In Vivo Electrophysiology - Science topic

I am afraid that is not ideal. Using a single electrode to record in vivo local field potentials (LFPs) in the hippocampus is generally not recommended because: 1. Lack of Reference Point: LFP recordings measure the summed electrical activity of populations of neurons. Without a separate reference electrode, it’s hard to distinguish between true neural activity and background noise or artifact. A single electrode does not provide a way to differentiate local changes in electrical potential from broader shifts in the electrical field. 2. Volume Conduction: The hippocampus is embedded in a complex neural network, and the electrical signals recorded by a single electrode can include not just local activity but also signals that are conducted from other brain regions. This "volume conduction" effect makes it difficult to isolate hippocampal activity with a single electrode. 3. Spatial Resolution: A single electrode does not provide information about spatial variations in the neural activity across different regions of the hippocampus. This is especially problematic because the hippocampus has distinct anatomical and functional regions (e.g., CA1, CA3, dentate gyrus) that can have different activity patterns. Multi-electrode arrays or tetrodes allow for simultaneous recordings from multiple locations, improving spatial resolution and allowing for more precise mapping of hippocampal activity. 4. Difficulty in Signal Interpretation: LFPs represent a mixture of signals from excitatory and inhibitory neurons, synaptic potentials, and even glial cell activity. With only a single electrode, it is much harder to interpret which sources are contributing to the recorded signal, making the analysis less reliable. 5. Signal-to-Noise Ratio (SNR): Single electrodes are prone to picking up noise, which can be difficult to filter out when there is no comparison or control signal from nearby regions. Using multiple electrodes allows for better noise cancellation and more accurate isolation of the signal of interest. In summary, using a single electrode for LFP recordings in the hippocampus is limited because of issues with reference point clarity, signal contamination from volume conduction, reduced spatial resolution, and difficulties in interpreting the resulting signal. Multi-electrode setups help overcome many of these issues, providing a more accurate representation of hippocampal activity.

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However, it's still technically possible if necessary for specific experimental constraints. Here is a draft protocol: 1. Preparation - Anesthetize the animal with an appropriate anesthetic (e.g., isoflurane or ketamine/xylazine) and place it in a stereotaxic frame to secure the head. Ensure that the animal is fully anesthetized by monitoring vital signs (e.g., absence of the pedal reflex). - Shave the scalp and sterilize the area with ethanol and iodine. Make a midline incision to expose the skull. - Use a stereotaxic atlas to determine the precise coordinates for the hippocampus. Mark the area on the skull. 2. Craniotomy and Electrode Placement - Carefully drill a small hole in the skull over the target location using a micro-drill. - Insert the reference electrode in a distant brain region (e.g., the cerebellum or a peripheral muscle). Fix the ground/reference electrode securely. - Lower the recording electrode: Using the stereotaxic manipulator, slowly lower the single recording electrode into the hippocampus. Advance the electrode slowly to avoid damaging the tissue. 3. Signal Amplification and Recording - Attach the recording electrode to a pre-amplifier and connect the reference electrode to the ground of the system. - Adjust filtering parameters: Set up the amplifier to filter the signals between 1 Hz and 300 Hz (the typical frequency range of LFPs). Adjust the gain of the amplifier to ensure the signal is visible but not saturated. - Monitor the signal: Observe the incoming LFP signal in real-time on the data acquisition system. Adjust the electrode depth if necessary to optimize the quality of the signal. LFPs are typically visible as slow oscillations in the frequency range of 0.5 to 100 Hz, depending on brain state. 4. Recording - Once the electrode is properly placed, begin recording LFP data. Record for a suitable time period depending on your experimental question (e.g., several minutes to hours). - Monitor physiological parameters: Ensure the animal remains properly anesthetized throughout the procedure and check for any signs of discomfort or distress. Continuously monitor vital signs. 5. Post-recording - Electrode removal: Once recording is complete, carefully remove the electrode without damaging the tissue. - Close the craniotomy: Clean the exposed area of the skull and apply dental cement to cover the craniotomy. Use sutures to close the scalp. - Postoperative care: Administer analgesics (e.g., buprenorphine) and monitor the animal until it recovers from anesthesia. Place the animal in a warm recovery area. 6. Data Analysis - Filtering: Apply additional digital filtering if necessary to isolate LFPs from noise. - Artifact removal: Identify and remove artifacts from movement, heartbeat, or respiration. - Spectral analysis: Perform frequency-domain analysis (e.g., power spectral density) to examine oscillations like theta (4-8 Hz), gamma (30-80 Hz), or sharp wave-ripples (~100-200 Hz). - Signal-to-noise analysis: Evaluate the quality of the recording and signal-to-noise ratio.

Hi Luca,

From what I understand, the problem is that when you switch between the two objectives, the field-of-view may sometimes change. However, this does not happen all the time.

First I want to rule out some simple errors that may cause this.

1) When you screw the objectives back in, did they go in straight and smoothly? Sometimes one can screw threads in a little bit at an angle.

2) When switching between the objectives, did you make sure that the slider is completely in position of the slots?

3) When switching between the objectives, could it be possible that the stage was moved accidentally?

Let me know if these already helped, or maybe there is indeed a more serious problem with the light path.

Best,

Aaron

Could you not just transfect your cells with two viruses? Also, I'm not a fan of the hM4Di, it just never seems to really do much, no matter how good the transfection (and it's possible this is caused by accumulation in the cytoplasm https://blog.addgene.org/tagging-optogenetics-and-chemogenetics-receptors-fluorescent-proteins-and-other-options).

There is an alternative opsin that can produce activation or inhibition of a neurone depending on the wavelength of light used. I've never used it, but it looks good: https://www.addgene.org/154951/

Assuming you have a thesis or research advisor, can he/she not coordinate for you to visit a lab that does rodent brain electrophysiology and for you to learn the technique there? If you don't have such an advisor, you will have to be the one contacting the labs and asking for permission to visit. Be prepared to offer to help the lab on a project while you learn the technique.

Dear , thank you for your answer.

I find the following result when I search the frequency band in Google.

" LFP frequency bands: The classification of the LFP into distinct frequency bands, known as delta (approx. 0-4 Hz), theta (4-10), alpha (8-12 Hz), beta (15-30 Hz) and gamma (30-90 Hz) has been adopted from the EEG literature. It is based on the strong correlation of each band with a distinct behavioral state. "

Is this classification also valid for rats?

Why do frequencies 8 to 10 belong to the two classes and why do frequencies from 12 Hz to 15 Hz not belong to any class? I also found different results in the articles. Anyway, what is the valid classification for frequency bands for mice?

Hi Claire,

In what aspects did your surgery technique improved?

Taping the joint by 3M 8979 duct tape（3M ID 70006709318）

I’ve done patch clamp recordings in awake mice. The craniotomy was surrounded by a stainless-steel ring cemented to the skull. By dripping some ACSF into the ring, over the craniotomy, I made a little well of saline into which I could dip a chlorided silver wire. That was my reference. And it worked nicely: stable, low noise.

To my knowledge, no one has done a systematic study of referencing methods, but there are a lot of them: press a silver-chloride pellet against the skull near the craniotomy and cover it in saline – sometimes adding a bit of agar so that the saline doesn’t dry up; make a second craniotomy away from the region of interest and insert a wire; sew the pellet into the muscles of the animal’s neck. I knew one guy who, for the sake of extracellular recording (just spikes), clipped a wire to the animal’s tail (I’m still surprised that that worked).

It seems that a lot of things will work and (my two cents) this isn’t a thing one should really worry about. In-vivo recording has plenty of other things to worry about!

As far as an agar bridge goes, I’m not clear on why that would help. When working in vitro, people use an agar bridge when they’re doing solution changes – washing in something or changing the ionic composition of the bath. The agar bridge minimizes effects associated with junction potential; it also, when the bath volume is small, keeps the reference/ground from polluting the bath. But in vivo, these considerations don’t really apply. You’re not going to change the ionic composition of CSF and the effect of the reference electrode (whatever it is) on CSF should be tiny.

So – again, my two cents – if you find a referencing format that gives you stability and low noise, you should be good to go.

The question asks about action potentials, dear Bernard.

Dear Lora if you want to be a neuroscientist, learn the jargon and think before asking a question, orthodromic and antidromic propagation are jargon words. and yes it valid for synaptic transimission also.

We use a micro drill from cell point scientific:

Andrew Chao no the project we were working on this for is currently stalled but will likely pick back up come May/June so if you figure this out it would be greatly appreciated. Likewise if we figure it out I'll send what ever we end up doing over your way.

possibly change to something from here...?

I'm afraid there are to many open questions to give a reasonable answer: Where are your electrodes seated? What kind of stimulus did you use? Are the spikes of same polarity or reverse polarity? What's the latency between the spikes? How do you control the positioning of electrodes?

Yes, a heating blanket inside the Faraday cage can produce substantial noise. Because of that, when I was doing anesthetized recordings some years ago, I used this isothermal pad (see below) instead. It worked well.

> Is there a trick to keep the series resistance low/stable for a period of at least 10-15 mins for a in vivo patch recording?

No, not really. You really want to avoid doing voltage clamp if you can avoid it because your success rate will be very low.

But some tips that might help a little bit

1) Don't advance the pipette much once you hit a cell. If you drive your pipette hard into a neuron, you almost guarantee a high series resistance recording.

2) Try to get your pipette tips as large as possible. In my experience 5-7 meg pipettes give you the best rate of getting a cell, but if you want to get low access, stick in that 3 meg range.

3) Use a HYPERtonic intracellular solution. 330mOsm will cause the cell to swell, which will, on average, give you better access for a longer time.

Both tips 1 and 2 will reduce your chance of getting a cell, but will decrease the Rs of the cells you do get.

Assume there is a headcap with dental acrylic.

Can add placement for a metal hook that can be manufactured from tinning wire or other tensile material.

Then use dental elastics to hold stage in place.

Dental elastics are cheap and will give sufficient olding power and easily manipulatple with forceps.

May need more than one anchoring position, but ground loop on Omnetics chip ideal for anchoring cabled headstage.

See powerpoint below for a quick mockup.

You can try coating your electrodes with DiI. These are fluorescent crystals that can be disolved in organic solvents (methanol and acetone). You can dip your electrodes in the solution before the experiment for 30 seconds, let evaporate the solvent and then put the electrodes in the brain. DiI is lipophilic, so it will stain the fat on the brain leaving a nice track of the electrode position that you will be able to see with naked eye (depending on the amount of DiI) while cutting the brain for histological processing, or under an epifluorescence microscope. You can see how I used it in Figure 1 of my Plos one paper 2011:

It's certainly the case that a high resistance ground will cause noisy recordings. However, I suspect that doing that measurement (ground pin to tail) would be a pretty silly way to measure your "ground resistance", as nearly all of that resistance is due to the resistance of the skin, and would be far more sensitive to "is the tail a little wet/what is the ambient humidity" than "is my ground good".

That said, if you get 1 resistance of >1 Meg, then you're ground is probably bad. That is to say, that measurement will tell you if your ground isn't working, but it wont tell you if it is average, good or great.

I'm confident there is lots of literature showing the importances of low ground resistance, not that I can find any with a trivial search (the Axon Guide talks about the importance of having a low resistance ground, but not in terms of noise).

Hi Ashley,

Another option is to just use cyanoacrylate glue only. Clean the skull, score it lightly, and then use a thin layer of glue. This is both faster and more secure.

Try Open Ephys.org for a flex drive core.

There is also a Jove video from Matt Wilson's lab that provides the code to print the 3D core of their drives. 

3D Systems, On Demand Manufacturing can print 25 3D cores for about $1000.  Make sure to use Accura60 for a polymer if you go this route. 

In both cases you still have to perform construction of microdrives of the array and make your own tetrodes out of Nickel-Chrome RO800, Redi Ohm 800 wire.

Your other option is to switch to silicon probes and use a Buzsaki style drive. Turn around on orders of silicon probes from NeuroNexus is pretty fast. The Buzsaki 32 is a nice array.

Hi, what type of analysis you are intending to do? Is that something to do with spatial behavior that you are looking for?I developing one such

You can email the first author of this study directly. We are collaborating with him, and he is very responsive and helpful. http://syntheticneurobiology.org/publications/publicationdetail/279/25

We do have this line and currently characterizing it.  This is the line from Allen brain institute.

I imagine it is pathologic as with Human

Hello Lars,

Good to know back from you. If you are sure that it's the local(?) radio station talking to you, well, the LPF I suggested was meant to filter out these RF signals and I believe, you are not interested monitoring those high frequencies in the MHz range where as your biomedical signals would be in the sub-kHz ranges (<1kHz or so, I believe!)

Therefore I suggested to put a LPF in line to cater those RADIO signals so that it does not corrupt the biosignals since the cut-off of this filter will be in the hundreds of kHz range.

Another point I suspect is the sampling speed of your data acquisition system that you  are recording the signals at the moment. If that is TOO HIGH, the ADC system itself act as a demodulator to your Radio station due to UNDERSAMPLING. So try pay attention and some trials with the sampling rate as well. I believe a sampling rate of around 1000samples per second (1ksps) to 10ksps would be just adequate to you.

Now, while reading your question again what I noticed in your last part is - "Can I still use the animal if I reference from another channel?"

Now, if your data acquisition system has isolated channels, i.e. GROUND for each channels is separate and isolated with each individual channel, then it would make a huge impact on your GROUND system in which case, yes you are likely to get such pick-up.

Hope this helps you further to reduce the noise problem.

Best wishes!

-Prasanna Waichal

I'm assuming you're referring to doing field recordings with single electrodes? If so, either can work, if impedance is equal, but the thicker ones will be more precise. A human hair is about 12-20 microns, so you probably have 25 micron for that one and possibly 125 micron+ for the other? Both work for recording brain sites like the hippocampus, To go deeper we even use a larger sharpened tungsten rod. The stability helps us achieve our target more reliably. A big concern with "old" electrodes is how good the shielding is. Old spools of wire can have cracked shielding and then your signal will be much poorer. An old case of pre-made electrodes can be fine however. (I never trust anything I didn't make myself though) If you are talking multi-unit recordings, then it's a different ball game than for induced EPSPs.

Good answer Debanjan,

We've also used short higher frequency trains (40Hz). I'll upload a recent publication on some of our work. 

Dear colleague, I think that we may use old stereotaxic atlases as inspiration. For example, the Pellegrino & Cushing 1979 atlas (for rats) uses a circa 27º downward inclination. Although I don't know any atlas that uses an inclinated reference plan for mice, you can create your own solution starting from the FSP-referenced coordinates provided by Paxinos & Franklin atlas and transpose those coordinates through axis rotation by means of analytical geometry. Let us know when you do it. I hope I helped you. Best regards.  

I have no personal experience with this chemical, but I was told that 5-fluorouracil can do wonderful things for chronic recording preps.

Spinks et al. (2003) Problem of dural scarring in recording from awake, behaving monkeys: a solution using 5-fluorouracil. J Neurophysiol 90(2) 1324-1332. DOI: 10.1152/jn.00169.2003

Hi Kristie,

these electrodes are so thin, that you need to do an electric lesion via the electrode to mark the recording site. We have used those electrodes and have been successful doing so.

Best,  Christoph 

Hi, the attached link will provide a paper published in 2008 by my group, showing that isoeugenol activates the TRPA1 ion channel expressed in human and rat sensory nerves.  This ion channel is involved in nociception or pain signalling.

Are you sure the noise is coming from the foot sensor? We have the same system and found that the heating mat was producing noise. We found that wrapping the mat in an adhesive, conductive material and then grounding it eliminated any noise. Hope this helps.

Dear Tianyang, thank you for your fast response and the additional information. You told me that the range of impedance of the tetrodes is between 100kOhm and 2MOhm. This is a relatively large range if we consider the fact that the wires have all the same diameter. I assume that the coating is stripped off the tips of some tetrode contacts when the tetrodes are introduced in the brain and this causes the impedance increase. As you know the tetrode basically is a multi-unit electrode and it should pick up the signals of several units at a time. The higher the impedance the smaller the recording sphere of a tetrode contact and the smaller also the number of units that the tetrode contact detects.    

Although many people use the twisted wire tetrodes and seem to get satisfactory results with these electrodes it is a matter of fact that the tips of these twisted wire tetrodes have sharp cutting edges that cause tissue damage when introduced in the brain. You can see a photo of a twisted wire stereo-electrode in the paper of McNaughton et al. (McNaughton B, Barnes CA, O´Keefe J. The stereotrode: a new technique for simultaneous isolation of several single units in the central nervous system from multiple unit records. J Neurosci Methods 1983; 8: 391-397.).

To see if you are able to get stable long term recordings with your flexdrive I would recommend to load your flexdrive with several single core electrodes with a conical tip shape (impedance 200-500kOhm, without plating at the tip) and try to make long term multi-unit recordings. If you will get stable multi-unit recordings over some days you can be sure that the tetrode itself causes the problem.

Basically it is difficult to hold cells over some days without readjusting the electrode position. Drift in the amplitudes and shapes of the spikes recorded at a site is often observed. This drift smears the clusters and increases the size of the clusters which can cause cluster overlap and thus makes it impossible to separate them. Details about this effect are published by Emondi et al. (see Journal of Neuroscience Methods 135 (2004) 95–105, Tracking neurons recorded from tetrodes across time, Emondi et al.). I think this effect is increased in the freely moving animal application that you have.

If the plating is removed from the tetrode tip, the impedance increases and the recording sphere of this tetrode contact decreases. Beside tissue movement this can be a reason that neurons disappear after some time of recording. I am not sure that the tissue damage caused by the tetrode would cause a gliosis around the tetrode tip after this short time. In long term recordings this is a problem, because then the electrode contact is insulated by glia cells. Twisted wire tetrodes have a slightly smaller recording radius as other tetrodes presently used in neuroscience. For example Mechler reported: “…Thomas tetrodes, the R50 recording radius was ≈80–85 μm for the smaller …tetrodes  and ≈95–100 μm for the larger tetrode. These ranges of tetrode sensitivity are generally consistent with the characteristic linear span (can be >200 μm) over which polytrodes reportedly register signals from cortical single units…but remain significantly larger than the ≈50- to 70-μm recording radii reported for twisted wire tetrodes…” (for details see: Three-dimensional localization of neurons in cortical tetrode recordings, Ferenc Mechler, Jonathan D. Victor, Ifije Ohiorhenuan, Anita M. Schmid, Qin Hu, Journal of Neurophysiology Published 1 August 2011 Vol. 106 no. 2, 828-848 DOI: 10.1152/jn.00515.2010).

I would recommend the following procedure: 1) first load the drive with low impedance not plated single electrodes (impedance between 100kOhm and 400kOhm, well suited for multi-unit recording) as mentioned before and make a long term test recording to make sure that the stability of the drive is good enough for this application. If not you should improve the stability, if yes, go to the next step 2) Consider using a different tetrode technology that causes less tissue damage and has a larger recording sphere. Speak with somebody who has made long term recordings with other tetrodes to get some user feedback (e.g. see: Saccades during visual exploration align hippocampal 3–8 Hz rhythms in human and non-human primates; Kari L. Hoffman, Michelle C. Dragan, Timothy K. Leonard, Cristiano Micheli, Rodrigo Montefusco-Siegmund and Taufik A. Valiante; Front. Syst. Neurosci., 30 August 2013 | http://dx.doi.org/10.3389/fnsys.2013.00043).

I hope this information was helpful. Let me know if you have any further question or if you need additional information. I would be pleased to help you. Best wishes, Dirk

In our experiments we used to ground under the open skin as close as possible to the animal nose

Hi Janet, the white spot usually disappears gradually during the recovery stage. I later find a complete wetting of the cornea during the whole surgery dramatically diminishes the chance of having white spots. In addition, it is good to avoid a direct surgical lighting towards the eyes.

Thanks for putting forward a possible cause by the hypotension. The cardiac output diminishes for sure when the mouse is anesthetized. 

Good luck to your research!

Jiahao

you will be welcome !

Dear Jérôme, I agree with you that if one uses constant current stimulation mode the voltage at the stimulation electrode will change due to electrode tissue impedance variation. Therefore one will not get a constant and reproductive potential at the stimulation electrode. This would only be the case if the electrode-tissue impedance always would be constant. Anyway I would recommend the following review paper for details: Tehovnik EJ. Electrical stimulation of neural tissue to evoke behavioral responses. J Neurosci Methods, 1995; 65: 1-17. Also please have a look at the paper published by Histed and others: Histed MH, Bonin V, Reid RC. Direct Activation of Sparse, Distributed Populations of Cortical Neurons by Electrical Microstimulation. Neuron, 2009; 63: 508-22. The authors reported that microstimulation sparsely activates neurons around the electrode, sometimes millimeters away, even at low currents. This very interesting paper raises several issues to consider for the design and interpretation of microstimulation studies! I hope my answer was helpful. Best wishes, Dirk

I want to solve your problem by presenting amazing article, which was very helpful for me when i had the same problem few months ago !!! in this article ,you will find out the exact way to locate electrodes .

 Hi Tianyang,

A picture says a thousand words. I didn't really get your question, but actually a really easy way to know that you haven't removed dura, is by seeing how the light shines off the tissue, if its pretty reflective, you probably haven't removed dura. In this pic I would say the dura is still on (though it is a bit out of focus).

Here is what to do, take a very fine needle (30 gauge) and slightly bend the tip of the needle so it resembles a hook of sorts. Then under the scope, poke around outside your area of interest (obviously carefully). You should be able to stick the point through the dura (and maybe a bit of cortex) and then lift the dura so it separates from the brain tissue. That is how you will know what to remove. From there you can use the ripped dura as an entry point to start cutting dura 2. Bleeding; blood vessles are every surgeons' nightmare, so you are not alone. Just try to avoid breaking it when you remove the dura, sometimes you can't avoid it. Have a syringe with saline prepared in a bucket of ice, and gently spray the area with saline until it stops bleeding (I also recommend non-toxic absorbant gelfoam). Good luck! 

The amplification you need is dependent on the ADC resolution. For example in the Mosers' lab we used 20000x amplification because the ADCs were only 8 bits. With a 16 bit ADC the resolution is 256 times better, which means amplification can be reduced accordingly. So when I used 3000x amplification in Johan F. Storm's lab the spike waveforms were actually measured an order of magnitude more precise than in the Moser's lab, although the amplification was reduced by an order of magnitude. An 8 bit ADC does not have sufficient resolution to measure spikes and LFP simultaneously. A 12 bit ADC is borderline. 16, 24 or 32 bit ADCs can easily do this. [The ADCs I used in the Mosers' lab were actually 16 bit, but the software decided to throw away the lower 8 bits, storing the signal at 8 bit per sample.]

Check the dynamic range of the ADC board (typically about 3 V) and its resolution (bits per sample) and then calculate what measurement precision you get with your level of amplification. The trick when recording spikes and LFP simultaneously is to keep the amplification so low that the LFP does not saturate the ADC. This requires lower amplification than what you would use to record spikes without LFP (by a factor of 10 or so) and thus higher ADC resolution is required to get the same precision on the measured spike waveforms.

Dear Mani:

You managed to put a quite straightforward question in a complicated way - ) What you would like to do, as I understood, is to measure the LTP of commissural inputs to CA1 neurons of the dorsal hippocampus. And you want to know where to place your stimulation electrode.

There are numerous papers on this topic and best will be for you to check them and read some of them as there are details that you need to be aware. As a starting point I would suggest to check the paper by Bliss, Lancaster and Wheal in JP (see below). I hope this will give you some background and will serve as a good starting point. 

 http://www.ncbi.nlm.nih.gov/pubmed/6620191. Hope this will set you on the right path. 

Good luck,

SV  

Hi Christian,

usually VEP amplitudes are more sensitive to experimental condition (e.g electrode location and depth), whereas latencies measurements are less affected.

Variables that I've found affect latency are:

Try to record VEP at the same hour of the day and mouse body temperature (with circulating water heating pad t 37° C).

Hi,

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!

Thanks, Michel

I recently received a comment on the subject, it seem that some people that regularly do Juxtacellular labelling in vivo, have found that it is easier to succed in labelling  if they use NB instead of BDA. as i can see, the  only advantage is that BDA have a longer half life.

Regarding your post-laser LFP observations:

I would expect the broad silencing of an entire population of important inhibitory neurons in V1, such as you describe here, to rapidly generate massive excitation across most or all layers. Due to recurrent connections across layers and within layers, and depending somewhat on the duration of the laser exposure, perhaps the PV-silencing leads to local seizures.

Depending on the duration of the laser exposure, and how many times this was repeated, it is possible that repeated, prolonged light-induced seizures have resulted in cell death and/or detrimental plasticity, essentially making V1 epileptic. Then what you are seeing in the post-laser LFP could be "spontaneous", local seizures.

Regarding the "pre-Laser" differences in LFP with respect to other mouse lines:

I wonder whether - depending on expression level and method - there might be some low-level activation (and hence proton-pumping) occurring even without exposure to green light, or perhaps at ambient light levels? At first this seems unlikely, but even a very small effect would have time to accumulate over weeks. But in any case you could compare whole-cell recordings from ArchT-expressing neurons and PV+ neurons in wild type, or by crossing PV-Cre with e.g. a tdTomato reporter line, and look for any differences in intrinsic properties (such as greater excitability, lower AP threshold in the ArchT mice).

Hi Diego, as you want to focus on chemical mGluR2/3 dependent LTD, together with all suggestions from our colleagues, I suggest you to have a look on the papers below. Both papers explore the roles of mGluR2/3 in the cortical – Temporo-ammonic path to CA1 region.

1.    Ceolin L et al., Study of Novel Selective mGlu2 Agonist in the Temporo-Ammonic Input to CA1 Neurons Reveals Reduced mGlu2 Receptor Expression in a Wistar Substrain with an Anxiety-Like Phenotype. The Journal of Neuroscience, May 4, 2011 • 31(18):6721– 6731 • 6721

2.    Hanna L et al., Differentiating the roles of mGlu2 and mGlu3 receptors using LY541850, an mGlu2 agonist/mGlu3 antagonist. Neuropharmacology 66 (2013) 114-121

Good luck.- Zuner

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).

I agree with Michael Okun; chloride your electrodes, set up your recording situation without the animal prep and determine whether you still get drift. If you do, then use the model cell circuit that comes with the Axoclamp amplifier to test for the same thing. If you get drift with the model cell, I'd suspect leakage current in the amplifier is causing the problem. You should be able to zero this out.

Hi Roberto. During my experiments on rabbits, I usually record inspiratory activity from the phrenic nerve using bipolar electrodes. The phrenic nerve (C3 or C5 roots) should be isolated in the neck when the animal is in a supine position. Cut the skin along the midline at the level of the trachea. Gently remove the muscle to expose vagus nerve; below it, I can easily recognize the phrenic nerve. Isolate it and put a wire around it. Then, put the animal on a stereotaxic frame and fix it. Cut the skin in the neck, catch the wire and put the nerve on the bipolar electrode. I usually cut the nerve and desheat from the myelin to record the efferent inspiratory activity, but you can also left intact. Remember to cover with vaseline to prevent drying the nerve.

Good work!

I'm glad you found somebody. Carlos Belmonte and Juan Lerma at University of Alicante in Spain work with DRG.

How do you get pass the satellite glial cell if you don't dissociate the DRG?

Dear Dr. Marwa, I am not sure if understand well. You want to apply AC EMF i.e AC electric field and voltage to an animal. There are numerous ways to provoke an AC el. Field and you have to specify more about your scope. For example you could have 2 wires from the grid, 230V, 50Hz, or a 3ph system 400V, 50Hz, or a small voltage (say 12V) with smaller frequency from a supply unit. Do you need a homogenous field or not? If yes, you would need Rogowski plates (not just parallel plates) and place inside your rat. If you dont need necessary a homogenous field you could use 2 wire terminals or two plates. If you need magnetic field instead of electric, THEN you could use coils and in case of homogenous, you should use Helmholtz coils. If not you could use a simple wire. A current flux into the wire provokes a magnetic field. Between two conductors (or electrodes) connected to a voltage source (not necessary with a flowing current) provoke an electric field. If I understand well you wont need Helmholtz coil, if you want an AC EMF, i.e an AC Voltage, an AC el. Field. 

I think the method for the 200 are explained above - but much more important is the question you are asking.  The fact is that membrane potentials measured with whole-cell mode are rather meaningless except at the time that you break in because the intracellular solution is perfused and, hence, the membrane potential will be set primarily by the concentration of the potassium in your patch solution!  Vito is correct: if you want to measure the membrane potential of the cell under the conditions that its in (say a slice) the only sure way is to use a sharp electrode filled with 3-5 M KCl.  This avoids not only the perfusion problem (replacing the intracellular solution) but also the junction potentials at the tip of a patch electrode (caused by the uneven rates of diffusion of your cations and anions).  Of course any damage to the cell results in a reduced potential. 

You could explore patching using the perforated patch method where you do not break into the cell, and hence, to not perfuse its contents, but gain access using ionophores - typically something like nystatin (e.g. http://www.ncbi.nlm.nih.gov/pubmed/18998090 ).  But there are hazards here too.. e.g. http://jn.physiology.org/content/90/5/2964

Another option is to explore potential-sensitive dyes - but that's an entirely different field...http://www.ncbi.nlm.nih.gov/pubmed/10707808

Thank you, I might do that.  I didnt think plastics one supplied electrodes.  FH-Co finally made contact with me today.  Their response was short but helpful.

1) I would recommend an outer diameter <200 um to reduce tissue damage, especially when your target I less than 2500 um in mice.

2) Also, our standard 75mm length should be fine. The is the entire length of the electrode. 4,5,6 could be any length you would like. There are three numbers because the length could be three digits if you prefer (ie 100, 150). 75 should be fine for what you are doing, though.

3) Pencil Point Tip Profiles are ideal for in vivo stimulation and recording. The is the tip configuration I would recommend

Im now thinking of the logisitcs of placement of the electrodes and recording in anaesthesied animals on the same day.  Any issues with recording so soon after electrode placement i should think about?  Are guide cannula's necessary?.  Anyone know of issues with using ketamine/xylazine mix while recording for anaesthesia? 

I have used AAV2/1 for brain injections in rats and perfused after 8 weeks and we also had the cell loss problem.  In our case the dose or titre of the virus was assumed to be one of the problem and optimizing the virus dose or titre would help in this case. Another assumption is some virus strains can have target specific toxicity. For example same virus can be good in cortex where as can kill cells in striatum or SNr.  Hope this can help you a bit.

If you are using a 50 micron wire, it should be stiff enough to penetrate the brain once you have removed the dura.  If you extend the wire about 1.5mm the cannula can set on the top of the brain with the wire going into the hippocampus.  If the cannula goes into the brain, that's ok, but you do not want to damage the brain too close to recording site.  Use a silver ball as a reference.

I think you may need to go to a different pressure-sensing technology, i.e., one that senses pressure closer to the tip of the implanted catheter. I am attaching a paper that describes one such unit that seems to work well in mice. Here is also the link for the company (Data Sciences International) that commercialized the unit (I think). I have never used these units so buyer beware.

Since stereotaxic surgeries generally involve targeting brain regions that lie ventral to other important brain regions, these other areas are necessarily disrupted or distroyed during the approach to the target region. Orienting the skull at a particular angle can help to avoid damage to these areas. Ultimately, it is up to the researcher to decide which areas are least problematic to disrupt for the particular study. That should clarify which setup is the most reasonable for a particular experiment.

From which parts of M1 do you wish to record? Vibrissal (whisker) M1? Or somatic (e.g. forelimb, hindlimb) M1?

Vibrissal M1 is medial to somatic M1. Secondary motor cortex (M2) appears to be anterior to somatic M1, in frontal cortex. This is according to a variety of recent studies (although how to define M2 functionally is not entirely clear). This is different from some atlas, e.g. Paxinos, which shows M2 medial to M1: the region labeled "M2" in many an atlas appears functionally to be vibrissal M1 (vM1). See for example papers by Svoboda and Petersen groups for location of vM1.

If you wish to record from vibrissal M1, rough coordinates are 1.0 anterior, 0.9 lateral to bregma. For somatic M1, you'd need to be more lateral, say 1.5 lateral to bregma.

If you get a high gigaseal and very low leak current after brake-in you can be sure you have a good cell. You will never get a good gigaseal and low leak with unhealthy cells.

Hi,

Maybe a good thing to do is to fill a neuron by putting biocytin or Alexa antibody in your internal solution when you patch and confirm the locaton afterwards by making slices and histological processing to reveal the neuron´s location. Then you check the slice that contains the neuron in an atlas -Paxinos, for example- and confirm the area of recording.

Another thing you can do in your first trials is to locate the area by means of an injection of a dye in the center of your recording area, for example DiI or pontamine sky blue. DiI is a fluorescent dye in the orange-red range and I guess pontamine sky blue can be seen directly in your slices after cutting, as it acts as an ink -this last statement would require confirmation by an expert-.

I guess you could even deep  with your fingers the tip of a pulled capillary  -I mean if you don´t have a stereotaxic tower-, in the area of recording and filled with the dye.

Good luck!!

thank you so much ;) it is what i was looking

I work with Female Wistar rats (230-250g). I usually measure the distance between bregma and lamba which is always in between 8 to 8.6 mm and rarely it is above 8.6mm. I used to clibrate the coordinates according to bregma-lambda distance. It was working quite good in my case. I used to calibrate like this..bregma-lambda distance/9 * x-coordinate (X=AP; ML; DL). Hope this helps you.

Dear Jose,

The majority of the commercially available bipolar concentric electrodes are designed to provide the best performance in terms of spatial charge distribution. I usually work with electrodes from FHC (http://www.fh-co.com/products/research-electrodes/concentric-bipolar-microelectrodes#research-electrodes), and I can tell that they all work very well. I would specially recommend those with a beveled or pencil point shape and outer diameter ≤200 µm to reduce the tissue damage, specially when you target CA3 (~2500 µm depth in mice).

Best,

1) apply a very small positive pressure in the pipette (if the pressure it too high, you won't be able to see any change in current; see below)

2) in voltage clamp mode, make a test pulse of + 10 mV lasting 50 ms repeated every 200-300 ms

3) bring the pipette down in the tissue with steps of 2 micrometers

4) read the current trace: this is your eyes. Each time the current drops a bit, this might indicate that you touch a membrane (i.e. increase in input resistance). If it happens, withdraw the pipette. The current should go back to control value. Bring the pipette down again. If the current drops again, then you can consider you are touching a cell.

5) release the positive pressure. The current trace should ideally collapse to 0. If not, try to help it by applying a gentle negative pressure while monitoring the current. Get the feeling that the negative pressure bring the current trace down to 0. 

6) when you have a stable gigaseal, move the holding potential to -60 to -70mV and apply a brief sharp small negative pressure to rupture the membrane. Then you are in whole cell.

If you train with this procedure, you should manage to get nice recordings within 1 or 2 weeks.

good luck

Hi Tanya, 

we work in rats, so I do not quite know where and how deep the mPP is in a mouse. You might want to give some more details on your experiment. In particular, do you want to record spikes or only LFPs? Which stereotax frame do you use? How do you cement the probes in? Is the connector stable? How thick a wire do you use for your probes? 

Usually, if you want to record spikes, you need small exposed tip area which comes at the price of high impedances (around 1MOhm is usual...). These small probes are not useful for stimulation, since even an average current will destroy the surface due to the high charge density - and probably the tissue to.

So, stimulation needs bigger electrode area with less impedance - but then they are not specific enough for spikes anymore....    So, I guess you have to decide. 

We use rats and did stimulation with an FHC concentric bipolar CBC in the Striatum. Parameters are in Hiller, A., S. Loeffler, C. Haupt, M. Litza, U. G. Hofmann and A. Moser (2007). "Electrical High Frequency Stimulation of the Caudate Nucleus Induces GABA Outflow in Freely Moving Rats." J. Neurosci. Methods 159(2): 286-290. Recording only provides LFP, though, due to the big ø75µm stimulation core...

I have two alternatives for you, one in the attached paper which we call Niotrode and you can make them yourself and the other is a lithographically produced one in the pic. 

I could send you a couple if you want to try them... 

The problem with chronic implants is always the same: They will become encapsulated by Microglia (first) and  Astroglia (later). I am convinced that all rigid probes will show that effect, so we moved on to flexible multisite probes. The encapsulation will both change the electrical connection of your probe to the tissue and will move or even kill neurons in the surrounding: No more neurons to record from within the 100µm recording horizon.... 

A very common way to visualize a patch-clamped cell is to include a fluorophore in the intra-cellular solution. For example adding 100 uM Alexa 488 that emits green when excited by blue light (around 488nm, hence the name). After going whole-cell, the fluorophore will distribute in the whole cell by diffusion, making it easily recognisable. You will need an excitation light source, and appropriate  excitation and emission filters to see the fluorescence, though this is quite standard equipment in most patch-clamp rigs.

For blind patch-clamp:

Margrie TW, Brecht M, Sakmann B (2002). In vivo, low-resistance, whole-cell recordings from neurons in the anaesthetized and awake mammalian brain. Pflugers Arch. 444(4):491-8.

Hello Aaron,   

          A similar trend seems to exist in autistic spectrum disorders where the prevalence  in males is 4-5 times more than females. Disturbances in oxytocin levels are commonly implicated in autistic spectrum disorders and therefore increased levels of oxytocin in females is thought to be protective against developing autism spectrum disorders. This could be the case with epilepsy as well. Infact, epilepsy seems to be quite common in people with autistic spectrum disorders. In the hippocampus, oxytocin can enhance inhibition through its action on interneurons. Thinking on this line, difference in oxytocin levels could be one of the reasons for your observations.

Deepak.

We routinely use 0.1um fluospheres from molecular probes at a 1:200 dilution in the injectate. 

Hi Jose,

I have done such experiments before successfully. As Mer-Shahram Safari mentioned in his fourth point, i have used multi barrel pipettes for such experiments. Gluing two electrodes is hard and it form two different tips, using multi barrel glass pipettes with separate filaments is of great advantage. When you pull them it forms a common tip so that you can infuse your drug around the recording site. The method of infusion of drug is also very important. I used micro-iontophoresis technique, it is a bit complicated and also may alter the quality of the recordings. I have published a paper describing such techniques. Please find "An in vivo technique for investigating electrophysiological effects of centrally administered drugs on single neurons and network behaviour". Hope it helps.

Hello

It may be that you trigger an LTD with your low frequency stimulation. I pulse each 15s is not much but I would try to switch to one pluse each minute, or wait 15 minute to test if the response still decrease

Good luck!!!!

1. For chronic LFP recording metal electrode would be the first choice. Depending on how much you can spend - platinum-iridium or tungsten :-). The size (surface) of an active tip would determine what would you have in the signal. With a small tip (10-25 µm) you can see also multi-unit spikes. With larger tips spikes would not be visible (due to signal averaging on the electrode large surface).

2. As concern reference - you need to remember that the recorded signal is differential : reference signal is subtracted from 'active' electrode signal. So - whatever you record on reference would be present in you recorded data. If the reference is close to 'active' electrode and its signal is similar - the effect of this subtraction can be almost zero. It is useful in single unit recording - if the reference is recording LFP but not spikes it reduces slow wave component in a signal from electrode with spikes.

However - it is not so good for LFP which you want to record - here you want to have the reference in a place as neutral as possible - it should record all the general fluctuations of the field around the brain but not the neural signal. In a perfect situation it can eliminate all the noise without adding any neural activity from it's location.

Bipolar recording might be sometimes interesting - when you want to see the difference between two recording locations. But today it can be easily obtained with an off-line analysis and 'rereferencing' in the software.

3. Ground - for monopolar recording the same place (like screw in a nasal bone) can be used for connected reference and ground.

regards

ewka

Some authors discussing the presence of OT-ergic neurons in the LA refer to the paper by Sofroniew (1983). However, in his original paper Sofroniew reports the presence of OT-ergic projections in the CeA, CoA, LA, BA altogether without distinguishing among the nuclei. That may have caused assumptions that the LA and BA have OT-ergic fibers. To my knowledge it has not been really confirmed.

Hi! The input-output relationship is usually a measure of the efficacy of transmitter release. The "input" is measured from the size of the pre-synaptic fiber volley, while the "output" is usually the slope of your field EPSP. In acute slices, it is often used as an indirect measure of how viable the tissue is.

It involves both pre- and post-synaptic mechanisms, because it is a ratio between both. The size of the fiber volley is related to the number of fibers you are stimulating, i.e. pre-synapses. On the other hand, the size of the fEPSP depends on how much neurotransmitter is released, how many receptors are bound by it, and how efficient the signalling mechanism is on the post-synaptic side.

By itself it does not really tell much. But it is useful when you compare two conditions, for example tissue from normal or epileptic animals. You can quantify the size of fEPSPs at the same "input" levels (or fiber volley size) and see if your pathological conditions make any difference.

Hope this helps!