- Toshio Murase added an answer:1Does anyone know how I can get precise data about ion pumps in cell membrane?
I am searching for data about ion pumps in cell membrane to improve my model on my e-cell. My problem is lacking data of numbers of ion pumps(for instance, how many Na+ pumps in one cell) and their coefficient of ion diffusion. Would you please provide me some resources about these data，or just telling me the reference or database to search on?
I have no idea.Following
- Konstantin Loganovsky added an answer:1Has anyone separated (cut) the hippocampus in vivo whilst recording LFPs?
Does anyone have experience in cutting the hippocampus (to separate dorsal from ventral hippocampus) in vivo (anesthetized rat) whilst stimulating and/or recording electrophysiology? If so, would you recommend me which coordinates and methods are less damaging? (scalpel, blade...?)
Thank you so much in advance!
You will laugh, at present we are conducting such experiments. However me is neuropsychiatrst and cannot cut hipocampus from humans. But my colleague - Prof. Victoria Talko, Director of our Institute of Experimental Radiology and her team do this. Contakt them = firstname.lastname@example.org
You may say her that it's me for forwarding.
- Saak V. Ovsepian added an answer:7With my In vitro electrophysiology, where is FV, and PS?
I work on LTP recording, I have figure, see attached, but I do not see any FV, Could anyone know where is FV, and PS? or not show both.
A simple test with paired-pulse stimulation at ISI 40 ms should reveal what actually your recordings represent. If the slower negative deflection becomes bigger, then your recordings represent a mixture of FV and post-synaptic response (the slower tail). If the response remains unchanged or becomes smaller, then you are dealing with the fiber valley only. Or perhaps you are recording the population discharge of pyramidal neurons in response to their direct stimulation (this can happen if the stimulus is too strong and slices are bad). The positive deflection of your traces indicates that the current sink is located far from the tip of your recording electrode, hence mis-location of the recording electrode. Try to keep your electrodes far apart, use weaker stimuli and place the electrodes further down from the s.pyr.
Hope this helps,
- Jonathan David Lippiat added an answer:8How can I measure fine changes in cell capacitance?
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.
It is possible to do this without lock-in. See this paper, but I am afraid that I cannot offer any guidance on the contents or implementation. I have EPC10s in my lab so never needed to go into its detail.Following
- Ruta Vosyliute added an answer:3How to measure Monophasic action potential?
In Langendorff apparatus (controlled or constant flow), what devices are necessary to acquire monophasic ventricular action potential in rat heart?
I'm also working on Langendorff-perfused hearts (mostly rabbits) using optical mapping and standard glass microelectrodes. I'm attaching a paper, maybe you will find some info there. If you need more information, let me know. Best regards!
- Norbert Weiss added an answer:3How do I dissolve heptanol into my physiological solution?
Hello, I am trying to perform gap-junction blocking experiments with Heptanol, which is not miscible with water. I have a 1M solution of Heptanol dissolved in 100% ethanol that I am adding to my perfusate to yield a (heptanol) concentration of 1.5mM. However, I am concerned that this is yielding the same result as simply adding raw heptanol to my perfusate (i.e. that the heptanol separates into its own phase). I should say that I have seen the method I am currently using in papers trying to achieve the same gap junction blockade.
Am I doing this right? If someone with experience of using heptanol for electrophysiology or calcium imaging could help me out, that would be much appreciated.
The only think you should check is that your 0.15% final ethanol has not effect by itself on your preparation (but it should be fine).Following
- Michael C Wiest added an answer:4Has anybody used Utah Array in their research? Is it a good option for rodent subjects?
I am contemplating about using them on mice
Utah arrays are about ten times more expensive than microwire arrays, so that's another reason you might want microwire arrays for rodents.Following
- Mao Yeh added an answer:9Why is my baseline drifting dramatically after offset and before whole-cell patch with the Cs-gluconate internal, but not the K-gluconate internal?
There is no other difference besides K+ and Cs+.
Is this your situation? Every time you change your pipette, the holding current is more than 2nA and gradually decrease.Following
- Sergio Martínez-Bellver added an answer:2How do I calculate theta index?
I want to analysis of theta wave of Hippocampal place cell. Any one explain or Share some good reference about " how to calculate theta index"?
I am not sure about what kind of Index you want to calculate. However, in one of my papers I used some index in order to evaluate the relation between single units and the hippocampal theta oscillation.
Hope it helps.Following
- Tomoki Kazawa added an answer:4Which cesium-based intracellular medium to use for calcium current measurements?
I want to do some calcium current measurements in dissociated hippocampal cultures, to make an I-V plot. I suspect the mutated protein I express, influences the VGCC-conductance. In order to isolate the calcium currents I plan on blocking the sodium-currents with TTX and use a cesium based intracellular to block the potassium currents.
My problem is choosing the cesium based intracellular medium. After doing a quick the literature I have seen several mixtures. I have seen recipes with either: Cs-glutamate, Cs-sulfate or Cs-methanesulfonate. Can someone help me by explaining these compounds?
Furthermore, I have seen some use barium instead of calcium to assess the conductance of the VGCC. Why use another ion?
Thanks in advance,
When not using Cl ion, the experimenter wants to care about Cl concentration or cl activated processes( Cl channel Cl activated Ca channel and etc.).
As a replacement of cl, probably methanesulfonate is newest (improvement from sulfate) among candidates you shown.Following
- Marco Mainardi added an answer:6I'm moving to record LFP in awake rats, but I have problem to figure out how to build the chronic implant. Do you have some suggestions for me?
I want o record the LFP in different place, but I don't have a clear Idea how to do it, can you suggest me some protocol or paper about this subject?
Daniel has already provided you several essential infos... If I may add:
- you can also consider using stainless steel wire electrodes (Advent Research Materials # FE632211) instead of screws, although the latter can provide additional stability (which is desirable in rats);
- if you plan to detect "subtle" events, such as inter-area LFP differences, performing bipolar recording can increase spatial specificity;
- the whole implant should be secured in place by using, e.g., dental acrylic cement (Paladur);
- you can check PMID 24478697, 22577757, 21966482 papers I published with experiments on mice.
Hope this helps,
- Saak V. Ovsepian added an answer:4When graphing IO curves from field recordings, how do I convert my stimulation parameters to µA?
I see papers graph IO curves as "fEPSP (mV/ms)" on the y-axis and "Stimulation Intensity (µA)" on the x-axis. How do I convert my stimulation parameters (ex: 2V at 40 µs duration) into µA? I'm using the following kind of stimulator. Any help would be greatly appreciated!
Thijs point is a good one! You can calculate the current intensity (based on the V and R) at the output of your unit but you will not be able to find out precisely the current intensity applied to the tissue due to mentioned variables.Following
- Andrey D Ivanov added an answer:3What is the best practice to perform daily maintenance of vibratome?
I am an electrophysiologist, and I prepare rat brain slices (not fixed, but directly from live animal) as a daily routine. I know that the vibratome is supposed to vibrate only in the horizontal direction, not in the vertical direction. However, every two weeks or so, the vibration in vertical direction gets really bad, and we have to recalibrate the vibratome. I have asked several labs who use vibratome in my department, and they seem to follow quite different practices to maintain and clean the vibratome.
In our lab, after cutting the slices, we unscrew the blade holder off the vibratome head stage and clean it separately, while keeping the ice tray (our post-doc believes that taking off the ice tray would make the vertical vibration worse). Another lab I asked does the opposite, where they take off the ice tray and keep the blade holder on the head stage. Yet another lab takes off neither, and they check the vibration on a daily basis.
I am trying to get collective wisdom from ResearchGate. How do you clean and maintain the vibratome? How often do you perform vibra-check?
I regularly use a Leica vibratome for acute rat brain slices preparation.
I also prefer regular razor blades, but I use one blade per day for 2 or 3 sectionings.
Of course, vibro-check is done after every blade replacement.
I do not normally remove the head stage for cleaning, but I always check all the screws before cutting.
I clean the head stage and the blade with bi-distilled sterile water and immediately gently dry it with a paper towel.Following
- Shakil Patel added an answer:9Can someone help as I am trying to fit the activation phase of my whole cell currents using function used in the attached paper but I am struggling?
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
Hello Guys. Thanks for your suggestions. The reason I wanted to apply the same fitting equation to Cahalan et al. was because the currents I see are the same currents as the currents he found. By applying the same function to my currents it just adds another paramater to compare. When I applied the equations I was just fitting the activation phase, I may just use the single exponential function after all.
Thanks for all your help guys.Following
- Jill Miotke added an answer:8Any advice dealing with extracellular matrix/perineuronal net in acute slice patching?
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.
First, I wanted to let you know how much I have enjoyed reading your posts. You have a great humorous writing style that captures the frustration we all have at the bench. Now I will always view micropipette placement as jousting.
I am not an electrophysiologist (I just do enough to attempt in vivo electroporation and helped map visual responses with fEPSPs), but there have been several in my mentor's lab that have tried whole cell patching on the retinorecipient neurons at the bottom of the SFGS in the goldfish tectum. They are pretty small (~ 10 um), so they finally went for a perforated patch setup that used gramacidine in the pipette (evidently nystatin didn't work well). So I was following this thread to see what others had found that might work in our lab.
My question is for you and Elise. Since mammalian perineuronal net is usually rich in CSPG, has anyone tried chondroitinase ABC to clear out the ECM? I didn't have time to do an exhaustive Pub Med search, but no one seems to have mentioned using it. Unfortunately, CSPG is involved in modulating neuroplasticity, so I am not sure if it would be a good idea. But then, all the other enzymes have effects, too. Just curious.
Jonny, I am glad something finally worked!
- Joseph Valentino Raimondo added an answer:23Can anyone provide advice on purchasing a vibratome for patch-clamp electrophysiology?
I'm looking for advice on purchasing a vibratome for patch-clamp electrophysiology. We'd be making cortical and hippocampal brain slices. Currently we use a Vibratome Series 1000 which works remarkably well for its age, but it could give up the ghost at any second.
I realise that z-axis deflection is important and that the Leica VT1200S is the top of the range instrument. That said it seems rather expensive and we won't be doing fancy dendritic recordings etc...
Does anyone know what happened to The Vibratome Company?: is the Vibratome 3000 Plus still made?
Has anyone tried the pelco easiSlicer: http://www.tedpella.com/easislicer.htm?
Any thoughts or recommendations would be highly appreciated.
Hi Kim, we still haven't bought the slicer yet so I'd be interested in how you find the EasiSlicer, and how much z-axis movement you observe. I'm still leaning toward the Compresstome but not sure...Following
- Arjun A Bhaskaran added an answer:4Which is the best way to avoid an inflammation?
Hi, I am doing Patch clamp, in vivo. Many times I am encountering an inflammation at the site of patch after a day or two!! Can anyone help me with a good solution to avoid inflammation? As I am going to try a virus, I need to keep the animal for atleast six days!
Thanks in advance
Thank you very much for the kind reply. I am doing my experiment in layer 2/3 neocortex. After the experiment I used to cover it with silicon. Still you can see an entire damage from Superficial cranitomy till layer 4-5. I felt it is because I use many pipettes, and then I reduced it into even single pipette, and again there was inflammation!Following
- Joseph Valentino Raimondo added an answer:7Which USB camera for IR-DIC?
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!
These are awesome for the price - I think the code is open too but not sure how much work it would take to get it working for your application.
- Celestino Sardu added an answer:48Young Investigators Research in Cardiology, Electrophysiolohy - Are you interested in collaboration?
I was just wondering if young investigators from Europe and the US would be interested in collaborating on multicenter clinical trials in the field of cardiology and electrophysiology. This may help those at the beginning of their career establishing them as potent partners in clinical cardiology research.
Looking forward to hearing your thoughts.
ok guys, it may be an important idea, but now it is the moment to start research together. Let me know please when we could start, I really appreciate this. regards,
- Sayantani Sikder added an answer:12What kind of anode to use for electroplating of silver electrodes?I was coating silver electrodes with bleach for patch clamp recordings. I was told that electroplating gives better and long lasting coating. I found protocols for making the electrode positive in a concentrated KCl bath, but I couldn't find any information about what the anode should be made of. Classic copper cable doesn't seems to be right.
We are using a fresh silver wire for electroplating. I couldn't find any information about the duration of the reverse polarity as well as the reason behind reversing the polarity. Moreover if we are using a silver wire for anode , will reversing the polarity not destroy the already coated wire?Following
- Manuel Castellano-Muñoz added an answer:5Has anyone measured the response of calcium indicator dyes to single neuronal spikes?
I've been scouring PubMed for a figure showing spiking in an electrophysiological trace recorded simultaneously with optical imaging of a calcium indicator dye (specifically fura and fluo-4) to no avail. The closest I've seen is a response to a single electrical field stimulus in Akerboom's GCaMP5 paper. I've also seen simultaneous electrophysiology and calcium dyes in non-spiking cells. Do we know what the single-spike response of these dyes looks like?
It is not exactly what you are looking for, but maybe be interesting for you:
Two-photon excitation of potentiometric probes enables optical recording of action potentials from mammalian nerve terminals in situ. Fisher JA, Barchi JR, Welle CG, Kim GH, Kosterin P, Obaid AL, Yodh AG, Contreras D, Salzberg BM. J Neurophysiol. 2008 Mar;99(3):1545-53.Following
- Bernd Fritzsch added an answer:33Dextran, Alexa Fluor retrograde labelling timecourseI have used dextran Alexa 488 3,000 MW from Life Technologies/Molecular Probes for tracing nerve fibres innervating the paws of mice and found that 3 days post-injection into the foot pad is usually long enough to identify labelled cells in dorsal root ganglion neurone cultures.
Does anyone have any advice on how stable such a labelling process would be over time? For example, if I inject into the foot pad, and performed a dorsal root ganglion neurone culture 1 month later, could I expect to see approximately the same number of labelled cells as 3 days post-injection?
I am not tied to using dextran Alexa 488, just something that has no substantial effect on cell viability and allows for easy identification of cells in culture.
when I worked out with Molecular Probes the production of 3k dextran amines (Fritzsch, 1993) we had already established that dextran amines are excellently suited for regeneration experiments. Here is a early reference that gives you on overview of different techniques using fluorophores. (Fritzsch, Sonntag, J Neurosci Method 39:9-17).Following
- Anping Chai added an answer:10How can we analyze the weighted decay time constant of NMDA receptor EPSCs?
Hi, I am struggling to analyze the decay time constant for NMDA receptor function using Clampfit. The way I want to analyze is the method used in the paper, "Developmental profile of the changing properties of NMDA receptors at cerebellar mossy fiber-granule cell synapses", J Neurosci, 2000 by Cathala L, Misra C, Cull-Candy S.
Thye averaged the traces and then normalised the traces so NMDAR EPSC recordings done in different conditions can be compared. However, I am stuck where they use double exponential fit with fast and slow component of amplitudes and so on. I have no idea how this is done (or even able to be done) in clampfit.
I know clampfit has its own category of decay time and slope (under selected % of current given like between 90% to 10%) under analyze > statistics botton but I just want to follow the method of the paper that I mentioned and really struggling.
Please anyon who has done it or knows how to do it, enlighten me and your help would be much appreciated!
Thank you William. I can get what you get by using "Chebyshev" rather than "Levenberg". Still do not know why.
- Wei Y added an answer:7What‘s the reason for this strange phenomenon?
I’m recording fEPSP in hippocampal schaffer—CA1 pathway. The potential was normal at the begining shown as picture 1. After a few minutes recording, the potential turn into a strange one as picture 2 shows. Has anyone encountered such a phenomenon? what has happened to the poor slice?
Sorry for late reply! I have been engaged in something else. With your helps, I‘ve solved the problem. Thank you all for your answers.Following
- Binu Ramachandran added an answer:8How do I keep brain slices alive and get stable recordings and long term exponentiation in the hippocampus (CA1) of mice using the med64 system?
I am a PhD student and i have been performing long term potentiation assays for the better part of 3 years now and i feel there may be something I'm missing as i get recordings (not necessarily good ones) from about 2 out of 8 slices. some of the main issues i face on a daily basis are;
after placing the slice in the recording chamber, i have issues getting a stable baseline. most times, the resulting signal strength starts and keeps increasing and then eventually dies off along with the rest of the slice. sometimes this happens after i do an input/output trace and other times, it happens while im still looking for good networks in the slice.
much more recently, even when i get a stable baseline, i have had problems inducing long term potentiation in slices from even 6 month old animals. i use a 4x 100hz stimulation with 30 secs interval which i believe should be very strong
im at the end of my wits here and was hoping someone could tell me what im doing wrong and is there a way to improve my method?
the mice i work with vary in age from 3 months to 2 years depending on the project im working on and my ACSF recipe is the standard recipe. I use sucrose-ACSF as the media in which i slice the hippocampus. my cuts are horizontal (not coronal). i put the slices in regular ACSF (oxygenated) to recover after cutting at abput 32-34 degrees for half an hour and then at room temp for at least 30 more minutes. i am using the p5002A probe which hasnt given me too much trouble apart from electrodes getting occasionally scratched. i use oxygenated ACSF (bubbled for at least 2 hours before being used for the experiment). the solution is heated keeping the slice in the probe at around 32 degrees throughout the recording. i use a gravity system to perfuse the slice with a rate of about 4-5ml/min i have tried a slower flow rate but that hasn't shown much difference. after performing a noise test, i proceed to finding a good forward network from the CA3 to CA1 regions or just in the CA1 region as long as its the right direction and not too close. i use a stimulation strength of -50 micro amps when looking for a good connection. then I perform an I/O recording choosing a stim strength of about half the maximal and then begin my recording for a stable baseline fr at least 10 minutes before inducing LTP as stated above. i usually get an immediate increase in response but it decays back to baseline in most cases. in cases where i have induced LTP, it is hardly ever stable and oscillates back and forth. as stated above, i am using the med64 system with 64 electrodes, and i can provide any more information you think is important. please help, im trying to figure out if there is something im doing wrong or if i just suck at this experiment.
thanks for any tips or advice you could give me.
the methodology for slice recording is very important. I am still getting very stable recordings from mouse and we keep the slice at 32 degree just after cutting (we use the same buffer for cutting and recordings) and keep it bit long-term like 3 hr this is something special because we record long term LTP for 6 hr even up to 10 h and use 400 micron thickness slice. I think better to go through the articles from Frey´s Lab.
Julietta Uta Frey, Magdeburg, Germany.
All the bestFollowing
- Mina Afhami added an answer:15Can you recommend a good protocol for an in vivo mouse preparation for 2-photon imaging and targeted whole-cell recording?
I am looking for any protocol related to in vivo preparations (head-restraint, anesthetized) for 2-photon imaging + electrophysiological recording (extracellular, patch-clamp).
Thank you for your advice
I have same question too. Does anybody have experience in using isoflurane in SJL mice?Following
- Erica Jung added an answer:6Why do I get a V reads -5mV in I=0 mode (zero current mode) while the pipette tip is submerged in tyrode, not patching a cell?
I am trying to patch HEK cells and often times cells don't have right resting potentials (-10~-5mV). I also found that when I=0 mode (zero current mode) while a pipette tip is submerged in tyrode solution, V reads -5mV. Is that just because the potential difference in tyrode solution and pipette solution? Or is my rig setting wrong?
Thank you so much for your reply. References are really helpful. I will look into it and find a way to improve my patching skill. Seems the offset problem is solved but still having the low membrane resistance. As you explained, that might be because of improper patching. Let me read those references and horn my patching skill.
Thank you so much!Following
- Mason Best added an answer:22Why is a change of the pipette resistance normal , when I give a positive pressure?
Hi, I used to work with Cs-based internal solution(Cs-IS). Now, I change to K-Glu based internal solution (K-IS) to record action potential. When I use the K-IS, I found the pipette resistance can change from 3.5M to 4.3M after a positive pressure, but not Cs-IS. When release the positive pressure, the pipette resistance change back to 3.5. I checked the pipette there is no air bubbles or dirty things clog the pipette.
I also checked the osmolarity of K-IS, it is about 285. It should be OK.
I am not familiar with K-IS. Is this normal?
Thank you for your reply. I patch pyramidal neurons. I usually like to use a big tip of pipptit to get a low Ra and good holding.Following
- Paul Farrow added an answer:49Recording resting membrane potentialI am trying to record resting membrane potential from hippocampal pyramidal neurons in whole-cell current-clamp mode. The problem I've been having is that the RMP observed is much less negative than it should be. I'm expecting around -70 mV but keep getting around -50 mV. I've been using internal solution described in publications which recorded RMP from hippocampal neurons and a standard ACSF.
The cells in the organotypic preparation appear healthy in all other regards and the seals and break-ins are fine. I am not new to patch-clamping but I am inexperienced with current-clamp recordings. Any suggestions?
You may not be able to measure RMP, but you should be able to get close to your theoretical MP based on the ionic compositions of your solutions and the resulting LJP. In my experience, a depolarised membrane potential is indicative of an unhealthy cell selection, given that all other methods mentioned above have been rigorously adhered to. Try switching up the cells you are going for and go as deep as you possibly can.Following
- Sutarmo Vincentius Setiadji added an answer:14How do I get whole cell patch clamp in cells deep within the slice?
A small, scattered sub-population of cells in the slice are labeled, and I'm trying to record from them using WC patch clamp. The problem is most of them are deep within the slice, and while I can identify them by DIC, I can hardly see their exact outline or the pipette tip (when going this deep). I think that's why so far I didn't manage to get any stable WC configuration from these cells. I have no problem with patching random superficial cells. Does anybody know any tricks for recording from visually identified cells deep within a slice?
Usuakky I used a blind patch clamp techniques
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.