- Wei Y added an answer:What‘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
- P. A. Ferchmin added an answer:How 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.
We prepare slices from both hippocampi of 2 rats (~150-200 g) and place them in the chamber within minutes of killing the poor rats. They are kept at the interface and are stable. You can have 20-30 slices to choose. We use them all. Check our methods hope it will inspire you. Remember practice makes perfect.Following
- Mina Afhami added an answer:Can 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
- Mahyar Janahmadi added an answer:Can 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
I am agree with Maximiliano. It would be easy to divide the whole current into two phases: one is the activation phase which starts from the beginning of the current initiation (after the capacitive current) to the peak and the second phase is the decay phase of the current which starts from the peak to the end (before the capacitive) current. Both can be fitted with a standard exponential function which gives you the time constant of activation or the decay, but you need to try different number of term (n) in order to get the best fitFollowing
- Erica Jung added an answer:Why 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:Why 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
- Neil V Marrion added an answer:Recording 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?
Some interesting answers, but the bottom line is you can't ever measure rmp properly, and certainly not with whole-cell. As soon as you change the ionic composition of the cytosol, the rmp will change. The only way is to do cell-attached patch and use the reversal of a potassium channel current to estimate rmp.Following
- Sutarmo Vincentius Setiadji added an answer:How 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
- Deepak Subramanian added an answer:Fixing noise issues from temperature controller during electrophysiological recordings?
My rig has recently started picking up some [really frustrating] noise, which I've narrowed down to the temperature-controller (Luigs & Neumann, TCIII). The machine seems to put out a pulse at ~2.5 Hz, which causes a 1-3 mV blip in the recording (picture attached). Has anyone had/fixed this problem in the past?
Have you grounded your perfusion tubes? We usually clip the perfusion tube as it enters the rig and insert a metal sleeve (a needle would do) between the cut ends of the tubing(see fig below) . Grounding this metal sleeve to the rigs main ground (the one point where all your grounds are connected to inside the rig, "a star configuration") would eliminate noise created by the temperature controller or perfusion pump. We do this for both the inlet and outlet. Also, grounding your control unit ,as mentioned by Friedrich, is very important.
======-----------------======= (ground the needle)
tube needle tube
Hope this helps!.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!
Yes, in both cases is Cre-recombinase.Following
- Jitendra Singh added an answer:Why are there current fluctuations in hERG patch clamp assay using port a patch (Nanion)?
I am doing hERG voltage clamp protocol using Port a patch but I am unable to get a steady current. The current fluctuations are so high that I can't even test a single conc. A brief about Port a Patch- its a semi automated patch clamp instrument where solutions, test compounds/ vehicle/ positive control all are added manually. This patch clamp has been recently installed in our lab and has passed the initial validation tests.
I have checked the pulse protocol and other settings but somehow I am not able to rectify this problem. Also I am using inducible CHO-TREx hERG expressing cell line. Kindly suggest..
who is the supplier of CHO-TREx hERG expressing cell line. make sure that it is expressing the herg channel by some florescence methods.
Proto col suggested by the Marry is ok u can try making following changes to your protocol
Protocol Voltage (mili-volts) Duration (mili-seconds)
Holding potential -80 50
depolarization +40 2000
depolarization +40 2000
Holding potential -80 200
Measure the current in 4th segment (+40 mV to -40 mV).
Also write in details how you are culturing the cell specifically media and antibiotic anti mycotic.Following
- Birger Brodin added an answer:Any advice on TER measurement with WPI EVOM2?
The resistance reading from WPI EVOM2 is ohm x cm2 or just ohm? I read the manual very carefully and thought it's ohm x cm2 and we don't need to convert it. But the TER value I got for my RPE monolayer on a 12-well transwell is much lower than expected, which makes me feel confused. Anyone has an idea?
the output is in ohm..so your values will depend on the size of the inserts, and the actual electrodes. so if youre using a cup electrode and a t12 insert with 1,12 cm2 support, you should multiply your values with 1,12 cm2Following
- Eamonn Fahy 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.
A big question for RGC patchers. Not sure if you are still struggling with this but thought I would contribute any others who are struggling. I've tried collagenase, which works like a treat to dissolve the ILM and hyaluronidase on its own and in combo with collagenase. Either way you deliver the hyaluronidase, I found it to be too harsh for the retina and compromised integrity of the flatmount. Indeed, this is the issue with collagenase and hyaluronidase - do they impact on the patching data?
Because of this, I stick to using an empty patch pipette to "rip" through the ILM. This is a technique that does require practice - from "catching a wave" where you catch the ILM with the pipette tip and move forward, to then "ripping" a hole in the ILM by lifting the pipette slightly and then moving from side to side, eventually finishing by going so far to the side one way or the other that the pipette breaks its contact from the ILM (can also move back and to the side to achieve this). Then you hope that the ILM doesn't float back over to obscure the hole you've just created.
Having said all this, actually being able to do this "ripping" is very much reliant on the quality of your retinal dissection. Leaving any vitreous behind will result in a pipette that glides along the surface of the retina without catching any ILM. The presence of vitreous seems random to me as my dissection technique has become highly consistent but with varying degrees of vitreous left depending on the retina. But all the nuances of retinal dissection is probably best left for another thread...
Hope this helps.Following
- Pedro J Romero added an answer:How do you improve seals with erythrocytes and how do you break their membrane to get the whole-cell configuration?I have started doing electrophysiology on human erythrocytes (whole-cell voltage clamp) and I have been unsuccessful with seals and breaking the membrane as well, as the cell either unseals or is sucked in the pipette even with 13 MOhms resistance of the pipette.
Glad to know things are becoming better. Trust you will succeed in your work. Many thanks for your comments of my work on voltage-dependent Ca channels. Kind regards. Pedro.Following
- Diego Fernández added an answer:How can I get stable and perdurable LTD in hippocampal slices from adult mice?
I have seen through publications that some labs have been routinaly able to induce very long-lasting in vitro LTD in young and adult animals. I would like to ask you if you can give me some details of the methods to achieve it.
I am trying to get LTD in my hippocampal slices from ~10 weeks old mice, but so far it has been almost impossible. I have been using both electrical stimulation (single 900 pulses at 1 Hz) or the application of mGluR2/3 agonist LY354740. I am stimulating the medial perforant path and recording fEPSPs in the dentate gyrus, at room temperature.
Does anybody have some advice to get stable LTD?
Thank you very much in advance for your contributions.
Thank you everybody for all your comments.
It seems clear that slice health is a critical factor for the induction of LTD, so we will focus on improving our slice preparation and also we will modify our setup to be able to work at more physiological temperatures, following your advice.
These contributions and the papers you've suggested have been very helpful.
Thank you all so much again.
- Karin R Aubrey added an answer:Rs badly increases and synaptic currents don't change during LTP induction into whole cell configuration. Bad recordings or not?
I am new into electrophysiology. I am studying LTP via a STDP protocol in whole-cell configuration, recording currents. After the LTP induction my Rs badly increases (more than 30%) but there is no change into synaptic currents. As far as i know, when Rs increases the currents should decrease following ohm law. If my currents don't decrease can be due to a potentiation into synaptic transmission?
I am using clampex 10.2 and multiclamp 700B to record from the cell, with a pipette resistance of 5 mOhm.
I hope I explained myself well,
There is a clear explanation of SR and how it effects currents written by Boris Barbour.
You can find it here.
- Einar Eftestøl 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 ?
And you are sure that this increase in mass is not due to inflammation or other type of damage resulting from cutting of the blood supply or massive bleeding from the gastroc, extensive forming of connective tissue etc? Is this larger increase in mass also reflected in an increased force production and/or muscle fiber cross-sectional area?Following
- Jorge Parodi added an answer:Why are neurons depolarized but not firing during whole cell recordings?
I am doing blind whole cell patch recording to record firing patterns in current clamp mode.
The cell fires action potentials at the beginning of recording when I give current injection (50pA to 100pA), but slowly reduces firing frequency then not fire at all after about 2 to 5 min into recording. Resting membrane potential (at 0mv holding), series resistance, input resistance and AP amplitude (if there is any) doesn't seem to change during this period.
I am using regular K-gluconate (135mM) based internal solution (295Osmol)and no blocker is used during recordings.
Since I am doing blind patch, I can not tell the health of the cell. Only thing I can rely on is the parameters that I can measure (resistances and membrane potentials).
Does anyone know what could be the reason for reducing the firing rate after break in?
Also, what parameters should I look to tell the health of the cell when I do blind patch?
One more not related questions is that..
Do I have to use bridge balance button (?) in multiclamp 700b when I am in current clamp mode? It doesn't seem to do anything whether I use it or not.
I appreciate any answers or suggestions.
finally observation the quality of your slice are important!!! mayde you are lose the circuitFollowing
- Jonathan T Ting added an answer:How can I get a stable fEPSP?
I am currently trying to get a stable I/O curve in the CA3-CA1 circuit. I am working with 400µm horizontal slices from Bl6 mice that are 4-8 weeks old. I am cutting in normal ice cold ACSF and can store my slices differently afterward. I can store them at 33°C in an interface chamber or submerged at RT (or warmer, but I havent tryed that). I also tryed NMDG aCSF for cutting and 10 min recovery with NMDG aCSF after cutting. I let the slices rest for at least 1 hour after cutting in normal ACSF bevor I start the recording.
My problem is, that if I get a fEPSP that is bigger than my FV, it drops after a few stimuli and is mostly gone after abot 5 minutes. Does anybody have an idea what I am doing wrong? How should the slices recover? What ACSF solutions are you using?
I had similar rundown problems when I first started learning how to do hippocampal field recordings. As Alexander suggested, my rundown was rapidly reversed by the A1AR antagonist DPCPX, in fact, the response was even larger than expected suggesting the presence of basal adenosine tone. As a second note, I also had run-down related to the home-made holding slice chamber because my volumes were too low and the metabolites like adenosine could rapidly build up over the experiment. This was improved by refreshing the solution in the holding slice chamber, or eliminated altogether by using a commercial slice holding chamber with large volume (I prefer the Brain Slice Keeper-4 sold by Automate Scientific).Following
- Valentina Kutyifa added an answer:Young 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.
Thank you all for your responses.
It seems that there is a need for collaboration among young EPs.Following
- Caroline Waters added an answer:Are there any data about electrophysiological features of tanycytes?
I try to patch them, but I have no idea what protocol should I use.
Correlated electrophysiology and morphology of the ependyma in rat hypothalamus.
Jarvis CR1 PMID: 3193176
Good review paper ATP-mediated glucosensing by hypothalamic tanycytes
Cameron Frayling DOI: 10.1113/jphysiol.2010.202051Following
- Mohsen Mosayebi Samani added an answer:Why do DC currents have opposite effects when interacting to cortical or peripheral neurons?
Anodal DC stimulation often leads to increased brain excitability when electrodes are positioned over the primary motor cortex. Cathodal stimulation, oppositely, leads to decreased neuronal excitability. However, when monophasic currents interact with the axon in peripheral nerves, the effects are reversed, with Cathodal stimulation leading to increased excitability. Does anyone have a reasonable explanation for this phenomenon? Are there any experimental data to support this neuronal behavior?
With respect to Prof. Hofmann. I recommend you to take a look at these freely available online references:
1. for cellular mechanism of tDCS (DC Stimulation):
Professor Bikson lecture: (Cellular Mechanism of tDCS)
2. for Cathodal/Anodal prepheral nerve stimulation:
Book: Bioelectromagnetism---chapter 21--- Functional electrical stimulation:
- Serguei N Skatchkov added an answer:Do I need to chloride a concentric electrode for electrophysiology?
I am putting together a stimulating electrode for electrophys. recording. Do I need to chloride a concentric electrode the same way I would a bipolar electrode?
if you do not cover your Ag-wire (reference electrode or in patch pipette), you will have tremendous and unpredictable shifts of electric potential which will produce potentially wrong data in electrophysiological experiments measuring DC currents/voltages. So, it is necessary to use stable electrodes. Ag-AgCl is mostly used, however there others such as (i) copper-copper sulfate electrode (Cu-CuSO4) or (ii) mercury electrode (Hg-HgCl2, called calomel electrode) that we used as well. All depends on particular design of the experiments.
Dr. Akopian suggested simple method how to cover rapidly your silver wire making Ag-AgCl elecrode. We also used successfully not electrochemical (cathode covering, using electric current) but just chemical procedures as Dr. Akopian suggested, but different methods:
1. CuSO4+NaCl solution method
2. fire melted AgCl coating
Both methods produce harder surface of coated wire and thus more mechanically stable.
Very cordially, Serguei.Following
- Olga R. Ayala added an answer:Does anybody know why the most of MEC/MFC studies tested are at applied voltage range of 0.2V to 1.1V ?
I am working on single -chamber MEC/MFC studies , from the literature review I found that all MEC/MFC have tested at applied voltage range of 0.2V<Eap<1.1V. Are there any reasons for using that range of applied voltage. How about <0.2V or >1.1V?
Well it depends of the product objective of your cells and the materials for anode and cathode used, for example to produce hydrogen in a cell of Graphite - Pt, the electrochemical reaction tells us that minimum we need 0.14 V in theory...in practice many articles shows that are higher, I am attaching one interesting article.
And for the maximum many articles used 1 or 1.1 V because is the “practical” sustainable voltage if you want to apply your research in future applications.Following
- Saikat Chakraborty added an answer:Can somebody recommend a lab specialized in acute dorsal root ganglia recording located in Europe?
I have started a project which requires electrophysiological recordings (intracellular or patch-clamp) from small neurons (<30 uM) in intact dorsal root ganglia (no culture involved, not dissociated, with suction electrode). I have so far manage to dissect correctly the DRG, prepared and verified the aCSF pH (7.3-7.4) and osmolarity (310 mOsm).
The membrane potential of the cells is between -60 and -55 mV which seems ok). So far, MO (20-100uM) or CAPS (100nM) application haven`t elicit any current or potential variation. I must be doing something wrong (or not quite right). It is hard through internet to get an adequate. Observing someone doing it would be the best.
Therefore, I want to get some training in a European lab for a week or so in order to solve this.
Does anyone have an idea?
Thanks a lot
You have any specific technical issue?Following
- Henrique P. von Gersdorff 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.
Try changing the batch of your pipette glass. The glass must be very clean to form a giga seal. Also filter your internal with a 0.2 micron filter. Use positive pressure (make sure the pressure in the holder and tubing is not leaking with a monomitor), see the dimple in the cell membrane and then let the pressure off and apply some suction and then hyper polarize the membrane patch to -80 mV... this should work.
Good luck, HenriqueFollowing
- 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
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.