• Huong Ha added an answer:
    How to increase the frequency/amplitude of AMPAR mediated mini EPSCs in culture neurons?

    Hi all,

    I am interested in AMPAR mediated mini EPSCs in hippocampal neurons. My current recording configuration (hibernate E as bath solution and Cs gluconate as internal) seem to allow me to record from them up to day in vitro (DIV) 25 for about 30 min with fairly stable access. The sad story is that I do not see a lot of mini events (1 - 2 events every 3 - 5 seconds --> much less than 1 Hz). I can see quite a lot spontaneous events starting at DIV 11 already (1 - 2 event every second or so). Does it sound like something you experience before? How would you recommend troubleshooting it? Maybe, like, changing the recording condition or culture condition to have more mini AMPAR EPSCs? Thanks a lot!!!



    Below are some more information if you would like to know....

    When the neurons are younger (Div 12 - 15), there are a lot of action potential driven EPSCs [huge events, > 100 pA]. And when they get to Div 25, there are mostly very small events (20 pA, more or less). The small events decay time is approximately from 4 - 30 ms. 

    Regarding the culture, Coverslips are coated with 1 mg/ml Poly D lysine. I plate the neurons from E17 - E 18 hippocampi at 1.4 millions neurons per 100 mm dish containing 6 coverslips. The coverslips are submerged in serum containing media. The coverslips have wax feet so I can flip them up side down into 60 mm dishes with neurobasal + B27 + glutamax + 20 % media conditioned by astrocytes [which facilitates the growth of a lot of astrocytes underneath the neurons].

    Sometimes I also co-culture the neuron coverslips in dishes with astrocyte feeder layers [in which the cells are not touching each other  i.e. Banker culture]. The viability and development look fairly good. I can see a fairly dense network of dendrites already at DIv 11 and it just gets denser over time. Cells are evenly distributing across the coverslip, not much fasciculation or any sign of substrate problem. I feed them twice a week after the first week.  

    Huong Ha · Stanford University

    @Thomas: As always, many ideas! Thank you! :) It is very helpful indeed!

    - Regarding your concern for LTD, I did not see any reduction of spon events within once cells. About 2 weeks ago, I recored from ~ 4 week old neurons (who were not co-cultured with astrocytes) and they did not have much of the big EPSCs. This week we started to have Banker cultured neurons at ~ 3 week old and these all seem to have very robust spontaneous EPSCs and look very nice (3D-looking, smooth membrane, nice processes).

    - I have not analyzed the data but looks like these new neurons have higher mini frequency already (still lower than 1 Hz though). I might still need to heat them up or try your suggestion with the KCl to get more response and reduce recording time. Thank you for this! 

    - The density is about as high as we can do considering other factors. I tried higher density before but the media just got acidic so quickly and the neurons were not healthy...

    -For your question about cortical and hippocampal neurons, no, they did not look like they had the same final density. The cortical neurons usually looked slightly denser than the hippocampal neurons even if we plated them at same density. I don't have any quantification on this yet though. This was mostly from my quick glance at the traces

    @Saak: That is super insightful! I do use Cs gluconate as internal with all of the voltage clamp experiment but did not really think about the geography of culture processes that hard. :) From my staining experiments, culture neurons do have synapses everywhere, and their dendrites stretch quite far. I have not measured and compared these numbers to in vivo neurons. Would be interesting to learn more.

    I did not know that culture neurons have mainly GABAergic synaptic activity... I saw quite a lot of mini-like activity (events <~ 20 pA, frequency is way higher than 1 Hz) when recording w/o drugs.  And when I added APV, picrotoxin and TTX, the frequency of tthese is like... below 1 Hz. I was thinking the activity was mostly driven by AMPAR and NMDAR since I am holding the cells at -70 mV (close to Cl- reverse potential). Did not at all think of GABA... Did you do some measure with this? 

  • Bruno Truchet added an answer:
    I have been facing noise problem while doing extracellular field recordings. Can anyone help me?

    I have checked the suction too. Its fine. I have also grounded the major areas, still it doesn't help.

    Bruno Truchet · Aix-Marseille Université


    if the noise frequency is around 50 hz, or a multiple (harmonic) of this frequency, you should try to borrow an apparatus like the HumBug and check whether it could fix your problem. It's way much better than a plain 50hz filter, and very helpful when performing extracellular recording in a noisy environment.

    All Best


  • John Hildyard added an answer:
    Why do I get blurred bands for low MW proteins but good bands for high MW ones?

    I use 4%/12% acrylamide gel, running at tris/ glycine buffer, initially 50V/ 20 min and then 125V, 90 min, constant voltage. Transfer in semi-dry system (TransBlot, Bio Rad) 25V (constant)/ 15 min.

    John Hildyard · Royal Veterinary College

    Smaller proteins are inherently more mobile, so will always tend to diffuse to a greater extent than larger proteins. Plus they travel through a lot more of the gel length: every bit of gel travel distance has a small chance of lateral diffusion, so you'll always see small proteins spread out more than large ones. Getting large proteins to move at ALL is pretty tricky. :)

    A gradient gel is a great way of minimising this effect, but it's still essentially inevitable.

  • Fouad Lemtiri-Chlieh added an answer:
    How can I accurately test access resistance during whole cell?

    Hi all,

    When I record EPSC, I give a 5 mv voltage to test access resistance and input resistance. I found lowpass bessel filter can significantly affect the access resistance. The access resistance is totally different in 2k Hz and 5k Hz.  The sampling rate is 10K Hz. Gain is set at 5 or 10.

    What do you think lowpass bessel filter is OK for whole cell recording?

    Fouad Lemtiri-Chlieh · King Abdullah University of Science and Technology

    Increase your sampling rate too

  • Huong Ha added an answer:
    What bath solutions do you prefer to use for cell culture recording?

    Hi all, 

    Would you mind sharing some of your experience with hippocampal cell culture recording? I have been testing out Hibernate E (a defined media from Brain Bit) and tyrodes (similar formula to ACSF but has only 100 mM of NaCl, and uses HEPES to buffer instead of HCO3). The recipe of Hibernate E is below. My experience is that the Hibernate E keeps the cells for much much longer. I can recording up to 2 hour from the same coverslips, and each recording can last nicely for 30 min (for tyrodes, they usually last for 15 min or so) . The cells also seem to be very active in this media. There are a lot of spontaneous activity. Have any of you used this solution? What would you think could explain for the high activity level? Is it due to all of the supplement like vitamins or amino acids in this media that is very similar to that of neurobasal? Would it be a problem to publish using this pre-made media instead of tyrodes/ACSF?

    I am really curious why people use tyrodes or ACSF because the sodium concentration in neurobasal is like... 51 mM (recipe below), which is much lower than a normal standard ACSF recipe. Maybe to compensate for the fact that we add only a few salts into ACSF, hence we need to dose up the sodium chloride to maintain the osmolarity? 

    Hope to hear from you! 

    Thanks very much!

    Hibernate E Media


    Neurobasal media


    Huong Ha · Stanford University

    Thank you Ewa for the answer! :) 

    Hi Victor, Yes, neuronal cells are super sensitive to the difference. The other day I made a mistake with the osmolarity pairing and the cells just crapped out after 10 min or so. I also thought it is fairly strange that my neurons like the internal to be hyper-osmotic in comparison to the external... My new lab is also an intensive ephys lab and everyone else is doing slice recording with 300 mOs external ACSF and 280 - 290 mOs internal. I just analyzed the access resistance from several cells I recorded recently and they either stayed very stable up to 30 min or somewhat improved after the first 3 min. And the values usually below 20 mega ohm. I was never able to have an access as good as this when using hypo-osmotic internal.

  • Randy Stout added an answer:
    Does anyone have any information about how to repair Siskiyou 3axis hydraulic micromanipulators?

    Model MX630R Siskiyou micromanipulator for electrophysiology. All 3 of the axes no longer work. The control knob turns OK but produces no movements in the manipulator at the the other end of the hydraulic line. None of the parts seem to be stuck. If there needs to be more water? hydaulic fluid? introduced to the line then where and how can I inject it? If there is a bubble, how can I get rid of it? Does anyone have knowledge that something else tends to go wrong with these things that I might have overlooked? I don't want to have to send it back to manufacturer and have downtime and cost of repair.

    Randy Stout · Albert Einstein College of Medicine

     Thank you all for your answers.

  • Troy A Hornberger added an answer:
    Why don't I see hypertrophy of plantaris after ablation of synergistic muscles?

    I removed gastrocnemius and soleus muscles from two months old WT mice (n = 4) but apparently two weeks of overload had no effect on plantaris size, compared to sham-operated mice. The wounds healed nicely and the mice were active so inactivity is out of the question. All mice are young and in growth phase. I identified plantaris correctly next to the bone. Literature reports 50-100% increase after two weeks of overload but I see no change in CSA.

     Any suggestion or explanation ?   

    Troy A Hornberger · University of Wisconsin, Madison


       I'm curious to know if you ever solved the problems you were having with this model. In C57 mice we typically see a very large increase in muscle weight, but fiber CSA does not change very much (type IIa increase, but type IIb do not, type IIx response is moderate). Instead we see a very large increase in fiber number. We also only remove about 1/2 of the gastroc, basically up to the point of where it becomes firmly attached to the plantaris. Would love to hear a response.

    Troy Hornberger

  • Matthew Van Hook added an answer:
    Patch clamping cultured neurons - can anyone help?
    I'm a fairly experienced slice physiologist but a current short project requires me to patch neurons in cortical / hippocampal cultures (I need whole cell VC recordings to be specific). I only gave it a couple of tries so far but it did not go as well as I originally expected.

    Forming a gigaseal is no problem but my cells all appear to be depolarised upon break in. Shortly afterwards they usually start blebbing (osmolarity issue I presume). The cells are cultured in neurobasal media with the usual b27 supplement. I have tried both my "normal" ACSF - internal combination (osmolarity 300 and 280 respectively) and some "special ACSF" with Na concentration and osmolarity matched to the media (and low osmolarity internal of course), no luck.

    Could someone point me towards some good literature or tutorials in whole cell recordings in culture? Any tips and tricks would be appreciated.
    Matthew Van Hook · University of Nebraska Medical Center

    I've had success with cultured hippocampal cells bathed in a HEPES-based extracellular solution bubbled with 100% O2 and warmed to 35degC. The recipe is 125 NaCl, 5 KCl, 3 CaCl2, 2 MgCl2, 10 Glucose, 10 HEPES. I make sure to adjust to ~285-290 mOsm and pH to 7.4. My intracellular solution is Cs-Gluconate-based with pH = 7.2 and Osm = ~275 mOsm.

  • Maryam Momtazan added an answer:
    Are there any studies comparing different stimulation frequencies in paired associative stimulation (PAS)?

    I'm planning a study about PAS in Huntington's Disease and have already found information about effective ISIS and number of stimuli. Now I'm just missing some info whether there are frequency effects and if they can be modulated.


    Maryam Momtazan · Ahvaz Jondishapour University of Medical Sciences

    Hi, Do you utilize rtms for off lable applications like panic attacks, mania, tinnitus ... can you explain your protocols?

  • Saivishal Daripelli added an answer:
    Which region is best for implanting reference electrode for measuring theta oscillations in CA1? How can I get peak frequency of 7-8Hz in CA1 region?

    I want to measure modulation of theta oscillation in CA1 (Hippocampus) region by stimulating NPO (Nucleus Pontis Oralis) in rats. I have gone through the literature where researchers used Frontal bone or ear bar or cerebellum as reference. Can anyone suggest me, which region is best for reference electrode to get synchronized theta waves?

    In literature it was mentioned 0.5 to 3.0V current is required to get synchronized theta. But we are getting synchronized theta in CA1 (cerebellum as reference), only when we stimulate NPO by 3.0V and peak frequency was 4-5Hz. Is there any possibility to get peak frequency of 7-8Hz with low voltage stimulation?

    Thank you

    Saivishal Daripelli · Suven Life Sciences Limited

    Thanks Bali and Molden for your replies. My apologies for the delay in acknowledging the same.

    Your suggestions are really helpful for us.

    By using constant current  (60 uA) for stimulation now we are able to get theta oscillations of 5-6Hz with cerebellar reference, but after Donepezil treatment we are observing decrease in amplitude of theta (which was reported to increase). What could be the reason for this?

    thank you.

  • Amir Ghayoor added an answer:
    Can someone explain how to calculate the cell capacitance?

    Can someone explain how to get the dielectric permittivity and conductivity of a neuron cell?

    I am making the voxel of a neuron cell to investigate affect of electromagnetic wave on it.So I need to have biological parameters of it.In my model there are 3 parts and I want to assign every part with a permittivity and conductance .These three parts are:1) extracellular side, 2) membrane of the cell  and  3) inner of the cell or cytoplasm.

    Thank you

    Thanks Gary
    I am working with pyramidal and motor neuron.All the property are in their simplest forms at the first.I think this special topic is directly related to biophysics.

  • Sergio Pinski added an answer:
    Is there an established reason why some patients show evidence of dual pathway electrophysiology in the AV node, and others not?

    Dual pathway electrophysiology in the AV node is found in 50-90% of patients with AVNRT, and around 10% without AVNRT (see link #1). Since it is the differences in refractory period between the fast and slow pathways which give rise to a discontinuous conduction curve, it is tempting to say that patients with a continuous AV conduction curve have no difference in refractoriness between the fast and slow pathways. However, it has also been suggested that a discontinuous AV conduction curve may be suggestive of longitudinal dissociation of the AV node i.e. a structural change. Is there a definitive answer as to whether it is differences in structure, electrophysiology, or both, which give rise to continuous and discontinuous AV conduction curves? Any references to recent articles would be appreciated.

    Sergio Pinski · Cleveland Clinic

    Morphology of coronary sinus has been associated with AV nodal reentry tachycardia

  • Sakthivel Govindaraj added an answer:
    Advice on Choice of Anesthesia for in vivo electrophysiology - can anyone help?
    I am performing in vivo electrophysiology on Sprague-Dawley rats, recording local field potentials and spiking activity in the VTA, SN and PFC.

    I have narrowed my choices down to Chloral hydrate, Isoflurane and Urethane. Aside from differences in route of administration, are there any real advantages of one over the other? Is there a better anesthetic agent apart from these three I mentioned?
    Sakthivel Govindaraj · University of Madras

    Isoflurane is the best choice of anesthesia mild effect and recover soon but for this need special gaseous unit.. in this case urethane is next choice of anesthesia this has the minimal cardiopulmonary suppression effect..

  • Rabih Moshourab asked a question:
    Anyone knows whether multielectrode arrays are used in to study the electrophysiologic characteristics of delirium?

    I need any study or reference that might be of help explaining the technique. Thanks

  • Michael Country added an answer:
    In patch-clamp, my cells appear constantly depolarized (Vm = +10-30 mV!). What could be going on?

    I'm patch-clamping horizontal cells from goldfish retinas, and the membrane potential is constantly reading 10-30 mV (from "I = 0" setting on my AxoPatch 200B amplifier). I don't know if it's physiological, pathological, or a problem with the equipment. When I put on glutamate, the cells repolarize towards the glutamate reversal potential (0 mV), so I think it's not an equipment problem - but I'm all ears if you have advice/ideas.

    I've tried less papain during dissociation; this didn't change anything. I've tried whole cell and perforated patch over and over - no difference. I've replaced solutions and tried different ones. I've checked my osmolarity and tried different ones. I've tried new batches of fish. Any ideas?

    Please help! You'll be my hero!

    Michael Country · University of Ottawa

    Hi everyone; thank you for the help. I've been troubleshooting in a number of ways, recently. Here's the story about my junction potential troubleshooting. So, after rechloriding the wires and changing the AgCl pellet, I still had high (and drifting) junction potentials.  I cleaned my old wires (cleaned under flame to remove Cl-, then cleaned with ethanol, and then soaked in bleach for 30 min to rechloride them), which then showed JPs from 15 mV (ECS to whole cell internal) to 35 mV (ECS to perforated patch internal with amphotericin B). Both JPs drifted strongly. I found a new spool of silver wire and that helped a lot; I guess the wire I had wasn't good anymore or something? The new JPs were roughly 4 mV and 8 mV, respectively - but the perforated patch internal JP drifted. Every JP reading after that drifted, too. So here's a question for you all: is there something other than the wires that would cause drift? I'll soon try another lab's equipment and see what happens.

    Eduardo; how would I check for [K+]? I remade my internal/external solutions and double-checked my calculations/osmolarity, if that's what you mean. Is there a better way? Values are below.

    Francisco; all in mM, my normal extracellular solution is:

    • 120 NaCl
    • 5 KCl
    • 2.5 CaCl2
    • 2 MgCl2
    • 10 HEPES
    • 10 glucose
    • buffered with NaOH to pH 7.8
    • osmolality checked to be 280 mmol/kg

    My whole-cell internal is:

    • 10 NaCl
    • 120 KCl
    • 0.5 CaCl2
    • 2 MgATP
    • 5 EGTA
    • 10 HEPES
    • buffered with KOH to pH 7.4
    • osmolality checked to be 276 mmol/kg

    My perforated patch internal is:

    • 110 K-gluconate
    • 10 KCl
    • 10 NaCl
    • 0.5 CaCl2
    • 5 EGTA
    • 10 HEPES
    • 2 MgCl2
    • buffered with KOH to pH 7.4
    • 0.26 mM amphotericin B (0.24 g/L), added just before patching by premade aliquots
    • osmolality checked to be 336 mmol/kg (!) - but the amphotericin's sticky and I don't completely trust that I can properly use this in our osmometer. I've gotten recordings with this high osmolality, consistently. Please let me know if this would be a separate issue!

    I've also tried a number of other externals, including ones with K channel blockers (Cs, TEA, and 4-AP). My cells looked depolarized with all internals and externals, and I've gotten Vm readings from +15-40 mV depending on the day, the solutions, etc. I haven't gotten negative readings yet.

    Martin: Yes, I'm new at patching (6 months now). I've been reading like my life depended on it and trying to be meticulous, though. I have good I-V curves that match the literature for the cells I'm working on, but the I=0 data (the membrane potentials) seem off. My cells aren't spiking cells; they're fish horizontal cells, which don't have voltage-gated Na channels as far as I know. I've been doing mostly voltage-clamp.

    Pipettes are ~ 3MOhms, which matches some literature values. I'll check this out next time I patch (tomorrow or the weekend.

    Yes, there is drift in the bath. Even after rechloriding and replacing the ground, and then changing the wire, there's a drift even in the bath - albeit slower than before.

    Yes, I only "count" GOhm seal recordings. Yes, my seals generally remain constant when I get them. Sometimes it's harder with amphotericin B, but once I get > 1GOgm resistance, it stays or I move on to another cell.

    I usually do voltage-clamp, but always start with a -60 mV clamp. Then, I test Vm by switching to I=0. Recently, I've been turning off "external commands" on my AxoPatch 200B, before testing the Vm - although it's the same as my I=0 data, I get superstitious, yeah? I've been switching to I=0 right after getting a gigaOhm seal, and occasionally throughout the recordings. No, I haven't tried checking Vm stability at a given, injected current. I'll try it out - although I haven't done much current clamp stuff yet. I'll have to read a bit first!

    William: I've done both recordings. I generally use perforated patch but I've tried whole cell as part of my troubleshooting process. Perforated patch was generally a bit more positive.

    Next, I plan to try another lab's electrophysiology setup to make sure my setup isn't the issue. One last question: when you rechloride the wire, do you rechloride the whole thing? Or do you leave the contact, where the wire would touch the gold pin?

    Thank you all.

  • Hiroki Yasuda added an answer:
    Why is the frequency of mEPCS of CA1 pyramidal neuron so low?

    Hi, I am trying to record the mEPSC in CA1 pyramidal neurons. I found the frequency is so low, at 0.1Hz, The acsf is normal, with 1.3mM MgCl2 and 2.5mM CaCl, 2.5mM KCl. I also added 1uM TTX.

    I am OK with whole cell. The access resistance was less than 20. 

    How to improve the frequency of mEPSC?

    Thank you very much.

    Hiroki Yasuda · Gunma University

    If you got cells with visual guidance, why not try a blind method?

    Typically mini EPSC frequency in the CA1 in developing mice is higher (-0.8 Hz) than that in rat (<0.5 Hz) when I used the blind patch.

  • Raiko Blondonnet added an answer:
    Does an assay exist to measure ENaC functional activity without the use electrophysiology ?

    I search an easy test to measure in vitro ENaC activity on lung but without use electrophysiology. Does fluorescent kit exist ?

    Raiko Blondonnet · Centre Hospitalier Universitaire de Clermont-Ferrand

    Dear Oliver,

    Thank you for your help.

  • John N J Reynolds added an answer:
    Can someone point to a reference characterizing the electrophysiological properties of the MOUSE dopaminergic neurons in the SNc?

    All of the papers I've found so far are on rats. I need some reference for the electrophysiological properties of the dopaminergic neurons in the substantia nigra pars compacta in the MOUSE. Thanks!

    John N J Reynolds · University of Otago

    I agree - Jochen Roeper is your man.

  • Justin P Rodriguez added an answer:
    Has anyone patch-clamped neurons with Hoechst in the pipette?

    I am performing patch-clamp in whole brains in vivo and including a fixable intracellular dye

    In order to locate the cell I have recorded from more easily, I figured if I add some Hoechst to the intracellular solution, then; as it is highly fluorescent, membrane-permeable and does not interfere with living cellular functions, it would serve as a general locational marker.

    Has anyone else tried this? I am expecting to see a nice UV patch in the cortex I have recorded from that should guide me to where the cell I have patched is :D

    Justin P Rodriguez · Stony Brook University

    I agree with Alexander. Use need to use a cell non-permiable dye. If you do not want to do a post-stain to see your cell, AlexaFlour dyes are nicely biocompatable. Very little Hoescht will stain any cell your pipette got close to and make your pipette solution into a potent mutagen/carcinogen/teratogen. 

  • Michael Risner added an answer:
    How do you remove the inner limiting membrane (ILM) for whole cell patch clamp of retinal ganglion cells?
    I usually patch bipolar and amacrine cells in retinal slices, but I am trying to do some wholemount stuff at the moment. I have varying degrees of success with either clearing the membrane with an empty, broken pipette first, or just applying a lot of positive pressure on the recording pipette, punching through, and then recording. The latter technique seem to work reasonably well but if I have neurobiotin in the pipette it labels all the surrounding Muller cells. I know everyone does this differently, so I'd really like to hear different suggestions.
    Michael Risner · Vanderbilt University

    I use the same concentration as shown in the Schmidt & Kofuji Jove video

  • Qing Yang added an answer:
    How can I increase spontaneous activity during Patch Clamp recordings?

    I am performing whole-cell patch clamp experiments on cultures of dissociated cortical neurons (DIV ~14-30) grown on microelectrode arrays (MEAs). Inside the incubator, I keep the cells in Neurobasal medium, where they exhibit nice spontaneous firing, which I can measure with the MEA. When I change the media to HEPES-buffered external bath solution containing (in mM:) NaCl 149, KCl 3.25, CaCl2 2, MgCl2 2, HEPES 10, Glucose 11 (pH: 7.35 adjusted by using NaOH 1M) for doing the patch clamp experiment (at room temperature), the spontaneous activity immediately goes down and most of the times disappears. The cells however remain healthy for ~4-6 hours. I can induce spikes by increasing the extracellular potassium concentration, but this generally leads to strong bursts, which are unfavorable for my experiments. Any suggestions/experiences which medium I can use to get spontaneous activity during patch clamp experiment?

    Thanks a LOT for any help!


    Qing Yang · University of Texas Health Science Center at Houston

    Did you adjust the Osm fro both external and internal solution? Because you did not mention it  here.

  • Mason Best added an answer:
    How can I get a input output curve (fiver volley vs fEPSP) like this ?

    Hi all,

    I want to know how get a input output figure like the attached. There is no error bar in fiber volley.

    Thank you so much.

    Mason Best · Zhejiang University

    Hi Olivier,

    Thank you very much. I got it.

    In the second figure, they just averaged the fiber volley and didn't show the erorr bar.

    Thanks again.

  • Fouad Lemtiri-Chlieh added an answer:
    Are NaH2PO4 and NaHCO3 necessary when HEPES is used in the external solution?

    I am searching for solutions to be used in perforated patch experiments in pancreatic beta cells. People use HEPES as well NaH2PO4 and NaHCO3 in non-oxygenized external solution. Are NaH2PO4 NaHCO3 meant to act as buffers? Are they really necessary when HEPES is used in the same solution?

    Fouad Lemtiri-Chlieh · King Abdullah University of Science and Technology

    To my knowledge. the straightforward answer is NO. Yo don't need both buffers (i.e., Hepes and the phosphate/bicarbonate in the same time) in non-oxygenatd EMs. The sodium phosphate and bicarbonate buffer will only be efficient if you use carbogen bublling.

  • Mason Best added an answer:
    Can anyone help with the difference of hippocampal slices oientation for electrophysiology?
    Hippocampal slices for patch clamp and field potential, are prepared from mice using coronal, horizontal, sagital, parasagittal or transverse section. A few papers show that slices orientation can affect synaptic plasticity. (Bartlett et al., Mol Brain. 2011 Nov 15;4:41. doi: 10.1186/1756-6606-4-41.,Sherwood et al., Neurochem Int. 2012 Sep;61(4):482-9. doi: 10.1016/j.neuint.2012.04.021. Epub 2012 Apr 27.)
    It is convenient and quick to get hippoampal slices from coronal, horizontal, sagital, parasagittal method. Some researchers said that cutting in a parallel way of the apical dendrites growth,transverse section, make slices health. But transverse section method is a little tricky and need more practice.
    1. Which orientation are used in your lab to get hippocampus slices? And why?
    2. I found that I got a smaller field EPSP response in horizontal and parasagittal slice than transvers slice.
    Mason Best · Zhejiang University

    Hi guys,  

    Most people wrote they got transverse sections of hippocampal slices. But they didn't say how.

    Can anyone tell me how to get transverse sections of hippocampus?

    Thank you.

  • Seyed Mousavi added an answer:
    How we can detect electrical signals in plants?


    I am research about communication between parts of plant. for example how plant close stomata in drought or heat stress.  I want know how we can detect electrical signals in plant. is there any equipment and instrument??

    thank you



    You may also find detailed method for surface electrical signals measurement on Arabidopsis leaves from the articles that recently published in Nature 




  • Eric L Hargreaves added an answer:
    What is an acceptable amount of variability in fEPSP peak and slope between sweeps in hippocampal recordings?

    I'm assessing LTP by stimulating the CA3 schaffer collaterals and recording for CA1 and I'm wondering what is an acceptable amount of variability in the peak and slope between each sweep from a single slice. As of now my slope varies by as little as 0.15 mV/ms to as large as 0.4 mV/ms (calculated by subtracting the smallest from the largest slope value during baseline). Are there any standard criteria to determine an acceptable amount of variability in a slice?

    Thanks for any and all help!

    Eric L Hargreaves · Robert Wood Johnson University Hospital

    Between slice runs can vary more than 5%, due to dissection health, vibratome slicing protocol and slice number within sequence (discard first few ans last few), temperature of Acsf, age of the pefrusate etc... within a single slice once recording fEPSp measures started should vary less than 5% and baselines should be carried out for minimum of 30minutes prior to tetanization. Also need to be careful when running baseline that the number of sweeps recorded per minute does not exceed 6 (1/10s; and 4/min or 1/15s is preferable), as there is a little known phenomena known as low frequency potentiation, which can appear based upon other variables ie bath temperature Acsf composition etc... Can check out figures and details from Cliff Abraham's lab. Paper listed below can be downloaded from my research gate or my website "Eric L Hargreaves PageONeuroplasticity". Other papers from that lab around that era, should also be very consistent on methods and recordings etc. Particularly slope measurement and duration of slope measurement segment and whether segment is closer to inital roll off or slid closer to the peak. most canned packages should have rules of thumb for measurements, or reference a number of papers that utilized the system and its measurments Its pretty amazing what can be done with the same sweeps in different hands and thats why its important to document document document.


    J. L. SWANSON-PARK,*† C. M. COUSSENS,† S. E. MASON-PARKER,† C. R. RAYMOND,†E. L. HARGREAVES,† M. DRAGUNOW,‡ A. S. COHEN†§ and W. C. ABRAHAM†  Neuroscience,  1999, Vol92, p485.

    Can also checkout fEPSP as % of max I/O value and its impact upon degree of potentiation. Where you are on the I/O curve also impacts the variability of measures. the closer you are to max the less variable are the measures. The trade off of course the closer you are to max the less LTP you will observe (which is more complex than just playing with percentages versus absolute differences). Can check out Ambrose Au's thesis in Stan Leung's lab. Brain Research Bulletin in 1994 (when BRB was a reputable journal).

    Long-term potentiation as a function of test pulse intensity: A study using input/output profilesOriginal Research Article

    Brain research Bulletin, 1994 Vol33, Pages 453-460
    L.Stan Leung, Ambrose S. Au

  • Rose M. Richardson added an answer:
    What are the gold standards to diagnose fit, seizure and syncope?

    There are thin lines amongst the three maladies. Could someone assist to define the gold standard in one's country in diagnosing any of such maladies thus making it easier to start treatment regime?

    Rose M. Richardson · Indiana University-Purdue University Indianapolis

    Exactly.  That is his diagnosis.  

  • Octavio Gonzalez-Lugo added an answer:
    What is the biological explanation for the Triple exponential fit?

    Some calcium inactivation currents can only be properly fit with a triple exponential. What is the biological explanation for this?

    Octavio Gonzalez-Lugo · Center for Research and Advanced Studies of the National Polytechnic Institute

    Dejar Manuel Castellano-Muñoz 

    A double   exponential fit would mean an activation-deactivation process, while a simple exponential fit it's more complicated, depending on the shape of the function could mean 2 different things, a function like {a*exp(b*T)}  where a and b are parameters ejemplifies a dissipative process like facilitated diffusion, but a function like {a*exp(b-T)} could represent an activation-deactivation process, but the change between one of this two states is not kinetic process, like an activation\deactivation fixed  threshold. 

  • Vini Gautam added an answer:
    What might cause cells to "burst" after taking them out of the incubator?

    I have been growing neuroblastoma B50 cells on glass coverslips and the cells appear healthy when I see them under the normal microscope. For electrophysiology, I take the coverslip out and place it on the stage of the recording microscope, and dip it with the same culture medium that it was inside the incubator. Then I see the cells under a 40X objective. To start with the cells look okay, and then suddenly their membranes start to rupture. At the end of 5 mins, they look like they have burst (I am attaching an image of a burst cell that I took with my mobile from the computer screen). If I wait longer, all of them burst completely. This effect is faster if I dip the coverslip in fresh DMEM (in this case they burst within 1 min).

    Few other details: Osmolarity of the growth medium is 320mOsmol. Another thing is the temperature of the recording stage is 27deg. 

    Can the drastic temperature difference be the cause? Or the pH of the medium changes so suddenly after taking it out from the incubator? Can someone please help me in this regard?

    Thanks heaps!

    Vini Gautam · Australian National University

    Thanks! To clarify, the coverslips are usually in a 24-well plate with 1 ml medium each. When I want to do electrophysiolgy, I take the coverslip out, put it on top of the recording stage, and then take the same 1 ml medium from the 24-well plate to cover it. So at the end, the cells and coverslips are dipped in the same medium that they were growing in...

  • Janina Kowalski added an answer:
    Is there a way to confirm the identity of the mouse M1 neuron?

    If I am going to do an in vivo patch clamp on mice M1 neuron (primary motor cortex) only, is there a way to identify/confirm the area?

    To the best of my knowledge, M1 is close to M2 and these 2 areas are quite "irregular" (a comma shape), If I don't identify these 2 areas, I may patch the wrong neurons.

    Thus far, I just know that I may confirm my neuron by: stimulating it and observe if the whiskers will move subsequently. However, just this method alone seems to me not sufficient to convince me that this is the M1.

    Thank you very much in advance.

    Janina Kowalski · IST Austria

    Hi Ping Kwan,
    as Marcel suggested, it is a good idea to stain the neuron that you patch for post hoc analysis. We had excellent results by adding 3 mg/ml biocytin to our internal solution. Find more details in: Pernía-Andrade and Jonas. Neuron. 2014 Jan 8;81(1):140-52

    Good luck!

About Electrophysiology

The study of the generation and behavior of electrical charges in living organisms particularly the nervous system and the effects of electricity on living organsims.

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