? We do provide support for our products, and in virtually all cases we can get the customer to the point they want to be.
Both Niraj and Harald correctly state that the P-2000 Puller is not optimized for capillaries with more than 1.2 mm diameter. That simply has to do with the diameter of the laser beam. Too thick capillaries absorb most of the beam on the first pass, and it becomes increasingly difficult to reflect and focus the remainder of the beam on the back of the capillary. Off-center taper and tip are the results. If you already own a P-1000 and do not want to use quartz for low-noise single-channel recordings, you should try to optimize that before spending the money on a P-2000.
I hope this helps,
Jan
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Hi all,
I've been attempting to get some action potentials out of my MEA setup for organotypic cortical slice cultures but have had little success so far. Long story short is that we follow the preparation described in Stoppini (1991), and have narrowed the problem down to something biological (the electronics/MEA system are working).
Basically we are running into the problem that we record many neurons firing if we apply various amounts of Picrotoxin - otherwise the recording is completely quiet. Just wondering if anyone has run into similar issues or has some ideas on where we should specifically be looking to troubleshoot.
Thanks!
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Frankly speaking, I dont have any experience in MEA recording. But let me clarify. Do you apply any stimulation protocol except for picrotoxin?
I mean most of neurons in slice/culture won't fire untill they are not stimulated, cause their main afferent paths have been interrupted during prep.
Although in some places you can catch pacemaker neurons activity (see 10.3389/neuro.01.1.1.009.2007 for further info). But most of neurons in slice/culture would make one single spontaneous spike in 5-10 minutes without any stimulation (eather chemical or electrical). Most of chemical stimulation protocols (low Mg2+, high K+, 4-AP, picrotoxin, etc.) are widely used as ex vivo epilepsy models. (see 10.3791/57548).
Hope this will help. Good luck!
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I've been working with field electrophysiology in the hippocampus of mice for 10+ years. I recently started electrophysiology in rats and for the life of me cannot create reproducible LTP! This is by either 2 trains 100Hz, 1 train 100Hz, or theta burst stimulation. I am beyond frustrated. If I tell you what I'm doing, perhaps you could shed some light on what I may be doing wrong?
-Cervical decapitation on awake Sprague Dawley rats (2 to 12 months)-no anesthesia.
-Brain removed within 30-60 seconds after decapitation into ice-cold oxygenated sucrose cutting solution.
-Brain chills for approximately 30-45 seconds before cutting 400um horizontal sections on Vibratome while remaining chilled.
-Dissected hippocampi rest at RT 50/50 Oxygenated ACSF/Cutting.
I thought it was my ACSF recipe since my d-glucose concentration was significantly higher than most other publications, but that hasn't changed even with lower glucose:
ACSF: (in mM) 125 NaCl, 3.5 KCl, 1.25 NaH2PO4, 25 NaHCO3, 10 d-glucose, 2.5 CaCl2, 1.3 MgCl2
-Slices mounted on interface chamber by Automate Scientific (https://www.autom8.com/bsc1-interface-submerged/) for 1 hour 20 minutes to 2 hours. ACSF is oxygenated and heated by TCU to 34 C (temperature of the slices ends up being 30 C by the time it flows around the interface netting). This chamber allows for a humid environment to reduce drying out the slices. Flow rate is ~1.5 ml/min by peristaltic pump.
-Stimulating with bipolar nichrome formvar electrodes at the edge of CA3
-Recording at CA1 with ACSF-filled recording electrodes (I use pClamp software for signal acquisition).
-Stable and strong signals are achieved before running input-output curve.
-Stimulus intensity level determined by 1/2 max fEPSP attained from I/O curve.
-Record for 20 minutes before initiating high-frequency stimulation (either 100Hz or TBS).
It should be noted that I get reproducible LTD with chemically-induced LTD with DHPG (haven't tried LFS).
-My signals will either remain the same intensity, reduce, or completely disappear during or after HFS.
Some slices will dry out before I even record.
Perhaps my electrode position is wrong? I usually look for optimal fEPSP slopes at about 5 ms past stimulation. This is what I normally do for mice. Perhaps the distance needs to be greater?
I would love some assistance with this. I have aged animals that need electrophysiology done now, but I cannot euthanize them knowing that they will not produce LTP.
Thanks in advance!
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here is my recipe for choline chloride cutting and slicing if you want to try it.
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Hello, all!
I'm inducing/measuring LTP in in-vitro mouse brain slices. My stimulating electrode is supposed to be in CA3 and my recording electrode is supposed to be in CA1 of the hippocampus. I was having trouble finding the Schaffer collaterals using the usual top lighting from a dissecting microscope, so I've set up a couple different ways to get the backlighting I prefer on the slice (involving such common items as a flashlight on a cell phone and a closet light consisting of a bar of LEDs). I haven't come up with anything ideal yet. I was wondering, what does everyone else use for lighting?
Thanks in advance!
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As I anderstand you wish to see Schaffer collaterals. You can't see them
All you need to see is cell layers and stratum radiatum, good anouth to be able to place electrodes. First place stimulation electrode in strat. rad. clouser to cell layer, were SC more dense, and start to place recording electrode somewhere in the middle of strat. rad., where dendrits is, (this is for field potential recording).
Than try to get EPSP (field potential) by increasing stimulation intensity. You may need to replace recording electrode several times to get good shaped potential.
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while chloride concentration is low inside cell , some paper recorded action potential in iPS-CM with Chloride based Internal cellular solution. How can this work? I think the high concentration could distort the RMP, what rational is it to use this ?
many thanks
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Dear Zhang,
In adiition to above comments, I would like to add that one can use high Cl- concentrations (500 mM to 3M KCl) with high impedance electrodes (60-100 megaOhm) during sharp electrode intracellular recordings. For examples, you can see my earlier publication.
Best wishes,
Refik
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The resistance reading from WPI EVOM2 is ohm x cm2 or just ohm? I read the manual very carefully and thought it's ohm x cm2 and we don't need to convert it. But the TER value I got for my RPE monolayer on a 12-well transwell is much lower than expected, which makes me feel confused. Anyone has an idea?
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Our company develops 3D cell barrier culture systems for in vitro study of virtually any cell barrier, and the system comes equipped with both a Fluid Perfusion Unit and a Trans-Endothelial/Epithelial Electrical Resistance (TEER) Measurement Unit! The use of a vessel-shaped 3D environment has been proven more effective than 2D systems like Transwell (
Article Santaguida S, Janigro D, Hossain M, Oby E, Rapp E, Cucullo L...
), and our advanced TEER Measurement Unit allows for frequency sweeps between 0.1-1,000 Hz, automated time point sampling, logging of data to Excel, and additional measurement of phase angle for cell capacitance calculations! For those that are committed to Transwell use, we also have a TEER Measurement Unit that is compatible with nearly all Transwell products (Endohm cell cup chamber, STX2 "chopstick" electrodes, etc) and allows the user to greatly expand their testing capabilities with Transwell equipment. Additionally, we have smaller modular systems that connect to microscope-friendly cell culture modules: i.e., a miniaturized 3D cell culture system that you can use right under your microscope! If you or anybody on this thread is interested in learning more, I encourage you to visit www.flocel.com or email me at djanigro@flocel.com, thank you!
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We are looking to reconstruct biocytin filled neurons from confocal image stacks. I realise Neurolucida is the gold standard but even the Neurolucida 360 lite version is hideously expensive (~$15 000). Is Neuronstudio a viable alternative despite not being updated since 2009. Are their suitable plugins for Fiji? It would be really great to reconstruct in 3D. Any thoughts would be appreciated.
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For reconstructions from 3D image stacks, we have been quite satisfied with the Neuromantic freeware:
It does what it is supposed to, with no bells and whistle that complicate matters more than they help. With Neuromantic, I have put dozens of undergraduate students on the manual reconstruction work, they get their lab experience, we get our reconstructions, everybody's happy. We've done >150 reconstructions in the lab this way. Works great, but support is nil and updates rare.
I have heard good things about Fiji/Simple Neurite Tracer but never tried it. There is also Herman Cuntz's TREES toolbox (http://www.treestoolbox.org), which is more advanced. Finally, you might benefit from the option not to reconstruct at all (Ferreira et al: Neuronal morphometry directly from bitmap images. Nature Methods 2014 DOI:10.1038/nmeth.3125.).
All of this requires 3D image stacks though, so you do not work "live" like people often do with Neurolucida. However, working offline comes with advantages too.
For our work with Neuromantic, see below, in particular Blackman et al:
Blackman AV, Grabuschnig S, Legenstein R, & Sjöström PJ: A comparison of manual neuronal reconstruction from biocytin histology or 2-photon imaging: morphometry and computer modeling. Frontiers in Neuroanatomy (2014) 8:65, DOI: 10.3389/fnana.2014.00065.
Lalanne T, Abrahamsson T, & Sjöström PJ: Using Multiple Whole-Cell Recordings to Study Spike-Timing-Dependent Plasticity in Acute Neocortical Slices. Cold Spring Harb Protoc (2016) 10.1101/pdb.prot091306
Buchanan KA, Blackman AV, Moreau AW, Elgar D, Costa RP, Lalanne T, Tudor Jones AA, Oyrer J, & Sjöström PJ: Target-Specific Expression of Presynaptic NMDA Receptors in Neocortical Microcircuits. Neuron (2012) 75:451-466.
Lalanne T, Oyrer J, Mancino A, Gregor E, Chung A, Huynh L, Burwell S, Maheux J, Farrant M, and Sjöström PJ: Synapse-specific expression of calcium-permeable AMPA receptors in neocortical layer 5. The Journal of Physiology (2016) 594(4):837-861.
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Iam looking for good papers with the protocol but I cant find papers showing the regions for stimulation like those of hippocampal recording.
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Thank you very much Kelvin Broad for the help.
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Hello,
I've been trying to measure NMDA currents in the BLA. I've been using magnesium free acsf (122 NaCl, 2 KCl, 1.25 KH2PO4, 26 NaHCO3, 2.5 CaCl2, 10 D-glucose with nbqx (5uM) and bicuculine (5uM) and stimulating external capsule while principal cell was held at -70 mV, but I can't get any baseline and they are very unstable. I was wondering about changing it to normal acsf with 1.3 MgSO4 and measure it at +40 mV or maybe try to stimulate locally or in central amygdala? Do you have any good practice to measure NMDA currents?
I would be very grateful for answers.
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I would test to make sure you're getting stable EPSCs before trying to measure the NMDA component of the EPSC. If you're not reliably evoking stable EPSCs, playing with the conditions to isolate the NMDA currents will just be frustrating. Using a Cs based internal solution with normal aCSF and measuring at +40 mV would be one way, and would probably be the easiest. That way, you can ensure you have a stable EPSC at -60mV and then block AMPA-R's with NBQX and shift to +40mV to measure the NMDA currents. Alternatively, reducing the Mg in your bath and measuring at -60 mV is another method people have used. You might also consider adding a low concentration of glycine to the bath to help facilitate the NMDA currents, even though it shouldn't be limiting.
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Please, can someone help me? I am trying to create a dv/dt graph after I apply the ramp protocol in the current clamp, but I am making a mistake somewhere. Can you help on how to do this graph. I created graph and ramp current recording was added. thanks a lot of.
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Dear Ercan,
Nice recordings.
From what I understand here you need to plot dV (voltage difference in mV) in the y axis and the dT (time difference-ms) on the x axis. It all seems to happen within 7 ms or so, as slowers are not able to generate any action potential-but only similar subthreshold potential changes. For example, 1ms long ramp at 1nA current is giving you about 70 mV difference (dV = 70 mV and dT = 1ms); 2 ms long 1 nA ramp giving you 65 mV (dV =65 and dT =2 ms); say 8 ms long ramp is giving you dV of 30 mV-the longer ramps are pretty much staying at that level and therefore you may need all of the longer ramps. So you plot them that way it should give you an exponential decline. If you like you can fit those with an exponential(s). These values you may compare among individual cells or groups of cells depending on what you are aiming at. For example some people use it/these values for comparing excitability.
Best wishes,
Refik
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I'm trying to perform patch-clamp recordings in the rabbit retina, but holding the section in the chamber is a problem. I use a grid currently, however there are always parts of the section that are not flat, therefore I cannot do proper recordings in those areas. I've tried  coating the chamber with polylycine but that is ineffective. 
Does anyone have any method that works well?
Thanks
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Do you want to perform patch clamp in slices or whole mount retina? For whole mount you can use small pieces of filter paper containing a small hole in the center. You can then attach the pieces of retina by applying gentle pressure (for more info see Marla Feller's papers, UC Berkeley). For slices, I think a grid would work fine, but be careful with the size of the slice. If it is too big, probabilities are that they will "fly away". 
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Hi everyone, I am new to voltage-dependent current measurement and I need help. I am applying a standard 100 ms V-step protocol from -100 to + 60 mV (holding level -70 mV and no channel blockers in the ACSF) to measure sodium and potassium currents. The typical trace that I obtain is shown in the image. My questions are: is this trace normal? And how do I calculate the peak sodium current, since the peaks are above the holding potential (see the red sweep)? Is there something wrong in these experiments?
Thank you very much for your help!!
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Inn addition to blocking all non-sodium currents, shorten your test pulse to 10 ms. 100 ms is too long and is suitable for calcium and/or potassium currents.
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Hi,
in several articles, the electrotaxis chamber that is used to electrically stimulate the cells, is isolated from the electrodes by agar salt bridges. Apparently, it is to limit the amount of electrochemical product inside the medium. However, recent studies have not used these agar salt bridges to stimulate the cell in order the miniaturize the experimental setup. I also have some spatial constraints and am thinking about trying without the agar salt bridges. Have you more information about the role of the agar salt bridges and are the cell viability and behavior affected without it ?
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Dear Jerome, thank you for your interest.
Yes, you are right, the idea of building our chamber was based on need of reproducibility, easy handling  and pretty big working surface (we search a lot of devices availible on market or published). We have before in our lab also setup with agar bridges and did nice observation of cell migration, but it is really not easy for handling, especially for long (2-3 weeks) -term stimulation of some samples in parallel.  
After 150 (200) mV with our setup we start having problem with medium and we were pretty happy with the effect of 100 mV. 
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Hi everyone
Would anybody be able to tell me if it is possible to culture cystic fibrosis bronchial epithelial (CFBE) cells at ALI on transwell inserts? I have found plenty of literature that describes the successful culture of CFBE polarised monolayers using liquid-liquid interface (LLI) but nothing to date for ALI. 
I have attempted to grow CFBE monolayers using 3 days under LLI and then switching to ALI. There is a monolayer of cells visible but the TEER is very low, even after 14 days of culture. Has anyone experience with this? Specifically, the optimal culture time under LLI before switching to ALI would be very helpful.
Thanks in advance,
Alan
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Not too late at all Matt, many thanks for your help!
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Hi everyone,
I do whole cell patch clamp recordings from layer V neurons. In current clamp mode (with zero holding current), I check spontaneous action potential firing in vitro in acute coronal rats slices using K-gluconate-based solution (in mM 140 K-Gluconate, 10 KCl, 0.1 CaCl2, 10 HEPES, 10 EGTA, 5 MgATP, 0.5 NaGTP, 10 Phosphocreatine, 280-290 mosmol, pH 7.3) at room temperature. ACSF is 295-300 mosmol, pH 7.3 and no ions omitted or modified. I mostly get quite low firing rates or silent neurons (compared to neuroelectro data - www.neuroelectro.org-though data here is mostly from in vivo experiments) in ~2 h-long recordings. Could you give me some hints about the factors affecting the spontaneous firing rate? Any link or reference would also be highly appreciated. Thank you in advance.
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Yes. Sometimes, under current clamp conditions, seal resistance may be unstable.  The baseline level of resting potential may fluctuate and hence generate action potential if potential change reaches the threshold level. "Spontaneous" action potentials could be virtually overestimated.
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I am doing extracellular single unit recordings and I am looking for a quantitative approach to show that spike clusters recorded on the same electrode over multiple recording sessions belong to the same neuron.
Is there a good quantitative or statistical test to show similarity between clusters or waveforms?
I see that some publications report the correlation between average waveforms. Would this be meaningful if the number of spikes in my clusters is small?
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De Vanja,
Please see the below given link.
Regards
Krishna
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I am testing the effects of a drug, soluble in DMSO, on electrophysiological properties of dopamine neurons. The experimental protocol consists of 110 min of incubation. Since this drug is only soluble in DMSO, I first check whether the sole incubation with DMSO for 110 min may causes changes of neuronal intrinsic properties. Following this incubation, my first impression is that neuronal membranes are more fragile; it is harder to obtain a cell-attached recording, and the membrane easily goes to rupture after gaining the seal.
Does anyone have experience with this kind of issue? Are there any useful tips to handle a prolonged incubation with a drug soluble in DMSO?
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DMSO can seriously damage  membranes and I think it could contribute to what you experienced. Are you using > 0.1% DMSO in the final solution? Some of DMSO soluble drugs can be dissolved in EtOH. Have you tried that?
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In preparing the ACSF solution, is it a mandatory and crucial to evaluate the osmolarity of ACSF solution?
Also whether it will have influence on hippocampal slice viability?
Further, what would be the ideal incubation period in bubble chamber for the best possible slice viability.
Please, share your expertise in this regard.
Thanking you,
Best Regards,
Grandhi V Ramalingayya
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It is mandatory to ascertain the osmolarity of the solution, otherwise the concentrtion may not support neuron survival. This article may help:Slice preparation Solutions ACSF: (in mM) 125 NaCl, 2.5 KCl, 2 CaCl2 ...
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Background: I am interested in studying the electrophysiological properties of PV interneurons in brain slices of adult mice. I bought a PV-eGFP line (CB6-Tg(Gad1-EGFP)G42Zjh/J) from Jackson to perform this experiment and patiently aged the mice. Unfortunately, there seems to be some epigenetic silencing of the eGFP with age (I blame Jackson for not properly documenting this, even though apparently many people have complained about this. Be wary of this line)! So I have all of these aged transgenic mice, but almost fluorescence anywhere!
Question:
Instead of wasting my efforts and sacrificing these aged mice, I would like to see if blind patching may be a viable alternative. Do any of you guys have any advice on how to identify PV interneurons using strictly DIC? We will ultimately be validating the identity with current injections to see spiking patterns, but I want to increase our chances of getting the right cells with DIC. I was told by some that PV interneurons tend to have smaller and rounder somas. Can anyone validate this? Or direct me to papers where they do blind patching on PV interneurons?
Thanks!
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Hi Ruoqi,
The answer is yes. It is definitely doable to patch PV+ Fast-spiking interneurons(FSIN) without fluorescence.But the difficulty is depend on the brain region. In dentate gyrus, you can Identify FSIN by there soma size, which is much larger than granule cell. However, in CA1 region, the FSIN soma size and shape are very similar to pyramidal cell, so it is much harder to find them with DIC. You should find out whether FSIN have special shape/size/location in your target region.
About the age, PV is not expressed in certain age. You can try to use lhx6 or GAD as promoter to drive GFP. So you can identify interneuron first, and further identify PV neurons with their intrinsic electrophysiological properties.
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We use PatchMaster with a HEKA EPC 10 amplifier and do whole cell recordings. In about 90% of the cases, when we go into whole cell mode and compensate for Cs, PatchMaster shows "wonky" values for both Cs and Rs (say 250 and 4), together with a red "E" lighting up beside the numbers. Usually, this situation yields bad recordings. What could be causing this?
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Cs stands for the membrane capacitance of neurons; when asked for compensation, the amplifier uses series of small currents to get the Cs values based on tau and membrane resistance. Based on these measurements, extra current is passed during recordings to load the Cs.
This automatic procedure however is run in a way to make sure that Cs is not over-compensated as the latter can cause damage of the membrane and deterioration of the patch quality. Thus, typically upon request for Cs compensation, the Cs is underestimated and under-compensated. This is especially the case with larger neurons.
If your settings are right, you should be able to repeat Cs compensation several times (with no harm) to get the Cs properly compensated yielding good recordings. Hope this helps, SV     
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I am working on N2A cells grown on 35 mm dishes and would like to generate action potential in these cells. I have come across methods like photo-uncaging from papers of Callaway, Kartz etc but that has been used for brain slices and i am not sure if this would be apt for cells grown in in-vitro. Any suggestion would be much appreciated.
Edit (12/10/2016): I do not have any electrical stimulation facility (like MEAs, tungsten electrodes etc) right now and i am looking for standard protocols for chemical and light based stimulation approach here. 
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You could try using multi electrode arrays embedded in glass bottom Petri dishes for electrical activation
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Hi everyone
I'm currently studying the function of gap junctions, and would like to be able to patch a cell and specifically block its gap junctional output. 
Does anyone know of a drug I can include in the patch pipette solution to block gap junctions from the inside? It doesn't have to be a particular-connexin-specific drug, can be a global connexin drug. 
Thanks!
Simon 
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Just  CO2 to the medium
Nat Methods. 2007 Apr;4(4):353-8.
Unitary permeability of gap junction channels to second messengers measured by FRET microscopy.
Hernandez VH1, Bortolozzi M, Pertegato V, Beltramello M, Giarin M, Zaccolo M, Pantano S, Mammano F.
Nat Cell Biol. 2005 Jan;7(1):63-9. 
Impaired permeability to Ins(1,4,5)P3 in a mutant connexin underlies recessive hereditary deafness.
Beltramello M1, Piazza V, Bukauskas FF, Pozzan T, Mammano F.
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I am trying to record synaptic plasticity in cortical acute slices (300um) over 1h however I always get a very dramatic LTD that result in almost no synaptic response at the end of the recording, even in control trials where I only record baseline synaptic responses with no additional stimulation. The patched cell is still alive and firing action potentials upon depolarization and the access resistance has not changed much.
I do have ATP/GTP in my internal solution, which is K-gluconate based. My baseline stimulation is every 30s. I tried both IC and VC mode which gave me similar results. I'm not blocking inhibition.
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There might be many reasons for that, it is quite common. I suppose you are talking about stimulated activity, how are you stimulating ? And what is the behavior of the spontaneous activity ?
What is your chloride reversal potential ? If you are not blocking GABA receptors, it is possible that you are recording mainly GABA responses, and with the diffusion of your intracellular solution you slowly shift them to shunting.
If you are stimulating too strongly, you may just exhaust your synapse. Try to be in minimal conditions.
It could be a too high temperature as well, exhausting your synapse. 31 - 32 °C is fine, higher can be trickier.
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I am using a prototypical hippocampal slice preparation. Slices are prepared in high Mg2+ and low Ca2+ and sucrose before being incubated in normal acsf.
I am able to get very good population spike recordings however my fepsps are very small and contaminated with population spikes even at very low currents.
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For acute slices it might be better to use a vibratome rather than McIlwain tissue chopper (more intersting for organotypic hippocampal slices from p5-7 pups). Is the ACSF in which you perform dissection and slicing cold enough to get the tissue dense to avoid crushing when you cut with the chopper ?
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I have been trying to patch from interneurons in the spinal cord but have had trouble getting healthy slices. I have been dissecting and slicing the spinal cord in low Ca2+ Ringer's solution as described in Ole Kiehn's paper and then allowing the slices to recover in normal ringer's solution for 40 mins before recording. I have tried altering the vibrotome setting and still I don't consistently get healthy slices. Any input on the matter will be greatly appreciated !
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How I can block NKG2A receptor signaling for in vitro experiments?
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You might consider to convert your anti-NKG2A antibody of choice (and the respective isotype control) into Fab/F(ab)2-fragments - there are several commercial kits for that purpose that are easy to use and reliable. By using blocking Fab/F(ab)2-fragments you circumvent the potential problem of Fc-receptor-mediated activation. Good luck!
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We have been done visual cortical brain slices of mice (300 μm, 20-25 days) for electrophysiology. We´ve been doing it successfully for years and never had this kind of problem. Since the last three weeks, the slices are always completely dead. Not even a single cell survived. For dead I mean that we can clearly see the dead cells (large cell body and clearly visible nucleus). The slice is already dead even if we put it under the microscope immediately after cut.
 We´re using the following:
-       Cooled (0°C) oxygenated (5% CO2-95% O2) cutting solution (in mM): 206 sucrose, 25 NaHCO3, 2.5 KCl, 10 MgSO4, 1.25 NaH2PO4, 0.5 CaCl2, and 11 D-glucose.
-       ACSF; in mM: 125 NaCl, 25 NaHCO3, 3 KCl, 1.25 NaH2PO4, 1 MgCl2, 2 CaCl2, and 25 D-glucose).
-       Leica vibratome 1100S
We´ve already tried the following:
We checked osmolarity and ph of both solutions many times with different machines and they are ok (300 mOsm-7,3).
We took distilled water from other laboratories.
We replaced all salts with brand new products.
We autoclaved all glass and then cleaned it with hydrochloric acid, then with alcohol and then with tons of distilled water.
We verified with weights samples that the balance is calibrated.
We thought that the problem could be the vibratome. However, it seems to be working normally. It vibrates horizontally and shows no visible sign of malfunction. The slices don´t look to have passed throught any kind of unusual mechanical damage.
We made a test cutting a brain with solutions from another laboratory, and the neurons were healty, so we suspect it is a contamination problem occurring during the preparation of the solutions in our lab. If this is the most likely hypothesis, we are afraid to do tests involving other laboratories and risk to contaminate their stuff.
We are continuing to replace components to isolate the problem but with no luck. As far as I know, bacteria can kill slices, but that requires at least few hours. I cannot think of a chemical specie capable of killing everything this way and persist after several washing processes. If someone has an idea of what can be going on, we would be grateful to receive an opinion.
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Roberto, test with a voltmeter wherther you have a passing current through the vibratome when slicing (one pole on the blade, the other one in solution)....
this happened to me in the past.... we were electrocuting the slices in the slicing chamber.... cheers
a ground got disconnected inside the slicer... it was supposed to ground the chasis..... yeah crazy shit!
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Hello,everyone! it's the first year of my graduate study, and i am starting to do whole cell recording on the brain slice  of aged mice.
i have got some help from the website:http://www.brainslicemethods.com/, but i am still not clear about the exact reason for the choice of modified ACSF, what's the effects of this chemicals (sucrose, HEPES, NMDG, Tris, Choline) and their differences on mice in different age (i only know they are the replacement of NaCl)?
In addition, i have used NMDG-acsf to get slices from 2M old mice, with perfusion(room temperature) and recovery in 33℃ for 12min. sometimes the slice seems healthy and the cell looks in normal morphology, and i can get "gigal cell" in the process of whole cell recording, but when you sucking, the cell membrane seems ruptured or else (i don't know what really happened), can you give me some advice and the trick you know about the methods to get slice from aged mice(1M-9M old)?    
Help please, and Thanks!
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Dear Sunney,
In aged mice, by replacing Na+ with sucrose, NMDG or choline, you are increasing the chances of neurons surviving the insult (dissection, slicing etc). I have tried most of these solutions to obtain good motoneurons, which are arguably the hardest neurons to obtain in adult mice/rat. For details of my ACSF solutions that I used for cutting brain slices at different stages of development, see my following paper. If you are cutting hippocampal, cortical, cerebellar or amygdala slices the modified sucrose solution works well with the old mice. 
best wishes, Refik
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I'm trying to patch-clamp cultured embryonic hippocampal neurons using ACSF, but I've had some problems with cells viability. Cells lose their smooth and birefringent surface relatively fast.
These cells are cultured in Neurobasal medium (~220mOsm) and I use ACSF (~310-330mOsm) during recordings. To not induce osmotic shock I gradually increase Neurobasal osmolarity using NaCl, but cells still die fast.
Can anybody help me?
Thank you in advance,
Diogo
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Well, I've tested in hippoc neurons from E17.5 mice also at DIVs 14-21 and in my best attempt the cells were "viable" only for 2h, but I'd say that after 1h the cells lose their smooth surface.
I've to do more tests and see if I can improve cell viability.
Thank you for your comments Alberto.
Regards!
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In my research, I have found that while kainate has been used in bath application for electrophys and quite extensively, there are not many recent publications using this model.  Instead, I find that it is very popular for intrahippocampal injections to simulate human TLE.  I know it is typically used for chronic models, but in the hc, in particular, these receptors play and important role in the CA3. If anyone can provide insight to this it would be much appreciated. Thanks!
P.S., I have generated epileptiform events using kainate. we are exploring excitatory models other than 0 Mg
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It's worth having a look at this: http://www.ncbi.nlm.nih.gov/pubmed/24184743
The advantage of the kainate model of epilepsy is that, after the first KA-induced seizure, there is a latent period after the initial insult which precedes the development of recurrent seizures in the animal. The kainate model also reproduces anatomical features of epilepsy, such as mossy fibre sprouting in the DG. These features of the in vivo KA model require time and anatomical changes in the brain, so are not appropriate to use in an in vitro slice model. Acute activation of KA receptors (don't forget that KA is also an agonist of AMPA receptors) in slices is quite different from the excitotoxic insult used in the in vivo model of TLE.
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 I injected AAV-ChR2 in rostralPAG and OT-Venus in oxytocin neuron in order to define synaptic connection between them, but none of oxytocin neuron showed light-evoked postsynaptic current.According to previous virus tracing result, they have synaptic connetions. ChR2 expression in axon terminals of PAG neurons seems mild surrounding oxytocin neuron, is that why I couldn't record any photo-responding oxytocin neuron? It wiil be very thankful if you can answer my question!
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In injecting the oxytocin neurons, where did you inject?  Not all OT neurons are similar in terms of the connectivity.
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Dear all,
I would be very happy if someone could help me with my question. We are using saggital hippocampal slices and try to record field potentials. We normally see the response of fiber volleys  followed by a synaptically elicited field potential (see attachment). As far as I know the field potentials from the dendrites should be much bigger than the response from the fiber volleys. Do you have any idea why the synaptically evoked field potential can be smaller than the response from the fiber volleys?
Thanks for your help,
Doris
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Out of the many possible explanations, I think the one that is most likely (and least satisfactory!) is that it's just a poor health region that you're recording from. This to a large extent depends on the age of animal you're slicing from - the younger, the healthier the slices would typically be. But also fEPSP recordings, can take a lot of "hunting around" for regions in the slice that will produce nice/healthy responses; sometimes you'll find that by moving your recording electrode a few micrometers laterally and/or deeper in the slice will give you dramatically improved post-synaptic response to pre-synaptic fibre volley ratio.
The other thing to consider is you mentioned you're using sagittal slices, which I don't think is the most common configuration (most people use whole brain coronal, or transverse cuts of isolated hippocampus), so it may be that in this configuration, it is less likely to obtain large post-synaptic responses.
There are lots of excellent resources in the literature that may be of use to you, this is an example of one that I found particularly helpful:
Bortolotto, Z.A., Anderson, W.W., Isaac, J.T., and Collingridge, G.L. 2001. Curr. Protocol Neurosci. Chapter 6: Unit 6.13.
Best of luck!
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I am recording fEPSP from sl-m of area CA1 with the stimulation electrode in the same layer around 1000 micron apart.  When I applied APV + CNQX with or without bicuculline, I got an enormousely increased fiber volley. Would you please tell me if the enlarged fiber volley implies a great preservation of perforant pathway fibers there? Thanks a lot!
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You could have at least a couple different things going on.
1) In the presence of bicuculline, you can get increased excitability and thus a larger fiber volley.
2) More likely, by blocking the excitatory response with APV + CNQX, you are unmasking the fiber volley from underneath the postsynaptic response. These can get tangled, especially when responses aren't as clean, like in a place like SLM.
A couple other caveats about PP-SLM recording. Your responses can also be contaminated by dentate gyrus pop. spikes because of your close proximity or via ring-around through CA3 and into Schaffer-Collateral. Some people place remove DG place a cut between CA3 and CA1 (see PubMed ID: 10085331, Figure 1 and Methods) . Also, make sure you check to make sure there isn't contamination of the response by the Schaffer Collaterals. Without moving your stimulating electrode, place your recording electrode into stratum radiatum. If you have a positive field potential then you're good. If you're field potential is negative, you're stimulating electrode is exciting Schaffer Collaterals as well. You'll have to move things around until the negative deflection in stratum radiatum goes away.
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What is appropriate Voltage-clamp protocol for measuring inward rectifier K+ currents?
Someone used Ramp protocol from -120mV to 10 mV with HP of -40mV and 1 sec duration, and some use Step pulse from -120 to 10mV with HP of -40mV and 500mec to 1sec
Which is better way to measure accurately?Any advantage/disadvantages of both techniques? 
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Dear Chandra,
In Figure 3, I measured the current at steady state or at the end of 1s long voltage step. The current at hyperpolarized levels was primarily a tertiapin sensitive K (Kir3.1 and Kir3.2) current.
best wishes, Refik
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Are there anyone with experience that moving a brain slice by the flow of ACSF under the slice?
In my new lab., I’m using a slice chamber (there was already it there, cartoon is attached) and a "harp" (U-shape slice anchor) made by platinum bar, however slices are gradually moving and lifted up. 
ACSF come to the chamber by gravity and these are aspirated with an air pump.  A slice is placed with harp and left at rest a few minutes, then I observe that slice surface is lifted.  Not surprisingly, if I performed patch-clamping, the cell has gone away... 
It is the first time to prove this problem, though  I have performed slice electrophysiology over a decade... I don't have ideas solving this problem but slower flow-rate improved a little (however, if I can, I don't do so because ACSF's temperature is kept at ~32 ℃). 
If anyone have a nice idea and/or same experience, I'm grad to share the good stuff!
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I really appreciate your useful tips!!
Then, now I report results so far and that  I was trying to.
1. Re-built a harp which has strings of a strong tension
RESULT: Although I seemed that slices were fixed stronger than before that, the motion drift was detected regretfully. The slices pressed by new harp's strings were DENTING, therefore I think that the pressure was sufficiently applied.
2. Additional anchor (small platinum bars)
RESULT: This was not effective in the problem.
3. Slower flow rate
RESULT: I set very slow flow rate (~ 0.5 ml/min), then the artifact was improved but not perfectly gone... This flow rate is too slow in my experimental conditions (acute slices from adult animals, keep the temp. of ACSF at ~32 degree celsius and keep a whole-cell condition over a hour, etc...). Needless to say, the condition of neurons was going to be poor visibly.
...Next I'll try to replace a glass at bottom of the chamber with an adhesive-coating glass as mentioned by Refik.
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In the litterature, people has reported using direct, alternative or pulsed currents, and so current generator. However, they give the value of the difference potential (in volt per mm) induced to the cells. Is there no contradiction here?
The current generator is generating a current that is independent of the resistance encountered because the voltage modulates (following the ohm’s law). How can they obtain a constant and reproductive difference potential in volt when using a current generator? 
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Dear Jérôme, I agree with you that if one uses constant current stimulation mode the voltage at the stimulation electrode will change due to electrode tissue impedance variation. Therefore one will not get a constant and reproductive potential at the stimulation electrode. This would only be the case if the electrode-tissue impedance always would be constant. Anyway I would recommend the following review paper for details: Tehovnik EJ. Electrical stimulation of neural tissue to evoke behavioral responses. J Neurosci Methods, 1995; 65: 1-17. Also please have a look at the paper published by Histed and others: Histed MH, Bonin V, Reid RC. Direct Activation of Sparse, Distributed Populations of Cortical Neurons by Electrical Microstimulation. Neuron, 2009; 63: 508-22. The authors reported that microstimulation sparsely activates neurons around the electrode, sometimes millimeters away, even at low currents. This very interesting paper raises several issues to consider for the design and interpretation of microstimulation studies! I hope my answer was helpful. Best wishes, Dirk
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Hello.
I used vibratome VT1000P for acute brain slice preparation and electrophysiological experiments such as extracellular field potential recordings and whole-cell recordings. But this one is too old and there were some problems to use. We thus purchased and have used Campden microtome 5100mz because this model is cheaper. Sometimes it is not bad, but my labmates argued that the slice quality is worse and sometimes I feel same. Does anyone use this microtome for E-phys? Do you have similar problems like me or do you use it well?
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I used Campden vibratome for spinal cord slices/patch clamp recordings. It was great. Maybe you need to adjust some of the cutting parameters for optimal results ( the speed, amplitude, frequency), fine-tune the z-axis oscillations, check the quality of the blade etc.
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Is it possible to isolate peak Ca2+ current using dihydropyridine on external solution with NaCl?
Is it compulsory to remove NaCl from Ext. solution or block by TTX to measure Ca2+ current ?
What is effect of Sodium current on calcium current ?
previous paper shows dihydropyridine block very less sodium current at +10mV (where less open probability of Na whereas higher open probability of Ca chanels )
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Dear Chandra,
In order to obtain Ca2+ current in cardiomyocytes first what you can do - keep them at holding potential of -40 mV and step them to 0 or +10 mV with rectangular pulse of 200-300 ms duration.
Then, to validate, that this is calcium current you can add dihydropyridine - verapamil in order to block ICa2+, or BayK8644 to increase it.
Patching at holding potential of -40 mV should help you to get rid of sodium current. In case if it does not happen, you can add 1 micromolar of TTX to your bath solution or reduce concentration of NaCl partially replacing it, for instance, with NMDG.
Best regards,
Miroslav
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Dear All, I am looking for patch-clamp researcher, who has already performed (or is able to perform) single channel patch-clamp experiments on Kv 1.2 channel.
My work concerns structure-based modelling of Kv 1.2 potassium channels. I have already formulated geometrical model of these channels, which describes the ranges of motions of functional subdomains by changes of cell membrane voltage. However, to verify model assumptions I need to compare my results with experimental data. In particular, open state probabilities and channel conductances for different voltages are obligatory for further analysis.
I would like to kindly invite you to cooperation and sharing scientific experience in this field.
Should you need any further information, please do not hesitate to contact me here 
I look forward to hearing from you soon.
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Dear Agata,
If I were you I would do the single channel recordings in expression systems that overexpress only your specific channel. And you can use native cells for studying whole-cell patch clamp properties of the channel. This way you can save your alot time and avoid frustration.
best wishes, Refik
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Dear all colleagues,
I have some questions about internal solution preparation. When I first learned patch clamping, I was taught to add all the chemicals together including EGTA, HEPS, ATP-Na and GTP-Na, and then titrated pH to 7.2-7.4 on ice.
Now I found someone prepared internal solution like this, he mixed all the chemicals except ATP and GTP, then titrated pH to 7.2-7.4. Then, he added ATP and GTP. 
My question is what way do you use to prepare internal solution.
Can ATP affect pH of internal solution and can pHing on ice affect the ph?
PS: Do you make a stock solution for EDTA? Or add EDTA power directly into internal solution?
Thank you
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ATP is actually quite stable in water, especially at pH 6.8-7.4 - adding freshly prepared ATP/GTP, or keeping the internal solution on ice is more like an habit which actually has little influence. But like always, better to play safe ;-)
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It is hard to get a good seal and low enough access resistance with slices  from old animals.  How to optimize the patch-procedure and what would be the ideal optimal series resistance (Rs) adjustment for CA1 cells? My electrodes are about 5-6 MOhm. The access resistance (Ra) is hard to get under 20 MOhms. I am using Axoclamp 700B and pClamp for acquisition.
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You should definitely explore optimizing your cutting and incubation conditions for slices from older animals.  Some prefer sucrose or NMDG-based substitution (see, for example, www.brainslicemethods.com for recipes).  We have found that cutting in NMDG-based aCSF, then storing in a modified higher (20 mM) HEPES aCSF can improve tissue viability dramatically.
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Can anybody tell how to measure the extracellular concentration of sodium and potassium ion of nerve cells ?
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Hi,  question is  how  fast and how accurately you want to do this and what equipment you have to hand ?  There are now K+ dyes available  for extracellular recording but they are not very accurate if your extracellular space is less than the Abbe limit and can't be resolved (less than 250 nm). There are of course a few 'traditional methods to do this. Depending on what you have availably in terms of equipment:
1. Extracellular ion sensitive micro electrodes. Pros good point accuracy measurement.  cons poor time resolution (response time slow)  and they tend to 'make' a several micron space so don't really reflect the real K concentration. Also there are iononophores availably for Na as well.
2.  If your cell has well characterised K currents and you have good whole cell voltage clamp,  you can use the change in reversal potential of the K current to measure the resting and dynamic changes in extracellular K concentration.  this is very accurate and has good time resolution but requires a bit of nifty calculation.  If they have prominent Na currents you could use them as well to determine the extracellular Na concentration. 
Hope that helps (I can send you refs for the 2nd option if you are interested.
good luck!
Euan
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Are there any ways to isolate smooth muscle cells from murine aorta with good membrane potential (ideally -60mV)?
For digestion I use collagenase, thermolysin and trypsin inhibitor. After digestion I use the set of polished glass pipettes to get cells in to suspension.
Normally I isolate a lot of nice looking rod shaped cells which are easy to patch but they have very low membrane potential. After isolation significant part of aorta remains solid. I use perforated patch to access the membrane potential, but in 100% cases it too low (0-20mV) and I cannot do any measurements.
Are there any tips which might help me to isolate smooth muscle cells with good membrane potential from mouse aorta?
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make sure that you are using the strong Collagenase (e.g. Worthington Biochem type 2) and try slicing the aorta so you get more surface area for digestion. You should also either use a fungicide during arota prep or do your prep in a biosafety cabinet to minimize fungal infection of the cells. Also, make sure to wash the cells in media at least twice and allow some recovery time in an incubator after digestion.
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Hey folks,
Can anyone suggest a good amplifier for extracellular field recordings from in vitro slices? I'll be setting up my lab in a couple of months and I'm starting to get quotes and equipment ordered. Normally when I've built an interface rig, I've put one together using old Axopatch 1Ds kicking our the various labs I've worked in but, starting a lab from scratch, that's not option and I'll need to buy something.
Obviously, getting a new Multiclamp to do this job is overkill , so I hoping to sample your collective wisdom and hopefully find a more economical solution.
Thanks in advance,
Mick Craig
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Hi Kathleen,
I've not ordered anything yet, but I'm reasonably certain I'm going to go for the A-M systems 1800 amplifier. I've not decided on an ADC yet, but it'll probably be either one of the NI systems, or an ITC16/18, with acquisition done through Igor Pro.
Although I've just got back from a collaboration doing in vivo field recordings, and I'm trying to work out if there's a way I could adapt the Intan RHD2000 evaluation system to double up as both an in vivo and in vitro amplifier, as these are cheap but really powerful, and I could dispense with the ADC altogether.
I hope this helps!
Mick
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If channel is potassium selective, Is it possible to block N-type inactivation with TEA in bath solution on  whole cell mode conventional patch clamp. If yes can anyone suggest any reference for the same.
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Perhaps my phrasing was unclear - I meant that TEA only blocks N-type inactivation when it is applied intracellularly as opposed to extracellularly, not that TEA blocks N-type inactivation and nothing else when applied intracellularly. You are correct that it blocks many K+ channels, I was assuming this would be in culture if the goal is to test a specific type of inactivation. 
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I am working with muscle type nicotinic acetylcholine receptors. I make transient transfection in HEK -293 cells. Does it make any difference in channel function specially in terms of kinetics , if I transfect them in CHO cells or fibroblasts etc(Providing same conditions as it is for HEK cells)?? 
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Hi Abhilasha,
I agree with both Teresa and Victor, and they both raised important points. In my experience I have always noticed difference in responses, as I think it was primarily due to intrinsic membrane and handling properties of individual cells.
best wishes,
Refik
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We would like to measure Local Field Potentials in hippocampus of P21-P23 mice. Is this a viable age, and what are optimal amplifier/stimulator settings for animal of such an age? 
Our goal is to stimulate in CA3/CA1 border stratum radiatum/schaffer collaterals and measure in CA1. However, at the moment we have to get luck to see anything resembling signals instead of stimulus artefacts. We stimulate with short (250-1000 us) strong (1-15V) stimluation.
Thanks in advance for your advice!
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Hi Jon,
That age should be no problem, though I would not go any younger as the hippocampus still developing.
For recording, I would recommend the Kerr Scientific Instruments mobile set up. It comes in a small suitcase including manipulators stimulating and recording electrodes and everything that you need. Extremely practical, cool, economic and highly recommended. I love it. You can check their products and all the details on line at:
cheers, Refik
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A few months back, the top screw used to adjust the blade angle during vibro-check (red circle shown in the picture attached) suddenly started to stick out (above the horizontal level of the headstage). It wasn't used to like it but now it seems that the 0 level of the blade is always associated with the screw above the horizontal level. I have completely unscrewed it and found nothing inside, except the central rod that the screw would screw onto, is also above the horizontal level. Does anyone have any idea what could have caused it and how to fix? Slice quality seems to have been affected.
Any suggestions will be appreciated!
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Hello MiaoMiao
Our lab also uses this vibratome and recently we have had the same problem. In our case, the spring at the bottom of the blade swing arm had become dislodged and resulted in the blade fitting becoming misaligned, hence the screw sticking up. Cutting under these conditions can put a lot of strain on the whole mechanism and result in further very expensive damage (as was our case). We rang our local Leica tech and had the machine serviced as soon as it was possible. As Refik suggested, best to have the company come out and have a look. Good luck!
Cheers Michelle
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I have noticed using CsF in my pipette solution even after filtering through 20 nM filters often leads to blockage of pipette as observed by higher pipette resistance >15 Mohms with a significant distortion too of the junction potentials. However, using CsCl seems to work just perfectly. Na currents were initially characterized using CsF in the cell line I am working with; TE671 cells. Has anyone had such experience before?
Can anyone also explain to me why I may be having higher (+130 mV)  than normal (+ 70 mV) reversal potential even though I am using same concentrations that was used to characterize these cells. 
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I agree with the above, you should make your pipettes have a wider aperture. Also of  note : your Cl ion concentration is going to be lower in your CsF internal compared with CsCl and as chloride ions are the mediators between ionic current in the bath and electrical current in your AgCl electrode the ability of it to pass current is diminished. What is your Cl internal concentration? I don't know why your reversal potential is so positive, what is the Nearsnt equation predicted Na reversal for your Na Gradient ? Perhaps if you post your internal and external solutions it might reveal something else.
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Hi,
In my experiment, the cells are treated with low temperature; when measuring electrolyte leakage, it is so high. I mean as the cells are mostly viable, it shows lets say 80% leakage. How come?
What parameter you think I should check or change? I am using a VWR conductivity meter and the suspension culture is used.
Best,
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True, ionic concentrations of media used for suspension culture may be the possible reason for such high value. You need to wash the cells maintaining it's osmolarity and then measure the EL.
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I am performing whole-cell patch clamping recordings on granule cells of the dentate gyrus and I am applying TEA (100 micro molar).
After a 5-min time application, cells are usually subjected to a massive drop in the resting membrane potential, of around 20 mV, sometimes more; and after the wash out there is not even a slight recovery.
Has anyone ever experience this? 
Thanks in advance,
Martina 
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Dear Martina:
What you see in your recordings is normal. As mentioned above, TEA blocks voltage-gated K+ channels and the end result of this is a drop of the membrane potential. Typically, after short exposure, TEA can be washed out quite well, which leads to  partial recovery of the membrane potential. Note, that when applied briefly, it blocks K+ channels from outside and only in their open state, i.e. if cells are clamped at  negative potentials, the potency of TEA drops dramatically. Although TEA is thought to be a non-permeable K+ channel blocker, during longer exposure some of it gets inside of cells (especially when the seal is not good).  When this happens, it also blocks K+ channels from inside, and as a result, its washout becomes very problematic. Hope this helps.  
Good luck,
SV
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If a synaptic channel was purely selective for potassium, how would you raising extracellular [K+] from 4 mM to 40 mM shift the reversal potential for the synaptic potential produced by opening this type of channel? 
This is a question in my class...I got lost...Thanks anybody who can answer this question...
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The reversal potential can be calculated using the Nernst equation:
EK = (RT)/F * ln ([K]o/[K]i)
where EK is the Nernst (or equilibrium or reversal) potential for potassium, R is the universal gas constant, T is the temperature (in Kelvin), F is the Faraday constant, and [K]o/i refers to the external and internal potassium concentration, respectively. Just plug in the numbers. For a HEK cell experiment, I keep my internal K concentration at 152 mM. If you use 4 mM external potassium (and we assume room temperature to be 25 C), then the EK comes to -93.4 mV. For 40 mM extracellular K, you get an EK of -34.3 mV.
To simplify: Potassium is high on the inside, which creates a chemical gradient toward the outside. Brownian motion, simply by statistics, will drive K toward the outside. As K leaves the cell, it takes with it one positive charge, making the inside more negative. With every K leaving the cell, the cell gets more polarized (negative on the inside); in other words, an electrical gradient is established that is opposite in direction compared to the chemical gradient (because the positive K+ is attracted to the negative inside). At some point, when the electrical and the chemical gradient are equal but opposite in sign, you have reached EK. There is no more *NET* movement of K toward the outside; there is still K exchange, but for each K leaving, one comes in.
When you raise your extracellular [K], then potassium has a harder time leaving the cell. Therefore, the reversal potential is not as negative.
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Hi,
I have been doing EPSC/IPSC recording from CA3 pyramidal neurons in acute slices using a CsGluconate internal solution (in mM): 
130 Cs-Gluconate, 10 HEPES, 0.6 EGTA, 3 MgCl2, 1 NaCl, 0.4 CsCl, 4/0.3  ATP/GTP, 10 phosphocreatine and 1 QX-314 (recording EPSCs at -70 mV in voltage clamp, IPSCs at 0 mV).
My colleague, who has been doing electrophysiology much longer than I, uses for the same experiment:
140 K-gluconate, 10 HEPES, 1 EGTA, 4 NaCl,  4/0.3  ATP/GTP, 10 phosphocreatine and 1 QX-314 (recording EPSCs at -65 mV in voltage clamp, IPSCs at 0 mV).
When I calculate the reversal potential for Cl-, mine comes to -73 mV and his to -89 mV (we both use the same ACSF with APV).  I can’t get a clear answer from him as to why he uses this solution and concentration of Cl-. 
Can anyone provide insight into this?  If so, I would greatly appreciate it.
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For reliable recording it is important to use K-gluconate. In addition to the problem of juction potential,  Cs blocks K channels and causes decrease of resting potential. So, parts of the cell, which you can't properly clamp (this is usual problem at whole-cell recording from neurons) will be artificially depolarised. 
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Hi, I new started patch clamp electrophysiology techniques and I m working in the cohlear nucleus neurons but I cant distinguish neurons and neuroglia in this area. I realize that most of cells which I did seal is glial cells. What could be reason for it?HowHow do I overcome it. Also would you give me some references /resources for current clamp analysis. thanks.
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Dear Caner, I agree with Henrique and Rogier as long as we consider astrocytes as the only glial cell you may encounter when patching in the CNS... Unfortunatelly for neurocentric oriented patch-clampers, but not for the health of your brain, you have also other types of glial cells that have quite different properties than astrocytes in terms of electrophysiological behavior. An easy one to distinghish is the microglia. In physiological conditions (even in slices) the input resistance of these cells is usually in the order of 1gigaohm. They express, no voltage activated currents and thus no action potential. The resting membrane potential is somewhere in between -20 and -70 mV but not so easy to measure due to the comparable membrane and seal resistances. They are not coupled through gap junctions. If you are in pathological conditions or during normal postal development, things may change (voltage activated potassium channels, chloride channels etc...) but still no action potential!. You can check the papers of Helmutt Kettenmann, Claudia Eder, or mine, (although none of us recorded these cells in current clamp I think). The most difficult ones are the NG2 cells of oligodendrocyte progenitors (OPCs). Those cells express voltage activated sodium channels and can fire small action portentials. David Attwell published a paper some years ago showing that some of these cells can fire repetitive action potentials just like neurons but I am not aware of any other report of this kind. What is sure though is that they can fire one or two-three small action potentials. And they also have actual synaptic inputs, so you will detect IPSPs and EPSPs from these cells, but usually at lower frequency than in most neurons. Also, the properties of NG2 cells change during postnatal development. You should check the papers of Dwight Bergles, Christian Steinhauser, Maria-Cecilia Angulo, David Attwell, Maria Kukley for instance to get a better idea of the behavior of these cells.  Finally, when NG2/OPCs differentiate into oligodendrocytes, they loose their ability to fire action potential, they don't show anymore synaptic currents and they express different potassium channels. Much easier to distinguish from neurons than NG2 cells.
I know this seems to be an awfull situation for beginers with an interest only in neurons but you will learn rapidly to distinguish neurons from glia. Using good optics (DIC or Dodt contrast) also helps to recognize them in slices where they usually have not the same aspect as neurons.
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Hello,
I work on LTP recording,
Thank you,
Matt
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Check your signal reversing stimulus polarity. If anything looks just mirrored vs. 0 level, it is all artifact. 
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I need to measure fine changes in cell capacitance. Does anybody know how to set pClamp 10 and axopatch 200b to have the lock-in amplifier function? I know it is possible on Heka amplifiers through the Pulse function.
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Will not work with lockin amplifier with pClamp 10.  Problem is that pCalmp 10 will not  run lockin function (If anyone knows how to I'd like to know as well).  Axon (Molecular devices) suggest using the sealtest function in pClamp 10 and this works OK.  When you say 'fine' what range are you thinking of sub fF? and at what sample rate?  
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Hi, Can anyone suggest that Lab bench and other settings for current clamp (Action potential) pClamp software. If I am setting the Lab bench Input settings mV for IN=0, the popup showing that "bad membrane signal". Analog IN units must be in Amps with or without a standard SI prefix to be compatible with this test (eg A, Amps, nA,pA the signal Imeb is in 'mV'. How can I set for action potential recordings? Here I attached the settings screenshots ppt slides for best understanding the problem, 
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Hi Rajesh,
I am assuming here that you have pClamp 10 and Axopatch 200B as per the question you submitted on LinkedIn. One last bit of useful information is, which Digidata model are you using?
The suggestion of Olivier to telegraph information from the Axopatch to the Digidata and Clampex is very good. I am just not sure that you can telegraph from the Axopatch 200B. That's a question you can ask Molecular Devices directly.
To configure pClamp for current clamp recording with an Axopatch without telegraphing I would use either Clampex or Axoscope software (both parts of pClamp). Historically I have used Axoscope because I didn't telegraph information from the amplifier (you can't receive telegraphed information on Axoscope whereas on Clampex you can) which in my case was also an Axopatch 200B. Then I would proceed as such:
  1. Plug in the amplifier to the digitizer (usually some version of Digidata).
  2. In Axoscope (or Clampex) configure the digitizer to receiver the data: Configure > Digitizer.
  3. Set the signals you receive: Configure > Lab bench. That's where you need to select which digitizer channel actually receives the signal ("Digitizer Channels" area). Then you configure the signal in terms of units, scale and gain.
  • I am assuming your reference to "IN=0" actually refers to "IN 0" a default signal created by pClamp for the Digitizer Channel "Analog IN #0".  So in this case, "IN 0" does not refer to "I=0" which you can find on the Axopatch 200B mode selector. Instead, "IN 0" in pClamp means "Input signal number zero", it's just the name of the signal created for "Analog IN #0" (Analog input number zero) by default in pClamp.
  • Anyhow, you may configure this default signal in current clamp, or you can create a new signal ("Add..." button in the "Signals" area of the Lab bench window). When you select the signal to be configured you then choose the unit (in your case most likely mV), the scale factor (I guess this should match the position of the "Output gain" selector on your Axopatch 200B amplifier) and the offset if any.
  • Then click OK and your signal should be set in current clamp mode. As confirmation, in your scope window the y axis should show the name of the signal (in this case "IN 0", unless you created a new signal) and the unit in mV.
One potential source of error might be if the scale factor you entered is not accepted by the software for some reason. In this case try using the "Scale Factor Assistant" (still in the Lab Bench window).
I hope it helps.
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I give a tetanus of 100 pulses at 100 Hz in extracellular field recordings from CA1 region of rat hippocampal slices. 
During the tetanus I get a decrease in the current intensity during successive pulses. Why is this the case and how can I resolve it?
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I presume you mean decreased current intensity within the slice? This sounds like simple habituation. If you're interested in cellular mechanism, the likely target would be voltage-gated ion channels. If voltage-gated ion channels are responsible, you could attempt to block them with a compound to "resolve" this.
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I am trying to set-up LTD experiments on acute rat brain slices in vitro.
I tried to prepare slices from P12-P14 and P18-P21 Wistar male rats and used low-frequency stimulation (1 Hz, 15 min) for LTD induction.
Both in the GD and in the CA1 the depression of fEPSPs was very weak and un-stable.
I would greatly appreciate an advice about the best age of rats and a good induction protocol.
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Okay then please check
Sajikumar and Frey, 2003, 2004
Synaptic tagging and cross-tagging: the role of protein kinase Mzeta in maintaining long-term potentiation but not long-term depression.
Sajikumar S1, Navakkode S, Sacktor TC, Frey JU.
Best
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Hello all! Surely those who have worked whole-cell with acute slices can commiserate with the impotent fury of seeing pipette after pipette slain, tangled in the sticky shielding of extracellular matrix.  I am also working with particularly old animals, and they seem to really pack it in densely as they age.
To be specific - even after cleaning I will approach a cell, perfectly in a position that I know will seal on the rare cells that have been given a shave by the vibrotome, using pipettes that have been pulled with a program I know to be stable and effective.... and upon releasing pressure I can get the resistance to a maximum 30-40MOhm with coaxing. Upon further movement it is clear that I am caught in extracellular matrix.
I have tried everything I can think of - traditional cleaning pipettes, clearing with positive pressure, grasping the matrix with suction and attempting to tear it/work a hole in it with the micromanipulator, pulling pipettes to a point and attempting to spear it, leaving tissues for a few extra moments on the vibrotome blade in an attempt to loosen it, and even breaking pipette tips manually with a diamond blade to make rough edges reminiscent of a broken beer bottle in an drastic attempt to rip it apart or just nuclearize the whole layer of cells. The only thing I haven't tried is any digestive enzyme for lack of funding and fear that it may render my prep excessively non-physiological, but at this point I'll try anything.
Has anyone found anything that works? Even for whole-cell my sense of sisyphus is reaching a breaking point.
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Appreciate the replies y'all. 
Agreed that tissue health is critical, Steven. I've found that keeping a low 'time from decapitation' seems to best guarantee that (in addition to the stated prep techniques). This not only leaves more cells alive from which to choose, but makes them better able to withstand the needed disruption of the matrix. 
Shockingly, this week I've had very good luck sending in a first pipette and *actually attempting a patch* on a cell of interest, then pulling that pipette back such that the cell is very slightly stretched and applying the short burst of suction typical of breakthrough at the same time as rapidly pulling the pipette back with the micromanipulator. This was initially done as frustration after a failed GOhm seal, but it seemed to rupture both the extracellular matrix and peri-neuronal net without killing the cell allowing another pipette a successful patch. Extremely resilient little dudes, these cells are.
I've also managed to break the matrix by lowering a relatively large (3-4MOhm) pipette that is unfilled into the bath and allowed it to passively attach to the matrix as it front-fills. An alteration between light suction and slow, low amplitude, but long repeated movement picks up a lot of the matrix - rinse and repeat until the cell is clean. For a particularly stubborn matrix I pulled a pipette to a point and jousted diagonally (it helps if you're playing medieval English music and fretting about the amount to which your glass is doped with chivalry) into the matrix until I made a hole and then proceeded with the above.
Certainly going to try enzymes in the future, but deadlines approach! I'd also love to try organotypic slice culture, but the same applies.
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Hell guys,
I am working on synaptic release triggered by field stimulation. We use the parallel platinum electrode. I can see some release using syt-pHluorin. But somehow my labmate could not see any response using FM dyes. Both FM1-43 or FM 4-64 have no response. But he get obvious release when using 70K ACSF (high potassium ACSF). 
I am confused now. Using the same stimulation protocol, syt-pHluorin could find some release but not FM dye. It seems the stimulation is working. But 70K could trigger some release but not electrical stimulation. It seems the problem is the electrical stimulation. 
I get a lot of pressure from my labmate these days. I wonder what is the difference between 70K and electrical stimulation. Is 70K more stronger than electrical stimulation? However, we using 1200 pulses with ~90V for amplitude and 1ms duration now. Still no responses. 
Can anyone give me some advises about the electrical stimulation to trigger synaptic release? We are using AMPI Iso-Flex isolator.
Thanks in advance.
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(continue) @Chenglong Yu
(roughly R is ~500-1000 Ohm, but it depends). I could not find the design map of the Iso-flex from website, then the internal resistance Ri could not be determined. But for the Max Output Power (MOP), we can estimate based on two parameters: Voltage output: 0-90V, Current output: 0-10mA, thus MOP is roughly 90Vx10mA =900mW. If you set the output voltage 90V, the expected current through electrodes will be around 90V/1KOhm = 90mA, then the power between electrode will be 90Vx90mA=8100mW>>900mW, means the Load is too high for the MOP, or say it is overloaded. In this case, the actual voltage between electrodes would be much smaller than 90V, which might not be adequate for inducing vesicle turnover.
2) If the liquid surface is very high, then the equivalent R of solution in chamber is very small, let's say 100-300 Ohm (but it depends). In this case, we could not neglect the internal resistance Ri of the isolator. The actual voltage between electrodes depends on the ratio of 90V x R/(R+Ri), which might be much smaller than 90V.
It should be noted that in both cases the key problem is the OVERLOAD of power.
In addition, the Iso-flex can only give monophasic but not biphasic pulse, leading to the ionization of solution near electrodes during high-frequency repetitive stimulation, such as 1200APs 10Hz stimulus. This might cause the change of local R and subsequently affect the local vesicle turnover.
Finally, small tip: always double-check the input signal generated by the stimulator and make sure the voltage of signal is > 5V, otherwise one can not even trigger the TTL circuit of isolator and no (or very weak) output pulse will be elicited.
Good luck! 
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Hey everyone!
I want to do e-phys recordings on mouse brain slices. Next to field-potential recordings and patch-clamp experiments (with IR-DIC) I also want to record an intrinsic optical signal. This should be recorded with a circular buffer that is running and updating during the experiment, so I don't miss the start of the event I want to record.
Therefore I am currently searching vor a USB camera that is compatible with the µManager softwarer.
I am not to familiar with camera qualities or technical features, but if you could tell me why the camera you suggest is the best, I'd be glad to look everything up I don't get.
Thanks a lot!
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We tend to use Retiga 200R cameras from QImaging for patching in slices. Thorlabs also sells a few less expensive cameras. Anecdotally I've heard of people even using the DCC1545M (about $350) for slice patching - these are supported by micromanager.
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I am using the pClamp 10 software to acquire and analyse whole cell currents. I am using the attached paper and trying to fit the whole cell currents to equation 2 found on page 207 and (its page 11 of 41 on the pdf document). I am struggling to find an equation similar to it using the same hodgkin and huxley type n4j fit used in the paper (link http://1drv.ms/1gySxVo) so I think it means we have to use the custom equation option but that is very difficult to use. Please can someone help me resolve this issue. I have linked some current traces for you to fit to if that helps (the link is http://1drv.ms/1gySclo).
I did email pClamp support but they have temporarily stopped the support until September.
Any help will be greatly appreciated
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Hi Shakil,
The exponential standard is not the same equation as the n4j equation of the H&H model. 
The approach suggested by Oscar is a convenient solution, but remember to follow his advise in carefully choosing the region to fit. 
The choice of the equation to use for fittings is very important and you must remember that it represents a model of how the physical phenomenon works. The n4j equation implies 4 activation gates and 1 inactivation gate. It must be fitted to the whole current to get the kinetics of both phenomena. 
Since you are interested only in the kinetics of the activation you may use a single or double standard exponential to fit the activation phase of your traces.
I would also use a P/N leak subtraction protocol in voltage-clamp experiments to get rid of the leak current.
For an explanation of the fitting to the activation phase of potassium currents you may refer to Liu & Bean (2014).
Best,
Maximiliano
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