Hi!

How about to try some HEPES-based saline solution with adjusted pH instead of using aCSF or neurobasal media which ususlly require oxigenation.

I use HBSS if I need to patch neuronal cell culture or heterologous cell culture and, of course, aCSF in case of patch clamp experiments in slices.

Best,

Miroslav

Hi!

How about to try some HEPES-based saline solution with adjusted pH instead of using aCSF or neurobasal media which ususlly require oxigenation.

I use HBSS if I need to patch neuronal cell culture or heterologous cell culture and, of course, aCSF in case of patch clamp experiments in slices.

Best,

Miroslav

I second Miroslav's answer of using HEPES buffered saline. I might also add, that a lot of cultures that people who never do whole-cell produce are not as robust as a slice physiologist would expect. Part of it is like working in a slice, and learning to pick the right cell (with phase contrast optics, this is a bit easier, as you look for the 'bright' (i.e. Tall) cells). I also suspect playing around with osmolarity might be a good idea. Try dropping it by 20 mOsm with some water, and see how you get on. And of course, try other ways of breaking in. A zap. Gentle suction + waiting. Different cells like different approaches. It also depends on what your definition of "depolarized" is. If they're resting at -50 mV, I wouldn't be terribly shocked.

I have always patched cultured cells in HEPES as Miroslav suggests.

Also, not sure if you use TEA in you solution but you could back from that perhaps, I recall this was one of the things I played with that helped with cell health and longevity once patched.

Perhaps post you internal and external solutions.

Are you recording calcium currents? Are you in Barium?

Are you using dissociated cultures or long-term embryonic cultures? What type of neurons are being used. If you are using acute dissociated cultures, the duration and digestion with the enzymes are critical. The cell should look healthy and hold their morphology (well-rounded) for at least 5 hrs. Use Kgluconate based intracellular solution.

LP

We normally use HEPES buffered extracellular solutions as well. ACSF required consistent bubling with O2/CO2 to keep the pH proper.

Well, I work mainly with epithelial cells, but recently have made a short project with cultured neurons (LUHMES cells). Indeed, it worked nice with HEPES-based solution, pH 7.4 and bath solution slightly hyperosmolar compared to pipette. As William noticed, their rest potential was near -50 mV; to elicit action potential I first hyperpolarised them injecting outward current of 5 pAmp, which lead to Vm=-90 mV.

I've done recordings from isolated retinal neurons and neurons in intact or sliced retina. My experience is that they tend to be especially fragile once isolated - while I could pretty aggressively dimple onto a neuron in a retinal slice, I need to approach much more gingerly with isolated cells.

You might also want to try using slightly higher resistance pipettes than you do in slices, as I think this can make things easier on isolated cells - either slowing dialysis or so that less surface area is being tugged into the pipette and thereby lessening some of the strain on the membrane.

Finally, what is the surface you're culturing them on? I think poly-lysine can be a bit rough on some cells, making them a less healthy and even more fragile. Something like Cell-Tak may be a bit gentler.

Assuming your osmolarities are accurate, I think it is likely to be the cells! Hippocampal or cortical cultures are difficult to do well. Do you seed them onto a glial cell layer? How old are the animals and how old are the cultures? Cells from older than embryonic animals are increasingly hard to culture, and even cells from prenates need a couple of weeks in culture before they are healthy enough to tolerate whole-cell dialysis. Recording with K-based internal solutions is the hardest thing to do with cultured neurons, it being far easier to use a Cs-based internal. Cells have to be really healthy to cope.

Hi Gyorgy,

I have been culturing neurons and recording (whole-cell) from them successfully using ACSF-based solution. Generally, I prefer using ACSF (although it is more of a pain to work with) because it reflects better the physiological environment. Assuming that you verified osomolarity and pH, it is possible that your neurons, while looking ok on the microscope, are not in optimal shape for recording.

Things you may want to consider:

1. As mentioned by others, try to find the best-looking neurons to record from. I usually go for middle-sized neurons (the very large ones do not patch well), with smooth-looking surface and ‘healthy’ morphology (pyramidal, no blebbing etc). There is some learning curve involved in finding the ‘perfect’ cell…

2. What age of neurons (DIV) are you using? Older neurons (>21 DIV) are usually more difficult to patch from compared to younger ones (10-17 DIV), probably due to reduced culture quality/ health with age.

3. What values of resting membrane potential do you get? Primary neurons have general more depolarized RMP than what you would expect to see in a slice. This is also age-dependent. At early developmental stages, I wouldn’t be surprised to get values as low as -30 to -40 mV; in a ‘mature’ neuron in a dish, values between -50 and -60 are not unusual.

4. Blebbing may indicate calcium-mediated toxicity. Did you make sure to buffer your intracellular solution? Perhaps if you provide the composition of your internal solution and ACSF it can give a clue as for what went wrong.

As mentioned by others, make sure to bubble your ACSF continuously, and use a K-gluconate based internal solution. Hope it works for you!

As far as I have experienced, cultured cells are more sensitive to some mechanical stress from patch pipette than cells in slices, because surrounding tissue serves as a shock absorber for the cell. As you know, mechanical stress causes depolarization.

Thanks for all the comments!

The cells were isolated at p0, transfected at p14, recorded at p16-18. I did not produce the cells, got them from a collegue. They grow on poly-d-lysin. This run was from hippocampus but I`ll also try cortex. I tried Cs-gluconate and CsMeSO4 internals with 100uM EGTA in the pipette. By "depolarised" I meant really dead, needing to inject over 500pA to hold -60mV and the overall recording being unacceptably noisy.

The idea would be recording GABA currents.

Bubbling to maintain pH is not a problem, my slice rig is set up for this kind of thing but it is possible that I could get more stable pH with HEPES, it`s worth a try.

Interestingly, it seems that the Neurobasal Media has about half the Na concentration (see Invitrogen website) as the ACSF I use for slices (that`s the generic stuff with 140 NaCl, 26 bicarbonate, 10 glucose, 2 Ca, 1Mg, etc), which can lead to obvious problems. Also, the media is not changed on the cultures so the osmolarity and salt content changes day by day as it evaporates (i have no idea if this is standard procedure for neuron cultures or not) which makes matching the solution tricky. Nevertheless, Na and osmolarity matched solutions were not that much help either. Part of it could be which cell to pick, they all look bad compared to slice and they rarely have clean surfaces, seem to be covered by fibres from other cells.

Either way, thanks for all the suggestions, I`ll post how it worked out when I get my hands on some more cells.

Hi Gyorgy,

I am more of an acute slice guy however I have used acutely dissociated and cultured hippocampal neurons. One thing I know for sure is they are more delicate than neurons in slices. Anyhow when I used K-gluconate internal solution with ACSF of the following composition(in mM): NaCl 150, KCl 3 and HEPES 10 I hade no issues. This solution also does not need bubbling and neurons stay pretty healthy for atleast 45 min or so. It might help!

Best,

Bopanna

Hi Gyorgi,

Not sure how you transfected your cells, but assuming you are using a lipocationic method (such as lipofectamine2000), this can be a real issue in trying to patch (even in non-transfected cells in the same dish). Perhaps you can try first with non-transfected cells and then move on to transfected ones. It may take much more patience (and luck...) to get good a recording from a transfected cell.

Good luck!

hi,

it might make a difference if you use antibiotics on the cultures or not. also, it's possible to adjust for evaporation by adding ddH2O every few days to keep osmolarity as constant as possible.

all the best!

I do not see any problems from your setting. Usually it is quite straight forward and easier than slices recording for me. what date after plating when you did recordings?

Dear Gyorgy,

we are recording hippocampal cultured neurons like you do starting about day 7 in vitro until about day 18 in vitro. At the beginning they have resting membrane potential around -40 mV, later about -50 mV or less, as was already mentioned. We do not use CsMeSO4, cells do not take very well these ions, we are using CsCl, but we are recording Ca currents. Did you chceck osmolarity of your pipette solution? It is important that it does not mdiffer from you bath solution by more than 1-2 mOsm, cell are very sensitive to this.

I would suggest trying Hibernate-A medium (from BrainBits, LLC) for your recordings. This medium is formulated to be nearly identical to Neurobasal but with MOPS buffer instead of NaHCO3 to maintain a stable pH in ambient air. I have had good luck with that in recent months. As the cultures mature the osmolarity of the medium will increase (unless you are using a frequent feeding schedule), so you may want to match the osmolarity of the recording solution by supplementation of additional sucrose or NaCl. Since Hibernate-A has a low NaCl level (~260 mOsm initially) it would be preferable to add NaCl to increase osmolarity as needed. My ideal osmolarity for cultured neuron recordings is 290-300 mOsm and my cultures usually reach this value around 11-14 days and it is good to check this before the recording experiment. Note: If you order a calcium free version of Hibernate-A be sure to add calcium for your recordings.

In addition I agree that neurons grown on astrocyte feeder layers or grown with astrocyte-conditioned media always are the best for electrophysiological recordings and will give more "normal" properties such as resting potential. In addition, if you have a choice between mouse or rat hippocampal/cortical neuron cultures, in general rat neurons are more robust. If you must use mouse neurons (e.g. transgenic animals) you may consider a special substrate to culture on such as Matrigel which contains several growth factors that can help keep the neurons healthy.

Max be you find some information in some publications by Karl Lynch III, University of Virginia. He did some work on anesthetic effects of anesthetics in patch clamp. Also Peter Weber, University of Bochum did some patch clamp work in neurons.

Gyorgy,

Are you overlooking something about the neuronal culture? I add fresh media on the 3rd day of culture without removing any and do medium changes (replace half a volume) every 5th day. I have been using poly-l-lysine for coating the plates and I have found that for patching purposes plastic bottom plates are better substratum for neurons in culture than glass bottom dishes unless you want it for imaging. I have also been using HEPES based solutions and had worked fine. I wash the cells gently in the extracellular solution before incubating in it for a few minutes before recording.

When I recorded from cultured hippocampal neurons we also used to change the media, as John already mentioned - our changes were twice a week. Cultured neurons are definitely a bit sensitive and I had occasions when they were really hard to get good recordings from, some virally-infected neurons in particular would be really hard to patch. I think the healthier the neurons are before you transfect/infect them then the better they survive. I also used HEPES buffered sline and that worked fine and could get good recordings for over an hour.

I have been patching hippocampal neurons in culture with no problem, in Ringer's solution (mM): NaCl 145, KCl 3, CaCl2 1.5, MgCl2 1 Hepes 10, Glucose10, ph7.4

Marker Gene Technologies this year launched a media that builds on the pioneering work of Chuck Stevens for neural recording, in which an HEPES solution is created with the preferred osmolarity between 280 and 320. It has additional components that protect from light induced toxicity and will show excellent reduction in background fluorescence if that is an eventually application. Search for Opti-Klear Live Cell Imaging Solution in 1x and 5x solutions for more information. I will claim commercial bias, but stand behind the product as we have done extensive in house and collaborative work with it.

i have also a similar background and went through similar problems. Yeah, you need to use hepes and u need to wait some time from the moment u get your cultures from that unknown pink stuff. after that is peanuts.

ah, and mixing solutions is as complicated as doing nescafe with sugar and whitening so, unless you have a lot of money to waste I wouldn't bother with premade commercial buffers.

Well, not quite Nescafe and sugar - unless you have an osmometer and several key ingredients not listed in on-line protocols. Yes commercial bias - but experience too 35 years and still counting). At 25 cents an experiment as cheap as Nescafe though.

Your request was for tips. I will start by giving you two: 1. Do Not use an enzyme to disassociate the cells from the cellular matrix. I've done research that showed second-messenger involvement with this protocol and presented data showing several different enzymes being used to check this at SFN meetings. The second-messenger involvement gave False data for anyone using the protocol. I used a different protocol (developed by me and a colleague at NIH) and found major differences in cell response for all Patch Clamp procedures (including whole-cell and single ion channels). That was 24 years ago and people still use an enzyme. Problem is, the protocol was developed for Cardiac cells and then just moved over to neurons. Big Mistake, and ego's seem to have kept it going; 2. Be very careful with the Carbogen in your incubator. I ran into a problem where the Co2 in it had been contaminated with Cyanide (it was found in All Co2 for the Eastern USA. Soda and otherwise). The company felt that it was harmless for people (ppb), but I had to tell them that it was killing my cells. They didn't seem to care but did slowly removed it. Good Luck! btw, Slice work is fun, I did 5HT effects on live Human Cortical Neurons with Slice, but only did one paper on it.

Hello,

I agree Miroslav. I also carried out patch clamp in both neurons of slice and cultured neuronal cells (both in primary neurons and neuronal cell lines). I never used O2/CO2 bubbled aCSF for the cultured neurons, rather HEPES-based extracellular solution. Try it! In addition, washing the culturing medium out off the cells might also be a critical point. Good luck!

Imre

Hi!!

From my experience I culture and let my neurons grow in NB media. I changed media every 3rd alternate day but do not aspirate whole media. Aspirate half of them and add fresh new half. Beside this you must check osmolarity because it can cause a difference. It also depend on which day you are patching post culture. May be this can help you. good luck !

This is a very important point!

Hi!

I work with neurons from enteric nervous system (patch clamp at whole cell configuration and voltage clamp), the electrophysiology solutions are HEPES based, as some people suggest, we use for culture and electrophysiology:

To culture neurons, you can use minimum essential medium 97.5%, containing 2.5% fetal bovine serum, L-glutamine 2 mM, penicillin 10 U/ml, streptomycin 10 µg/ml and glucose 15 mM, and incubated in O2/CO2 atmosphere at 37°C.

For electrophysiological recordings, we use external solution as follows (in mM): CaCl2•2H2O 2, CsCl 3, glucose 11, HEPES 4.8, NaCl 160 and pH was adjusted to 7.3-7.4 with NaOH. The internal solution is composed by (in mM): CsCl 150, EGTA 10, HEPES 5, NaCl 10, ATPMg 4.5, GTP 0.1 and pH was adjusted to 7.3-7.4 with KOH. The holding potential is -60 mV in enteric neurons.

NOTE: we use CsCl instead KCl to avoid potassium currents.

References:

Barajas-Lopez, C., Espinosa-Luna, R., Christofi, F.L. 2000. Changes in intracellular Ca2+ by activation of P2 receptors in submucosal neurons in short-term cultures. Eur J Pharmacol 409, 243-257.

Barajas-Lopez, C., Peres, A.L., Espinosa-Luna, R. 1996a. Cellular mechanisms underlying adenosine actions on cholinergic transmission in enteric neurons. Am J Physiol 271, C264-275.

Zhou, X., Ren, J., Brown, E., Schneider, D., Caraballo-Lopez, Y., Galligan, J.J. 2002. Pharmacological properties of nicotinic acetylcholine receptors expressed by guinea pig small intestinal myenteric neurons. J Pharmacol Exp Ther 302, 889-897.

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.

Hi! I have been patching iPSC-derived neural cells, and desipte a good success with mouse/rat slices, I find quite difficult to perform nice recordings in these cultured cells. They are more fragile and often don't survive after going to whole cell. Moreover, often I get very high membrane resistance (around 1 - 1,5 GigaOhm), and low probability to see EPSC or IPSC, despite normal firing. I use K-Gluconate based solution for EPSCs and KCL based solution for IPSCc (recording both at -80 mV).

Does someone have any suggestion?

Thanks!!!!!!

I feel the same. The maturity of the neuronal cells could be the key issue. I use aCSF. I do not think that solution is a problem.

Hello György,

I agree with the previous comments that HEPES-based extracellular solution (without bubbling with O2/CO2) should be used instead of aCSF. I worked with cultured neuronal cell lines and primary neuron cultures as well, and now I record in brain slices, but I never used aCSF in the cultures. There are many HEPES-based recipes in the literature, I do not want to cite them here.

Another aspect that might be important in the culture is the culturing medium itself. Cultured neurons are very vulnerable regarding detergents (soap, dish-washing liquids, etc.). Couple of years ago I lost half a year because the assistants washed the glass bottle in which they prepared the medium with detergent. The glass wall adsorbed this detergent which was slowly dissolved into the medium. This tiny amount of detergent made the membrane of the cells a bit fragile. I could achieve the gigaseal well, but when I broke the membrane to achieve whole-cell configuration, I lost the cell. Later I started preparing the culturing medium myself and I used single-use culturing flasks for the preparation. I never used glass bottle any longer and all of my problems disappeared. So, good luck!

Imre