Thor Eysteinsson added an answer:Why does seal resistance fall to less than 50 MOhms when I do whole cell patch clamp?
I am new to electrophysiology in general and I am doing whole cell patch clamp at the moment to investigate the impact of centipede venom on NaV 1.7 expressed by TE671 cells. For some couple of days after learning patch clamps I had everything going on so perfectly with brilliant seals and excellent patches coupled with low leakages ( picture a). However, for some time now my good days are gone as recently I hardly have good patches formed. It appears when I apply negative pressure after the tip of the pipette just touches the cell membrane, the seal resistance increases to somewhat 500-800 MOhms and then decreases to less than 50 MOhms ( picture d) making it impossible measure a sodium current coupled with large leakages (nA). Maybe the pictures may help explain the issue more NB: Pipette solution is 140mM CsCl, 1mM MgCl2, 11mM EGTA, and 5mM HERPES, pH 7.2. We do not fire polish the tips of the pipette in our lab as patch clamps have been achieved in the pass without fire polishing.
I agree with Elise, I don´t apply the Vhold until the seal has been formed, or if the resistance has increased but not all the way to GOhm, then sometimes applying Vhold ssems to help. But the best seals form without Vhold. Then you apply Vhold, and I do that gradually, from 0 to about -70 mV. So what I do is bring the pipette tip to the top center of the cell (very important), release the positive pressure (usually brings the resistance up), apply negative pressure gently until a gigaseal is formed, and then either shift the Vhold or compensate for capacitance until all transients are gone. Then make sure the Vhold is correct at -70 mV, and apply additional negative pressure to attempt for break-in. The last bit I find the most difficult and risky. You need to compensate for capacitance because pipette capacitance is irrelevant, and with whole cell capacitance you can get a value for the cell capacitance if you are interested in current density.Following
John Y Lin added an answer:Are the problems with recording neuronal action potentials using Di-3-ANEPPDHQ, filter or sampling rate issues?
Try to make this brief, currently trying to record action potentials across a neural network, using Di-3-ANEPPDHQ, a voltage sensitive dye which blue shifts during excitation of the cell. My current filter cube has a band pass of 485/20nm, a beam splitter of 510nm, and a long pass of 515nm (see Zeiss filter cube 16). As such, I would expect a reduction in intensity during depolarisation, as the blue shift will reduce the amount of exciting light hitting the sample. Due to limitations of my current camera, I am recording maximally at 500Hz.
I am trying to observe spontaneous action potentials in my neural cultures, however there seems to be no discernible difference between untreated and TTX treated (1uM) recordings (change of F/F0). I have also applied high concentrations of KCl to force depolarisation and that has only reduced signal intensity in a couple of experiments.
Does the problem stem from my current filter set up, or is it due to my low sampling frequency rate? I was planning on moving onto a faster camera next (>2KHz), however I'm anxious that I'm not seeing any change following the KCl. Any suggestions would be greatly appreciated, and if you need to know anything else, please ask.
Using the ANEPP dyes to pick up spontaneous potentials in single neurons are just difficult due to its small S/N as mentioned before. If you look at the literature, it is often used to looked at population response or to look at APs when you know there is one (e.g., when you are patching a cell). There are 'tricks' to maximize your S/N but you are probably not going to spend that much time and money developing your hardware to do so. Molecular Probe is selling a new class of dye made by Evan Miller at UC Berkeley (VoltageFluor) that will give you better S/N and would give you a better chance at this.Following
Gábor Hollandi added an answer:What may cause the facilitation in the fEPSP amplitude in a Haas type interface chamber?
I’m working with a new Haas type interface chamber, called BSC-HT Brain Slice Chamber System Haas Top (Harvard Apparatus). The problem I encountered is as follows: after calibrating the perfusion, the slice doesn’t look dry, yet the fEPSP amplitudes facilitate during the recording (increases by about 400 uV in an hour).
The ACSF is saturated with carbogen and is at room temperature, carbogen is also directly flown into the chamber to produce carbogen-steam. I tried different perfusion speeds (1-4 ml/min), but it didn’t make significant difference. I previously worked with a similar Haas type chamber, and managed to overcome similar difficulties.
I wonder what may couse the facilitation in the fEPSP amplitude - any suggestion is welcome.
Thanks in advance.
Anton Sheinin, I tried different water levels, but it did not help.
Robert A Pearce, I couldn't find a way to modify the perfusion in the manner you suggested, but the same idea occured to me too.
Eric Michael Prager, before placing the slice in the chamber, it can survive sufficient time, but in the chamber it doesn't. I didnt't modify the chamber itself (with pieces of plastic).
I would like to thank everybody for the tips and suggestions.Following
Edina Varga asked a question:Are there any data about electrophysiological features of tanycytes?
I try to patch them, but I have no idea what protocol should I use.Following
William M Connelly added an answer:Does someone have experience with blocking calcium channels on primary cortical culture by means of Cadmium/Nickel/Cobalt-Chloride?
I would like to perform some extracellular recordings on MEAs from primary cortical neurons, where I block Ca+ channels by using CdCl, NiCl, and CoCl.
My doubts are about proper concentrations and how toxic are these compounds.
Moreover, how long I could leave the compounds in the bath without compromising the cell?
Thanks for any help.
The concentrations ARE the concentrations you use in slices/culture.Following
Vicente Lumbreras added an answer:What is the best approach to analyze spike encoding collected using current clamp methods?
I am trying to analyze spike data collected in current clamp to study spike encoding of information in the calyceal hair cell synapse.
-Spontaneous activity data without any stimuli.
-Spike Data collected driven by steps of current.
-Spike Data collected driven by mechanical stimuli of the hair bundle.
Suggestions about which model to use or analysis to make? Thanks.
The data is recoded using an Axon and a Heka amplifier. Other postdocs in my lab use clampfit to analyze the data obtained in these amplifiers.
I am interested also in determining if I can use this data for input/output modeling using Volterra filtering for example or do some information theory analysis with it (calculate mutual information between stimulus and response for example).
Pepin Marshall 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
Thanks very much for the suggestions guys. I tried with Lucifer Yellow CH and it works pretty well, although I may get even better seals with neurobiotin in the pipette.
The Hoechst idea was just to make a UV patch to make locating the dye-filled cell easier, but actually it's not even necessary as I can see the electrode tracks.
Hoechst is non-toxic as far as I am aware though; it is used for sperm sorting in cytometry (if I recall correctly). Ethidium bromide, used for DNA staining in gels is the nasty DNA intercalator. Hoechst will wash off with time, it binds to the grooves of the helix.Following
Daniel M Cohen added an answer:Does pIRES2 express more EGFP than the mRNA before the IRES?
I've subcloned all of my mRNAs into pIRES2 (CMV:mRNA-IRES-EGFP) for expression in cell culture (CHO and TSA cells) and electrophysiology. I have been getting a lot of bright green cells with absolutely no measurable expression of my mRNA: about 50% of the green cells show the expected current, the other 50% react like an untransfected cell.
Is this a common problem with the pIRES?
The IRES-GFP cassette is designed to work in conjunction with a stop codon in your GOI. If this is absent, your GOI may not express properly. Based on what you describe, it sounds like the co-transfection experiment was a better setup. Perhaps the transfection efficiency of the IRES2-GFP plasmid is simply less good in your hands either due to increased plasmid size or due to variability in the quality of the purified DNA.Following
Alessandro Bilella added an answer:DREADD vs. optogeneticsI am thinking of employing both these techniques in my lab and was wondering if people would like to share their thoughts/experience.
From a behavioural point of view I suppose that a major advantage of optogenetics is that you have greater temporal control over stimulation: once a dose of CNO is administered to an animal expressing DREADD there is presumably a time-to-onset and later a decline in receptor occupation and effect, whereas with ChR2/NpHR simulation is phase-locked to light stimulation.
By contrast, I would imagine that light scattering (optic fibre in brain)/failure of light to penetrate tissue sufficiently (stimulation of peripheral nerves in skin) is a drawback of optogenetic stimulation compared to oral administration of CNO, which has known efficacy at different DREADDs.
Any thoughts/comments welcomed!
to use optogenetics instead of DREADD or viceversa depends from the behavioral experiment!
The optogenetic stimulation is time-dependent, while if you would like to record the behavior for a couple of hours the DREADD system is better.
Brad Karain 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.
You may well have "grounded the major areas", but have inadvertently introduced ground loops. The noise you describe could easily be caused by this phenomenon. If you're so inclined, let me know how your troubleshooting goes. Noise can be really tricky. I'm always interested to hear about some new source people finally eliminated.Following
Huong Ha added an answer:How to increase the frequency/amplitude of AMPAR mediated mini EPSCs in culture neurons?
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.
@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?Following
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.
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.Following
Fouad Lemtiri-Chlieh added an answer:How can I accurately test access resistance during whole cell?
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?
Increase your sampling rate tooFollowing
Huong Ha added an answer:What bath solutions do you prefer to use for cell culture recording?
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
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.Following
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.
Thank you all for your answers.Following
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 ?
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.
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.
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.Following
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.
Hi, Do you utilize rtms for off lable applications like panic attacks, mania, tinnitus ... can you explain your protocols?Following
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?
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?
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.
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.Following
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.
Morphology of coronary sinus has been associated with AV nodal reentry tachycardiaFollowing
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?
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..Following
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. ThanksFollowing
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!
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.Following
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.
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.Following
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 ?
Thank you for your help.Following
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!
I agree - Jochen Roeper is your man.Following
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.
I use the same concentration as shown in the Schmidt & Kofuji Jove videoFollowing
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!
Did you adjust the Osm fro both external and internal solution? Because you did not mention it here.Following
Mason Best added an answer:How can I get a input output curve (fiver volley vs fEPSP) like this ?
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.
Thank you very much. I got it.
In the second figure, they just averaged the fiber volley and didn't show the erorr bar.
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.